Deposition of Energy

60-35-5

DLFVBJFMPXGRIB-UHFFFAOYSA-N

Acetamide

Acetamid acetamida Acetic acid amide Acetimidic acid Ethanamide Ethanimidic acid Methanecarboxamide NSC 25945

DTXSID7020005

103-90-2

RZVAJINKPMORJF-UHFFFAOYSA-N

Acetaminophen

4-Acetamidophenol APAP Paracetamol 4-hydroxyacetanilide Acetamide, N-(4-hydroxyphenyl)- 4-(Acetylamino)phenol 4-(N-Acetylamino)phenol 4-Acetaminophenol 4'-Hydroxyacetanilide Abensanil Acetagesic Acetalgin ACETAMIDE, N-(4-HYDROXYPHENYL) Acetaminofen Acetanilide, 4'-hydroxy- ACETANILIDE, 4-HYDROXY- Algotropyl Alvedon Anaflon Apamide Banesin Ben-u-ron Bickie-mol Biocetamol Cetadol Citramon P Claratal Clixodyne Dafalgan Daphalgan Dial-a-gesic Disprol Doliprane Dolprone Dymadon Efferalgan Endophy Febrilex Febrilix Febro-Gesic Febrolin Fepanil Finimal Gattaphen T Gelocatil Gutte Enteric Homoolan Jin Gang Lestemp Liquagesic Lonarid Lyteca Syrup Minoset Momentum N-(4-Hydroxyphenyl)acetamide N-Acetyl-4-aminophenol N-Acetyl-4-hydroxyaniline N-Acetyl-p-aminophenol Napafen Naprinol Nobedon NSC 109028 NSC 3991 Ortensan p-(Acetylamino)phenol p-Aceaminophenol Pacemol p-Acetamidophenol p-Acetoaminophen P-ACETYLAMINOPHENOL Paldesic panadeine Panadol Panadol Actifast Panadol Extend Panaleve Panasorb Panodil Paracetamol DC Paracetamole Parageniol Paramol Paraspen Parelan Pasolind N Perfalgan Phenaphen Phendon p-Hydroxyacetanilide Prodafalgan Puerxitong Pyrinazine Resfenol Resprin Rhodapop NCR Salzone Tabalgin Tachipirina Tempanal Tralgon Tylenol TylolHot Valadol Valgesic Vermidon Vick Pyrena

DTXSID2020006

968-81-0

VGZSUPCWNCWDAN-UHFFFAOYSA-N

Acetohexamide

Benzenesulfonamide, 4-acetyl-N-[(cyclohexylamino)carbonyl]- 1-(p-Acetylbenzenesulfonyl)-3-cyclohexylurea 1-[(p-Acetylphenyl)sulfonyl]-3-cyclohexylurea Acetohexamid acetohexamida Dimelin Dimelor Dymelor Gamadiabet Hypoglicil Metaglucina Minoral N-(p-Acetylphenylsulfonyl)-N'-cyclohexylurea Ordimel Tsiklamid Urea, 1-[(p-acetylphenyl)sulfonyl]-3-cyclohexyl-

DTXSID7020007

67-66-3

HEDRZPFGACZZDS-UHFFFAOYSA-N

Chloroform

Trichloromethane Methane, trichloro- CARBON TRICHLORIDE Chloroforme cloroformo Formyl trichloride Methane trichloride Methane,trichloro- NSC 77361 Trichloroform UN 1888

DTXSID1020306

110-00-9

YLQBMQCUIZJEEH-UHFFFAOYSA-N

Furan

Divinylene oxide furanne Furfuran Oxacyclopentadiene Tetrole UN 2389

DTXSID6020646

7429-90-5

XAGFODPZIPBFFR-UHFFFAOYSA-N

AZDRQVAHHNSJOQ-UHFFFAOYSA-N

Aluminum

Aisin Metal Fiber Al 050P-H24 ALC Fine Alcan XI 1391 Almi-Paste SSP 303AR Aloxal 3010 Alpaste 00-0506 Alpaste 0100M Alpaste 0100MA Alpaste 0100M-C Alpaste 0200M Alpaste 0200T Alpaste 0230M Alpaste 0230T Alpaste 0241M Alpaste 0300M Alpaste 0500M Alpaste 0539X Alpaste 0620MS Alpaste 0625TS Alpaste 0638-70C Alpaste 0700M Alpaste 0780M Alpaste 0900M Alpaste 100M Alpaste 100MS Alpaste 100MSR Alpaste 1100M Alpaste 1100MA Alpaste 1100N Alpaste 1100NA Alpaste 1109MA Alpaste 1109MC Alpaste 1200M Alpaste 1200T Alpaste 1260MS Alpaste 1500MA Alpaste 1700NL Alpaste 1810YL Alpaste 1830YL Alpaste 1900M Alpaste 1900XS Alpaste 1950M Alpaste 1950N Alpaste 210N Alpaste 2172EA Alpaste 2173 Alpaste 240T Alpaste 241M Alpaste 417 Alpaste 46-046 Alpaste 4-621 Alpaste 4919 Alpaste 50-63 Alpaste 50-635 Alpaste 51-148B Alpaste 51-231 Alpaste 5205N Alpaste 5207N Alpaste 52-509 Alpaste 52-568 Alpaste 5301N Alpaste 5302N Alpaste 53-119 Alpaste 5422NS Alpaste 54-452 Alpaste 54-497 Alpaste 54-542 Alpaste 55-516 Alpaste 55-519 Alpaste 55-574 Alpaste 5620NS Alpaste 5630NS Alpaste 5640NS Alpaste 56-501 Alpaste 5650NS Alpaste 5653NS Alpaste 5654NS Alpaste 5680N Alpaste 5680NS Alpaste 60-600 Alpaste 60-760 Alpaste 60-768 Alpaste 62-356 Alpaste 6340NS Alpaste 6370NS Alpaste 6390NS Alpaste 640NS Alpaste 65-388 Alpaste 66NLB Alpaste 710N Alpaste 7130N Alpaste 7160N Alpaste 7160NS Alpaste 725N Alpaste 740NS Alpaste 7430NS Alpaste 7580NS Alpaste 7620NS Alpaste 7640NS Alpaste 7670M Alpaste 7670NS Alpaste 7675NS Alpaste 7679NS Alpaste 7680N Alpaste 7680NS Alpaste 76840NS Alpaste 7730N Alpaste 7770N Alpaste 7830N Alpaste 8004 Alpaste 8080N Alpaste 8260NAR Alpaste 891K Alpaste 91-0562 Alpaste 92-0592 Alpaste 93-0595 Alpaste 93-0647 Alpaste 94-2315 Alpaste 95-0570 Alpaste 96-0635 Alpaste 96-2104 Alpaste 97-0510 Alpaste 97-0534 Alpaste AW 520B Alpaste AW 612 Alpaste AW 9800 Alpaste F 795 Alpaste FM 7680K Alpaste FX 440 Alpaste FX 910 Alpaste FZ 0534 Alpaste FZU 40C Alpaste G Alpaste HR 8801 Alpaste HS 2 Alpaste J Alpaste K 9800 Alpaste MC 666 Alpaste MC 707 Alpaste MF 20 Alpaste MG 01 Alpaste MG 1000 Alpaste MG 1300 Alpaste MG 500 Alpaste MG 600 Alpaste MH 6601 Alpaste MH 8801 Alpaste MH 9901 Alpaste MR 7000 Alpaste MR 9000 Alpaste MS 630 Alpaste N 1700NL Alpaste NS 7670 Alpaste O 100N Alpaste O 2130 Alpaste O 300M Alpaste P 0100 Alpaste P 1950 Alpaste S Alpaste SAP 110 Alpaste SAP 414P Alpaste SAP 550N Alpaste SCR 5070 Alpaste TCR 2020 Alpaste TCR 2060 Alpaste TCR 2070 Alpaste TCR 3010 Alpaste TCR 3030 Alpaste TCR 3040 Alpaste TCR 3130 Alpaste TD 200T Alpaste UF 500 Alpaste WB 0230 Alpaste WD 500 Alpaste WJP-U 75C Alpaste WX 0630 Alpaste WX 7830 Alpaste WXA 7640 Alpaste WXM 0630 Alpaste WXM 0650 Alpaste WXM 0660 Alpaste WXM 1415 Alpaste WXM 1440 Alpaste WXM 5422 Alpaste WXM 760b Alpaste WXM 7640 Alpaste WXM 7675 Alpaste WXM-T 60B Alpaste WXM-U 75 Alpaste WXM-U 75C Altop X Aluchrome Ultrafin Super Alumat 1600 Alumet H 30 aluminio Aluminium Aluminium Flake Aluminum 27 Aluminum atom Aluminum element Aluminum Flake PCF 7620 Aluminum granules ALUMINUM METAL/GRANULE ALUMINUM PASTE ALUMINUM PIGMENT ALUMINUM TURNINGS Alumi-paste 640NS Alumipaste 91-0562 Alumipaste 98-1822T Alumipaste AW 620 Alumipaste CR 300 Alumipaste GX 180A Alumipaste GX 201A Alumipaste HR 7000 Alumipaste HR 850 Alumipaste MG 11 Alumipaste MH 8801 Aquamet NPW 2900 Aquapaste 205-5 Aquasilver LPW Astroflake 40 Astroflake Black N 020 Astroflake Black N 070 Astroflake LG 40 Astroflake LG 70 Astroflake Silver N 040 Astroshine NJ 1600 Astroshine T 8990 Atomizalumi VA 200 C.I. PIGMENT METAL 1 Chromal IV Chromal X Decomet 1001/10 Decomet 2018/10 Decomet High Gloss Al 1002/10 Ecka AS 081 Eckart 9155 Eterna Brite 301-1 Eterna Brite 601-1 Eterna Brite 651-1 Eterna Brite EBP 251PA Eterna Brite Primier 251PA Ferro FX 53-038 Friend Color F 500GR-W Friend Color F 500WT Friend Color F 700RE-W Friend Color F 701RE-W Hi Print 60T High Print 60T Hisparkle HS 2 Hydro Paste 8726 Hydrolac WHH 2153 Hydrolan 3560 Hydrolux Reflexal 100 Hydroshine WS 1001 JISA 51010P Kryal Z Lansford 243 LE Sheet 800 Leafing Alpaste LG-H Silver 25 Lunar Al-V 95 Metallux 161 Metallux 2154 Metallux 2192 Metalure Metalure 55350 Metalure L 55350 Metalure L 59510 Metalure W 2001 Metapor Metasheen 1800 Metasheen HR 0800 Metasheen KM 100 Metasheen KM 1000 Metasheen Slurry 1807 Metasheen Slurry 1811 Metasheen Slurry KM 100 Metax G Metax S Mirror Glow 1000 Mirror Glow 600 Mirrorsheen Noral Aluminium Noral Ink Grade Aluminium Obron 10890 Offset FM 4500 Puratronic Reflexal 145 Reynolds 400 Reynolds 4-301 Reynolds 4-591 Reynolds 667 SAP 260PW-HS SAP-FM 4010 SBC 516-20Z Scotchcal 7755SE Serumekku Setanium 50MIS-H8 Siberline ET 2025 Siberline ST 21030E1 Silvar A Silver VT 522 Silverline SSP 353 Silvex 793-20C Sparkle Silver 3141ST Sparkle Silver 3500 Sparkle Silver 3641 Sparkle Silver 5000AR Sparkle Silver 516AR Sparkle Silver 5242AR Sparkle Silver 5245AR Sparkle Silver 5271AR Sparkle Silver 5500 Sparkle Silver 5745 Sparkle Silver 7000AR Sparkle Silver 7005AR Sparkle Silver 7500 Sparkle Silver 960-25E1 Sparkle Silver E 1745AR Sparkle Silver L 1526AR Sparkle Silver Premier 751 Sparkle Silver SS 3130 Sparkle Silver SS 5242AR Sparkle Silver SS 5588 Sparkle Silver SSP 132AR Special PCR 507 Splendal 6001BG Spota Mobil 801 SSP 760-20C Stapa Aloxal PM 2010 Stapa Aloxal PM 3010 Stapa Aloxal PM 4010 Stapa Hydrolac BG 8n.1 Stapa Hydrolac BGH Chromal X Stapa Hydrolac PM Chromal VIII Stapa Hydrolac W 60NL Stapa Hydrolac WH 16 Stapa Hydrolac WH 66NL Stapa Hydrolux 2192 Stapa Hydrolux 8154 Stapa IL Hydrolan 2192-55900G Stapa Metallic R 607 Stapa Metallux 1050 Stapa Metallux 211 Stapa Metallux 212 Stapa Metallux 2196 Stapa Metallux 274 Stapa Mobilux 181 Stapa Offset 3000 Stapa PV 10 Stapa VP 46432G Starbrite 2100 Super Fine 18000 Super Fine 22000 Supramex 2022 Toyo Aluminum 02-0005 Toyo Aluminum 93-3040 Transmet K 102HE Tufflake 3645 Tufflake 5843 UN 1396 US Aluminum 809 Valimet H 2 Valimet H 3 White Silver 7080N White Silver 7130N

DTXSID3040273

7440-43-9

BDOSMKKIYDKNTQ-UHFFFAOYSA-N

Cadmium

Cadimium CADMIUM BLUE CADMIUM, IN PLATTEN, STANGEN, BROCKEN,KOERNER

DTXSID1023940

7439-97-6

QSHDDOUJBYECFT-UHFFFAOYSA-N

Mercury

Liquid silver Mercure MERCURIC METAL TRIPLE DISTILLED mercurio Mercury element Quecksilber Quicksilver UN 2024 UN 2809

DTXSID1024172

7440-61-1

JFALSRSLKYAFGM-UHFFFAOYSA-N

Uranium

Uranium, isotope of mass 238 238U Element UN 2979 (DOT) Uranium I

DTXSID1042522

7440-38-2

RQNWIZPPADIBDY-UHFFFAOYSA-N

Arsenic

As Arsenic black ARSENIC METAL arsenico Grey arsenic UN 1558

DTXSID4023886

7440-22-4

BQCADISMDOOEFD-UHFFFAOYSA-N

Silver

Ag Nanopaste NPS-J 90 Ag Sphere 2 Ag-C-GS Algaedyn Arctic Silver 3 Argentum Astroflake 5 Carey Lea silver Colloidal silver Dotite XA 208 Du Pont 4943 ECM 100AF4810 Enlight 600 Enlight silver plate 600 Epinall Finesphere SVND 102 Fordel DC FP 5369-502 Jelcon SH 1 Jungindai Takasago 300 KS (metal) LCP 1-19SFS Metz 3000-1 Nanomelt AGC-A Nanomelt Ag-XA 301 Nanomelt Ag-XF 301 Nanomelt Ag-XF 301H Nanopaste NPS-J 90 Perfect Silver Puff Silver X 1200 RT 1710S-C1 SD (metal) Shell Silver Silbest E 20 Silbest F 20 Silbest J 18 Silbest TC 12 Silbest TC 20E Silbest TC 25A Silbest TCG 1 Silbest TCG 7 Silcoat AgC 103 Silcoat AgC 2011 Silcoat AgC 209 Silcoat AgC 2190 Silcoat AgC 222 Silcoat AgC 2411 Silcoat AgC 74T Silcoat AgC-A Silcoat AgC-AO Silcoat AgC-B Silcoat AgC-BO Silcoat AgC-D Silcoat AgC-G Silcoat AgC-GS Silcoat AgC-L Silcoat AgC-O Silcoat GS Silcoat RF 200 Silflake 135 Silsphere 514 Silver atom Silver element Silver Flake 1 Silver Flake 25 Silver Flake 52 Silver Flake 7A SILVER FLAKES Silver metal Silvest TCG 11N Technic 299 Technic 450 Techno Alpha 175

DTXSID4024305

7439-96-5

PWHULOQIROXLJO-UHFFFAOYSA-N

Manganese

Colloidal manganese Cutaval Manganese element Manganese fulleride Manganese metal alloy Manganese-55 manganeso

DTXSID2024169

7440-02-0

PXHVJJICTQNCMI-UHFFFAOYSA-N

Nickel

Carbonyl 255 Carbonyl Ni 123 Carbonyl Ni 283 Carbonyl Nickel 123 Carbonyl Nickel 283 Carbonyl Nickel 287 Cerac N 2003 CNS 10 Micron Exmet 4 Ni X-4/0 Fibrex P Incofoam Nickel element NICKEL ROUND ANODES Nicrobraz LM:BNi 2 Ni-Flake 95 Novamet 123 Novamet 4SP Novamet 4SP10 Novamet 525 Novamet CNS 400 Novamet HCA 1 Novamet NI 255 Raney nickel Raney nickel 2800 UN 1325 UN 2881

DTXSID2020925

7440-66-6

HCHKCACWOHOZIP-UHFFFAOYSA-N

Zinc

Zn Asarco L 15 C.I. Pigment Black 16 Merrillite NC-Zinc Rheinzink Stapa TE Zinc AT UF (metal) UN 1436 Zinc dust Zinc Dust 3 Zinc Dust 500 mesh Zinc Dust LS 2 Zinc Dust MCS Zinc Flakes GTT ZINC METAL ZINC MOSSY ZINC STRIP ZINC, MOSSY Zincsalt GTT

DTXSID7035012

UBERON:0001986

UBERON endothelium

UBERON:0001981

UBERON blood vessel

MP:0003674

MP oxidative stress

GO:0002526

GO acute inflammatory response

GO:0033484

GO nitric oxide homeostasis

GO:0001974

GO blood vessel remodeling

WIKI increased

WIKI functional change

WIKI occurrence

Ionizing Radiation Ionizing radiation can vary in energy, dose, charge, and in the spatial distributions of energy transferred to other matter (linear energy transfer per unit length or LET) (ICRU 1970). At the same dose, low and high LET both generate energy deposition events, including many higher energy events (Goodhead and Nikjoo 1989). However, they differ in the spatial distribution and upper range of intensity of energy deposited. Lower LET such as gamma rays sparsely deposit many individual excitations or small clusters of excitations of low energy (Goodhead 1988). In contrast, high LET such as alpha particles have fewer tracks but readily transfer their energy to matter and therefore deposit their energy over a much smaller area (Goodhead 1994). Consequently, alpha and other high LET particles penetrate less deeply into tissue, interactions are densely focused on a narrow track, and individual energy depositions can be large (Goodhead 1988). These different energy deposition patterns can lead to differences in radiation effects including the pattern of DNA damage.

Exposure to ionizing radiation can come from natural and industrial sources. Space and terrestrial radiation includes a range of LET particles, while diagnostic radiation methods such as X-ray imaging, mammography and CT scans use low LET X-rays. Radiation therapy can use an external beam to direct radiation on a focused tissue area, or deposit solid or liquid radioactive materials in the body that release (mostly gamma) radiation internally. External radiotherapy typically uses X-rays but is moving towards higher LET charged particles such as protons and heavy ions (Durante, Orecchia et al. 2017).

2019-05-03T12:36:36

2019-05-07T12:12:13

Acetaminophen

2016-11-29T18:42:26

Chloroform

2016-11-29T18:42:27

furan

2020-05-01T14:35:22

Platinum

2022-02-04T14:36:54

Aluminum

2022-02-04T14:42:11

Cadmium

2017-10-25T08:33:12

Mercury

2016-11-29T18:42:19

Uranium

2021-08-05T14:28:50

Arsenic

2021-04-27T00:15:21

Silver

2022-02-03T11:20:11

Manganese

2022-02-04T14:47:23

Nickel

2022-02-04T14:47:59

Zinc

2022-02-04T15:05:00

nanoparticles

2016-12-21T09:40:06

Topoisomerase inhibitors

2019-05-19T20:21:24

Radiomimetic compounds

2019-05-19T20:21:42

WCS_9606

common toxicological species human

10116

NCBI rat

10090

NCBI mouse

6239

NCBI nematode

WCS_7955

common ecological species zebrafish

3702

NCBI thale-cress

3349

NCBI Scotch pine

WCS_35525

common ecological species Daphnia magna

3055

NCBI Chlamydomonas reinhardtii

WCS_6396

common ecological species common brandling worm

WCS_4472

common ecological species Lemna minor

8030

NCBI Salmo salar

WikiUser_26

ApacheUser rodents

9606

NCBI Homo sapiens

WikiUser_28

Vertebrates

WCS_9986

common toxicological species rabbit

WikiUser_25

Wikiuser: Cyauk human and other cells in culture

9913

NCBI bovine

WikiUser_24

Wikiuser:Migration Pig

9823

NCBI pigs

Deposition of Energy

Energy Deposition

Molecular

Deposition of energy refers to events where energetic subatomic particles, nuclei, or electromagnetic radiation deposit energy in the media through which they transverse. The energy may either be sufficient (e.g. ionizing radiation) or insufficient (e.g. non-ionizing radiation) to ionize atoms or molecules (Beir et al.,1999).   Ionizing radiation can cause the ejection of electrons from atoms and molecules, thereby resulting in their ionization and the breakage of chemical bonds. The energy of these subatomic particles or electromagnetic waves mostly range from 124 KeV to 5.4 MeV and is dependent on the source and type of radiation (Zyla et al., 2020). To be ionizing the incident radiation must have sufficient energy to free electrons from atomic or molecular electron orbitals. The energy deposited can induce direct and indirect ionization events and this can be via internal (injections, inhalation, or absorption of radionuclides) or external exposure from radiation fields -- this also applies to non-ionizing radiation. Direct ionization is the principal path where charged particles interact with biological structures such as DNA, proteins or membranes to cause biological damage. Photons, which are electromagnetic waves can also deposit energy to cause direct ionization. Ionization of water, which is a major constituent of tissues and organs, produces free radical and molecular species, which themselves can indirectly damage critical targets such as DNA (Beir et al., 1999; Balagamwala et al., 2013) or alter cellular processes. Given the fundamental nature of energy deposition by radioactive/unstable nuclei, nucleons or elementary particles in material, this process is universal to all biological contexts. The spatial structure of ionizing energy deposition along the resulting particle track is represented as linear energy transfer (LET) (Hall and Giaccia, 2018 UNSCEAR, 2020). High LET refers to energy mostly above 10 keV μm-1 which produces more complex, dense structural damage than low LET radiation (below 10 keV μm-1). Low-LET particles produce sparse ionization events such as photons (X- and gamma rays), as well as high-energy protons. Low LET radiation travels farther into tissue but deposits smaller amounts of energy, whereas high LET radiation, which includes heavy ions, alpha particles and high-energy neutrons, does not travel as far but deposits larger amounts of energy into tissue at the same absorbed dose. The biological effect of the deposition of energy can be modulated by varying dose and dose rate of exposure, such as acute, chronic, or fractionated exposures (Hall and Giaccia, 2018). Non-ionizing radiation is electromagnetic waves that does not have enough energy to break bonds and induce ion formation but it can cause molecules to excite and vibrate faster resulting in biological effects. Examples of non-ionizing radiation include radio waves (wavelength: 100 km-1m), microwaves (wavelength: 1m-1mm), infrared radiation (wavelength: 1mm- 1 um), visible light (wavelengths: 400-700 nm), and ultraviolet radiation of longer wavelengths such as UVB (wavelengths: 315-400nm) and UVA (wavelengths: 280-315 nm). UVC radiation (200-280 nm) is, in contrast to UVB and UVA, considered to be a type of ionizing radiation.

<table border="1" bordercolor="#ccc" cellpadding="5" cellspacing="0" style="border-collapse:collapse"> <tbody> <tr> <td style="background-color:#eeeeee; text-align:center">Radiation type</td> <td style="background-color:#eeeeee; text-align:center"> Assay Name </td> <td style="background-color:#eeeeee; text-align:center"> References </td> <td style="background-color:#eeeeee; text-align:center"> Description </td> <td style="background-color:#eeeeee; text-align:center"> OECD Approved Assay </td> </tr> <tr> <td style="text-align:center">Ionizing radiation</td> <td style="text-align:center"> Monte Carlo Simulations (Geant4) </td> <td style="text-align:center"> Douglass et al., 2013; Douglass et al. 2012; Zyla et al., 2020 </td> <td style="text-align:center"> Monte Carlo simulations are based on a computational algorithm that mathematically models the deposition of energy into materials. </td> <td style="text-align:center"> No </td> </tr> <tr> <td style="text-align:center">Ionizing radiation</td> <td style="text-align:center"> Fluorescent Nuclear Track Detector (FNTD) </td> <td style="text-align:center"> Sawakuchi, 2016; Niklas, 2013; Koaira & Konishi, 2015 </td> <td style="text-align:center"> FNTDs are biocompatible chips with crystals of aluminium oxide doped with carbon and magnesium; used in conjuction with fluorescent microscopy, these FNTDs allow for the visualization and the linear energy transfer (LET) quantification of tracks produced by the deposition of energy into a material. </td> <td style="text-align:center"> No </td> </tr> <tr> <td style="text-align:center">Ionizing radiation</td> <td style="text-align:center">Tissue equivalent proportional counter (TEPC)</td> <td style="text-align:center">Straume et al, 2015</td> <td style="text-align:center">Measure the LET spectrum and calculate the dose equivalent.</td> <td style="text-align:center">No</td> </tr> <tr> <td style="text-align:center">Ionizing radiation</td> <td style="text-align:center">alanine dosimeters/NanoDots</td> <td style="text-align:center"> Lind et al. 2019; Xie et al., 2022 </td> <td style="text-align:center"> </td> <td style="text-align:center">No</td> </tr> <tr> <td style="text-align:center">Non-ionizing radiation</td> <td style="text-align:center">UV meters or radiameters</td> <td style="text-align:center">Xie et at., 2020</td> <td style="text-align:center">UVA/UVB (irradiance intensity), UV dosimeters (accumulated irradiance over time), Spectrophoto meter (absorption of UV by a substance or material)</td> <td style="text-align:center">No</td> </tr> </tbody> </table>

Energy can be deposited into any substrate, both living and non-living; it is independent of age, taxa, sex, or life-stage. Taxonomic applicability: This MIE is not taxonomically specific.   Life stage applicability: This MIE is not life stage specific.  Sex applicability: This MIE is not sex specific.

Low Unspecific

High

All life stages

Moderate Moderate Moderate High High High Moderate High Moderate Moderate High Low Balagamwala, E. H. et al. (2013), “Introduction to radiotherapy and standard teletherapy techniques”, Dev Ophthalmol, Vol. 52, Karger, Basel, https://doi.org/10.1159/000351045  Beir, V. et al. (1999), “The Mechanistic Basis of Radon-Induced Lung Cancer”, in Health Risks of Exposure to Radon: BEIR VI, National Academy Press, Washington, D.C., https://doi.org/10.17226/5499  Douglass, M. et al. (2013), “Monte Carlo investigation of the increased radiation deposition due to gold nanoparticles using kilovoltage and megavoltage photons in a 3D randomized cell model”, Medical Physics, Vol. 40/7, American Institute of Physics, College Park, https://doi.org/10.1118/1.4808150  Douglass, M. et al. (2012), “Development of a randomized 3D cell model for Monte Carlo microdosimetry simulations.”, Medical Physics, Vol. 39/6, American Institute of Physics, College Park, https://doi.org/10.1118/1.4719963  Hall, E. J. and Giaccia, A.J. (2018), Radiobiology for the Radiologist, 8th edition, Wolters Kluwer, Philadelphia.   Kodaira, S. and Konishi, T. (2015), “Co-visualization of DNA damage and ion traversals in live mammalian cells using a fluorescent nuclear track detector.”, Journal of Radiation Research, Vol. 56/2, Oxford University Press, Oxford, https://doi.org/10.1093/jrr/rru091  Lind, O.C., D.H. Oughton and Salbu B. (2019), "The NMBU FIGARO low dose irradiation facility", International Journal of Radiation Biology, Vol. 95/1, Taylor & Francis, London, https://doi.org/10.1080/09553002.2018.1516906. Sawakuchi, G.O. and Akselrod, M.S. (2016), “Nanoscale measurements of proton tracks using fluorescent nuclear track detectors.”, Medical Physics, Vol. 43/5, American Institute of Physics, College Park, https://doi.org/10.1118/1.4947128  Straume, T. et al. (2015), “Compact Tissue-equivalent Proportional Counter for Deep Space Human Missions.”, Health physics, Vol. 109/4, Lippincott Williams & Wilkins, Philadelphia, https://doi.org/10.1097/HP.0000000000000334  Niklas, M. et al. (2013), “Engineering cell-fluorescent ion track hybrid detectors.”, Radiation Oncology, Vol. 8/104, BioMed Central, London, https://doi.org/10.1186/1748-717X-8-141  UNSCEAR (2020), Sources, effects and risks of ionizing radiation, United Nations.  Xie, Li. et al. (2022), "Ultraviolet B Modulates Gamma Radiation-Induced Stress Responses in Lemna Minor at Multiple Levels of Biological Organisation", SSRN, Elsevier, Amsterdam, http://dx.doi.org/10.2139/ssrn.4081705 . Zyla, P.A. et al. (2020), Review of particle physics: Progress of Theoretical and Experimental Physics, 2020 Edition, Oxford University Press, Oxford.      AOPWiki 2019-08-22T09:44:23

2023-08-09T15:06:01

Oxidative Stress

Molecular

Oxidative stress is defined as an imbalance in the production of reactive oxygen species (ROS) and antioxidant defenses. High levels of oxidizing free radicals can be very damaging to cells and molecules within the cell.  As a result, the cell has important defense mechanisms to protect itself from ROS. For example, Nrf2 is a transcription factor and master regulator of the oxidative stress response. During periods of oxidative stress, Nrf2-dependent changes in gene expression are important in regaining cellular homeostasis (Nguyen, et al. 2009) and can be used as indicators of the presence of oxidative stress in the cell. In addition to the directly damaging actions of ROS, cellular oxidative stress also changes cellular activities on a molecular level. Redox sensitive proteins have altered physiology in the presence and absence of ROS, which is caused by the oxidation of sulfhydryls to disulfides (2SH àSS) on neighboring amino acids (Antelmann and Helmann 2011). Importantly Keap1, the negative regulator of Nrf2, is regulated in this manner (Itoh, et al. 2010). ROS also undermine the mitochondrial defense system from oxidative damage. The antioxidant systems consist of superoxide dismutase, catalase, glutathione peroxidase and glutathione reductase, as well as antioxidants such as α-tocopherol and ubiquinol, or antioxidant vitamins and minerals including vitamin E, C, carotene, lutein, zeaxanthin, selenium, and zinc (Fletcher, 2010). The enzymes, vitamins and minerals catalyze the conversion of ROS to non-toxic molecules such as water and O2. However, these antioxidant systems are not perfect and endogenous metabolic processes and/or exogenous oxidative influences can trigger cumulative oxidative injuries to the mitochondria, causing a decline in their functionality and efficiency, which further promotes cellular oxidative stress (Balasubramanian, 2000; Ganea & Harding, 2006; Guo et al., 2013; Karimi et al., 2017). However, an emerging viewpoint suggests that ROS-induced modifications may not be as detrimental as previously thought, but rather contribute to signaling processes (Foyer et al., 2017).  Protection against oxidative stress is relevant for all tissues and organs, although some tissues may be more susceptible. For example, the brain possesses several key physiological features, such as high O2 utilization, high polyunsaturated fatty acids content, presence of autooxidable neurotransmitters, and low antioxidant defenses as compared to other organs, that make it highly susceptible to oxidative stress (Halliwell, 2006; Emerit and al., 2004; Frauenberger et al., 2016). Sources of ROS Production Direct Sources: Direct sources involve the deposition of energy onto water molecules, breaking them into active radical species. When ionizing radiation hits water, it breaks it into hydrogen (H*) and hydroxyl (OH*) radicals by destroying its bonds. The hydrogen will create hydroxyperoxyl free radicals (HO2*) if oxygen is available, which can then react with another of itself to form hydrogen peroxide (H2O2) and more O2 (Elgazzar and Kazem, 2015). Antioxidant mechanisms are also affected by radiation, with catalase (CAT) and peroxidase (POD) levels rising as a result of exposure (Seen et al. 2018; Ahmad et al. 2021). Indirect Sources: An indirect source of ROS is the mitochondria, which is one of the primary producers in eukaryotic cells (Powers et al., 2008).  As much as 2% of the electrons that should be going through the electron transport chain in the mitochondria escape, allowing them an opportunity to interact with surrounding structures. Electron-oxygen reactions result in free radical production, including the formation of hydrogen peroxide (H2O2) (Zhao et al., 2019). The electron transport chain, which also creates ROS, is activated by free adenosine diphosphate (ADP), O2, and inorganic phosphate (Pi) (Hargreaves et al. 2020; Raimondi et al. 2020; Vargas-Mendoza et al. 2021). The first and third complexes of the transport chain are the most relevant to mammalian ROS production (Raimondi et al., 2020). The mitochondria have its own set of DNA and it is a prime target of oxidative damage (Guo et al., 2013). ROS are also produced through nicotinamide adenine dinucleotide phosphate oxidase (NOX) stimulation, an event commenced by angiotensin II, a product/effector of the renin-angiotensin system (Nguyen Dinh Cat et al. 2013; Forrester et al. 2018). Other ROS producers include xanthine oxidase, immune cells (macrophage, neutrophils, monocytes, and eosinophils), phospholipase A2 (PLA2), monoamine oxidase (MAO), and carbon-based nanomaterials (Powers et al. 2008; Jacobsen et al. 2008; Vargas-Mendoza et al. 2021).

Oxidative Stress. Direct measurement of ROS is difficult because ROS are unstable. The presence of ROS can be assayed indirectly by measurement of cellular antioxidants, or by ROS-dependent cellular damage. Listed below are common methods for detecting the KE, however there may be other comparable methods that are not listed <ul> <li>Detection of ROS by chemiluminescence (https://www.sciencedirect.com/science/article/abs/pii/S0165993606001683)</li> <li>Detection of ROS by chemiluminescence is also described in OECD TG 495 to assess phototoxic potential.</li> <li>Glutathione (GSH) depletion. GSH can be measured by assaying the ratio of reduced to oxidized glutathione (GSH:GSSG) using a commercially available kit (e.g., http://www.abcam.com/gshgssg-ratio-detection-assay-kit-fluorometric-green-ab138881.html). </li> <li>TBARS. Oxidative damage to lipids can be measured by assaying for lipid peroxidation using TBARS (thiobarbituric acid reactive substances) using a commercially available kit. </li> <li>8-oxo-dG. Oxidative damage to nucleic acids can be assayed by measuring 8-oxo-dG adducts (for which there are a number of ELISA based commercially available kits),or  HPLC, described in Chepelev et al. (Chepelev, et al. 2015).</li> </ul> Molecular Biology: Nrf2. Nrf2’s transcriptional activity is controlled post-translationally by oxidation of Keap1. Assay for Nrf2 activity include: <ul> <li>Immunohistochemistry for increases in Nrf2 protein levels and translocation into the nucleus</li> <li>Western blot for increased Nrf2 protein levels</li> <li>Western blot of cytoplasmic and nuclear fractions to observe translocation of Nrf2 protein from the cytoplasm to the nucleus</li> <li>qPCR of Nrf2 target genes (e.g., Nqo1, Hmox-1, Gcl, Gst, Prx, TrxR, Srxn), or by commercially available pathway-based qPCR array (e.g., oxidative stress array from SABiosciences)</li> <li>Whole transcriptome profiling by microarray or RNA-seq followed by pathway analysis (in IPA, DAVID, metacore, etc.) for enrichment of the Nrf2 oxidative stress response pathway (e.g., Jackson et al. 2014)</li> <li>OECD TG422D describes an ARE-Nrf2 Luciferase test method</li> <li>In general, there are a variety of commercially available colorimetric or fluorescent kits for detecting Nrf2 activation</li> </ul>   <table border="1" cellpadding="1" cellspacing="1"> <tbody> <tr> <td>Assay Type & Measured Content</td> <td>Description</td> <td>Dose Range Studied</td> <td> Assay Characteristics (Length / Ease of use/Accuracy) </td> </tr> <tr> <td> ROS Formation in the Mitochondria assay (Shaki et al., 2012) </td> <td>“The mitochondrial ROS measurement was performed flow cytometry using DCFH-DA. Briefly, isolated kidney mitochondria were incubated with UA (0, 50, 100 and 200 μM) in respiration buffer containing (0.32 mM sucrose, 10 mM Tris, 20 mM Mops, 50 μM EGTA, 0.5 mM MgCl2, 0.1 mM KH2PO4 and 5 mM sodium succinate) [32]. In the interval times of 5, 30 and 60 min following the UA addition, a sample was taken and DCFH-DA was added (final concentration, 10 μM) to mitochondria and was then incubated for 10 min. Uranyl acetate-induced ROS generation in isolated kidney mitochondria were determined through the flow cytometry (Partec, Deutschland) equipped with a 488-nm argon ion laser and supplied with the Flomax software and the signals were obtained using a 530-nm bandpass filter (FL-1 channel). Each determination is based on the mean fluorescence intensity of 15,000 counts.”</td> <td>0, 50, 100 and 200 μM of Uranyl Acetate</td> <td> Long/ Easy High accuracy </td> </tr> <tr> <td> Mitochondrial Antioxidant Content Assay Measuring GSH content (Shaki et al., 2012)</td> <td>“GSH content was determined using DTNB as the indicator and spectrophotometer method for the isolated mitochondria. The mitochondrial fractions (0.5 mg protein/ml) were incubated with various concentrations of uranyl acetate for 1 h at 30 °C and then 0.1 ml of mitochondrial fractions was added into 0.1 mol/l of phosphate buffers and 0.04% DTNB in a total volume of 3.0 ml (pH 7.4). The developed yellow color was read at 412 nm on a spectrophotometer (UV-1601 PC, Shimadzu, Japan). GSH content was expressed as μg/mg protein.”</td> <td> 0, 50, 100, or 200 μM Uranyl Acetate </td> <td> </td> </tr> <tr> <td> H2O2 Production Assay Measuring H2O2 Production in isolated mitochondria (Heyno et al., 2008)</td> <td>“Effect of CdCl2 and antimycin A (AA) on H2O2 production in isolated mitochondria from potato. H2O2 production was measured as scopoletin oxidation. Mitochondria were incubated for 30 min in the measuring buffer (see the Materials and Methods) containing 0.5 mM succinate as an electron donor and 0.2 µM mesoxalonitrile 3‐chlorophenylhydrazone (CCCP) as an uncoupler, 10 U horseradish peroxidase and 5 µM scopoletin.” (</td> <td> 0, 10, 30  μM Cd2+ 2  μM antimycin A</td> <td> </td> </tr> <tr> <td> Flow Cytometry ROS & Cell Viability (Kruiderig et al., 1997)</td> <td>“For determination of ROS, samples taken at the indicated time points were directly transferred to FACScan tubes. Dih123 (10 mM, final concentration) was added and cells were incubated at 37°C in a humidified atmosphere (95% air/5% CO2) for 10 min. At t 5 9, propidium iodide (10 mM, final concentration) was added, and cells were analyzed by flow cytometry at 60 ml/min. Nonfluorescent Dih123 is cleaved by ROS to fluorescent R123 and detected by the FL1 detector as described above for Dc (Van de Water 1995)”</td> <td> </td> <td> Strong/easy medium</td> </tr> <tr> <td> DCFH-DA Assay Detection of hydrogen peroxide production (Yuan et al., 2016) </td> <td> Intracellular ROS production was measured using DCFH-DA as a probe. Hydrogen peroxide oxidizes DCFH to DCF. The probe is hydrolyzed intracellularly to DCFH carboxylate anion. No direct reaction with H2O2 to form fluorescent production.    </td> <td>0-400 µM</td> <td> Long/ Easy High accuracy </td> </tr> <tr> <td> H2-DCF-DA Assay Detection of superoxide production (Thiebault et al., 2007) </td> <td>This dye is a stable nonpolar compound which diffuses readily into the cells and yields H2-DCF. Intracellular OH or ONOO- react with H2-DCF when cells contain peroxides, to form the highly fluorescent compound DCF, which effluxes the cell. Fluorescence intensity of DCF is measured using a fluorescence spectrophotometer.</td> <td>0–600 µM</td> <td> Long/ Easy High accuracy </td> </tr> <tr> <td>CM-H2DCFDA Assay</td> <td>**Come back and explain the flow cytometry determination of oxidative stress from Pan et al. (2009)**</td> <td> </td> <td> </td> </tr> </tbody> </table> Direct Methods of Measurement <table cellspacing="0" class="Table" style="border-collapse:collapse; width:623px"> <tbody> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:1px solid black; vertical-align:top; width:141px"> Method of Measurement  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; vertical-align:top; width:151px"> References  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; vertical-align:top; width:255px"> Description  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; vertical-align:top; width:76px"> OECD-Approved Assay </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; vertical-align:top; width:141px"> Chemiluminescence  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:151px"> (Lu, C. et al., 2006;  Griendling, K. K., et al., 2016) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:255px"> ROS can induce electron transitions in molecules, leading to electronically excited products. When the electrons transition back to ground state, chemiluminescence is emitted and can be measured. Reagents such as uminol and lucigenin are commonly used to amplify the signal.  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:76px"> No   </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; vertical-align:top; width:141px"> Spectrophotometry  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:151px"> (Griendling, K. K., et al., 2016) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:255px"> NO has a short half-life. However, if it has been reduced to nitrite (NO2-), stable azocompounds can be formed via the Griess Reaction, and further measured by spectrophotometry.  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:76px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; vertical-align:top; width:141px"> Direct or Spin Trapping-Based Electron Paramagnetic Resonance (EPR) Spectroscopy  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:151px"> (Griendling, K. K., et al., 2016) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:255px"> The unpaired electrons (free radicals) found in ROS can be detected with EPR, and is known as electron paramagnetic resonance. A variety of spin traps can be used.  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:76px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; vertical-align:top; width:141px"> Nitroblue Tetrazolium Assay  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:151px"> (Griendling, K. K., et al., 2016) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:255px"> The Nitroblue Tetrazolium assay is used to measure O2•– levels. O2•– reduces nitroblue tetrazolium (a yellow dye) to formazan (a blue dye), and can be measured at 620 nm.  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:76px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; vertical-align:top; width:141px"> Fluorescence analysis of dihydroethidium (DHE) or Hydrocyans  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:151px"> (Griendling, K. K., et al., 2016) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:255px"> Fluorescence analysis of DHE is used to measure O2•– levels. O2•–  is reduced to O2 as DHE is oxidized to 2-hydroxyethidium, and this reaction can be measured by fluorescence. Similarly, hydrocyans can be oxidized by any ROS, and measured via fluorescence.  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:76px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; vertical-align:top; width:141px"> Amplex Red Assay  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:151px"> (Griendling, K. K., et al., 2016) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:255px"> Fluorescence analysis to measure extramitochondrial or extracellular H2O2 levels. In the presence of horseradish peroxidase and H2O2, Amplex Red is oxidized to resorufin, a fluorescent molecule measurable by plate reader.  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:76px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; vertical-align:top; width:141px"> Dichlorodihydrofluorescein Diacetate (DCFH-DA)  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:151px"> (Griendling, K. K., et al., 2016) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:255px"> An indirect fluorescence analysis to measure intracellular H2O2 levels. H2O2 interacts with peroxidase or heme proteins, which further react with DCFH, oxidizing it to dichlorofluorescein (DCF), a fluorescent product.  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:76px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; vertical-align:top; width:141px"> HyPer Probe  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:151px"> (Griendling, K. K., et al., 2016) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:255px"> Fluorescent measurement of intracellular H2O2 levels. HyPer is a genetically encoded fluorescent sensor that can be used for in vivo and in situ imaging.  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:76px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; vertical-align:top; width:141px"> Cytochrome c Reduction Assay  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:151px"> (Griendling, K. K., et al., 2016) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:255px"> The cytochrome c reduction assay is used to measure O2•– levels. O2•–  is reduced to O2 as ferricytochrome c is oxidized to ferrocytochrome c, and this reaction can be measured by an absorbance increase at 550 nm.  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:76px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; vertical-align:top; width:141px"> Proton-electron double-resonance imagine (PEDRI) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:151px"> (Griendling, K. K., et al., 2016) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:255px"> The redox state of tissue is detected through nuclear magnetic resonance/magnetic resonance imaging, with the use of a nitroxide spin probe or biradical molecule.  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:76px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; vertical-align:top; width:141px"> Glutathione (GSH) depletion  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:151px"> (Biesemann, N. et al., 2018)  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:255px"> A downstream target of the Nrf2 pathway is involved in GSH synthesis. As an indication of oxidation status, GSH can be measured by assaying the ratio of reduced to oxidized glutathione (GSH:GSSG) using a commercially available kit (e.g., <a href="http://www.abcam.com/gshgssg-ratio-detection-assay-kit-fluorometric-green-ab138881.html">http://www.abcam.com/gshgssg-ratio-detection-assay-kit-fluorometric-green-ab138881.html</a>).   </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:76px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; vertical-align:top; width:141px"> Thiobarbituric acid reactive substances (TBARS)  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:151px"> (Griendling, K. K., et al., 2016) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:255px"> Oxidative damage to lipids can be measured by assaying for lipid peroxidation with TBARS using a commercially available kit.   </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:76px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; vertical-align:top; width:141px"> Protein oxidation (carbonylation) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:151px"> (Azimzadeh et al., 2017; Azimzadeh etal., 2015; Ping et al., 2020) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:255px"> Can be determined with enzyme-linked immunosorbent assay (ELISA) or a commercial assay kit. Protein oxidation can indicate the level of oxidative stress. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:76px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; vertical-align:top; width:141px">Seahorse XFp Analyzer   </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:151px">Leung et al. 2018   </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:255px">The Seahorse XFp Analyzer provides information on mitochondrial function, oxidative stress, and metabolic dysfunction of viable cells by measuring respiration (oxygen consumption rate; OCR) and extracellular pH (extracellular acidification rate; ECAR).   </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:76px">No </td> </tr> </tbody> </table> Molecular Biology: Nrf2. Nrf2’s transcriptional activity is controlled post-translationally by oxidation of Keap1. Assays for Nrf2 activity include:  <table cellspacing="0" class="Table" style="border-collapse:collapse; width:623px"> <tbody> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:1px solid black; vertical-align:top; width:154px"> Method of Measurement  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; vertical-align:top; width:139px"> References  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; vertical-align:top; width:256px"> Description  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; vertical-align:top; width:75px"> OECD-Approved Assay </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; vertical-align:top; width:154px"> Immunohistochemistry  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:139px"> (Amsen, D., de Visser, K. E., and Town, T., 2009) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:256px"> Immunohistochemistry for increases in Nrf2 protein levels and translocation into the nucleus   </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:75px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; vertical-align:top; width:154px"> Quantitative polymerase chain reaction (qPCR)  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:139px"> (Forlenza et al., 2012) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:256px"> qPCR of Nrf2 target genes (e.g., Nqo1, Hmox-1, Gcl, Gst, Prx, TrxR, Srxn), or by commercially available pathway-based qPCR array (e.g., oxidative stress array from SABiosciences)  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; vertical-align:top; width:75px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:46px; vertical-align:top; width:154px"> Whole transcriptome profiling via microarray or via RNA-seq followed by a pathway analysis </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:46px; vertical-align:top; width:139px"> (Jackson, A. F. et al., 2014) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:46px; vertical-align:top; width:256px"> Whole transcriptome profiling by microarray or RNA-seq followed by pathway analysis (in IPA, DAVID, metacore, etc.) for enrichment of the Nrf2 oxidative stress response pathway </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:46px; vertical-align:top; width:75px"> No </td> </tr> </tbody> </table>

Taxonomic applicability: Occurrence of oxidative stress is not species specific.   Life stage applicability: Occurrence of oxidative stress is not life stage specific.  Sex applicability: Occurrence of oxidative stress is not sex specific.  Evidence for perturbation by prototypic stressor: There is evidence of the increase of oxidative stress following perturbation from a variety of stressors including exposure to ionizing radiation and altered gravity (Bai et al., 2020; Ungvari et al., 2013; Zhang et al., 2009).

High Mixed

High

All life stages

High High

Ahmad, S. et al. (2021), “60Co-γ Radiation Alters Developmental Stages of Zeugodacus cucurbitae (Diptera: Tephritidae) Through Apoptosis Pathways Gene Expression”, Journal Insect Science, Vol. 21/5, Oxford University Press, Oxford, <a href="https://doi.org/10.1093/jisesa/ieab080" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1093/jisesa/ieab080</a> Antelmann, H. and J. D. Helmann (2011), “Thiol-based redox switches and gene regulation.”, Antioxidants & Redox Signaling, Vol. 14/6, Mary Ann Leibert Inc., Larchmont, <a href="https://doi.org/10.1089/ars.2010.3400" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1089/ars.2010.3400</a> Amsen, D., de Visser, K. E., and Town, T. (2009), “Approaches to determine expression of inflammatory cytokines”, in Inflammation and Cancer, Humana Press, Totowa, <a href="https://doi.org/10.1007/978-1-59745-447-6_5" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1007/978-1-59745-447-6_5</a>  Azimzadeh, O. et al. (2015), “Integrative Proteomics and Targeted Transcriptomics Analyses in Cardiac Endothelial Cells Unravel Mechanisms of Long-Term Radiation-Induced Vascular Dysfunction”, Journal of Proteome Research, Vol. 14/2, American Chemical Society, Washington, <a href="https://doi.org/10.1021/pr501141b" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1021/pr501141b</a> Azimzadeh, O. et al. (2017), “Proteome analysis of irradiated endothelial cells reveals persistent alteration in protein degradation and the RhoGDI and NO signalling pathways”, International Journal of Radiation Biology, Vol. 93/9, Informa, London, <a href="https://doi.org/10.1080/09553002.2017.1339332" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1080/09553002.2017.1339332</a> Azzam, E. I. et al. (2012), “Ionizing radiation-induced metabolic oxidative stress and prolonged cell injury”, Cancer Letters, Vol. 327/1-2, Elsevier, Ireland, https://doi.org/10.1016/j.canlet.2011.12.012  Bai, J. et al. (2020), “Irradiation-induced senescence of bone marrow mesenchymal stem cells aggravates osteogenic differentiation dysfunction via paracrine signaling”, American Journal of Physiology - Cell Physiology, Vol. 318/5, American Physiological Society, Rockville, <a href="https://doi.org/10.1152/ajpcell.00520.2019." style="color:#0563c1; text-decoration:underline">https://doi.org/10.1152/ajpcell.00520.2019.</a> Balasubramanian, D (2000), “Ultraviolet radiation and cataract”, Journal of ocular pharmacology and therapeutics, Vol. 16/3, Mary Ann Liebert Inc., Larchmont, <a href="https://doi.org/10.1089/jop.2000.16.285.%22%20/t%20%22_blank" rel="noreferrer noopener" target="_blank">https://doi.org/10.1089/jop.2000.16.285.</a>  Biesemann, N. et al., (2018), “High Throughput Screening of Mitochondrial Bioenergetics in Human Differentiated Myotubes Identifies Novel Enhancers of Muscle Performance in Aged Mice”, Scientific Reports, Vol. 8/1, Nature Portfolio, London, <a href="https://doi.org/10.1038/s41598-018-27614-8" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1038/s41598-018-27614-8</a>.  Elgazzar, A. and N. Kazem. (2015), “Chapter 23: Biological effects of ionizing radiation” in The Pathophysiologic Basis of Nuclear Medicine, Springer, New York, pp. 540-548 Fletcher, A. E (2010), “Free radicals, antioxidants and eye diseases: evidence from epidemiological studies on cataract and age-related macular degeneration”, Ophthalmic Research, Vol. 44, Karger International, Basel, <a href="https://doi.org/10.1159/000316476.%22%20/t%20%22_blank" rel="noreferrer noopener" target="_blank">https://doi.org/10.1159/000316476.</a>   Forlenza, M. et al. (2012), “The use of real-time quantitative PCR for the analysis of cytokine mRNA levels” in Cytokine Protocols, Springer, New York, <a href="https://doi.org/10.1007/978-1-61779-439-1_2" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1007/978-1-61779-439-1_2</a>  Forrester, S.J. et al. (2018), “Angiotensin II Signal Transduction: An Update on Mechanisms of Physiology and Pathophysiology”, Physiological Reviews, Vol. 98/3, American Physiological Society, Rockville, <a href="https://doi.org/10.1152/physrev.00038.201" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1152/physrev.00038.201</a> Foyer, C. H., A. V. Ruban, and G. Noctor (2017), “Viewing oxidative stress through the lens of oxidative signalling rather than damage”, Biochemical Journal, Vol. 474/6, Portland Press, England, https://doi.org/10.1042/BCJ20160814  Ganea, E. and J. J. Harding (2006), “Glutathione-related enzymes and the eye”, Current eye research, Vol. 31/1, Informa, London, <a href="https://doi.org/10.1080/02713680500477347.%22%20/t%20%22_blank" rel="noreferrer noopener" target="_blank">https://doi.org/10.1080/02713680500477347.</a>  Griendling, K. K. et al. (2016), “Measurement of reactive oxygen species, reactive nitrogen species, and redox-dependent signaling in the cardiovascular system: a scientific statement from the American Heart Association”, Circulation research, Vol. 119/5, Lippincott Williams & Wilkins, Philadelphia, <a href="https://doi.org/10.1161/RES.0000000000000110" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1161/RES.0000000000000110</a> Guo, C. et al. (2013), “Oxidative stress, mitochondrial damage and neurodegenerative diseases”, Neural regeneration research, Vol. 8/21, Publishing House of Neural Regeneration Research, China, <a href="https://doi.org/10.3969/j.issn.1673-5374.2013.21.009" style="color:#0563c1; text-decoration:underline">https://doi.org/10.3969/j.issn.1673-5374.2013.21.009</a> Hargreaves, M., and L. L. Spriet (2020), “Skeletal muscle energy metabolism during exercise.”, Nature Metabolism, Vol. 2, Nature Portfolio, London, <a href="https://doi.org/10.1038/s42255-020-0251-4" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1038/s42255-020-0251-4</a> Hladik, D. and S. Tapio (2016), “Effects of ionizing radiation on the mammalian brain”, Mutation Research/Reviews in Mutation Research, Vol. 770, Elsevier, Amsterdam, <a href="https://doi.org/10.1016/j.mrrev.2016.08.003" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1016/j.mrrev.2016.08.003</a> Itoh, K., J. Mimura and M. Yamamoto (2010), “Discovery of the negative regulator of Nrf2, Keap1: a historical overview”, Antioxidants & Redox Signaling, Vol. 13/11, Mary Ann Leibert Inc., Larchmont, <a href="https://doi.org/10.1089/ars.2010.3222" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1089/ars.2010.3222</a> Jackson, A.F. et al. (2014), “Case study on the utility of hepatic global gene expression profiling in the risk assessment of the carcinogen furan.”, Toxicology and Applied Pharmacology, Vol. 274/11, Elsevier, Amsterdam, <a href="https://doi.org/10.1016/j.taap.2013.10.019" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1016/j.taap.2013.10.019</a> Jacobsen, N.R. et al. (2008), “Genotoxicity, cytotoxicity, and reactive oxygen species induced by single-walled carbon nanotubes and C60 fullerenes in the FE1-MutaTM Mouse lung epithelial cells”, Environmental and Molecular Mutagenesis, Vol. 49/6, John Wiley & Sons, Inc., Hoboken, <a href="https://doi.org/10.1002/em.20406" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1002/em.20406</a> Karimi, N. et al. (2017), “Radioprotective effect of hesperidin on reducing oxidative stress in the lens tissue of rats”, International Journal of Pharmaceutical Investigation, Vol. 7/3, Phcog Net, Bengaluru, <a href="https://doi.org/10.4103/jphi.JPHI_60_17.%E2%80%AF" rel="noreferrer noopener" target="_blank">https://doi.org/10.4103/jphi.JPHI_60_17.</a> Leung, D.T.H., and Chu, S. (2018), “Measurement of Oxidative Stress: Mitochondrial Function Using the Seahorse System” In: Murthi, P., Vaillancourt, C. (eds) Preeclampsia. Methods in Molecular Biology, vol 1710. Humana Press, New York, NY. https://doi.org/10.1007/978-1-4939-7498-6_22  Lu, C., G. Song, and J. Lin (2006), “Reactive oxygen species and their chemiluminescence-detection methods”, TrAC Trends in Analytical Chemistry, Vol. 25/10, Elsevier, Amsterdam, <a href="https://doi.org/10.1016/j.trac.2006.07.007" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1016/j.trac.2006.07.007</a> Nguyen Dinh Cat, A. et al. (2013), “Angiotensin II, NADPH oxidase, and redox signaling in the vasculature”, Antioxidants & redox signaling, Vol. 19/10, Mary Ann Liebert, Larchmont, <a href="https://doi.org/10.1089/ars.2012.4641" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1089/ars.2012.4641</a> Ping, Z. et al. (2020), “Oxidative Stress in Radiation-Induced Cardiotoxicity”, Oxidative Medicine and Cellular Longevity, Vol. 2020, Hindawi, <a href="https://doi.org/10.1155/2020/3579143" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1155/2020/3579143</a> Powers, S.K. and M.J. Jackson. (2008), “Exercise-Induced Oxidative Stress: Cellular Mechanisms and Impact on Muscle Force Production”, Physiological Reviews, Vol. 88/4, American Physiological Society, Rockville, <a href="https://doi.org/10.1152/physrev.00031.2007" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1152/physrev.00031.2007</a> Raimondi, V., F. Ciccarese and V. Ciminale. (2020), “Oncogenic pathways and the electron transport chain: a dangeROS liason”, British Journal of Cancer, Vol. 122/2, Nature Portfolio, London, <a href="https://doi.org/10.1038/s41416-019-0651-y" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1038/s41416-019-0651-y</a> Seen, S. and L. Tong. (2018), “Dry eye disease and oxidative stress”, Acta Ophthalmologica, Vol. 96/4, John Wiley & Sons, Inc., Hoboken, <a href="https://doi.org/10.1111/aos.13526" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1111/aos.13526</a> Ungvari, Z. et al. (2013), “Ionizing Radiation Promotes the Acquisition of a Senescence-Associated Secretory Phenotype and Impairs Angiogenic Capacity in Cerebromicrovascular Endothelial Cells: Role of Increased DNA Damage and Decreased DNA Repair Capacity in Microvascular Radiosensitivity”, The Journals of Gerontology Series A: Biological Sciences and Medical Sciences, Vol. 68/12, Oxford University Press, Oxford, <a href="https://doi.org/10.1093/gerona/glt057." style="color:#0563c1; text-decoration:underline">https://doi.org/10.1093/gerona/glt057.</a>   Vargas-Mendoza, N. et al. (2021), “Oxidative Stress, Mitochondrial Function and Adaptation to Exercise: New Perspectives in Nutrition”, Life, Vol. 11/11, Multidisciplinary Digital Publishing Institute, Basel, <a href="https://doi.org/10.3390/life11111269" style="color:#0563c1; text-decoration:underline">https://doi.org/10.3390/life11111269</a> Wang, H. et al. (2019), “Radiation-induced heart disease: a review of classification, mechanism and prevention”, International Journal of Biological Sciences, Vol. 15/10, Ivyspring International Publisher, Sydney, <a href="https://doi.org/10.7150/ijbs.35460" style="color:#0563c1; text-decoration:underline">https://doi.org/10.7150/ijbs.35460</a> Zhang, R. et al. (2009), “Blockade of AT1 receptor partially restores vasoreactivity, NOS expression, and superoxide levels in cerebral and carotid arteries of hindlimb unweighting rats”, Journal of applied physiology, Vol. 106/1, American Physiological Society, Rockville, <a href="https://doi.org/10.1152/japplphysiol.01278.2007" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1152/japplphysiol.01278.2007</a>. Zhao, R. Z. et al. (2019), “Mitochondrial electron transport chain, ROS generation and uncoupling”, International journal of molecular medicine, Vol. 44/1, Spandidos Publishing Ltd., Athens, <a href="https://doi.org/10.3892/ijmm.2019.4188" style="color:#0563c1; text-decoration:underline">https://doi.org/10.3892/ijmm.2019.4188</a> AOPWiki 2017-05-30T13:58:17

2023-03-21T15:16:10

Increased Pro-inflammatory mediators

Increased pro-inflammatory mediators

Tissue

Inflammatory mediators are soluble, diffusible molecules that act locally at the site of tissue damage and infection, and at more distant sites. They can be divided into exogenous and endogenous mediators. Exogenous mediators of inflammation are bacterial products or toxins like endotoxin or LPS. Endogenous mediators of inflammation are produced from within the (innate and adaptive) immune system itself, as well as other systems. They can be derived from molecules that are normally present in the plasma in an inactive form, such as peptide fragments of some components of complement, coagulation, and kinin systems. Or they can be released at the site of injury by a number of cell types that either contain them as preformed molecules within storage granules, e.g. histamine, or which can rapidly switch on the machinery required to synthesize the mediators. Table1: a non-exhaustive list of examples for pro-inflammatory mediators <table border="1" cellpadding="0" cellspacing="0"> <tbody> <tr> <td style="width:253px"> Classes of inflammatory mediators </td> <td style="width:361px"> Examples </td> </tr> <tr> <td style="width:253px"> Pro-inflammatory cytokines </td> <td style="width:361px"> TNF-a, Interleukins (IL-1, IL-6, IL-8), Interferons  (IFN-g), chemokines (CXCL, CCL, GRO-α, MCP-1), GM-CSF </td> </tr> <tr> <td style="width:253px"> Prostaglandins </td> <td style="width:361px"> PGE2 </td> </tr> <tr> <td style="width:253px"> Bradykinin </td> <td style="width:361px">   </td> </tr> <tr> <td style="width:253px"> Vasoactive amines </td> <td style="width:361px"> histamine, serotonin </td> </tr> <tr> <td style="width:253px"> Reactive oxygen species (ROS) </td> <td style="width:361px"> O2-, H2O2 </td> </tr> <tr> <td style="width:253px"> Reactive nitrogen species (RNS) </td> <td style="width:361px"> NO, iNOS </td> </tr> </tbody> </table> The increased production of pro-inflammatory mediators can have negative consequences on the parenchymal cells leading even to cell death, as described for TNF-a or peroxynitrite on neurons (Chao et al., 1995; Brown and Bal-Price, 2003). Along with TNF-α, IL-1β and IL-6 have been shown to exhibit negative consequences on neurogenesis and neuronal precursor cell proliferation when overexpressed. IFN-γ is also associated with neuronal damage, although it is not as extensively studied compared to TNF-α, IL-1β and IL-6. In addition, via a feedback loop, they can act on the reactive resident cells thus maintaining or exacerbating their reactive state; and by modifying elements of their signalling pathways, they can favour the M1 phenotypic polarization and the chronicity of the inflammatory process (Taetzsch et al., 2015). Basically, this event occurs equally in various tissues and does not require tissue-specific descriptions. Nevertheless, there are some specificities such as the release of glutamate by brain reactive glial cells (Brown and Bal-Price, 2003; Vesce et al., 2007).The differences may rather reside in the type of insult favouring the increased expression and/or release of a specific class of inflammatory mediators, as well the time after the insult reflecting different stages of the inflammatory process. For these reasons, the analyses of the changes of a battery of inflammatory mediators rather than of a single one is a more adequate measurement of this KE. Regulatory examples using the KE CD54 and CD 86 as well as IL-8 expression is used to assess skin sensitization potential (OECD TG 442E). IL-2 expression is used to assess immunotoxicity (and will become an OECD test guideline); for the latter see also doi: 10.1007/s00204-018-2199-7.   LIVER: When activated, resident macrophages (Kupffer cells) release inflammatory mediators including cytokines, chemokines, lysosomal, and proteolytic enzymes and are a main source of TGF-β1 - the most potent pro-fibrogenic cytokine. Following the role of TGF-β is described in more detail. Transforming growth factor β (TGF-β) is a pleiotropic cytokine with potent regulatory and inflammatory activity [Sanjabi et al., 2009; Li and Flavell, 2008a;2008b]. The multi-faceted effects of TGF-β on numerous immune functions are cellular and environmental context dependent [Li et al., 2006]. TGF-β binds to TGF-β receptor II (TGF-βRII) triggering the kinase activity of the cytoplasmic domain that in turn activates TGF-βRI. The activated receptor complex leads to nuclear translocation of Smad molecules, and transcription of target genes [Li et al., 2006a]. The role of TGF-β as an immune modulator of T cell activity is best exemplified by the similarities between TGF-β1 knockout and T cell specific TGF-β receptor II knockout mice [Li et al., 2006b; Marie et al., 2006;Shull et al., 1992]. The animals in both of these models develop severe multi-organ autoimmunity and succumb to death within a few weeks after birth [Li et al., 2006b; Marie et al., 2006; Shull et al., 1992]. In addition, in mice where TGF-β signaling is blocked specifically in T cells, the development of natural killer T (NKT) cells, natural regulatory T (nTreg) cells, and CD8+ T cells was shown to be dependent on TGF-β signaling in the thymus [Li et al., 2006b; Marie et al., 2006]. TGF-β plays a major role under inflammatory conditions. TGF-β in the presence of IL-6 drives the differentiation of T helper 17 (Th17) cells, which can promote further inflammation and augment autoimmune conditions [Korn et al., 2009]. TGF-β orchestrates the differentiation of both Treg and Th17 cells in a concentration-dependent manner [Korn et al., 2008]. In addition, TGF-β in combination with IL-4, promotes the differentiation of IL-9- and IL-10-producing T cells, which lack suppressive function and also promote tissue inflammation [Dardalhon  et al., 2008; Veldhoen et al., 2008]. The biological effects of TGF-β under inflammatory conditions on effector and memory CD8+ T cells are much less understood. In a recent study, it was shown that TGF-β has a drastically opposing role on naïve compared to antigen-experienced/memory CD8+ T cells [Filippi et al., 2008]. When cultured in vitro, TGF-β suppressed naïve CD8+ T cell activation and IFN-γ production, whereas TGF-β enhanced survival of memory CD8+ T cells and increased the production of IL-17 and IFN-γ [Filippi et al., 2008]. TGF-β also plays an important role in suppressing the cells of the innate immune system. The transforming growth factor beta (TGF-β) family of cytokines are ubiquitous, multifunctional, and essential to survival. They play important roles in growth and development, inflammation and repair, and host immunity. The mammalian TGF-β isoforms (TGF-β1, β2 and β3) are secreted as latent precursors and have multiple cell surface receptors of which at least two mediate signal transduction. Autocrine and paracrine effects of TGF-βs can be modified by extracellular matrix, neighbouring cells and other cytokines. The vital role of the TGF-β family is illustrated by the fact that approximately 50% of TGF-1 gene knockout mice die in utero and the remainder succumb to uncontrolled inflammation after birth. The role of TGF-β in homeostatic and pathogenic processes suggests numerous applications in the diagnosis and treatment of various diseases characterised by inflammation and fibrosis. [Clark and Coker, 1998; Santibañez et al., 2011; Pohlers et al., 2009] Abnormal TGF-β regulation and function are implicated in a growing number of fibrotic and inflammatory pathologies, including pulmonary fibrosis, liver cirrhosis, glomerulonephritis and diabetic nephropathy, congestive heart failure, rheumatoid arthritis, Marfan syndrome, hypertrophic scars, systemic sclerosis, myocarditis, and Crohn’s disease. [Gordon and Globe,2008] TGF-β1 is a polypeptide member of the TGF-β superfamily of cytokines. TGF-β is synthesized as a non-active pro-form, forms a complex with two latent associated proteins latency-associated protein (LAP) and latent TGF- β binding protein (LTBP) and undergoes protolithic cleavage by the endopeptidase furin to generate the mature TGF-β dimer. Among the TGF-βs, six distinct isoforms have been discovered although only the TGF-β1, TGF-β2 and TGF-β3 isoforms are expressed in mammals, and their human genes are located on chromosomes 19q13, 1q41 and 14q24, respectively. Out of the three TGF-β isoforms (β1, β2 and β3) only TGF-β1 was linked to fibrogenesis and is the most potent fibrogenic factor for hepatic stellate cells. [Roberts, 1998; Govinden and Bhoola, 2003]. During fibrogenesis, tissue and blood levels of active TGF-β are elevated and overexpression of TGF-β1 in transgenic mice can induce fibrosis. Additionally, experimental fibrosis can be inhibited by anti-TGF-β treatments with neutralizing antibodies or soluble TGF-β receptors [Qi et al.; 1999; Shek and Benyon , 2004; De Gouville et al., 2005; Chen et al., 2009]. TGF-β1 induces its own mRNA to sustain high levels in local sites of injury. The effects of TGF-β1 are classically mediated by intracellular signalling via Smad proteins. Smads 2 and 3 are stimulatory whereas Smad 7 is inhibitory. [Parsons et al., 2013; Friedman, 2008; Kubiczkova et al., 2012] Smad1/5/8, MAP kinase (mitogen-activated protein) and PI3 kinase are further signalling pathways in different cell types for TGF-β1 effects. TGF-β is found in all tissues, but is particularly abundant in bone, lung, kidney and placental tissue. TGF-β is produced by many, but not all parenchymal cell types, and is also produced or released by infiltrating cells such as lymphocytes, monocytes/macrophages, and platelets. Following wounding or inflammation, all these cells are potential sources of TGF-β. In general, the release and activation of TGF-β stimulates the production of various extracellular matrix proteins and inhibits the degradation of these matrix proteins. [Branton and Kopp, 1999] TGF-β 1 is produced by every leukocyte lineage, including lymphocytes, macrophages, and dendritic cells, and its expression serves in both autocrine and paracrine modes to control the differentiation, proliferation, and state of activation of these immune cells. [Letterio and Roberts; 1998] In the liver TGF-β1 is released by activated Kupffer cells, liver sinusoidal endothelial cells, and platelets; in the further course of events also activated hepatic stellate cells express TGF-β1. Hepatocytes do not produce TGF-β1 but are implicated in intracellular activation of latent TGF-β1. [Roth et al., 1998; Kisseleva and Brenner, 2007; Kisseleva and Brenner, 2008; Poli, 2000; Liu et al., 2006] TGF-β1 is the most established mediator and regulator of epithelial-mesenchymal-transition (EMT) which further contributes to the production of extracellular matrix. It has been shown that TGF-β1 mediates EMT by inducing snail-1 transcription factor and tyrosine phosphorylation of Smad2/3 with subsequent recruitment of Smad4. [Kolios et al., 2006; Bataller and Brenner, 2005; Guo and Friedman,2007; Brenner,2009; Kaimori et al., 2007; Gressner et al., 2002; Kershenobich Stalnikowitz and Weisssbrod, 2003; Li et al., 2008; Matsuoka and Tsukamoto, 1990; Kisseleva and Brenner, 2008; Poli, 200; Parsons et al., 2007; Friedman 2008; Liu et al., 2006] TGF-β1 induces apoptosis and angiogenesis in vitro and in vivo through the activation of vascular endothelial growth factor (VEGF) High levels of VEGF and TGF-β1 are present in many tumors. Crosstalk between the signalling pathways activated by these growth factors controls endothelial cell apoptosis and angiogenesis. [Clark and Coker; 1998]

The specific type of measurement(s) might vary with tissue, environment and context and will need to be described for different tissue contexts  as used within different AOP descriptions. In general, quantification of inflammatory markers can be done by: <ul> <li>qRT-PCR (mRNA expression)</li> <li>ELISA</li> <li>Immunocytochemistry</li> <li>Immunoblotting</li> </ul> For descriptions of techniques, see Falsig 2004; Lund 2006 ; Kuegler 2010; Monnet-Tschudi et al., 2011; Sandström et al., 2014; von Tobel et al.,  2014   LIVER: There are several assays for TGB-β1 measurement available. e.g. Human TGF-β1 ELISA Kit. The Human TGF-β 1 ELISA (Enzyme –Linked Immunosorbent Assay) kit is an in vitro enzyme-linked immunosorbent assay for the quantitative measurement of human TGF-β1 in serum, plasma, cell culture supernatants, and urine. This assay employs an antibody specific for human TGF-β1 coated on a 96-well plate. Standards and samples are pipetted into the wells and TGF-β1 present in a sample is bound to the wells by the immobilized antibody. The wells are washed and biotinylated anti-human TGF-β1 antibody is added. After washing away unbound biotinylated antibody, HRP- conjugated streptavidin is pipetted to the wells. The wells are again washed, a TMB substrate solution is added to the wells and colour develops in proportion to the amount of TGF-β1 bound. The StopSolution changes the colour from blue to yellow, and the intensity of the colour is measured at 450 nm [Mazzieri et al., 2000] Listed below are common methods for detecting the KE, however there may be other comparable methods that are not listed. <table border="1"> <tbody> <tr> <td> Assay  </td> <td> Reference  </td> <td> Description  </td> <td> OECD Approved Assay  </td> </tr> <tr> <td> <ul> <li> RT-qPCR  </li> <li> Q-PCR  </li> </ul> </td> <td> (Veremeyko et al., 2012; Alwine et al, 1977; Forlenza et al., 2012)  </td> <td> Measures mRNA expression of cytokines, chemokines and inflammatory markers   </td> <td> No  </td> </tr> <tr> <td> Immunoblotting (western blotting)  </td> <td> (Lee et al., 2008)  </td> <td> Uses antibodies specific to proteins of interest, can used to detect presence of pro-inflammatory mediators in samples of cell or tissue lysate  </td> <td> No  </td> </tr> <tr> <td> Whole blood stimulation assay  </td> <td> (Thurm & Halsey, 2005)  </td> <td>  Detects inflammatory cytokines in blood  </td> <td> No  </td> </tr> <tr> <td> Imaging tests  </td> <td> (Rollins & Miskolci, 2014)  </td> <td> A qualitative technique using a cytokine specific antibodies and fluorophores can be used to visualize expression patterns, subcellular location of the target and protein-protein interactions.   Common examples include double immunofluorescence confocal microscopy or other molecular imaging modalities.  </td> <td> No  </td> </tr> <tr> <td> Flow-cytometry  </td> <td> (Karanikas et al., 2000)  </td> <td> Detects the intracellular cytokines with stimulation.  </td> <td> No  </td> </tr> <tr> <td> Immunoassays (ex. enzyme-linked immunosorbent assay (ELISA), enzyme-linked immunospot (ELISpot), radioimmunoassay)  </td> <td> (Amsen et al., 2009; Engvall & Perlmann, 1972; Ji & Forsthuber, 2016; Goldsmith, 1975)  </td> <td> Plate based assay technique using antibodies to detect presence of a protein in a liquid sample.   Can be used to identify presence of an inflammatory cytokine of interest especially when in low concentrations.   </td> <td> No  </td> </tr> <tr> <td> Inflammatory cytokine arrays    </td> <td> (Amsen et al., 2009)    </td> <td> Similar to the ELISA, except using a membrane-based rather than plate-based approach. Can be used to measure multiple cytokine targets concurrently.   </td> <td> No  </td> </tr> <tr> <td> Immunohistochemistry (IHC)  </td> <td> (Amsen et al., 2009; Coons et al., 1942)  </td> <td> Immobilized tissue or cell cultures are stained using antibodies for specificity of ligands of interest. Versions of the assays can be used to visualize localization of inflammatory cytokines.   </td> <td> No  </td> </tr> </tbody> </table>

LIVER: Human [Santibañez et al., 2011] Rat [Luckey and Petersen, 2001] Mouse [Nan et al., 2013] BRAIN: Falsig 2004; Lund 2006 ; Kuegler 2010; Monnet-Tschudi et al., 2011; Sandström et al., 2014; von Tobel et al.,  2014   Taxonomic applicability: The inflammatory response and increase of the pro-inflammatory mediators has been observed across species from simple invertebrates such as Daphnia to higher order vertebrates (Weavers & Martin, 2020). Life stage applicability: This key event is not life stage specific (Kalm et al., 2013; Veeraraghan et al., 2011; Hladik & Tapio, 2016).   Sex applicability:  Most studies conducted were on male models, although sex-dependent differences in pro-inflammatory markers have been previously reported (Cekanaviciute et al., 2018; Parihar et al., 2020).   Evidence for perturbation by a prototypic stressor: There is evidence of the increase of pro-inflammatory mediators following perturbation from a variety of stressors including exposure to ionizing radiation. (Abdel-Magied et al., 2019; Cho et al., 2017; Gaber et al., 2003; Ismail et al., 2016; Kim et al. 2002; Lee et al., 2010; Parihar et al., 2018).

Not Specified Unspecific

Not Specified

All life stages

Not Specified Not Specified

Brown GC, Bal-Price A (2003) Inflammatory neurodegeneration mediated by nitric oxide, glutamate, and mitochondria. Mol Neurobiol 27: 325-355 Dong Y, Benveniste EN (2001) Immune Function of Astrocytes. Glia 36: 180-190 Falsig J, Latta M, Leist M. Defined inflammatory states in astrocyte cultures correlation with susceptibility towards CD95-driven apoptosis. J Neurochem. 2004  Jan;88(1):181-93. Falsig J, Pörzgen P, Lund S, Schrattenholz A, Leist M. The inflammatory transcriptome of reactive murine astrocytes and implications for their innate immune function. J Neurochem. 2006 Feb;96(3):893-907. Falsig J, van Beek J, Hermann C, Leist M. Molecular basis for detection of invading pathogens in the brain. J Neurosci Res. 2008 May 15;86(7):1434-47. Hamadi N, Sheikh A, Madjid N, Lubbad L, Amir N, Shehab SA, Khelifi-Touhami F, Adem A: Increased pro-inflammatory cytokines, glial activation and oxidative stress in the hippocampus after short-term bilateral adrenalectomy. BMC Neurosci 2016, 17:61. Kuegler PB, Zimmer B, Waldmann T, Baudis B, Ilmjärv S, Hescheler J, Gaughwin P, Brundin P, Mundy W, Bal-Price AK, Schrattenholz A, Krause KH, van Thriel C, Rao MS, Kadereit S, Leist M. Markers of murine embryonic and neural stem cells, neurons and astrocytes: reference points for developmental neurotoxicity testing. ALTEX. 2010;27(1):17-42 Lund S, Christensen KV, Hedtjärn M, Mortensen AL, Hagberg H, Falsig J, Hasseldam H, Schrattenholz A, Pörzgen P, Leist M. The dynamics of the LPS triggered inflammatory response of murine microglia under different culture and in vivo conditions. J Neuroimmunol. 2006 Nov;180(1-2):71-87. Monnet-Tschudi, F., A. Defaux, et al. (2011). "Methods to assess neuroinflammation." Curr Protoc Toxicol Chapter 12: Unit12 19. Sandstrom von Tobel, J., D. Zoia, et al. (2014). "Immediate and delayed effects of subchronic Paraquat exposure during an early differentiation stage in 3D-rat brain cell cultures." Toxicol Lett. DOI : 10.1016/j.toxlet.2014.02.001 Taetzsch T, Levesque S, McGraw C, Brookins S, Luqa R, Bonini MG, Mason RP, Oh U, Block ML (2015) Redox regulation of NF-kappaB p50 and M1 polarization in microglia. Glia 5, 63:423-440. Vesce S, Rossi D, Brambilla L, Volterra A (2007) Glutamate release from astrocytes in physiological conditions and in neurodegenerative disorders characterized by neuroinflammation. Int Rev Neurobiol. 82 :57-71.  LIVER: <ul style="list-style-type:circle"> <li>Bataller, R. and D.A. Brenner (2005), Liver Fibrosis, J.Clin. Invest, vol. 115, no. 2, pp. 209-218.</li> <li>Branton, M.H. and J.B. Kopp (1999), TGF-beta and fibrosis, Microbes Infect, vol. 1, no. 15, pp. 1349-1365.</li> <li>Brenner, D.A. (2009), Molecular Pathogenesis of Liver Fibrosis, Trans Am Clin Climatol Assoc, vol. 120, pp. 361–368.</li> <li>Cheng, K., N.Yang and R.I. Mahato (2009), TGF-beta1 gene silencing for treating liver fibrosis, Mol Pharm, vol. 6, no. 3, pp. 772–779.</li> <li>Clark, D.A. and R.Coker (1998), Transforming growth factor-beta (TGF-beta), Int J Biochem Cell Biol, vol. 30, no. 3, pp. 293-298.</li> <li>Dardalhon V, Awasthi A, Kwon H, Galileos G, Gao W, Sobel RA, Mitsdoerffer M, Strom TB,</li> <li>De Gouville, A.C. et al. (2005), Inhibition of TGF-beta signaling by an ALK5 inhibitor protects rats from dimethylnitrosamine-induced liver fibrosis, Br J Pharmacol, vol. 145, no. 2, pp. 166–177.</li> <li>Filippi CM, Juedes AE, Oldham JE, Ling E, Togher L, Peng Y, Flavell RA, von Herrath MG, Transforming growth factor-beta suppresses the activation of CD8+ T-cells when naive but promotes their survival and function once antigen experienced: a two-faced impact on autoimmunity. Diabetes 2008;57:2684–2692.</li> <li>Friedman, S.L. (2008), Mechanisms of Hepatic Fibrogenesis, Gastroenterology, vol. 134, no. 6, pp. 1655–1669.</li> <li>Gordon, K.J. and G.C. Blobe (2008), Role of transforming growth factor-β superfamily signalling pathways in human disease, Biochim Biophys Acta, vol. 1782, no. 4, pp. 197–228.</li> <li>Govinden, R. and K.D. Bhoola (2003), Genealogy, expression, and cellular function of transforming growth factor-β, Pharmacol. Ther, vol. 98, no. 2, pp. 257–265.</li> <li>Gressner, A.M. et al. (2002), Roles of TGF-β in hepatic fibrosis. Front Biosci, vol. 7, pp. 793-807.</li> <li>Guo, J. and S.L. Friedman (2007), Hepatic fibrogenesis, Semin Liver Dis, vol. 27, no. 4, pp. 413-426.</li> <li>Kaimori, A. et al. (2007), Transforming growth factor-beta1 induces an epithelial-to-mesenchymal transition state in mouse hepatocytes in vitro, J Biol Chem, vol. 282, no. 30, pp. 22089-22101.</li> <li>Kershenobich Stalnikowitz, D. and A.B. Weisssbrod (2003), Liver Fibrosis and Inflammation. A Review, Annals of Hepatology, vol. 2, no. 4, pp.159-163.</li> <li>Kisseleva T and Brenner DA, (2008), Mechanisms of Fibrogenesis, Exp Biol Med, vol. 233, no. 2, pp. 109-122.</li> <li>Kisseleva, T. and Brenner, D.A. (2007), Role of hepatic stellate cells in fibrogenesis and the reversal of fibrosis, Journal of Gastroenterology and Hepatology, vol. 22, Suppl. 1; pp. S73–S78.</li> <li>Kolios, G., V. Valatas and E. Kouroumalis (2006), Role of Kupffer cells in the pathogenesis of liver disease, World J.Gastroenterol, vol. 12, no. 46, pp. 7413-7420.</li> <li>Korn T, Mitsdoerffer M, Croxford AL, Awasthi A, Dardalhon VA, Galileos G, Vollmar P, Stritesky GL, Kaplan MH, Waisman A, Kuchroo VK, Oukka M., IL-6 controls Th17 immunity in vivo by inhibiting the conversion of conventional T cells into Foxp3+ regulatory T cells, Proceedings of the National Academy of Sciences Nov 2008, 105 (47) 18460-18465; DOI: 10.1073/pnas.0809850105</li> <li>Korn T, Bettelli E, Oukka M, Kuchroo VK. IL-17 and Th17 Cells. Annu Rev Immunol. 2009</li> <li>Kubiczkova, L. et al, (2012), TGF-β - an excellent servant but a bad master, J Transl Med, vol. 10, p. 183.</li> <li>Letterio, J.J. and A.B. Roberts (1998), Regulation of immune responses by TGF-beta, Annu Rev Immunol, vol.16, pp. 137-161.</li> <li>Li MO, Flavell RA. Contextual regulation of inflammation: a duet by transforming growth factor-beta and interleukin-10. Immunity 2008a;28:468–476.</li> <li>Li MO, Flavell RA. TGF-beta: a master of all T cell trades. Cell 2008b;134:392–404.</li> <li>Li MO, Sanjabi S, Flavell RA. Transforming growth factor-beta controls development, homeostasis, and tolerance of T cells by regulatory T cell-dependent and -independent mechanisms. Immunity 2006b;25:455–471.</li> <li>Li MO, Wan YY, Sanjabi S, Robertson AK, Flavell RA. Transforming growth factor-beta regulation of immune responses. Annu Rev Immunol 2006a;24:99–146.</li> <li>Li, Jing-Ting et al. (2008), Molecular mechanism of hepatic stellate cell activation and antifibrotic therapeutic strategies, J Gastroenterol, vol. 43, no. 6, pp. 419–428.</li> <li>Liu, Xingjun et al. (2006), Therapeutic strategies against TGF-beta signaling pathway in hepatic fibrosis. Liver Int, vol.26, no.1, pp. 8-22.</li> <li>Luckey, S.W., and D.R. Petersen (2001), Activation of Kupffer cells during the course of carbon tetrachloride-induced liver injury and fibrosis in rats, Exp Mol Pathol, vol. 71, no. 3, pp. 226-240.</li> <li>Marie JC, Liggitt D, Rudensky AY. Cellular mechanisms of fatal early-onset autoimmunity in mice with the T cell-specific targeting of transforming growth factor-beta receptor. Immunity 2006;25:441–454.</li> <li>Matsuoka, M. and H. Tsukamoto, (1990), Stimulation of hepatic lipocyte collagen production by Kupffer cell-derived transforming growth factor beta: implication for a pathogenetic role in alcoholic liver fibrogenesis, Hepatology, vol. 11, no. 4, pp. 599-605.</li> <li>Mazzieri, R .et al. (2000), Measurements of Active TGF-β Generated by Culture Cells, Methods in Molecular Biology, vol. 142, pp. 13-27.</li> <li>Nan, Y.M. et al. (2013), Activation of peroxisome proliferator activated receptor alpha ameliorates ethanol mediated liver fibrosis in mice, Lipids Health Dis, vol. 12, p.11.</li> <li>Parsons, C.J., M.Takashima and R.A. Rippe (2007), Molecular mechanisms of hepatic fibrogenesis. J Gastroenterol Hepatol, vol. 22, Suppl.1, pp. S79-S84.</li> <li>Pohlers , D. et al. (2009), TGF-β and fibrosis in different organs – molecular pathway imprints, Biochim. Biophys. Acta, vol. 1792, no. 8, pp.746–756.</li> <li>Poli, G. (2000), Pathogenesis of liver fibrosis: role of oxidative stress, Mol Aspects Med, vol. 21, no. 3, pp. 49 – 98.</li> <li>Qi Z et al.(1999),Blockade of type beta transforming growth factor signalling prevents liver fibrosis and dysfunction in the rat, Proc Natl Acad Sci USA, vol. 96, no. 5, pp. 2345-2349.</li> <li>Roberts, A.B. (1998), Molecular and cell biology of TGF-β, Miner Electrolyte Metab, vol. 24, no. 2-3, pp. 111-119.</li> <li>Roth, S., K. Michel and A.M. Gressner (1998), (Latent) transforming growth factor beta in liver parenchymal cells, its injury-dependent release, and paracrine effects on rat HSCs, Hepatology, vol. 27, no. 4, pp. 1003-1012.</li> <li>Sanjabi S, Zenewicz LA, Kamanaka M, Flavell RA. Anti- and Pro-inflammatory Roles of TGF-β, IL-10, and IL-22 In Immunity and Autoimmunity. Current opinion in pharmacology. 2009;9(4):447-453.</li> <li>Santibañez J.F., M. Quintanilla and C. Bernabeu (2011), TGF-β/TGF-β receptor system and its role in physiological and pathological conditions, Clin Sci (Lond), vol. 121, no. 6, pp. 233-251.</li> <li>Shek, F.W. and R.C. Benyon (2004), How can transforming growth factor beta be targeted usefully to combat liver fibrosis? Eur J Gastroenterol Hepatol, vol. 16, no. 2, pp.123-126.</li> <li>Shull MM, Ormsby I, Kier AB, Pawlowski S, Diebold RJ, Yin M, Allen R, Sidman C, Proetzel G,  Calvin D, et al. Targeted disruption of the mouse transforming growth factor-beta 1 gene results in multifocal inflammatory disease. Nature. 1992 Oct 22;359(6397):693-9.</li> <li>Veldhoen M, Uyttenhove C, van Snick J, Helmby H, Westendorf A, Buer J, Martin B, Wilhelm C, Stockinger B. Transforming growth factor-beta 'reprograms' the differentiation of T helper 2 cells and promotes an interleukin 9-producing subset. Nat Immunol 2008;9:1341–1346.</li> </ul>    Abdel-Magied, N., S. M., Shedid and Ahmed, A. G. (2019), “Mitigating effect of biotin against irradiation-induced cerebral cortical and hippocampal damage in the rat brain tissue”, Environmental Science and Pollution Research, Vol. 26/13, Springer, London, <a href="https://doi.org/10.1007/S11356-019-04806-X" rel="noreferrer noopener" target="_blank">https://doi.org/10.1007/S11356-019-04806-X</a>.    Alwine, J. C., D. J. Kemp and G. R. Stark (1977), “Method for detection of specific RNAs in agarose gels by transfer to diazobenzyloxymethyl-paper and hybridization with DNA probes”, Proceedings of the National Academy of Sciences of the United States of America, Vol. 74/12, United States National Academy of Sciences, Washington, D.C., <a href="https://doi.org/10.1073/pnas.74.12.5350" rel="noreferrer noopener" target="_blank">https://doi.org/10.1073/pnas.74.12.5350</a>  Amsen, D., de Visser, K. E., and Town, T. (2009), “Approaches to determine expression of inflammatory cytokines”, in Inflammation and Cancer, Humana Press, Totowa, <a href="https://doi.org/10.1007/978-1-59745-447-6_5" rel="noreferrer noopener" target="_blank">https://doi.org/10.1007/978-1-59745-447-6_5</a> Cekanaviciute, E., S. Rosi and S. Costes. (2018), "Central Nervous System Responses to Simulated Galactic Cosmic Rays", International Journal of Molecular Sciences, Vol. 19/11, Multidisciplinary Digital Publishing Institute (MDPI) AG, Basel,  https://doi.org/10.3390/ijms19113669.   Cho, H. J. et al. (2017), “Role of NADPH Oxidase in Radiation-induced Pro-oxidative and Pro-inflammatory Pathways in Mouse Brain”, International Journal of Radiation Biology, Vol. 93/11, Informa, London, <a href="https://doi.org/10.1080/09553002.2017.1377360" rel="noreferrer noopener" target="_blank">https://doi.org/10.1080/09553002.2017.1377360</a>.    Coons, A. H. et al. (1942), “The Demonstration of Pneumococcal Antigen in Tissues by the Use of Fluorescent Antibody”, The Journal of Immunology, Vol. 45/3, American Association of Immunologists, Minneapolis, pp. 159-169  Engvall, E., and P. Perlmann (1972), “Enzyme-Linked Immunosorbent Assay, Elisa”, The Journal of Immunology, Vol. 109/1, American Association of Immunologists, Minneapolis, pp. 129-135  Fan, L. W. and Y. Pang. (2017), "Dysregulation of neurogenesis by neuroinflammation: Key differences in neurodevelopmental and neurological disorders", Neural Regeneration Research, Vol. 12/3, Wolters Kluwer, Alphen aan den Rijn, <a href="https://doi.org/10.4103/1673-5374.202926" rel="noreferrer noopener" target="_blank">https://doi.org/10.4103/1673-5374.202926</a>.   Forlenza, M. et al. (2012), “The use of real-time quantitative PCR for the analysis of cytokine mRNA levels” in Cytokine Protocols, Springer, New York, https://doi.org/10.1007/978-1-61779-439-1_2   Gaber, M. W. et al. (2003), “Differences in ICAM-1 and TNF-alpha expression between large single fraction and fractionated irradiation in mouse brain”, International Journal of Radiation Biology, Vol. 79/5, Informa, London, <a href="https://doi.org/10.1080/0955300031000114738" rel="noreferrer noopener" target="_blank">https://doi.org/10.1080/0955300031000114738</a>.    Goldsmith, S. J. (1975), "Radioimmunoassay: Review of basic principles", Seminars in Nuclear Medicine, Vol. 5/2, https://doi.org/10.1016/S0001-2998(75)80028-6.  Hladik, D. and S. Tapio. (2016), "Effects of ionizing radiation on the mammalian brain", Mutation Research/Reviews in Mutation Research, Vol. 770, Elsevier B. b., Amsterdam, https://doi.org/10.1016/j.mrrev.2016.08.003.   Ismail, A. F. M., A.A.M. Salem and M.M.T. Eassawy (2016), “Modulation of gamma-irradiation and carbon tetrachloride induced oxidative stress in the brain of female rats by flaxseed oil”, Journal of Photochemistry and Photobiology B: Biology, Vol. 161, Elsevier, Amsterdam, <a href="https://doi.org/10.1016/J.JPHOTOBIOL.2016.04.031" rel="noreferrer noopener" target="_blank">https://doi.org/10.1016/J.JPHOTOBIOL.2016.04.031</a>.  Ji, N. and T. G. Forsthuber. (2014), "ELISPOT Techniques" (pp. 63–71), https://doi.org/10.1007/7651_2014_111.  Kalm, M., K. Roughton and K. Blomgren. (2013), "Lipopolysaccharide sensitized male and female juvenile brains to ionizing radiation", Cell Death & Disease, Vol. 4/12, Nature Publishing Group, Berlin, https://doi.org/10.1038/cddis.2013.482.  Karanikas, V. et al. (2000), “Flow cytometric measurement of intracellular cytokines detects immune responses in MUC1 immunotherapy”, Clinical Cancer Research, Vol. 6/3, American Association for Cancer Research, Philadelphia, pp. 829–837   Kim, S. H. et al. (2002), “Expression of TNF-alpha and TGF-beta 1 in the rat brain after a single high-dose irradiation”, Journal of Korean Medical Science, Vol. 17/2, Korean Medical Association, Seoul, <a href="https://doi.org/10.3346/JKMS.2002.17.2.242" rel="noreferrer noopener" target="_blank">https://doi.org/10.3346/JKMS.2002.17.2.242</a>.    Lee, J. W. et al. (2008), “Neuro-inflammation induced by lipopolysaccharide causes cognitive impairment through enhancement of beta-amyloid generation”, Journal of Neuroinflammation, Vol. 5/1, BioMed Central, London, <a href="https://doi.org/10.1186/1742-2094-5-37" rel="noreferrer noopener" target="_blank">https://doi.org/10.1186/1742-2094-5-37</a>  Lee, W. H. et al. (2010), “Irradiation induces regionally specific alterations in pro-inflammatory environments in rat brain”, International Journal of Radiation Biology, Vol. 86/2, Informa, London, https://doi.org/10.3109/09553000903419346.  Parihar, V. K. et al. (2018), “Persistent nature of alterations in cognition and neuronal circuit excitability after exposure to simulated cosmic radiation in mice”, Experimental Neurology, Vol. 305, Elsevier, Amsterdam, https://doi.org/10.1016/J.EXPNEUROL.2018.03.009.  Parihar, V. K. et al. (2020), "Sex-Specific Cognitive Deficits Following Space Radiation Exposure", Frontiers in Behavioral Neuroscience, Vol. 14, https://doi.org/10.3389/fnbeh.2020.535885.  Rollins, J. and V. Miskolci (2014), “Immunofluorescence and subsequent confocal microscopy of intracellular TNF in human neutrophils” in Cytokines Bioassays, Springer, London, <a href="https://doi.org/10.1007/978-1-4939-0928-5_24" rel="noreferrer noopener" target="_blank">https://doi.org/10.1007/978-1-4939-0928-5_24</a> Thurm, C. W. and J. F. Halsey (2005), “Measurement of Cytokine Production Using Whole Blood”, in Current Protocols in Immunology, John Wiley & Sons, Inc., Hoboken, <a href="https://doi.org/10.1002/0471142735.im0718bs66" rel="noreferrer noopener" target="_blank">https://doi.org/10.1002/0471142735.im0718bs66</a>  Veeraraghavan, J. et al. (2011), "Low-dose γ-radiation-induced oxidative stress response in mouse brain and gut: Regulation by NFκB–MnSOD cross-signaling", Mutation Research/Genetic Toxicology and Environmental Mutagenesis, Vol. 718/1–2, Elsevier, Amsterdam, https://doi.org/10.1016/j.mrgentox.2010.10.006.  Veremeyko, T. et al. (2012), “Detection of microRNAs in microglia by real-time PCR in normal CNS and during neuroinflammation”, Journal of Visualized Experiments: JoVE, Vol. 65, MyJove Corporation, Cambridge, <a href="https://doi.org/10.3791/4097" rel="noreferrer noopener" target="_blank">https://doi.org/10.3791/4097</a> Weavers, H. and P. Martin (2020), “The cell biology of inflammation: From common traits to remarkable immunological adaptations”, Journal of Cell Biology, Vol. 219, Rockefeller University Press, New York, <a href="https://doi.org/10.1083/jcb.202004003" rel="noreferrer noopener" target="_blank">https://doi.org/10.1083/jcb.202004003</a>  AOPWiki 2017-11-28T09:00:54

2023-03-21T15:50:49

Altered Signaling Pathways

Altered Signaling

Molecular

Cells receive, process, and transmit signals to respond to their environment via signaling pathways. Signaling pathways are groups of molecules that work together in a cell to control physiological and metabolic processes. Kinases, for example, are important signaling molecules that can phosphorylate other proteins (Svoboda & Reenstra, 2002). Initiation of signaling pathways is an important component of cellular homeostasis including normal cell development and the response to cellular damage from exposure to external stressors (Esbenshade & Duzic, 2006). Signaling pathways are themselves activated by signals and the same signal can lead to different responses depending on the tissue type (Hamada, et al. 2011; Svoboda & Reenstra, 2002). Examples of signals include the activation of receptors to activate transcriptional targets, induction of receptor-ligand interactions and the initiation of cell-cell contact, or cell-extracellular matrix contact (Hunter, 2000). Many signalling pathways are crucial to intercellular communication via membrane receptors that transduce signals into the cell, while others are activated in an intracellular manner (Svoboda & Reenstra, 2002). Altered signalling (i.e., increase/decrease) can lead to different physiological outcomes, meaning that the directionality of the signaling response, determines the end outcome. For example, increase of the PI3K/Akt/mTOR pathway, which under physiological conditions is responsible for regulating the cell cycle, can lead to increased proliferation and decreased apoptosis. However, a decrease expression of this pathway can lead to an increase in apoptosis and decreased proliferation (Porta et al., 2014; Venkatesulu et al., 2018).

<table cellspacing="0" class="Table" style="border-collapse:collapse; border:none; width:685px"> <tbody> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:1px solid black; height:19px; vertical-align:top; width:144px"> Method of Measurement </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; height:19px; vertical-align:top; width:148px"> Reference </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; height:19px; vertical-align:top; width:255px"> Description </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; height:19px; vertical-align:top; width:137px"> OECD Approved Assay </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:144px"> Kinase assays  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:148px"> (Svoboda & Reenstra, 2002)  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:255px"> Block kinase with inhibitors to monitor the activity of a kinase of interest.  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:137px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:144px"> Cell behaviour assays  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:148px"> (Svoboda & Reenstra, 2002)  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:255px"> Signal transduction events of cells are monitored. Cells are exposed to varying levels of signaling proteins and the resulting actions of a cell are observed (changes in structure, cell shape, matrix binding etc.). </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:137px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:144px"> Ratiometric or single-wavelength dyes  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:148px"> (Svoboda & Reenstra, 2002)  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:255px"> Detects alterations in signal-transduction activities via monitoring changes in detectable wavelengths.  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:137px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:144px"> Fluorescence microscopy/spectroscopy  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:148px"> (Oksvold et al., 2002)    </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:255px"> Measures cell localization, protein interactions, signal propagation, amplification, and integration in the cell in real-time, or upon stimulation.  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:137px"> Yes </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:144px"> Green Fluorescent Protein (GFP)   </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:148px"> (Zaccolo and Pozzan, 2000)  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:255px"> GFP assays act as fluorescent reporters but also as a marker of intracellular signalling events i.e. second messengers Ca2+ and cAMP, or for pH in different various cell compartments  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:137px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:144px"> Fluorescence Resonance Energy Transfer (FRET)  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:148px"> (Bunt and Wouters, 2017)  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:255px"> Assay helps illuminate the interactions between biological molecules   </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:137px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:144px"> Fluorescence recovery after photobleaching (FRAP)  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:148px"> (Svoboda & Reenstra, 2002)  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:255px"> Determines mobility and diffusion of small molecules.  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:137px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:144px"> Immunoprecipitation  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:148px"> (Svoboda & Reenstra, 2002)  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:255px"> Involves isolating and concentrating a particular protein from mixed samples to detect changes in signalling molecule activity.  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:137px"> Chromatin immunoprecipitation approved for analyzing histone modifications </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:144px"> Immunohistochemistry  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:148px"> (Kurien et al., 2011; Svoboda & Reenstra, 2002)  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:255px"> Northern, western and southern blotting techniques can be used to visualize signal transduction events. For example, antibodies with recognition epitopes can be used to locate active configurations or phosphorylated proteins within a cell or cell lysate. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:137px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:144px"> Reverse transcription-quantitative polymerase chain reaction (RT-qPCR) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:148px"> (Veremeyko et al., 2012; Alwine et al, 1977) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:255px"> Measures mRNA expression of the gene of interest. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:137px"> No </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:144px"> Enzyme-linked immunosorbent assay (ELISA) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:148px"> (Amsen et al., 2009; Engvall & Perlmann, 1972) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:255px"> Plate-based assay technique using antibodies to detect presence of a protein in a liquid sample. Can be used to identify presence of a protein of interest especially in when in low concentrations </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:37px; vertical-align:top; width:137px"> No </td> </tr> </tbody> </table>

Taxonomic applicability: Altered signaling is applicable to all animals as cell signaling occurs among animal cells. This includes vertebrates such as humans, mice and rats (Nair et al., 2019). Life stage applicability: This key event is not life stage specific. Sex applicability: This key event is not sex specific. Evidence for perturbation by a stressor: Multiple studies show that signaling pathways can be disrupted by many types of stressors including ionizing radiation and altered gravity (Cheng et al., 2020; Coleman et al., 2021; Su et al., 2020; Yentrapalli et al., 2013).

Low Unspecific

Moderate

All life stages

Moderate Moderate Moderate Alwine, J. C., D. J. Kemp and G. R. Stark (1977), “Method for detection of specific RNAs in agarose gels by transfer to diazobenzyloxymethyl-paper and hybridization with DNA probes”, Proceedings of the National Academy of Sciences of the United States of America, Vol. 74/12, United States National Academy of Sciences, Washington, D.C., <a href="https://doi.org/10.1073/pnas.74.12.5350" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1073/pnas.74.12.5350</a> Amsen, D., de Visser, K. E., and Town, T. (2009), “Approaches to determine expression of inflammatory cytokines”, in Inflammation and Cancer, Humana Press, Totowa, <a href="https://doi.org/10.1007/978-1-59745-447-6_5" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1007/978-1-59745-447-6_5</a> Bunt, G., and F. S. Wouters (2017), “FRET from single to multiplexed signaling events”, Biophysical reviews, Vol. 9, Springer, London, <a href="https://doi.org/10.1007/s12551-017-0252-z" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1007/s12551-017-0252-z</a> Cheng, Y. P. et al. (2017), “Acid sphingomyelinase/ceramide regulates carotid intima-media thickness in simulated weightless rats”, Pflugers Archiv European Journal of Physiology, Vol. 469, Springer, New York, <a href="https://doi.org/10.1007/s00424-017-1969-z" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1007/s00424-017-1969-z</a> Coleman, M. A. et al. (2015), “Low-dose radiation affects cardiac physiology: gene networks and molecular signaling in cardiomyocytes”,  American Journal of Physiology - Heart and Circulatory Physiology, Vol. 309/11, American Physiological Society, Rockville, <a href="https://doi.org/10.1152/ajpheart.00050.2015" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1152/ajpheart.00050.2015</a> Engvall, E., and P. Perlmann (1972), “Enzyme-Linked Immunosorbent Assay, Elisa”, The Journal of Immunology, Vol. 109/1, American Association of Immunologists, Minneapolis, pp. 129-135 Esbenshade, T. A., and E. Duzic (2006), “Overview of signal transduction”, Current Protocols in Pharmacology, Vol. 31/1, John Wiley & Sons, Ltd., Hoboken, <a href="https://doi.org/10.1002/0471141755.ph0201s31" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1002/0471141755.ph0201s31</a> Hamada, N. et al. (2011), “Signaling pathways underpinning the manifestations of ionizing radiation-induced bystander effects”, Current Molecular Pharmacology, Vol. 4/2, Bentham Science Publishers, Sharjah UAE, <a href="https://doi.org/10.2174/1874467211104020079" style="color:#0563c1; text-decoration:underline">https://doi.org/10.2174/1874467211104020079</a> Hunter, T. (2000), “Signaling - 2000 and beyond”, Cell, Vol. 100/1, Cell Press, Cambridge, <a href="https://doi.org/10.1016/s0092-8674(00)81688-8" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1016/s0092-8674(00)81688-8</a> Kurien, B. T. et al. (2011), “An overview of Western blotting for determining antibody specificities for immunohistochemistry”, in Signal Transduction Immunohistochemistry Methods and Protocols, Springer, London, <a href="https://doi.org/10.1007/978-1-61779-024-9_3" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1007/978-1-61779-024-9_3</a> Nair, A. et al. (2019), “Conceptual Evolution of Cell Signaling”, International journal of molecular sciences, Vol. 20/13, Multidisciplinary Digital Publishing Institute, Basel, <a href="https://doi.org/10.3390/ijms20133292" style="color:#0563c1; text-decoration:underline">https://doi.org/10.3390/ijms20133292</a> Oksvold, M. P. et al. (2002), “Fluorescent histochemical techniques for analysis of intracellular signaling”, The Journal of Histochemistry and Cytochemistry, Vol. 50/3, SAGE Publications, Thousand Oaks, <a href="https://doi.org/10.1177/002215540205000301" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1177/002215540205000301</a> Porta, C., C. Paglino and A. Mosca (2014), “Targeting PI3K/Akt/mTOR Signaling in Cancer”, Frontiers in Oncology, Vol. 4, Frontiers Media SA, Lausanne, <a href="https://doi.org/10.3389/fonc.2014.00064" style="color:#0563c1; text-decoration:underline">https://doi.org/10.3389/fonc.2014.00064</a> Su, Y. T. et al. (2020), “Acid sphingomyelinase/ceramide mediates structural remodeling of cerebral artery and small mesenteric artery in simulated weightless rats”, Life Sciences, Vol. 243, Elsevier, Amsterdam, <a href="https://doi.org/10.1016/j.lfs.2019.117253" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1016/j.lfs.2019.117253</a> Svoboda, K. K. and W. R. Reenstra (2002), “Approaches to studying cellular signaling: a primer for morphologists”, The Anatomical record, Vol. 269/2, John Wiley & Sons, Ltd., Hoboken, <a href="https://doi.org/10.1002/ar.10074" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1002/ar.10074</a> Venkatesulu, B. P. et al. (2018), “Radiation-Induced Endothelial Vascular Injury: A Review of Possible Mechanisms”, JACC: Basic to Translational Science, Vol. 3/4, Elsevier, Amsterdam, <a href="https://doi.org/10.1016/j.jacbts.2018.01.014" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1016/j.jacbts.2018.01.014</a> Veremeyko, T. et al. (2012), “Detection of microRNAs in microglia by real-time PCR in normal CNS and during neuroinflammation”, Journal of Visualized Experiments: JoVE, Vol. 65, MyJove Corporation, Cambridge, <a href="https://doi.org/10.3791/4097" style="color:#0563c1; text-decoration:underline">https://doi.org/10.3791/4097</a> Yentrapalli, R. et al. (2013), “The PI3K/Akt/mTOR pathway is implicated in the premature senescence of primary human endothelial cells exposed to chronic radiation”, PloS one, Vol. 8/8, PLOS, San Francisco, <a href="https://doi.org/10.1371/journal.pone.0070024" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1371/journal.pone.0070024</a> Zaccolo, M. and T. Pozzan (2000), “Imaging signal transduction in living cells with GFP-based probes”, IUBMB life, Vol. 49/5, John Wiley & Sons, Ltd., Hoboken, https://doi.org/10.1080/152165400410218 AOPWiki 2022-09-28T11:42:43

2023-03-22T10:18:32

Altered, Nitric Oxide Levels

Cellular

Nitric oxide (NO) is a diffusible molecule produced by many cell types, including endothelial cells, and is responsible for vasodilation (Schulz, Gori & Münzel, 2011; Soloviev & Kizub, 2019). The source of endogenous NO is L-arginine (Burov et al., 2022). Production of NO in the body can occur through nitric oxide synthase (NOS), an enzyme that degrades L-arginine in the presence of oxygen and nicotinamide adenine dinucleotide phosphate (NADPH) (Luiking, Engelen & Deutz, 2010). Tetrahydrobiopterin (BH4) is an important cofactor of NOS, allowing the enzymatic production of NO. A non-enzymatic method to produce NO includes the reduction of nitrite (Luiking, Engelen & Deutz, 2010). NO is constitutively produced by endothelial nitric oxide synthase (eNOS) and neuronal NOS (nNOS), and can be increased by inducible NOS (iNOS) (Powers & Jackson, 2008). Changes in the expression or activity of NOS enzymes can cause changes in NO levels. For example, iNOS is mainly regulated through transcription and its upregulation can result in increased production of NO (Farah, Michel & Balligand, 2018). Also, eNOS can be regulated by Ca2+ concentrations and blood flow shear stress through phosphorylation at Ser1177 (activating) and Thr495 (inhibiting) (Förstermann, 2010).

Without measuring NO levels directly, NOS levels can be used as a proxy to measure NO production. eNOS and iNOS are common points for assessing NO levels indirectly. Decreased NOS protein expression often corresponds to a decrease in NO. However, it is important to note that NOS levels do not perfectly correlate with NO levels. Increased NOS can also decrease NO if paired with a simultaneous increase in ROS, which, through oxidizing the enzyme’s cofactor BH4, causes NOS uncoupling (Forstermann, 2010; Zhang et al., 2009). Uncoupled NOS produces additional ROS that react with NO and reduce its overall abundance. Therefore, in this case, higher levels of NOS correlate to increased quantity of uncoupled NOS and a subsequent drop in NO bioavailability (Soloviev & Kizub, 2019).  <table border="1"> <tbody> <tr> <td> Assay  </td> <td> Reference  </td> <td> Description  </td> <td> OECD Approved Assay  </td> </tr> <tr> <td> Western blotting/immunoblotting  </td> <td> (Hong et al., 2013; Baker et al., 2009; Yan et al., 2020; Zhang et al., 2009; Zhang et al., 2008; Shi et al., 2012; Azimzadeh et al., 2017; Azimzadeh et al., 2015)  </td> <td> Western blotting/immunoblotting is used to determine levels of inducible and endothelial NOS (NO synthesizing enzyme) in its phosphorylated and unphosphorylated forms, as well as nitrotyrosine (an indicator of NO). NOS and nitrotyrosine are detected by antibodies of each protein, visualized using chemiluminescence, and quantified using densitometry.  </td> <td> No  </td> </tr> <tr> <td> Nitric oxide/nitrate/nitrate (NOx) assay kit (Griess assay)  </td> <td> (Azimzadeh et al., 2017; Adbel-Magied & Shedid, 2019; Yan et al., 2020; Cervelli et al., 2017; Siamwala et al., 2010)   </td> <td> Levels of nitrite/nitrate (NOx) are determined using the NO assay kit. Nitrate reductase is used to convert nitrate into nitrite and the Griess reagent is then used to quantify levels of nitrite.  </td> <td> No  </td> </tr> <tr> <td> Immunohistochemical staining  </td> <td> (Fuji et al., 2016)  </td> <td> Uses an antibody to detect and measure levels of eNOS.  </td> <td> No  </td> </tr> <tr> <td> Immunofluorescence  </td> <td> (Hamada et al., 2019)  </td> <td> Uses fluorescent dye-labeled eNOS antibodies to visualize and determine eNOS levels.  </td> <td> No  </td> </tr> <tr> <td> ELISA kit  </td> <td> (Hasan et al., 2020; Azimzadeh et al., 2015)  </td> <td> Used to determine levels of NO and iNOS in serum by immobilizing the target antigen and binding it to associated antibodies linked to reporter enzymes. The activity of the reporter enzymes is then measured to determine levels of NO and iNOS.  </td> <td>   </td> </tr> <tr> <td> 4-amino-5-methylamino-2′,7′-difluorofluorescein diacetate (DAF-FM) fluorescent probe  </td> <td> (Soucy et al., 2011; Soucy et al., 2010)  </td> <td> Used to detect low concentrations of NO by reacting with it to become a fluorescent benzotriazole that can then be visualized and measured.  </td> <td> No  </td> </tr> </tbody> </table>

Taxonomic applicability: Altered nitric oxide is applicable to vertebrates only, as endothelial NO synthase (eNOS) is required for the formation of NO from the amino acid, L-arginine, and only vertebrates have a true endothelial lining (Yano et al., 2007). Life stage applicability: This key event is not life stage specific. Sex applicability: This key event is not sex specific (Soucy et al., 2011; Takeda et al., 2003). Evidence for perturbation by a stressor: Current literature provides ample evidence of external stressors, including ionizing radiation exposure and altered gravity, inducing significant changes to levels of nitric oxide, nitrate, and NO synthase (Soucy et al., 2011; Zhang et al., 2009).

High Male Low Female Moderate Unspecific

Moderate

Adult

Moderate

Not Otherwise Specified

High High Moderate Moderate

Abdel-Magied, N. and S. M. Shedid (2019), “Impact of zinc oxide nanoparticles on thioredoxin-interacting protein and asymmetric dimethylarginine as biochemical indicators of cardiovascular disorders in gamma-irradiated rats”, Environmental Toxicology, Vol. 35/4, John Wiley & Sons, Inc., Hoboken,  <a href="https://doi.org/10.1002/tox.22879" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1002/tox.22879</a> Azimzadeh, O. et al. (2015), “Integrative Proteomics and Targeted Transcriptomics Analyses in Cardiac Endothelial Cells Unravel Mechanisms of Long-Term Radiation-Induced Vascular Dysfunction”, Journal of Proteome Research, Vol. 14/2, American Chemical Society, Washington, <a href="https://doi.org/10.1021/pr501141b" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1021/pr501141b</a> Azimzadeh, O. et al. (2017), “Proteome analysis of irradiated endothelial cells reveals persistent alteration in protein degradation and the RhoGDI and NO signalling pathways”, International Journal of Radiation Biology, Vol. 93/9, Informa, London, <a href="https://doi.org/10.1080/09553002.2017.1339332" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1080/09553002.2017.1339332</a> Baker, J. E. et al. (2009), “10 Gy total body irradiation increases risk of coronary sclerosis, degeneration of heart structure and function in a rat model”, International Journal of Radiation Biology, Vol. 85/12, Informa, London, <a href="https://doi.org/10.3109/09553000903264473" style="color:#0563c1; text-decoration:underline">https://doi.org/10.3109/09553000903264473</a> Burov, O. N. et al. (2022), “Mechanisms of nitric oxide generation in living systems”, Nitric Oxide, Vol. 118, Elsevier, Amsterdam, <a href="https://doi.org/10.1016/j.niox.2021.10.003" rel="noreferrer noopener" target="_blank">https://doi.org/10.1016/j.niox.2021.10.003</a>. Cervelli, T. et al. (2017), “A new natural antioxidant mixture protects against oxidative and DNA damage in endothelial cell exposed to low-dose irradiation”, Oxidative Medicine and Cellular Longevity, Vol. 2017, Hindawi, London, <a href="https://doi.org/10.1155/2017/9085947" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1155/2017/9085947</a> Farah, C., L. Y. M. Michel and J.-L. Balligand. (2018), "Nitric oxide signalling in cardiovascular health and disease", Nature Reviews Cardiology, Vol. 15/5, Springer Nature, London, <a href="https://doi.org/10.1038/nrcardio.2017.224" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1038/nrcardio.2017.224</a>. Förstermann, U. (2010), "Nitric oxide and oxidative stress in vascular disease", Pflügers Archiv - European Journal of Physiology, Vol. 459, Springer Nature, London, <a href="https://doi.org/10.1007/S00424-010-0808-2" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1007/S00424-010-0808-2</a>. Fuji, S. et al. (2016), “Association between endothelial function and micro-vascular remodeling measured by synchrotron radiation pulmonary micro-angiography in pulmonary arterial hypertension”, General Thoracic and Cardiovascular Surgery, Vol. 64/10, Springer, London, <a href="https://doi.org/10.1007/s11748-016-0684-6" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1007/s11748-016-0684-6</a> Hamada, N. et al. (2020), “Ionizing Irradiation Induces Vascular Damage in the Aorta of Wild-Type Mice”, Cancers, Vol. 12/10, Multidisciplinary Digital Publishing Institute, Basel, <a href="https://doi.org/10.3390/CANCERS12103030" style="color:#0563c1; text-decoration:underline">https://doi.org/10.3390/CANCERS12103030</a> Hamada, N. et al. (2022), “Temporal Changes in Sparing and Enhancing Dose Protraction Effects of Ionizing Irradiation for Aortic Damage in Wild-Type Mice”, Cancers, Vol. 14/14, Multidisciplinary Digital Publishing Institute, Basel, https://doi.org/10.3390/cancers1414331 Hasan, H. F., R. R. Radwan and S. M. Galal (2020), “Bradykinin‐potentiating factor isolated from Leiurus quinquestriatus scorpion venom alleviates cardiomyopathy in irradiated rats via remodelling of the RAAS pathway”, Clinical and Experimental Pharmacology and Physiology, Vol. 47/2, Wiley-Blackwell, Hoboken, <a href="https://doi.org/10.1111/1440-1681.13202" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1111/1440-1681.13202</a> Hong, C. W. et al. (2013), “Involvement of inducible nitric oxide synthase in radiation-induced vascular endothelial damage”, Journal of Radiation Research, Vol. 54/6, Oxford University Press, Oxford, <a href="https://doi.org/10.1093/JRR/RRT066" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1093/JRR/RRT066</a> Powers, S. K. and M. J. Jackson. (2008), "Exercise-Induced Oxidative Stress: Cellular Mechanisms and Impact on Muscle Force Production", Physiological Reviews, Vol. 88/4, The American Physiological Society, Rockville, <a href="https://doi.org/10.1152/physrev.00031.2007" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1152/physrev.00031.2007</a>. Schulz, E., T. Gori and T. Münzel. (2011), "Oxidative stress and endothelial dysfunction in hypertension", Hypertension Research, Vol. 34/6, Nature Portfolio, London, <a href="https://doi.org/10.1038/hr.2011.39" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1038/hr.2011.39</a>. Shi, F. et al. (2012), “Effects of Simulated Microgravity on Human Umbilical Vein Endothelial Cell Angiogenesis and Role of the PI3K-Akt-eNOS Signal Pathway”, PLoS ONE, Vol. 7/7, PLOS, San Francisco, <a href="https://doi.org/10.1371/journal.pone.0040365" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1371/journal.pone.0040365</a> Siamwala, J. H. et al. (2010), “Simulated microgravity perturbs actin polymerization to promote nitric oxide-associated migration in human immortalized Eahy926 cells”, Protoplasma, Vol. 242/1, Springer, London, <a href="https://doi.org/10.1007/S00709-010-0114-Z" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1007/S00709-010-0114-Z</a> Soloviev, A. I. and I. V. Kizub. (2019), "Mechanisms of vascular dysfunction evoked by ionizing radiation and possible targets for its pharmacological correction", Biochemical Pharmacology, Vol. 159, Elsevier, Amsterdam, <a href="https://doi.org/10.1016/J.BCP.2018.11.019" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1016/J.BCP.2018.11.019</a> Soucy, K. G. et al. (2010), “Dietary inhibition of xanthine oxidase attenuates radiation-induced endothelial dysfunction in rat aorta”, Journal of Applied Physiology, Vol. 108/5, American Physiological Society, Rockville, <a href="https://doi.org/10.1152/japplphysiol.00946.2009" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1152/japplphysiol.00946.2009</a>. Soucy, K. G. et al. (2011), “HZE 56Fe-ion irradiation induces endothelial dysfunction in rat aorta: Role of xanthine oxidase”, Radiation Research, Vol. 176/4, Radiation Research Society, Bozeman, <a href="https://doi.org/10.1667/RR2598.1" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1667/RR2598.1</a>. Yan, T., et al. (2020), “Ionizing radiation induces BH4 deficiency by downregulating GTP-cyclohydrolase 1, a novel target for preventing and treating radiation enteritis”, Biochemical Pharmacology, Vol. 180, Elsevier, Amsterdam, <a href="https://doi.org/10.1016/J.BCP.2020.114102" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1016/J.BCP.2020.114102</a> Takeda, I., et al. (2013), “Possible Role of Nitric Oxide in Radiation-Induced Salivary Gland Dysfunction”, Radiation Research, Vol. 159/4, BioOne, <a href="https://doi.org/10.1667/0033-7587(2003)159%5b0465:PRONOI%5d2.0.CO;2" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1667/0033-7587(2003)159[0465:PRONOI]2.0.CO;2</a> Yano, K., et al. (2007), “Phenotypic heterogeneity is an evolutionarily conserved feature of the endothelium”, Blood, Vol. 109/2, American Society of Hematology, Washington, D.C., <a href="https://doi.org/10.1182/blood-2006-05-026401" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1182/blood-2006-05-026401</a> Yao, L. et al. (2010), "The role of RhoA/Rho kinase pathway in endothelial dysfunction", Journal of Cardiovascular Disease Research, Vol. 1/4, Elsevier, Amsterdam, <a href="https://doi.org/10.4103/0975-3583.74258" style="color:#0563c1; text-decoration:underline">https://doi.org/10.4103/0975-3583.74258</a> Zhang, R. et al. (2009), “Blockade of AT1 receptor partially restores vasoreactivity, NOS expression, and superoxide levels in cerebral and carotid arteries of hindlimb unweighting rats”, Journal of Applied Physiology, Vol. 106, American Physiological Society, Rockville, <a href="https://doi.org/10.1152/japplphysiol.01278.2007" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1152/japplphysiol.01278.2007</a> AOPWiki 2022-09-28T12:00:33

2023-04-13T09:00:04

Increase, Endothelial Dysfunction

Tissue

The endothelium is the innermost lining of blood vessels consisting of a single layer of endothelial cells. As the layer separating blood and vessel walls, the endothelium controls the flow of molecules, fluid, and circulating blood cells between the two. However, the specific functions and even the structure of endothelial cells vary greatly depending on the organ (Ricard et al., 2021). Dysfunction to the vascular endothelium can age arteries and is the result of increased proliferation and apoptotic behaviour of cells including an increased response to endothelial constrictors. It is also represented by an imbalance between vasodilators and vasoconstrictors which are produced by the endothelium. The dysfunction can encompass vasospasm, thrombosis, penetration of immune cells (i.e macrophage) and an increase in cyclooxygenase. These processes can activate the endothelium and a prolonged state of activation is problematic and is referred to as endothelial dysfunction (Sitia et al., 2010; Deanfield et al., 2005; Konukoglu & Uzun, 2017; Korpela & Liu, 2014). Other factors leading to endothelial dysfunction are loss in endothelial function leading to cell senescence and a low proliferative capacity of endothelial progenitor cells.

Endothelial cell senescence  <table border="1"> <tbody> <tr> <td> Assay  </td> <td> Reference  </td> <td> Description  </td> <td> OECD Approved Assay  </td> </tr> <tr> <td> Senescence-associated beta-galactosidase staining (SA-beta-gal)  </td> <td> (Farhat et al., 2008; González-Gualda et al., 2021; Hooten et al., 2017)  </td> <td> Can be used to measure senescence-associated β-galactosidase activity, a marker for senescent cells.  </td> <td> No  </td> </tr> <tr> <td> Bromodeoxyuridine (BrdU) detected with staining incorporation  </td> <td> (González-Gualda et al., 2021)  </td> <td> Reduced BrdU incorporation can indicate a lack of DNA synthesis.  </td> <td> No  </td> </tr> <tr> <td> Immunohistochemistry to detect senescence markers.  </td> <td> (González-Gualda et al., 2021)  </td> <td> Markers include Ki67 and Lamin B1. Reduced Ki67 can indicate reduced proliferation. Reduced Lamin B1 indicates impaired structural integrity of the nucleus.  </td> <td> No  </td> </tr> <tr> <td> Cell morphology and size measured with light microscopy or flow cytometry.  </td> <td> (González-Gualda et al., 2021)  </td> <td> Senescent cells exhibit an enlarged and flattened morphology.  </td> <td> No  </td> </tr> </tbody> </table>   Cell death:  See the <a href="https://aopwiki.org/events/1825" rel="noreferrer noopener" target="_blank">increase, cell death KE</a> for methods to measure endothelial cell death.    Impaired vasomotion <table border="1"> <tbody> <tr> <td> Assay  </td> <td> Reference  </td> <td> Description  </td> <td> OECD Approved Assay  </td> </tr> <tr> <td> Concentration-response curves to vasodilators/vasoconstrictors  </td> <td> (Deanfield et al., 2005; Verma et al., 2003)  </td> <td> Measurement of endothelial relaxation/contraction of blood vessels can give insight into endothelial dysfunction. This can be induced by endothelium-independent stimuli to stimulate vasodilation or vasoconstriction. A decreased stimuli response can be indicative of endothelial dysfunction.   </td> <td> No  </td> </tr> <tr> <td> Detection of contractile factors (eg. endothelin) using enzyme-linked immunosorbent assay (ELISA).  </td> <td> (Abdel-Sayed et al., 2003)  </td> <td> Endothelin is an endothelium-derived vasoconstrictor.  </td> <td> No  </td> </tr> </tbody> </table>

Taxonomic applicability: Endothelial dysfunction is applicable to vertebrates as only vertebrates have a true endothelial lining (Yano et al., 2007). Life stage applicability: Although endothelial dysfunction may occur due to aging (Hererra et al., 2010), this key event can occur at any life stage (Chang et al., 2017; Lee et al., 2020). Sex applicability: This key event is not sex specific (Hughson et al., 2018; Lee et al., 2020). Evidence for perturbation by a stressor: Multiple studies show that endothelial dysfunction can be triggered by many types of stressors including ionizing radiation and altered gravity (Cheng et al., 2017; Soucy et al., 2011; Su et al., 2020; Yentrapalli et al., 2013).

UBERON:0001986

UBERON endothelium

Moderate Unspecific

Moderate

All life stages

Moderate Moderate Moderate Abdel-Sayed, S. et al. (2003), “Measurement of plasma endothelin-1 in experimental hypertension and in healthy subjects”, American Journal of Hypertension, Vol. 16/7, Oxford University Press, Oxford, <a href="http://doi.org/10.1016/S0895-7061(03)00903-8">https://doi.org/10.1016/S0895-7061(03)00903-8</a> Chang, P. Y. et al. (2017), “MSC-derived cytokines repair radiation-induced intra-villi microvascular injury”, Oncotarget, Vol. 8/50, Impact Journals, Orchard Park, <a href="https://doi.org/10.18632/oncotarget.21236" style="color:#0563c1; text-decoration:underline">https://doi.org/10.18632/oncotarget.21236</a> Cheng, Y. P. et al. (2017), “Acid sphingomyelinase/ceramide regulates carotid intima-media thickness in simulated weightless rats”, Pflugers Archiv European Journal of Physiology, Vol. 469, Springer, New York, <a href="http://doi.org/10.1007/s00424-017-1969-z">https://doi.org/10.1007/s00424-017-1969-z</a> Deanfield, J. et al. (2005), “Endothelial function and dysfunction”, Journal of hypertension, Vol. 23/1, Lippincott Williams & Wilkins, Philadelphia, <a href="https://doi.org/10.1097/00004872-200501000-00004" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1097/00004872-200501000-00004</a> Farhat, N. et al. (2008), “Stress-induced senescence predominates in endothelial cells isolated from atherosclerotic chronic smokers”, Canadian Journal of Physiology and Pharmacology, Vol. 86/11, Canadian Science Publishing, Ottawa, <a href="https://doi.org/10.1139/Y08-082" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1139/Y08-082</a> González-Gualda, E. et al. (2021), “A guide to assessing cellular senescence in vitro and in vivo”, The FEBS Journal, Vol. 288, FEBS press, <a href="http://doi.org/10.1111/febs.15570">https://doi.org/10.1111/febs.15570</a>  Herrera, M. D. et al. (2010), “Endothelial dysfunction and aging: An update”, Ageing Research Reviews, Vol 9/2, Elsevier, Amsterdam, <a href="https://doi.org/10.1016/j.arr.2009.07.002" rel="noreferrer noopener" target="_blank">https://doi.org/10.1016/j.arr.2009.07.002</a> Hooten, N. N. and M. K. Evans (2017), “Techniques to Induce and Quantify Cellular Senescence”, Journal of Visualized Experiments: JoVE, Vol. 123, MyJove Corporation, Cambridge,  <a href="https://doi.org/10.3791/55533" style="color:#0563c1; text-decoration:underline">https://doi.org/10.3791/55533</a> Hughson, R. L., A. Helm and M. Durante (2018), “Heart in space: effect of the extraterrestrial environment on the cardiovascular system”, Nature Reviews Cardiology, Vol. 15/3, Nature Portfolio, London, <a href="https://doi.org/10.1038/nrcardio.2017.157" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1038/nrcardio.2017.157</a> Konukoglu, D., and H. Uzun (2017), “Endothelial Dysfunction and Hypertension”, in Hypertension: from basic research to clinical practice, Springer, London, <a href="https://doi.org/10.1007/5584_2016_90" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1007/5584_2016_90</a> Korpela, E., and S. K. Liu (2014), “Endothelial perturbations and therapeutic strategies in normal tissue radiation damage”, Radiation Oncology, Vol. 9, BioMed Central, London, <a href="https://doi.org/10.1186/s13014-014-0266-7" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1186/s13014-014-0266-7</a> Lee, S. et al.  (2020), “Arterial structure and function during and after long-duration spaceflight”, Journal of Applied Physiology, Vol. 129/1, American Physiological Society, Rockville, <a href="https://doi.org/10.1152/japplphysiol.00550.2019" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1152/japplphysiol.00550.2019</a> Ricard, N. et al. (2021), “The quiescent endothelium: signalling pathways regulating organ-specific endothelial normalcy”, Nature reviews cardiology, Vol. 18/8, Springer Nature, <a href="http://doi.org/10.1038/s41569-021-00517-4 ">https://doi.org/10.1038/s41569-021-00517-4 </a> Sitia, S. et al. (2010), “From endothelial dysfunction to atherosclerosis”, Autoimmunity Reviews, Vol. 9/12, Elsevier, Amsterdam, <a href="http://doi.org/10.1016/j.autrev.2010.07.016">https://doi.org/10.1016/j.autrev.2010.07.016</a> Soucy, K. G. et al. (2011), “HZE 56Fe-ion irradiation induces endothelial dysfunction in rat aorta: Role of xanthine oxidase”, Radiation Research, Vol. 176/4, Radiation Research Society, Bozeman, <a href="https://doi.org/10.1667/RR2598.1" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1667/RR2598.1</a> Su, Y. T. et al. (2020), “Acid sphingomyelinase/ceramide mediates structural remodeling of cerebral artery and small mesenteric artery in simulated weightless rats”, Life Sciences, Vol. 243, Elsevier, Amsterdam, <a href="http://doi.org/10.1016/j.lfs.2019.117253">https://doi.org/10.1016/j.lfs.2019.117253</a> Verma, S., M. R. Buchanan and T. J. Anderson (2003), “Endothelial function testing as a biomarker of vascular disease”, Circulation, Vol. 108/17, Lippincott Williams & Wilkins, Philadelphia, <a href="https://doi.org/10.1161/01.CIR.0000089191.72957.ED" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1161/01.CIR.0000089191.72957.ED</a> Yano, K. et al. (2007), “Phenotypic heterogeneity is an evolutionarily conserved feature of the endothelium”, Blood, Vol. 109/2, American Society of Hematology, Washington, D.C., <a href="https://doi.org/10.1182/blood-2006-05-026401" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1182/blood-2006-05-026401</a> Yentrapalli, R. et al. (2013), “The PI3K/Akt/mTOR pathway is implicated in the premature senescence of primary human endothelial cells exposed to chronic radiation”, PloS one, Vol. 8/8, PLOS, San Francisco, <a href="https://doi.org/10.1371/journal.pone.0070024" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1371/journal.pone.0070024</a> AOPWiki 2022-09-28T12:39:53

2023-03-21T12:17:07

Occurrence, Vascular Remodeling

Organ

The vascular wall is composed of endothelial, smooth muscle and fibroblast cell interactions (Gibbons & Dzau, 1994; Renna, Heras & Miatello, 2013). The vasculature is capable of detecting changes in its surroundings and maintaining homeostasis (Gibbons & Dzau, 1994; Renna, Heras & Miatello, 2013). The functionality of blood vessels is highly dependent on their structure, with changes in arterial morphology being associated with downstream impacts (Gibbons & Dzau, 1994). Vascular remodeling is a term for many histological changes, including increased vascular stiffness, wall shear stress, intima-media thickening (IMT), increased intima-media section area, and increased vessel diameter (Herity et al., 1999). As blood vessels stiffen, this impacts systolic and diastolic pressure and pulse which can be indicators of vascular remodeling. Cellular level changes characterized by processes of growth, death, migration and production or degradation of the extracellular matrix (ECM) result in inflammation (increase in VCAM, ICAM, cytokines, chemokines) and calcification (changes in ratios of collagen and elastin) (Gibbons & Dzau, 1994). Initial tissue injury and resulting remodeling can also lead to turbulent blood flow causing further structural changes like increased vessel fibrosis.  Increased vascular remodelling is often associated with a build-up of plaque in the arteries (known as atherosclerosis) due to impaired healing, which forces the vessel walls to attempt to remodel to maintain blood flow (Sylvester et al., 2018).

Taxonomic applicability: Vascular remodelling is applicable to all species with a closed circulatory system where blood is transported throughout the body via blood vessels with corresponding vessel walls (Renna, Heras & Miatello, 2013). Closed circulatory systems are present in most vertebrates and some invertebrates. Life stage applicability: This key event is not life stage specific. However, advancing age is a risk factor for vascular remodeling (Harvey, Montezano & Touyz, 2015). Sex applicability: This key event is not sex specific. However, men are shown to develop vascular remodeling younger than women (Kessler et al., 2019). Evidence for perturbation by a stressor: Current literature provides ample evidence of vascular remodelling being induced by stressors including ionizing radiation exposure and altered gravity (Shen et al. 2018; Su et al., 2020; Delp et al., 2000, Cheng et al., 2017, Yu et al., 2011; Soucy et al., 2007; Soucy et al., 2010; Soucy et al., 2011).

UBERON:0001981

UBERON blood vessel

Moderate Male Low Female Moderate Unspecific

Moderate

Adult

Moderate

Not Otherwise Specified

Moderate Moderate Moderate

Cheng, Y. P. et al. (2017), "Acid sphingomyelinase/ceramide regulates carotid intima-media thickness in simulated weightless rats", Pflugers Archiv European Journal of Physiology, Vol. 469, Springer Nature, London, <a href="https://doi.org/10.1007/s00424-017-1969-z" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1007/s00424-017-1969-z</a> Delp, M.D. et al. (2000), “Structural and functional remodeling of skeletal muscle microvasculature is induced by simulated microgravity”, American Journal of Physiology - Heart and Circulatory Physiology, Vol. 278, American Physiological Society, Rockville, <a href="https://doi.org/10.1152/ajpheart.2000.278.6.h1866" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1152/ajpheart.2000.278.6.h1866</a> Gibbons, G. H., and V. J. Dzau (1994), “The Emerging Concept of Vascular Remodeling”, New England Journal of Medicine, Vol. 330/20, Massachusetts Medical Society, Waltham, <a href="https://doi.org/10.1056/NEJM199405193302008" rel="noreferrer noopener" target="_blank">https://doi.org/10.1056/NEJM199405193302008</a> Harvey, A., A. C. Montezano, & R. M. Touyz. (2015), “Vascular biology of ageing-Implications in hypertension”, Journal of molecular and cellular cardiology, Vol. 83, Elsevier, Amsterdam, <a href="https://doi.org/10.1016/j.yjmcc.2015.04.011" rel="noreferrer noopener" target="_blank">https://doi.org/10.1016/j.yjmcc.2015.04.011</a> Herity, N.A. et al. (1999), “Review: Clinical Aspects of Vascular Remodeling”, Journal of Cardiovascular Electrophysiology, Vol.10/7, Wiley, <a href="http://doi.org/10.1111/j.1540-8167.1999.tb01273.x">https://doi.org/10.1111/j.1540-8167.1999.tb01273.x</a> Kessler, E. L. et al. (2019), “Sex-specific influence on cardiac structural remodeling and therapy in cardiovascular disease”, Biology of Sex Differences, Vol. 10, Springer Nature, <a href="http://doi.org/10.1186/s13293-019-0223-0">https://doi.org/10.1186/s13293-019-0223-0</a> Lee, S. M. C. et al. (2020), “Arterial structure and function during and after long-duration spaceflight”, Journal of Applied Physiology, Vol. 129, American Physiological Society, Rockville, https://doi.org/10.1152/japplphysiol.00550.2019 Patel, S. (2020), "The effects of microgravity and space radiation on cardiovascular health: From low-Earth orbit and beyond", IJC Heart and Vasculature, Vol. 30, Elsevier, Amsterdam, <a href="https://doi.org/10.1016/j.ijcha.2020.100595" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1016/j.ijcha.2020.100595</a>. Renna, N. F., N. Heras, R. M. Miatello (2013), “Pathophysiology of Vascular Remodeling in Hypertension”, International Journal of Hypertension, Vol. 2013, Hindawi, London, <a href="http://doi.org/10.1155/2013/808353" style="color:#0563c1; text-decoration:underline">http://doi.org/10.1155/2013/808353</a> Sárközy, M. et al. (2019), "Selective heart irradiation induces cardiac overexpression of the pro-hypertrophic miR-212", Frontiers in Oncology, Vol. 9, Frontiers Media S.A., Lausanne, <a href="https://doi.org/10.3389/FONC.2019.00598/FULL" style="color:#0563c1; text-decoration:underline">https://doi.org/10.3389/FONC.2019.00598/FULL</a> Shen, Y. et al. (2018), “Transplantation of bone marrow mesenchymal stem cells prevents radiation-induced artery injury by suppressing oxidative stress and inflammation”, Oxidative Medicine and Cellular Longevity, Vol. 2018, Hindawi, London, <a href="https://doi.org/10.1155/2018/5942916" style="color:#0563c1; text-decoration:underline">https://doi.org/10.1155/2018/5942916</a>. Sofronova, S. I. et al. (2015), “Spaceflight on the Bion-M1 biosatellite alters cerebral artery vasomotor and mechanical properties in mice”, Journal of Applied Physiology, Vol. 118/7, American Physiological Society, Rockville, https://doi.org/10.1152/japplphysiol.00976.2014. Soucy, K. G. et al. (2011), “HZE 56Fe-ion irradiation induces endothelial dysfunction in rat aorta: Role of xanthine oxidase”, Radiation Research, Vol. 176/4, Radiation Research Society, Bozeman, https://doi.org/10.1667/RR2598.1. Soucy, K. G. et al. (2010), “Dietary inhibition of xanthine oxidase attenuates radiation-induced endothelial dysfunction in rat aorta”, Journal of Applied Physiology, Vol. 108/5, American Physiological Society, Rockville, https://doi.org/10.1152/japplphysiol.00946.2009. Soucy, K. G. et al. (2007), “Single exposure gamma-irradiation amplifies xanthine oxidase activity and induces endothelial dysfunction in rat aorta”, Radiation and Environmental Biophysics, Vol. 46, Springer, New York, https://doi.org/10.1007/s00411-006-0090-z. Sridharan, V. et al. (2020), "Effects of single-dose protons or oxygen ions on function and structure of the cardiovascular system in male Long Evans rats", Life Sciences in Space Research, Vol. 26, Elsevier, Amsterdam, https://doi.org/10.1016/j.lssr.2020.04.002. Su, Y. T. et al. (2020), "Acid sphingomyelinase/ceramide mediates structural remodeling of cerebral artery and small mesenteric artery in simulated weightless rats", Life Sciences, Vol. 243, Elsevier, Amsterdam, https://doi.org/10.1016/j.lfs.2019.117253. Sylvester, C. B. et al. (2018), "Radiation-Induced Cardiovascular Disease: Mechanisms and Importance of Linear Energy Transfer", Frontiers in Cardiovascular Medicine, Vol. 5, Frontiers Media SA, Lausanne, https://doi.org/10.3389/fcvm.2018.00005. Tarasova, O. S. et al. (2020), "Simulated Microgravity Induces Regionally Distinct Neurovascular and Structural Remodeling of Skeletal Muscle and Cutaneous Arteries in the Rat", Frontiers in Physiology, Vol. 1, Frontiers Media SA, Lausanne, https://doi.org/10.3389/fphys.2020.00675. Yu, T. et al. (2011), "Iron-ion radiation accelerates atherosclerosis in apolipoprotein E-Deficient mice", Radiation Research, Vol. 175/6, Radiation Research Society, Bozeman, https://doi.org/10.1667/RR2482.1. AOPWiki 2022-10-17T16:49:01

2023-03-21T12:24:23

Increase, DNA strand breaks

Molecular

DNA strand breaks can occur on a single strand (SSB) or both strands (double strand breaks; DSB). SSBs arise when the phosphate backbone connecting adjacent nucleotides in DNA is broken on one strand. DSBs are generated when both strands are simultaneously broken at sites that are sufficiently close to one another that base-pairing and chromatin structure are insufficient to keep the two DNA ends juxtaposed. As a consequence, the two DNA ends generated by a DSB can physically dissociate from one another, becoming difficult to repair and increasing the chance of inappropriate recombination with other sites in the genome (Jackson, 2002). SSB can turn into DSB if the replication fork stalls at the lesion leading to fork collapse. Strand breaks are intermediates in various biological events, including DNA repair (e.g., excision repair), V(D)J recombination in developing lymphoid cells and chromatin remodeling in both somatic cells and germ cells. The spectrum of damage  can be complex, particularily if the stressor is from large amounts of deposited energy which can result in complex lesions and clustered damage defined as two or more oxidized bases, abasic sites or starnd breaks on opposing DNA strands within a few helical turns. These lesions are more difficult to repair and have been studied in many types of models  (Barbieri et al., 2019 and Asaithamby et al., 2011). DSBs and complex lesions  are of particular concern, as they are considered the most lethal and deleterious type of DNA lesion. If misrepaired or left unrepaired, DSBs may drive the cell towards genomic instability, apoptosis or tumorigenesis (Beir, 1999).

Please refer to the table below for details regarding these and other methodologies for detecting DNA DSBs.   <table cellspacing="0" class="Table" style="border-collapse:collapse; border:none; margin-left:15px"> <tbody> <tr> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:1px double #2b2b2b; border-right:1px double #2b2b2b; border-top:1px double #2b2b2b; height:37px; vertical-align:top; width:85px"> Assay Name </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:1px double #2b2b2b; height:37px; vertical-align:top; width:89px"> References </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:1px double #2b2b2b; height:37px; vertical-align:top; width:228px"> Description </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #7f7f7f; border-top:1px double #2b2b2b; height:37px; vertical-align:top; width:46px"> OECD Approved Assay </td> </tr> <tr> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:1px double #2b2b2b; border-right:1px double #2b2b2b; border-top:none; height:61px; vertical-align:top; width:85px"> Comet Assay (Single Cell Gel Eletrophoresis - Alkaline) </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:61px; vertical-align:top; width:89px"> Collins, 2004; Olive and Banath, 2006; Platel et al., 2011; Nikolova et al., 2017 </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:61px; vertical-align:top; width:228px"> To detect SSBs or DSBs, single cells are encapsulated in agarose on a slide, lysed, and subjected to gel electrophoresis at an alkaline pH (pH >13); DNA fragments are forced to move, forming a "comet"-like appearance </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #7f7f7f; border-top:none; height:61px; vertical-align:top; width:46px"> Yes (No. 489) </td> </tr> <tr> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:1px double #2b2b2b; border-right:1px double #2b2b2b; border-top:none; height:61px; vertical-align:top; width:85px"> Comet Assay (Single Cell Gel Eltrophoresis - Neutral) </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:61px; vertical-align:top; width:89px"> Collins, 2014; Olive and Banath, 2006; Anderson and Laubenthal, 2013; Nikolova et al., 2017 </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:61px; vertical-align:top; width:228px"> To detect DSBs, single cells are encapsulated in agarose on a slide, lysed, and subjected to gel electrophoresis at a neutral pH; DNA fragments, which are not denatured at the neutral pH, are forced to move, forming a "comet"-like appearance </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #7f7f7f; border-top:none; height:61px; vertical-align:top; width:46px"> N/A </td> </tr> <tr> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:1px double #2b2b2b; border-right:1px double #2b2b2b; border-top:none; height:37px; vertical-align:top; width:85px"> γ-H2AX Foci Quantification - Flow Cytometry </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:37px; vertical-align:top; width:89px"> Rothkamm and Horn, 2009; Bryce et al., 2016 </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:37px; vertical-align:top; width:228px"> Measurement of γ-H2AX immunostaining in cells by flow cytometry, normalized to total levels of H2AX </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #7f7f7f; border-top:none; height:37px; vertical-align:top; width:46px"> N/A </td> </tr> <tr> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:1px double #2b2b2b; border-right:1px double #2b2b2b; border-top:none; height:37px; vertical-align:top; width:85px"> γ-H2AX Foci Quantification - Western Blot </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:37px; vertical-align:top; width:89px"> Burma et al., 2001; Revet et al., 2011 </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:37px; vertical-align:top; width:228px"> Measurement of γ-H2AX immunostaining in cells by Western blotting, normalized to total levels of H2AX </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #7f7f7f; border-top:none; height:37px; vertical-align:top; width:46px"> N/A </td> </tr> <tr> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:1px double #2b2b2b; border-right:1px double #2b2b2b; border-top:none; height:49px; vertical-align:top; width:85px"> γ-H2AX Foci Quantification - Microscopy </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:49px; vertical-align:top; width:89px"> Redon et al., 2010; Mah et al., 2010; Garcia-Canton et al., 2013 </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:49px; vertical-align:top; width:228px"> Quantification of γ-H2AX immunostaining by counting γ- H2AX foci visualized with a microscope </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #7f7f7f; border-top:none; height:49px; vertical-align:top; width:46px"> N/A </td> </tr> <tr> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:1px double #2b2b2b; border-right:1px double #2b2b2b; border-top:none; height:37px; vertical-align:top; width:85px"> γ-H2AX Foci Detection - ELISA and flow cytometry </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:37px; vertical-align:top; width:89px"> Ji et al., 2017; Bryce et al., 2016 </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:37px; vertical-align:top; width:228px"> Detection of γ-H2AX in cells by ELISA, normalized to total levels of H2AX; γH2AX foci detection can be high-throughput and automated using flow cytometry-based immunodetection. </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #7f7f7f; border-top:none; height:37px; vertical-align:top; width:46px"> N/A </td> </tr> <tr> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:1px double #2b2b2b; border-right:1px double #2b2b2b; border-top:none; height:61px; vertical-align:top; width:85px"> Pulsed Field Gel Electrophoresis (PFGE) </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:61px; vertical-align:top; width:89px"> Ager et al., 1990; Gardiner et al., 1985; Herschleb et al., 2007; Kawashima et al., 2017 </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:61px; vertical-align:top; width:228px"> To detect DSBs, cells are embedded and lysed in agarose, and the released DNA undergoes gel electrophoresis in which the direction of the voltage is periodically alternated; Large DNA fragments are thus able to be separated by size </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #7f7f7f; border-top:none; height:61px; vertical-align:top; width:46px"> N/A </td> </tr> <tr> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:1px double #2b2b2b; border-right:1px double #2b2b2b; border-top:none; height:73px; vertical-align:top; width:85px"> The TUNEL (Terminal Deoxynucleotidyl Transferase dUTP Nick End Labeling) Assay </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:73px; vertical-align:top; width:89px"> Loo, 2011 </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:73px; vertical-align:top; width:228px"> To detect strand breaks, dUTPs added to the 3’OH end of a strand break by the DNA polymerase terminal deoxynucleotidyl transferase (TdT) are tagged with a fluorescent dye or a reporter enzyme to allow visualization (We note that this method is typically used to measure apoptosis) </td> <td style="background-color:whitesmoke; border-bottom:1px double #2b2b2b; border-left:none; border-right:1px double #7f7f7f; border-top:none; height:73px; vertical-align:top; width:46px"> N/A </td> </tr> <tr> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:1px double #2b2b2b; border-right:1px double #2b2b2b; border-top:none; height:49px; vertical-align:top; width:85px"> In Vitro DNA Cleavage Assays using Topoisomerase </td> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:49px; vertical-align:top; width:89px"> Nitiss, 2012 </td> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:49px; vertical-align:top; width:228px"> Cleavage of DNA can be achieved using purified topoisomerase; DNA strand breaks can then be separated and quantified using gel electrophoresis </td> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:none; border-right:1px double #7f7f7f; border-top:none; height:49px; vertical-align:top; width:46px"> N/A </td> </tr> <tr> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:1px double #2b2b2b; border-right:1px double #2b2b2b; border-top:none; height:49px; text-align:center; vertical-align:top; width:85px">PCR assay </td> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:49px; text-align:center; vertical-align:top; width:89px">Figueroa‑González & Pérez‑Plasencia, 2017 </td> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:49px; text-align:center; vertical-align:top; width:228px">Assay of strand breaks through the observation of DNA amplification prevention. Breaks block Taq polymerase, reducing the number of DNA templates, preventing amplification </td> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:none; border-right:1px double #7f7f7f; border-top:none; height:49px; text-align:center; vertical-align:top; width:46px">N/A</td> </tr> <tr> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:1px double #2b2b2b; border-right:1px double #2b2b2b; border-top:none; height:49px; text-align:center; vertical-align:top; width:85px">Sucrose density gradient centrifuge </td> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:49px; text-align:center; vertical-align:top; width:89px">Raschke et al. 2009 </td> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:49px; text-align:center; vertical-align:top; width:228px">Division of DNA pieces by density, increased fractionation leads to lower density pieces, with the use of a sucrose cushion </td> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:none; border-right:1px double #7f7f7f; border-top:none; height:49px; text-align:center; vertical-align:top; width:46px">N/A</td> </tr> <tr> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:1px double #2b2b2b; border-right:1px double #2b2b2b; border-top:none; height:49px; text-align:center; vertical-align:top; width:85px">Alkaline Elution Assay </td> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:49px; text-align:center; vertical-align:top; width:89px">Kohn, 1991 </td> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:49px; text-align:center; vertical-align:top; width:228px">Cells lysed with detergent-solution, filtered through membrane to remove all but intact DNA </td> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:none; border-right:1px double #7f7f7f; border-top:none; height:49px; text-align:center; vertical-align:top; width:46px">N/A</td> </tr> <tr> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:1px double #2b2b2b; border-right:1px double #2b2b2b; border-top:none; height:49px; text-align:center; vertical-align:top; width:85px">Unwinding Assay </td> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:49px; text-align:center; vertical-align:top; width:89px">Nacci et al. 1992 </td> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:none; border-right:1px double #2b2b2b; border-top:none; height:49px; text-align:center; vertical-align:top; width:228px">DNA is stored in alkaline solutions with DNA-specific dye and allowed to unwind following removal from tissue, increased strand damage associated with increased unwinding </td> <td style="background-color:whitesmoke; border-bottom:1px double #7f7f7f; border-left:none; border-right:1px double #7f7f7f; border-top:none; height:49px; text-align:center; vertical-align:top; width:46px">N/A</td> </tr> </tbody> </table>

Taxonomic applicability: DNA strand breaks are relevant to all species, including vertebrates such as humans, that contain DNA (Cannan & Pederson, 2016).   Life stage applicability: This key event is not life stage specific as all life stages display strand breaks. However, there is an increase in baseline levels of DNA strand breaks seen in older individuals though it is unknown whether this change due to increased break induction or a greater retention of breaks due to poor repair (White & Vijg, 2016).  Sex applicability: This key event is not sex specific as both sexes display evidence of strand breaks. In some cell types, such as peripheral blood mononuclear cells, males show higher levels of single strand breaks than females (Garm et al., 2012).  Evidence for perturbation by a stressor: There are studies demonstrating that increased DNA strand breaks can result from exposure to multiple stressor types including ionizing & non-ionizing radiation, chemical agents, and oxidizing agents (EPRI, 2014; Hamada, 2014; Cencer et al., 2018; Cannan & Pederson, 2016; Yang et al., 1998).

High Unspecific

High

All life stages

Not Specified Ager, D. D. et al. (1990). “Measurement of Radiation- Induced DNA Double-Strand Breaks by Pulsed-Field Gel Electrophoresis.” Radiat Res. 122(2), 181-7. Anderson, D. & Laubenthal J. (2013), “Analysis of DNA Damage via Single-Cell Electrophoresis. In: Makovets S, editor. DNA Electrophoresis. Totowa.”, NJ: Humana Press. p 209-218. Asaithamby, A., B. Hu and D.J. Chen. (2011) Unrepaired clustered DNA lesions induce chromosome breakage in human cells. Proc Natl Acad Sci U S A 108(20): 8293-8298 . Barbieri, S., G. Babini, J. Morini et a l (2019). . Predicting DNA damage foci and their experimental readout with 2D microscopy: a unified approach applied to photon and neutron exposures. Scientific Reports 9(1): 14019 Bryce, S. et al. (2016), “Genotoxic mode of action predictions from a multiplexed flow cytometric assay and a machine learning approach.”, Environ Mol Mutagen. 57:171-189. Doi: 10.1002/em.21996. Burma, S. et al. (2001), “ATM phosphorylates histone H2AX in response to DNA double-strand breaks.”, J Biol Chem, 276(45): 42462-42467. doi:10.1074/jbc.C100466200 Cannan, W.J. and D.S. Pederson (2016), "Mechanisms and Consequences of Double-Strand DNA Break Formation in Chromatin.", Journal of Cellular Physiology, Vol.231/1, Wiley, New York, https://doi.org/10.1002/jcp.25048.   Cencer, C. et al. (2018), “PARP-1/PAR Activity in Cultured Human Lens Epithelial Cells Exposed to Two Levels of UVB Light”, Photochemistry and Photobiology, Vol.94/1, Wiley-Blackwell, Hoboken, https://doi.org/10.1111/php.12814.   Charlton, E. D. et al. (1989), “Calculation of Initial Yields of Single and Double Stranded Breaks in Cell Nuclei from Electrons, Protons, and Alpha Particles.”,  Int. J. Radiat. Biol. 56(1): 1-19. doi: 10.1080/09553008914551141. Collins, R. A. (2004), “The Comet Assay for DNA Damage and Repair. Molecular Biotechnology.”, Mol Biotechnol. 26(3): 249-61. doi:10.1385/MB:26:3:249 EPRI (2014), Epidemiology and mechanistic effects of radiation on the lens of the eye: Review and scientific appraisal of the literature, EPRI, California.  Figueroa‑González, G. and C. Pérez‑Plasencia. (2017), “Strategies for the evaluation of DNA damage and repair mechanisms in cancer”, Oncology Letters, Vol.13/6, Spandidos Publications, Athens, https://doi.org/10.3892/ol.2017.6002.  Garcia-Canton, C. et al. (2013), “Assessment of the in vitro p-H2AX assay by High Content Screening asa novel genotoxicity test.”, Mutat Res. 757:158-166.  Doi:  10.1016/j.mrgentox.2013.08.002 Gardiner, K. et al. (1986), “Fractionation of Large Mammalian DNA Restriction Fragments Using Vertical Pulsed-Field Gradient Gel Electrophoresis.”,  Somatic Cell and Molecular Genetics. 12(2): 185-95.Doi: 10.1007/bf01560665. Garm, C. et al. (2012), “Age and gender effects on DNA strand break repair in peripheral blood mononuclear cells”, Aging Cell, Vol.12/1, Blackwell Publishing Ltd, Oxford, https://doi.org/10.1111/acel.12019.  Hamada, N. (2014), “What are the intracellular targets and intratissue target cells for radiation effects?”, Radiation research, Vol. 181/1, The Radiation Research Society, Indianapolis, https://doi.org/10.1667/RR13505.1.  Herschleb, J. et al. (2007), “Pulsed-field gel electrophoresis.”,  Nat Protoc. 2(3): 677-684. doi:10.1038/nprot.2007.94 Iliakis, G. et al. (2015), “Alternative End-Joining Repair Pathways Are the Ultimate Backup for Abrogated Classical Non-Homologous End-Joining and Homologous Recombination Repair: Implications for the Formation of Chromosome Translocations.”,  Mutation Research/Genetic Toxicology and Environmental Mutagenesis. 2(3): 677-84. doi: 10.1038/nprot.2007.94 Jackson, S. (2002). “Sensing and repairing DNA double-strand breaks.”,  Carcinogenesis. 23:687-696. Doi:10.1093/carcin/23.5.687. Ji, J. et al. (2017), “Phosphorylated fraction of H2AX as a measurement for DNA damage in cancer cells and potential applications of a novel assay.”,  PLoS One. 12(2): e0171582. doi:10.1371/journal.pone.0171582 Kawashima, Y.(2017), “Detection of DNA double-strand breaks by pulsed-field gel electrophoresis.”,  Genes Cells 22:84-93. Doi: 10.1111/gtc.12457. Khoury, L. et al. (2013), “Validation of high-throughput genotoxicity assay screening using cH2AX in-cell Western assay on HepG2 cells.”, Environ Mol Mutagen, 54:737-746. Doi:  10.1002/em.21817. Khoury, L. et al. (2016), “Evaluation of four human cell lines with distinct biotransformation properties for genotoxic screening.”, Mutagenesis, 31:83-96. Doi: <a href="https://doi.org/10.1093/mutage/gev058" target="_blank">10.1093/mutage/gev058</a>. Kohn, K.W. (1991), “Principles and practice of DNA filter elution”, Pharmacology & Therapeutics, Vol.49/1, Elsevier, Amsterdam, https://doi.org/10.1016/0163-7258(91)90022-E.  Loo, DT. (2011), “In Situ Detection of Apoptosis by the TUNEL Assay: An Overview of Techniques. In: Didenko V, editor. DNA Damage Detection In Situ, Ex Vivo, and In Vivo. Totowa.”, NJ: Humana Press. p 3-13.doi: <a href="https://doi.org/10.1007/978-1-60327-409-8_1" target="_blank">10.1007/978-1-60327-409-8_1</a>. Mah, L. J. et al. (2010), “Quantification of gammaH2AX foci in response to ionising radiation.”,  J Vis Exp(38). doi:10.3791/1957. Nacci, D. et al. (1992), “Application of the DNA alkaline unwinding assay to detect DNA strand breaks in marine bivalves”, Marine Environmental Research, Vol.33/2, Elsevier BV, Amsterdam, https://doi.org/10.1016/0141-1136(92)90134-8.  Nikolova, T., F. et al. (2017), “Genotoxicity testing: Comparison of the γH2AX focus assay with the alkaline and neutral comet assays.”,  Mutat Res 822:10-18. Doi: <a href="https://doi.org/10.1016/j.mrgentox.2017.07.004" target="_blank">10.1016/j.mrgentox.2017.07.004</a>. Nitiss, J. L. et al. (2012), “Topoisomerase assays. ”, Curr Protoc Pharmacol. Chapter 3: Unit 3 3. OECD. (2014). Test No. 489: “In vivo mammalian alkaline comet assay.”  OECD Guideline for the Testing of Chemicals, Section 4 . Olive, P. L., & Banáth, J. P. (2006), “The comet assay: a method to measure DNA damage in individual cells.”,  Nature Protocols. 1(1): 23-29. doi:10.1038/nprot.2006.5. Platel A. et al. (2011), “Study of oxidative DNA damage in TK6 human lymphoblastoid cells by use of the thymidine kinase gene-mutation assay and the in vitro modified comet assay: Determination of No-Observed-Genotoxic-Effect-Levels.”,  Mutat Res 726:151-159. Doi: 10.1016/j.mrgentox.2011.09.003. Raschke, S., J. Guan and G. Iliakis. (2009), “Application of alkaline sucrose gradient centrifugation in the analysis of DNA replication after DNA damage”, Methods in Molecular Biology, Vol.521, Humana Press, Totowa, https://doi.org/10.1007/978-1-60327-815-7_18.  Redon, C. et al. (2010), “The use of gamma-H2AX as a biodosimeter for total-body radiation exposure in non-human primates.”,  PLoS One. 5(11): e15544. doi:10.1371/journal.pone.0015544 Revet, I. et al. (2011), “Functional relevance of the histone γH2Ax in the response to DNA damaging agents.” Proc Natl Acad Sci USA.108:8663-8667. Doi: 10.1073/pnas.1105866108 Rogakou, E.P. et al. (1998), “DNA Double-stranded Breaks Induce Histone H2AX Phosphorylation on Serine 139.” , J Biol Chem, 273:5858-5868. Doi: 10.1074/jbc.273.10.5858 Rothkamm, K. & Horn, S. (2009), “γ-H2AX as protein biomarker for radiation exposure.”,  Ann Ist Super Sanità, 45(3): 265-71. White, R.R. and J. Vijg. (2016), “Do DNA Double-Strand Breaks Drive Aging?”, Molecular Cell, Vol.63, Elsevier, Amsterdam, http://doi.org/10.1016/j.molcel.2016.08.004.  Yang, Y. et al. (1998), “The effect of catalase amplification on immortal lens epithelial cell lines”, Experimental Eye Research, Vol.67/6, Academic Press Inc, Cambridge, https://doi.org/10.1006/exer.1998.0560.   AOPWiki 2019-05-19T16:33:20

2023-05-15T08:39:51

<upstream-id>8d6a4475-5817-4386-be53-eb911186774f</upstream-id> <downstream-id>b813d3a8-c9cc-49b4-bd70-a0bb3e9e8efd</downstream-id> Energy deposited onto biomolecules stochastically in the form on ionizing and non-ionizing radiation can cause direct and indirect molecular-level damage. As energy is deposited in an aqueous solution, water molecules can undergo radiolysis, breaking bonds to produce reactive oxygen species (ROS) (Ahmadi et al., 2021; Karimi et al., 2017) or directly increase function of enzymes involved in ROS generation (i.e. catalaze). Various species of ROS can be generated with differing degrees of biological effects. For example, singlet oxygen, superoxide, and hydroxyl radical are highly unstable, with short half-lives and react close to where they are produced, while species like H2O2 are much more stable and membrane permeable, meaning they can travel from the site of production, reacting elsewhere as a much weaker oxidant (Spector, 1990). In addition, enzymes involved in reactive oxygen and nitrogen species (RONS) production can be directly upregulated following the deposition of energy (de Jager, Cockrell and Du Plessis, 2017). Although less common than ROS, reactive nitrogen species (RNS) can also be produced by energy deposition resulting in oxidative stress (Cadet et al., 2012; Tangvarasittichai & Tangvarasittichai, 2019), a state in which the amount of ROS and RNS, collectively known as RONS, overwhelms the cell’s antioxidant defense system. This loss in redox homeostasis can lead to oxidative damage to macromolecules including proteins, lipids, and nucleic acids (Schoenfeld et al., 2012; Tangvarasittichai & Tangvarasittichai, 2019; Turner et al., 2002).

The strategy for collating the evidence to support the relationship is described in Kozbenko et al 2022.  Briefly, a scoping review methodology was used to prioritize studies based on a population, exposure, outcome, endpoint statement.

Overal weight of evidence: High

A large body of literature supports the linkage between the deposition of energy and oxidative stress. Multiple reviews describe the relationship in the context of ROS production (Marshall, 1985; Balasubramanian, 2000; Jurja et al., 2014),  antioxidant depletion (Cabrera et al., 2011; Fletcher, 2010; Ganea & Harding, 2006; Hamada et al., 2014; Spector, 1990; Schoenfeld et al., 2012; Wegener, 1994), and overall oxidative stress (Eaton, 1994, Tangvarasittichai & Tangvarasittichain, 2019). This includes investigations into the mechanism behind the relationship (Ahmadi et al., 2021; Balasubramanian, 2000; Cencer et al., 2018; Eaton, 1994; Fletcher, 2010; Jiang et al., 2006; Jurja et al., 2014; Padgaonkar et al., 2015; Quan et al., 2021; Rong et al., 2019; Slezak et al., 2015; Soloviev & Kizub, 2019; Tian et al., 2017; Tahimic & Globus, 2017; Varma et al., 2011; Venkatesulu et al., 2018; Wang et al., 2019a; Yao et al., 2008; Yao et al., 2009; Zigman et al., 2000).   Water radiolysis is a main source of free radicals. Energy ionizes water and free radicals are produced that combine to create more stable ROS, such as hydrogen peroxide, hydroxide, superoxide, and hydroxyl (Eaton, 1994; Rehman et al., 2016; Tahimic & Globus, 2017; Tian et al., 2017; Varma et al., 2011; Venkatesulu et al., 2018). ROS formation causes ensuing damage to the body, as ~80% of tissues are comprised of water (Wang et al., 2019a). Ionizing radiation (IR) is a source of energy deposition, it can also interact with molecules, such as nitric oxide (NO), to produce less common free radicals, including RNS (Slezak et al., 2015; Tahimic & Globus, 2017; Wang et al., 2019a). Free radicals can diffuse  throughout the cell and damage vital cellular components, such as proteins, lipids, and DNA, as well as dysregulate cellular processes, such as cell signalling (Slezak et al., 2015; Tian et al., 2017).   ROS are also commonly produced by nicotinamide adenine dinucleotide phosphate (NADPH) oxidase (NOX). Deposition of energy can activate NOX and induce expression of its catalytic and cytosolic components, resulting in increased intracellular ROS (Soloviev & Kizub, 2019). Intracellular ROS production can also be initiated through the expression of protein kinase C, which in turn activates NOX through phosphorylation of its cytosolic components (Soloviev & Kizub, 2019). Alternatively, ROS are often formed at the electron transport chain (ETC) of the mitochondria, due to IR-induced electron leakage leading to ionization of the surrounding O2 to become superoxide (Soloviev & Kizub, 2019). Additionally, energy reaching a cell can be absorbed by an unstable molecule, often NADPH, known as a chromophore, which leads to the production of ROS (Balasubramanian, 2000; Cencer et al., 2018; Jiang et al., 2006; Jurja et al., 2014; Padgaonkar et al., 2015; Yao et al., 2009; Zigman et al., 2000).  Energy deposition can also weaken a cell’s antioxidant defense system through the depletion of certain antioxidant enzymes, such as superoxide dismutase (SOD) and catalase (CAT). Antioxidants are consumed during the process of neutralizing ROS, so as energy deposition stimulates the formation of ROS it begins to outpace the rate at which antioxidants are replenished; this results in an increased risk of oxidative stress when their concentrations are low (Belkacémi et al., 2001; Giblin et al., 2002; Ji et al., 2014; Kang et al., 2020; Karimi et al., 2017; Padgaonkar et al., 2015; Rogers et al., 2004; Slezak et al., 2015; Tahimic & Globus, 2017; Wang et al., 2019a; Wegener, 1994; Weinreb & Dovrat, 1996; Zhang et al., 2012; Zigman et al., 1995; Zigman et al., 2000). When the amount of ROS overwhelms the antioxidant defense system, the cell will enter oxidative stress leading to macromolecular and cellular damage (Tangvarasittichai & Tangvarasittichai, 2019).

The relationship between energy deposition and oxidative stress is strongly supported by primary research on the effects of IR on ROS and antioxidant levels (Bai et al., 2020; Cervelli et al., 2017; Hatoum et al., 2006; Huang et al., 2018; Huang et al., 2019; Karam & Radwan, 2019; Kook et al., 2015; Liu et al., 2018; Liu et al., 2019; Mansour, 2013; Philipp et al., 2020; Ramadan et al., 2020; Sharma et al., 2018; Shen et al., 2018; Soltani et al., 2016; Soucy et al., 2010; Soucy et al., 2011; Ungvari et al., 2013; Wang et al., 2016; Wang et al., 2019b; Zhang et al., 2018; Zhang et al., 2020). Of note is that the relationship is demonstrated across studies conducted using various cell types, models and using broad dose-ranges as summarized below.  Much evidence is available and described to help discern the quantitative understanding of the relationship, since it is well established.    Dose Concordance  It is well-accepted that any dose of radiation will deposit energy onto matter. Doses as low as 1 cGy support this relationship (Tseung et al., 2014). Following the deposition of energy, markers of oxidative stress are observed in the form of RONS, a change in levels of antioxidants, and oxidative damage to macromolecules. These effects have been shown across various organs/tissues and cell types as described below.    RONS  Cardiovascular tissue:  There is a considerable amount of evidence to support this relationship in cell types and tissues of relevance to the cardiovascular system. Recent studies have shown a linear increase in ROS in human umbilical vein endothelial cells (HUVECs) following 0-5 Gy gamma irradiation (Wang et al., 2019b). HUVECs irradiated with 0.25 Gy X-rays (Cervelli et al., 2017) and 9 Gy 250kV photons (Sharma et al., 2018) show increased ROS. Gamma ray irradiated rats at 5 Gy display increased ROS levels in the aorta (Soucy et al., 2010). A study using cerebromicrovascular endothelial cell (CMVECs) showed a dose-dependent increase in ROS from 0-8 Gy gamma irradiation (Ungvari et al., 2013). Additionally, telomerase-immortalized coronary artery endothelial (TICAE) and telomerase-immortalized microvascular endothelial (TIME) cells irradiated with 0.1 and 5 Gy of X-rays displayed increased ROS production (Ramadan et al., 2020). Gut arterioles of rats showed increased ROS following multiple fractions of 2.5 Gy X-ray rat irradiation (Hatoum et al., 2006). Additionally, rats irradiated with 1 Gy of 56Fe expressed increased ROS levels in the aorta (Soucy et al., 2011).      Brain tissue:   Markers of oxidative stress have also been consistently observed in brain tissue. Human neural stem cells subjected to 1, 2 or 5 Gy gamma rays showed a dose-dependent increase in RONS production (Acharya et al., 2010). A dose-dependent increase in ROS was observed in rat brains following 1-10 Gy gamma rays (Collins-Underwood et al., 2008). Neural precursor cells exposed to 0-10 Gy of X-irradiation showed increased ROS levels (Giedzinski et al., 2005; Limoli et al., 2004). Mice brain tissue displayed increased ROS following proton irradiation (Baluchamy et al., 2012; Giedzinski et al., 2005). Neural processor cells expressed linearly increased ROS levels following doses of 56Fe (Limoli et al., 2007). A dose-dependent increase in RONS was also observed after exposure to 1-15 cGy 56Fe irradiation in mice neural stem/precursor cell (Tseng et al., 2014). Human neural stem cells exposed to 5-100 cGy of various ions demonstrated a dose-dependent increase in RONs (Baulch et al., 2015).    Eye tissue:  The eye is also sensitive to the accumulation of free radicals, in a state of antioxidant decline. It has been shown in human lens epithelial cells (HLECs) and HLE-B3 following gamma irradiation of 0.25 and 0.5 Gy that ROS levels are markedly increased (Ahmadi et al., 2021). Exposure to non-ionizing radiation, such as ultraviolet (UV)-B, has also led to increased ROS in HLECs and mice lenses (Ji et al., 2015; Kubo et al., 2010; Rong et al., 2019; Yang et al., 2020)    Bone tissue:  Rat bone marrow-derived mesenchymal stem cell (bmMSCs) irradiated with 2, 5 and 10 Gy gamma rays and Murine MC3T3-E1 osteoblast cells irradiated with 2, 4, and 8 Gy of X-rays have shown a dose-dependent increase in ROS levels (Bai et al., 2020; Kook et al., 2015). Murine RAW264.7 cells and rat bmMSC irradiated with 2 Gy of gamma rays displayed increased ROS levels (Huang et al., 2019; Huang et al., 2018; hang et al., 2020). Human bone marrow-derived mesenchymal stem cell (hBMMSCs) irradiated with 2 or 8 Gy X-rays showed increased ROS (Liu et al., 2018; Zhang et al., 2018). Similarly, murine MC3T3-E1 osteoblast-like cells irradiated with 6 Gy of X-rays also displayed increased ROS (Wang et al., 2016). Finally, whole-body irradiation of mice with 2 Gy of 31.6 keV/mm LET 12C heavy ions showed increased ROS (Liu et al., 2019)    Antioxidants  Blood:   Workers exposed to X-rays at less than 1 mSv/year for an average of 15 years showed around 20% decreased antioxidant activity compared to unexposed controls (Klucinski et al., 2008). Similarly, adults exposed to high background irradiation of 260 mSv/year showed about 50% lower antioxidant activity power compared to controls (Attar, Kondolousy and Khansari, 2007).    Cardiovascular tissue:  Heart tissue of rats following gamma irradiation of rats at 5 and 6 Gy resulted in a decrease in antioxidant levels (Karam & Radwan, 2019; Mansour, 2013). Similarly, HUVECs (Soltani, 2016) and TICAE cells (Philipp et al., 2020) irradiated at 2 Gy and 0.25-10 Gy gamma rays, respectively, displayed decreased antioxidant levels. Mice exposed to 18 Gy of X-ray irradiation showed decreased antioxidants in the aorta (Shen et al., 2018).   Brain tissue:   Mice brain tissue following 2, 10 and 50 cGy whole-body gamma irradiation revealed a dose-dependent change in SOD2 activity (Veeraraghan et al., 2011). Mice brain tissue showed decreased glutathione (GSH) and SOD levels following proton irradiation (Baluchamy et al., 2012)    Eye tissue:  Rats exposed to 15 Gy gamma rays demonstrated decreased antioxidants in the lens tissue (Karimi et al, 2017).  Neutron irradiation of rats at 3.6 Sv resulted in a decrease in antioxidants in lens (Chen et al., 2021). A few studies found a dose concordance between UV irradiation and decreased antioxidant levels (Hua et al, 2019; Ji et al, 2015; Zigman et al., 2000; Zigman et al, 1995). HLECs following UVB exposure from 300 J/m2 to 14,400 J/m2 in HLECs showed linear decreases in antioxidant activity (Ji et al., 2015). Similarly, HLEC exposed to 4050, 8100 and 12,150 J/m2 found decreased antioxidant levels (Hua et al., 2019). Following UV irradiation of rabbit and squirrel lens epithelial cells (LECs) showed a linear decrease of antioxidant level, CAT (Zigman et al., 2000; Zigman et al., 1995). Mice exposed to UV irradiation found decreased antioxidant levels in lens (Zhang et al., 2012). Similarly, SOD levels decreased following 0.09 mW/cm 2 UVB exposure of HLECs (Kang et al., 2020).    Bone tissue:  Rat bmMSCs irradiated with 2, 5 and 10 Gy gamma rays and Murine MC3T3-E1 osteoblast cells irradiated with 2, 4, and 8 Gy of X-rays showed a dose-dependent decrease in antioxidant levels (Bai et al., 2020; Kook et al., 2015). hBMMSCs irradiated with 8 Gy X-rays also showed a decrease in antioxidant, SOD, levels (Liu et al., 2018).    Oxidative Damage  Cardiovascular tissue:  HUVECs and rat hearts irradiated by gamma rays at 2 and 6 Gy, respectively, resulted in increased levels of oxidative stress markers, such as malondialdehyde (MDA), and thiobarbituric reactive substances (TBARS) (Mansour, 2013; Soltani, 2016).     Brain tissue:   Mice brain tissue were shown to have increased lipid peroxidation (LPO) as determined by MDA measurements, following proton irradiation at 1 and 2 Gy (Baluchamy et al., 2012). Neural precursor cells from rat hippocampus exposed to 0, 1, 5 and 10 Gy of X-irradiation resulted in increased lipid peroxidation (Limoli et al., 2004).   Eye tissue:  Rats exposed to 15 Gy gamma rays demonstrated increased MDA in lens tissue (Karimi et al, 2017). Neutron irradiation of rats at 3.6 Sv resulted in an initial decrease, followed by an increase in MDA in lens (Chen et al., 2021). Following UV irradiation at 300 4050, 8100 and 12,150 J/m2, there was an increase in LPO in in human lens (Chitchumroonchokchai et al, 2004; Hua et al, 2019). Similarly, LPO increased following 0.09 mW/cm 2 UVB exposure of HLECs (Kang et al., 2020).    Time Concordance  It is well-accepted that deposition of energy into matter results in immediate vibrational changes to molecules or ionization events. Deposition of energy is therefore an upstream event to all follow-on latent events like oxidative stress.    RONS  Cardiovascular tissue:  In TICAE and TIME cells, ROS increased at 45 mins after X-ray irradiation (Ramadan et al., 2020). Superoxide and peroxide production were increased 1 day after 2-8 Gy of gamma irradiation in CMVECs (Unvari et al., 2013).    Bone tissue:  hBMMSCs irradiated with X-rays at 2 Gy showed peak ROS production at 2-8h post-irradiation (Zhang et al., 2018). Murine RAW264.7 cells (can undergo osteoclastogenesis) irradiated with 2 Gy of gamma rays showed increased ROS at 2-8h post-irradiation (Huang et al., 2018).    Brain tissue:  In human lymphoblast cells exposed to 2 Gy of X-rays, ROS were increased at various times between 13 and 29 days post-irradiation (Rugo and Schiestl, 2004). RONS were increased in human neural stem cells at 12-48h post-irradiation with 2 and 5 Gy of gamma rays (Acharya et al., 2010). ROS levels were increased in rat neural precursor cells at 6-24h after irradiation with 1-10 Gy of protons (Giedzinksi et al., 2005). Both 56Fe (1.3 Gy) and gamma ray (2 Gy) irradiation of mice increased ROS levels after 2 months post-irradiation in the cerebral cortex (Suman et al., 2013). ROS were also increased 12 months after 56Fe irradiation (Suman et al., 2013). RONS increased as early as 12h post-irradiation continuing to 8 weeks with 2-200 cGy doses of 56Fe irradiation of mouse neural stem/precursor cells (Tseng et al., 2014). The same cell type irradiated with 1 and 5 Gy of 56Fe irradiation showed increased ROS at 6h post-irradiation, with the last increase observed 25 days post-irradiation (Limoli et al., 2004).    Eye tissue:  Mice exposed to 11 Gy of X-rays showed increased ROS at 9 months post-irradiation in lenses (Pendergrass et al., 2010). In human lens cells, ROS were found increased at 1h after 0.25 Gy gamma ray irradiation (Ahmadi et al., 2021), 15 minutes after 30 mJ/cm2 UV radiation (Jiang et al., 2006), 2.5-120 minutes after 0.014 and 0.14 J/cm2 UV radiation (Cencer et al., 2018), and 24h after 30 mJ/cm2 UVB radiation (Yang et al., 2020).    Antioxidants  Cardiovascular tissue:  CAT antioxidant enzyme was decreased in mice aortas as early as 3 days post-irradiation, remaining decreased until 84 days after irradiation with 18 Gy of X-rays (Shen et al., 2018). The antioxidant enzymes peroxiredoxin 5 (PRDX5) and SOD were both shown to have the greatest decrease at 24h after 2 Gy gamma irradiation of TICAE cells (Philipp et al., 2020).    Eye tissue:  Bovine lenses irradiated with 44.8 J/cm2 of UVA radiation showed decreased CAT levels at 48-168h post-irradiation (Weinreb and Dovrat, 1996). UV irradiation of mice at 20.6 kJ/m2 led to decreased GSH at both 1 and 16 months post-irradiation in the lens (Zhang et al., 2012). Bovine lens cells exposed to 10 Gy of X-rays showed decreased levels of the antioxidant GSH at 24 and 120h after exposure (Belkacemi et al., 2001).    Oxidative damage markers  Cardiovascular tissue:  Oxidative damage markers 4-hydroxynonemal (4-HNE) and 3-Nitrotyosine (3-NT) were both significantly increased in the aorta of mice at 3 days post-irradiation, remaining increased until 84 days after irradiation with 18 Gy of X-rays (Shen et al., 2018).    Essentiality  Radiation has been found to induce oxidative stress above background levels. Many studies have shown that lower doses of ionizing radiation resulted in decreased  levels in markers of oxidative stress in multiple cell types (Acharya et al., 2010; Ahmadi et al., 2021; Bai et al., 2020; Baluchamy et al., 2012 Chen et al., 2021; Collins-Underwood et al., 2008; Giedzinski et al., 2005; Kook et al., 2015; Kubo et al., 2010; Philipp et al., 2020; Ramadan et al., 2020; Ungvari et al., 2013; Veeraraghan et al., 2011; Wang et al., 2019b; Zigman et al., 2000; Zigman et al., 1995). The essentiality of deposition of energy can be assessed through the removal of deposited energy, a physical stressor that does not require to be metabolized in order to elicit downstream effects on a biological system. Studies that do not deposit energy are observed to have no downstream effects.

There are several uncertainties and inconsistencies in this KER.   <ul> <li> Chen et al. (2021) found that radiation can have adaptive responses. The study used three neutron radiation doses, 0.4 and 1.2 Sv, and 3.6 Sv. After 0.4 and 1.2 Sv, the activity of antioxidant enzymes GSH and SOD increased, and the concentration of malondialdehyde, a product of oxidative stress, decreased. After 3.6 Sv, the opposite was true.  </li> </ul> <ul> <li> While the concentration of most antioxidant enzymes decreases after energy deposition, there is some uncertainty with SOD. Certain papers have found that its concentration decreases with dose (Chen et al., 2021; Hua et al., 2019; Ji et al., 2015; Kang et al., 2020) while others found no difference after irradiation (Rogers et al., 2004; Zigman et al., 1995). Several studies have also found that higher levels of SOD do not increase resistance to UV radiation (Eaton, 1994; Hightower, 1995).  </li> <li> At 1-week post-irradiation with 10 Gy of 60Co gamma rays, TICAE cells experienced a significant increase in levels of the antioxidant, PRDX5, contrary to the decrease generally seen in antioxidant levels following radiation exposure (Philipp et al., 2020).  </li> <li> Various studies found an increase in antioxidant SOD levels within the brain after radiation exposure (Acharya et al., 2010; Baluchamy et al., 2012; Baulch et al., 2015; Veeraraghan et al., 2011).  </li> </ul>

<table cellspacing="0" class="MsoTableGrid" style="border-collapse:collapse; border:none; width:620px"> <tbody> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:1px solid black; height:19px; vertical-align:top; width:107px"> Modulating Factors </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; height:19px; vertical-align:top; width:172px"> MF details </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; height:19px; vertical-align:top; width:161px"> Effects on the KER </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; height:19px; vertical-align:top; width:180px"> References </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:327px; vertical-align:top; width:107px"> Antioxidants </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:327px; vertical-align:top; width:172px"> CAT, GSH-Px, SOD, PRDX, vitamin E, C, carotene, lutein, zeaxanthin, selenium, zinc, alpha-lipoic acid, melatonin, gingko biloba leaf, fermented gingko biloba leaf, Nigella sativa oil, thymoquinone, and ferulic acid  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:327px; vertical-align:top; width:161px"> Adding or withholding antioxidants will decrease or increase the level of oxidative stress respectively  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:327px; vertical-align:top; width:180px"> Zigman et al., 1995; Belkacémi et al., 2001; Chitchumroonchokchai et al., 2004; Fatma et al., 2005; Jiang et al., 2006; Fletcher, 2010; Karimi et al., 2017; Hua et al., 2019; Kang et al., 2020; Yang et al., 2020; Manda et al., 2008; Limoli et al., 2007; Manda et al., 2007; Taysi et al., 2012; Ismail et al., 2016; Demir et al., 2020; Chen et al., 2021 </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:164px; vertical-align:top; width:107px"> Age </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:164px; vertical-align:top; width:172px"> Increased age </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:164px; vertical-align:top; width:161px"> Antioxidant levels are lower and show a greater decrease after radiation in older organisms. This compromises their defense system, resulting in ROS increases and therefore, an increased likelihood of oxidative stress  </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:164px; vertical-align:top; width:180px"> Marshall, 1985; Spector, 1990; Giblin et al., 2002; Kubo et al., 2010; Pendergrass et al., 2010; Zhang et al., 2012; Hamada et al., 2014; Tangvarasittichai & Tangvarasittichai, 2019 </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:56px; vertical-align:top; width:107px"> Oxygen </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:56px; vertical-align:top; width:172px"> Increased oxygen levels </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:56px; vertical-align:top; width:161px"> Higher oxygen concentrations increase sensitivity to ROS </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:56px; vertical-align:top; width:180px"> Hightower et al., 1992; Eaton, 1994; Huang et al., 2006; Zhang et al., 2010; Schoenfeld et al., 2012 </td> </tr> </tbody> </table>

The table below provides some representative examples of quantitative linkages between the two key events. It was difficult to identify a general trend across all the studies due to differences in experimental design and reporting of the data. All data is statistically significant unless otherwise stated.

Dose Concordance  <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Attar, Kondolousy and Khansari, 2007   </td> <td> In vivo. One hundred individuals between 20 and 50 years old in two villages in Iran exposed to background IR at 260 mSv/year had antioxidant levels measured. The control group was from two villages not exposed to the high background radiation. The total antioxidant levels in the blood were determined by the ferric reducing/antioxidant power assay.   </td> <td> The total antioxidant level was significantly reduced from 1187±199 µmol in the control to 686±170 µmol in the exposed group.   </td> </tr> <tr> <td> Klucinski et al., 2008   </td> <td> In vivo. A group of 14 men and 31 women aged 25–54 years working X-ray equipment (receiving doses of less than 1 mSv/year) for an average of 15.3 years (range of 2-33 years) were compared to a control group for antioxidant activity. Antioxidant activity of SOD, glutathione peroxidase (GSH-Px), and CAT in erythrocytes were measured in U/g of hemoglobin.  </td> <td> All three enzymes showed significantly decreased antioxidant activity in the workers.   In the controls (U/g of Hb):   <ul> <li> SOD: 1200 ± 300   </li> <li> GSH-Px: 39 ± 7   </li> <li> CAT: 300 ± 60   </li> </ul> In the workers (U/g of Hb):   <ul> <li> SOD: 1000 ± 200  </li> <li> GSH-Px: 29 ± 4   </li> <li> CAT: 270 ± 50   </li> </ul> </td> </tr> <tr> <td> Limoli et al., 2007  </td> <td> In vitro. Neural precursor cells isolated from rat hippocampi was exposed to 0.25-5 Gy of 56Fe irradiation at dose rates of 0.5-1.0 Gy/min. ROS were measured 6h post-irradiation.   </td> <td> At a low dose of 0.25 Gy and 0.5 Gy, relative ROS levels were significantly elevated and showed a linear dose response (from ~1 to ~2.25 relative ROS levels) until 1 Gy, where it reached its peak (~3 relative ROS levels). At higher doses, the relative ROS levels decreased.   </td> </tr> <tr> <td> Tseng et al., 2014  </td> <td> In vitro. Neural stem/precursor cells isolated from mouse subventricular and hippocampal dentate subgranular zones were exposed to 1-15 cGy of 56Fe irradiation at dose rates ranging from 5-50 cGy/min. RONS levels were measured.  </td> <td> A dose-dependent and significant rise in RONS levels was detected after 56Fe irradiation. 12 h post-irradiation, a steady rise was observed and reached a 6-fold peak after 15 cGy.   </td> </tr> <tr> <td> Limoli et al., 2004  </td> <td> In vitro. Neural precursor cells from rat hippocampus were exposed to 0, 1, 5 and 10 Gy of X-irradiation at a dose rate of 4.5 Gy/min. ROS levels were measured.    In vivo. MDA was used to quantify oxidative stress.       </td> <td> A dose-dependent increase in ROS levels was seen in the first 12 h post-irradiation, with relative maximums at 12 h after 5 Gy (35% increase) and 24 h after 1 Gy (31% increase). ROS levels measured 1 week after 5 Gy were increased by 180% relative to sham-irradiated controls. MDA levels increased significantly (approximately 1.3-fold) after exposure to 10 Gy.   </td> </tr> <tr> <td> Collins-Underwood et al., 2008  </td> <td> In vitro. Immortalized rat brain microvascular endothelial cells were exposed to 1-10 Gy of 137Cs-irradiation at a dose rate of 3.91 Gy/min. Intracellular ROS and O2- production were both measured.   </td> <td> Irradiation resulted in a significant dose-dependent increase in intracellular ROS generation from 1-10 Gy. At 5 Gy, there was an approximate 10-fold increase in ROS levels, and at 10 Gy there was an approximate 20-fold increase.    </td> </tr> <tr> <td> Giedzinski et al., 2005  </td> <td> In vitro. Neural precursor cells were irradiated with 1, 2, 5 and 10 Gy of 250 MeV protons (1.7-1.9 Gy/min) and X-irradiation (4.5 Gy/min). ROS levels were measured.   </td> <td> There was a rapid increase in ROS at 6, 12, 18 and 24h after proton irradiation, with an exception at the 1 Gy 18h point. Most notably, at 6h post-irradiation, a dose-dependent increase in relative ROS levels from 1 to 10 Gy was seen that ranged from 15% (at 1 Gy) to 65% (at 10 Gy). Linear regression analysis showed that at ≤2 Gy, ROS levels increased by 16% per Gy. The linear dose response obtained at 24h showed that proton irradiation increased the relative ROS levels by 3% per Gy.   </td> </tr> <tr> <td> Veeraraghan et al., 2011  </td> <td> In vivo. Adult mice were exposed to 2, 10 or 50 cGy of whole-body gamma irradiation at 0.81 Gy/min. Brain tissues were harvested 24h post-irradiation. SOD2 levels and activity were measured.   </td> <td> Compared to the controls, the levels of SOD2 expression increased in the brain after 2, 10 and 50 cGy. Analysis revealed a significant and dose-dependent change in SOD2 activity. More specifically, SOD2 activity showed significant increases after 10 (~25% increase above control) and 50 cGy (~60% increase above control), but not 2 cGy.    </td> </tr> <tr> <td> Baluchamy et al., 2012  </td> <td> In vivo. Male mice were exposed to whole-body irradiation with 250 MeV protons at 0.01, 1 and 2 Gy and the whole brains were dissected out. ROS, LPO, GSH and total SOD were measured.  </td> <td> Dose-dependent increases in ROS levels was observed compared to controls, with a two-fold increase at 2 Gy. A 2.5 to 3-fold increase in LPO levels was also seen at 1 and 2 Gy, respectively, which was directly correlated with the increase in ROS levels. Additionally, results showed a significant reduction in GSH (~70% decrease at 2 Gy) and SOD activities (~2-fold decrease) following irradiation that was dose-dependent.    </td> </tr> <tr> <td> Acharya et al., 2010  </td> <td> In vitro. Human neural stem cells were subjected to 1, 2 or 5 Gy of gamma irradiation at a dose rate of 2.2 Gy/min. RONS and superoxide levels were determined.  </td> <td> Intracellular RONS levels increased by approximately 1.2 to 1.3-fold compared to sham-irradiated controls and was found to be reasonable dose-responsive.    At 12h, levels of superoxide increased 2 and 4-fold compared to control for 2 and 5 Gy, respectively. At 24h and 48h, there was a dose-dependent increase in RONS levels. At 7 days, levels of RONS increased approximately 3 to 7-fold for 2 and 5 Gy, respectively.       </td> </tr> <tr> <td> Baulch et al., 2015  </td> <td> In vitro. Human neural stem cells were exposed to 5-100 cGy of 16O, 28Si, 48Ti or 56Fe particles (600 MeV) at 10-50 cGy/min. RONS and superoxide levels were determined.  </td> <td> 3 days post-irradiation, oxidative stress was found to increase after particle irradiation. Most notably, exposure to 56Fe resulted in a dose-dependent increase with 100% increase in RONS levels at 100 cGy. Dose-dependent increase was also seen in superoxide levels after 56Fe irradiation. At 7 days post-irradiation, 56Fe irradiation induced significantly lower nitric oxide levels by 47% (5 cGy), 55% (25 cGy) and 45% (100 cGy).    </td> </tr> <tr> <td> Bai et al., 2020  </td> <td> In vitro. bmMSCs were taken from 4-week-old, male Sprague-Dawley rats. After extraction, cells were then irradiated with 2, 5, and 10 Gy of 137Cs gamma rays. Intracellular ROS levels and relative mRNA expression of the antioxidants, SOD1, SOD2, and CAT2, were measured to assess the extent of oxidative stress induced by IR.   </td> <td> Cellular ROS levels increased significantly in a dose-dependent manner from 0-10 Gy. Compared to sham-irradiated controls, ROS levels increased by ~15%, ~55%, and ~105% after exposure to 2, 5, and 10 Gy, respectively. Antioxidant mRNA expression decreased in a dose-dependent manner from 0-10 Gy, with significant increases seen at doses 2 Gy for SOD1 and CAT2 and 5 Gy for SOD2. Compared to sham-irradiated controls, SOD1 expression decreased by ~9%, ~18%, and ~27% after exposure to 2, 5, and 10 Gy, respectively. SOD2 expression decreased by ~31% and ~41% after exposure to 5 and 10 Gy, respectively. CAT2 expression decreased by ~15%, ~33%, and ~58% after exposure to 2, 5, and 10 Gy, respectively.   </td> </tr> <tr> <td> Liu et al., 2018  </td> <td> In vitro. hBMMSCs were irradiated with 8 Gy of X-rays at a rate of 1.24 Gy/min. Intracellular ROS levels and SOD activity were measured to analyze IR-induced oxidative stress.   </td> <td> Compared to sham-irradiated controls, hBMMSCs irradiated with 8 Gy of X-rays experienced a significant increase to intracellular ROS levels. hBMMSCs irradiated with 8 Gy of X-rays experienced a ~46% reduction in SOD activity.   </td> </tr> <tr> <td> Kook et al., 2015  </td> <td> In vitro. Murine MC3T3-E1 osteoblast cells were irradiated with 2, 4, and 8 Gy of X-rays at a rate of 1.5 Gy/min. Intracellular ROS levels and the activity of antioxidant enzymes, including GSH, SOD, CAT, were measured to assess the extent of oxidative stress induced by IR exposure.   </td> <td> Compared to sham-irradiated controls, irradiated MC3T3-E1 cells experienced a dose-dependent increase in ROS levels, with significant increases at 4 and 8 Gy (~26% and ~38%, respectively). Antioxidant enzyme activity initially increased by a statistically negligible amount from 0-2 Gy and then decreased in a dose-dependent manner from 2-8 Gy. SOD activity decreased significantly at 4 and 8 Gy by ~29% and ~59%, respectively. GSH activity similarly decreased significantly at 4 and 8 Gy by ~30% and ~48%, respectively. CAT activity did not change by a statistically significant amount.   </td> </tr> <tr> <td> Liu et al., 2019  </td> <td> In vivo. 8–10-week-old, juvenile, female SPF BALB/c mice underwent whole-body irradiation with 2 Gy of 31.6 keV/µm 12C heavy ions at a rate of 1 Gy/min. ROS levels were measured from femoral bone marrow mononuclear cells of the irradiated mice to analyze IR-induced oxidative stress.   </td> <td> Compared to sham-irradiated controls, irradiated mice experienced a ~120% increase in ROS levels.   </td> </tr> <tr> <td> Zhang et al., 2020  </td> <td> In vitro. Murine RAW264.7 osteoclast precursor cells were irradiated with 2 Gy of 60Co gamma rays at a rate of 0.83 Gy/min. ROS levels were measured to determine the extent of oxidative stress induced by IR exposure.   </td> <td> Compared to sham-irradiated controls, ROS levels in irradiated RAW264.7 cells increased by ~100%.   </td> </tr> <tr> <td> Wang et al., 2016  </td> <td> In vitro. Murine MC3T3-E1 osteoblast-like cells were irradiated with 6 Gy of X-rays. Intracellular ROS production was measured to assess oxidative stress from IR exposure.   </td> <td> Compared to sham-irradiated controls, Intracellular ROS production increased by ~81%.   </td> </tr> <tr> <td> Huang et al., 2018  </td> <td> In vitro. Murine RAW264.7 osteoblast-like cells were irradiated with 2 Gy of gamma rays at a rate of 0.83 Gy/min. ROS levels were measured to analyze IR-induced oxidative stress.   </td> <td> Compared to sham-irradiated controls, ROS levels in RAW264.7 cells increased by ~138% by 2 h post-irradiation.   </td> </tr> <tr> <td> Zhang et al., 2018  </td> <td> In vitro. hBMMSCs were irradiated with 2 Gy of X-rays at a rate of 0.6 Gy/min. Relative ROS concentration was measured to assess the extent of oxidative stress induced by IR.   </td> <td> Compared to sham-irradiated controls, irradiated hBMMSCs experienced a maximum increase of ~90% to ROS levels at 3 h post-irradiation.   </td> </tr> <tr> <td> Huang et al., 2019  </td> <td> In vitro. Rat bmMSC were irradiated with 2 Gy of 60Co gamma rays at a rate of 0.83 Gy/min. ROS levels were measured to assess IR-induced oxidative stress.   </td> <td> Compared to sham-irradiated controls, ROS levels in irradiated bone marrow stromal cells increased by approximately 2-fold.            </td> </tr> <tr> <td> Soucy et al., 2011  </td> <td> In vivo. 7- to 12-month-old, adult, male Wistar rats underwent whole-body irradiation with 1 Gy of 56Fe heavy ions. ROS production in the aorta was measured along with changes in activity of the ROS-producing enzyme xanthine oxidase (XO) to assess IR-induced oxidative stress.   </td> <td> Compared to sham-irradiated controls, irradiated mice experienced a 74.6% increase in ROS production (from 4.84 to 8.45) and XO activity increased by 36.1% (6.12 to 8.33).   </td> </tr> <tr> <td> Soucy et al., 2010  </td> <td> In vivo. 4-month-old, adult, male Sprague-Dawley rats underwent whole-body irradiation with 5 Gy of 137Cs gamma rays. Changes in XO activity and ROS production were measured in the aortas of the mice to assess IR-induced oxidative stress.   </td> <td> Compared to sham-irradiated controls, irradiated mice experienced a ~68% increase in ROS production and a ~46% increase in XO activity.   </td> </tr> <tr> <td> Karam & Radwan, 2019  </td> <td> In vivo. Adult male Albino rats underwent irradiation with 5 Gy of 137Cs gamma rays at a rate of 0.665 cGy/s. Activity levels of the antioxidants, SOD and CAT, present in the heart tissue were measured to assess IR-induced oxidative stress.   </td> <td> Compared to the sham-irradiated controls, SOD and CAT activity decreased by 57% and 43%, respectively, after irradiation.   </td> </tr> <tr> <td> Cervelli et al., 2017  </td> <td> In vitro. HUVECs were irradiated with 0.25 Gy of X-rays at a rate of 91 mGy/min. ROS production was measured to analyze IR-induced oxidative stress.   </td> <td> Compared to the sham-irradiated controls, irradiated mice experienced a ~171% increase in ROS production (not significant).   </td> </tr> <tr> <td> Mansour, 2013  </td> <td> In vivo. Male Wistar rats underwent whole-body irradiation with 6 Gy of 137Cs gamma rays at a rate of 0.012 Gy/s. MDA was measured from heart homogenate, along with the antioxidants: SOD, GSH, and GSH-Px.   </td> <td> Compared to sham-irradiated controls, MDA increased by 65.9%. SOD, GSH-Px, and GSH decreased by 33.8%, 42.4%, and 50.0%, respectively.   </td> </tr> <tr> <td> Soltani, 2016  </td> <td> In vitro. HUVECs were irradiated with 2 Gy of 60Co gamma rays at a dose rate of 0.6 Gy/min. Markers of oxidative stress, including reduced GSH and TBARS, were measured to assess GSH depletion and LPO, respectively. </td> <td> Compared to non-irradiated controls, sham-irradiated cells experienced a ~28% decrease in GSH and a ~433% increase in TBARS. </td> </tr> <tr> <td> Wang et al., 2019b  </td> <td> In vitro. HUVECs were irradiated with 0.2, 0.5, 1, 2, and 5 Gy of 137Cs gamma rays. ROS production was measured to assess IR-induced oxidative stress.   </td> <td> Compared to sham-irradiated controls, ROS production saw a significant, ~32% increase at 5 Gy. While changes to ROS production were insignificant at doses <2 Gy, they followed a linear increase from 0-5 Gy.   </td> </tr> <tr> <td> Sharma et al., 2018  </td> <td> In vitro. HUVECs were irradiated with 9 Gy of photons. ROS production was measured to determine the effects of IR on oxidative stress.   </td> <td> Compared to sham-irradiated controls, irradiated HUVECs saw a significant, ~133% increase in ROS production.   </td> </tr> <tr> <td> Hatoum et al., 2006  </td> <td> In vivo. Sprague-Dawley rats were irradiated with 9 fractions of 2.5 Gy of X-rays for a cumulative dose of 22.5 Gy at a rate of 2.43 Gy/min. Production of the ROS superoxide and peroxide in gut arterioles were measured to determine the level of oxidative stress caused by irradiation.  </td> <td> ROS production started increasing compared to the sham-irradiated control after the second dose and peaked at the fifth dose. By the ninth dose, superoxide production increased by 161.4% and peroxide production increased by 171.3%.   </td> </tr> <tr> <td> Phillip et al., 2020  </td> <td> In vitro. Human TICAE cells were irradiated with 0.25, 0.5, 2, and 10 Gy of 60Co gamma rays at a rate of 0.4 Gy/min. Levels of the antioxidants, SOD1 and PRDX5 were measured to assess oxidative stress from IR exposure.   </td> <td> While SOD1 levels did not follow a dose-dependent pattern. At 2 Gy, SOD1 decreased about 0.5-fold. At 1-week post-irradiation, PRDX5 remained at approximately control levels for doses <2 Gy but increased by ~60% from 2-10 Gy. PRDX5 only decreased at 2 Gy and 24h post-irradiation.   </td> </tr> <tr> <td> Ramadan et al., 2020  </td> <td> In vitro. Human TICAE/TIME cells were irradiated with 0.1 and 5 Gy of X-rays at a dose rate of 0.5 Gy/min. Intracellular ROS production was measured to determine the extent of IR-induced oxidative stress.   </td> <td> ROS production saw a dose-dependent increase in both TICAE and TIME cells. By 45 mins post-irradiation, 0.1 Gy of IR had induced increases to ROS production of ~3.6-fold and ~8-fold in TICAE and TIME cells, respectively, compared to sham-irradiated controls. 5 Gy of IR caused more significant increases to ROS production of ~18-fold and ~17-fold in TICAE and TIME cells, respectively, compared to sham-irradiated controls.   </td> </tr> <tr> <td> Shen et al., 2018  </td> <td> In vivo. 8-week-old, female, C57BL/6 mice were irradiated with 18 Gy of X-rays. Levels of the oxidative markers, 4-HNE and 3-NT, and the antioxidants, CAT and heme oxygenase 1 (HO-1) were measured in the aortas of the mice.   </td> <td> Compared to sham-irradiated controls, irradiated mice saw maximum increases of ~1.75-fold on day 14 and ~2.25-fold on day 7 to 4-HNE and 3-NT levels, respectively. While CAT levels decreased up to 0.33-fold on day 7, HO-1 levels increased by ~1.9-fold on day 7.   </td> </tr> <tr> <td> Ungvari et al., 2013  </td> <td> In vitro. The CMVECs of adult male rats were irradiated with 2, 4, 6, and 8 Gy of 137Cs gamma rays. Production of the reactive oxygen species, peroxide and O2.-, were measured to assess the extent of IR-induced oxidative stress.   </td> <td> Compared to sham-irradiated controls, production of peroxide in CMVECs of irradiated mice 1 day-post exposure increased in a dose-dependent manner from 0-8 Gy, with significant changes observed at doses >4 Gy. At 8 Gy, peroxide production had increased ~3.25-fold. Production of O2.- followed a similar dose-dependent increase with significant observed at doses >6 Gy. At 8 Gy, O2.- production increased ~1.6-fold. 14 days post-exposure, IR-induced changes to ROS production were not significant for either peroxide or O2.- and did not show a dose-dependent pattern. ROS production progressively decreased from 0-4 Gy and then recovered from 6-8 Gy back to control levels.   </td> </tr> <tr> <td> Ahmadi et al., 2021  </td> <td> In vitro. HLEC and HLE-B3 cells were exposed to 0.1, 0.25 and 0.5 Gy of gamma irradiation at 0.3 and 0.065 Gy/min. Intracellular ROS levels were measured.   </td> <td> In HLE-B3 cells, there were about 7 and 17% ROS-positive cells 1 h after exposure to 0.25 and 0.5 Gy respectively at 0.3 Gy/min.   24 h after exposure there were about 10% ROS-positive cells after 0.5 Gy at 0.3 Gy/min.   1 h after exposure there were about 13 and 17% ROS-positive cells at 0.25 and 0.5 Gy and 0.065 Gy/min.   24 h after exposure there were 8% ROS-positive cells after 0.5 Gy and 0.065 Gy/min.   In human lens epithelial cells 1 h after exposure there were about 10 and 19% ROS-positive cells after 0.25 and 0.5 Gy at 0.3 Gy/min.   After exposure to 0.5 Gy at 0.065 Gy/min there were about 16 and 9% ROS-positive cells one and 24 h after exposure.      </td> </tr> <tr> <td> Ji et al, 2015  </td> <td> In vitro. HLECs were exposed to UVB-irradiation (297 nm; 2 W/m2) for 0 – 120 min. Total antioxidative capability (T-AOC), ROS levels, MDA, and SOD were measured at various time points at 5-120 min.   </td> <td> HLECs exposed to 1 W/m2 UVB for 0 - 120 min (representative of dose) showed a gradual increase in ROS levels that began to plateau 105 min post-irradiation at an ROS level 750 000x control.         </td> </tr> <tr> <td> Hua et al, 2019  </td> <td> In vitro. HLECs were exposed to 4050, 8100 and 12,150 J/m2 of UVB-irradiation at 1.5, 3.0 and 4.5 W/m2. MDA, SOD, GSH-Px, and GSH were measured.  </td> <td> MDA activity as a ratio of the control increased about 1.5 at 3.0 W/m2 and about 3 at 4.5 W/m2.   SOD activity as a ratio of the control decreased about 0.1 at 1.5 W/m2, 0.2 at W/m2, and 0.3 at 4.5 W/m2.   GSH-Px activity as a ratio of the control decreased about 0.02 at 3.0 W/m2 and 0.2 at 4.5 W/m2.   GSH activity .as a ratio of the control decreased about 0.2 at 3.0 W/m2 and 0.7 at 4.5 W/m2.      </td> </tr> <tr> <td> Chen et al, 2021  </td> <td> In vivo. Male rats were irradiated with 0, 0.4, 1.2 and 3.6 Sv of neutron-irradiation at 14, 45 and 131 mSv/h. In rat lenses, MDA, GSH, and SOD, were measured.  </td> <td> MDA concentration decreased by about 1.5 nmol/mg protein at 1.2 Sv and increased by about 7.5 nmol/mg protein relative to the control at 3.6 Sv.   GSH concentration increased by about 3.5 µg/mg protein and decreased by about 1 µg/mg protein relative to the control at 3.6 Sv (neutron radiation).   SOD activity decreased by about 0.08 U/mg protein relative to the control at 3.6 Sv.   It should be noted that Sv is not the correct unit when investigating animals and cultured cells, radiation should have been measured in Gy (ICRU, 1998).      </td> </tr> <tr> <td> Zigman et al., 2000  </td> <td> In vitro. Rabbit LECs were exposed to 3-12 J/cm2 of UVA-irradiation (300-400 nm range, 350 nm peak). CAT activity was assayed to demonstrate oxidative stress.    </td> <td> Rabbit LECs exposed to 3 – 12 J/cm2 UVA showed an approximately linear decrease in catalase activity (indicative of increased oxidative stress) with the maximum dose displaying a 3.8x decrease.   </td> </tr> <tr> <td> Chitchumroonchokchai et al, 2004  </td> <td> In vitro. HLECs were exposed to 300 J/m2 of UVB-irradiation at 3 mW/cm2. MDA and HAE were used to measure oxidative stress.  </td> <td> The concentration of MDA and HAE increased by about 900 pmol/mg protein compared to the control after irradiation with 300 J/m2 UVB.   </td> </tr> <tr> <td> Zigman et al, 1995  </td> <td> In vitro. Rabbit and squirrel LECs were exposed to 6, 9, 12, 15 and 18 J/m2 of UV-irradiation at 3 J/cm2/h (300-400 nm range, 350 nm peak). CAT was used to measure oxidative stress levels.   </td> <td> The CAT activity was 10% of the control activity at 6 J/cm2, and then decreased to 0% of the control activity at 18 J/cm2 (99.9% UV-A and 0.1% UV-B).   </td> </tr> <tr> <td> Karimi et al, 2017  </td> <td> In vivo. Adult rats were exposed to 15 Gy of gamma 60Co-irradiation at a dose rate of 98.5 cGy/min. In lens tissue, MDA, thiobarbituric acid (TBA), and GSH levels were used to indicate oxidative stress.  </td> <td> MDA concentration increased from 0.37 +/- 0.03 to 1.60 +/- 0.16 nmol/g of lens after irradiation.   GSH concentration decreased from 0.99 +/- 0.06 to 0.52 +/- 0.16 µmol/g of lens after exposure.      </td> </tr> <tr> <td> Rong et al., 2019  </td> <td> In vitro. HLECs were exposed to UVB-irradiation (297 nm; 2 W/m2 for 10 min). Intracellular H2O2 and superoxide levels were measured.   </td> <td> The amount of ROS was measured as the dicholofluoroscein (DCFH-DA) fluorescence density, which increased about 10-fold relative to the control.   A similar test but with dihydroethidium (DHE) staining showed a fluorescence density increase of about 3-fold relative to the control.      </td> </tr> <tr> <td> Kubo et al., 2010  </td> <td> In vitro. Lenses isolated from mice were exposed to 400 or 800 J/m2 of UVB-irradiation. ROS levels were measured.    </td> <td> The ratio of ROS level/survived LECs increased from about 175 to 250% after exposure to 400 and 800 J/m2 UVB respectively.   </td> </tr> <tr> <td> Kang et al., 2020  </td> <td> In vitro. HLECs were exposed to 0.09 mW/cm2 UVB-irradiation (275-400 nm range, 310 nm peak) for 15 mins. MDA and SOD activity were measured.  </td> <td> MDA activity increased about 30% compared to control after 15 mins of 0.09 mW/cm2 UVB exposure. SOD activity decreased about 50% compared to control under the same conditions.  </td> </tr> <tr> <td> Yang et al., 2020  </td> <td> In vitro. HLECs were irradiated with 30 mJ/cm2 of UVB-irradiation. ROS levels were determined.  </td> <td> The level of ROS production in HLECs increased approximately 5-fold as determined by 2’,7’-dichlorofluorescein diacetate after exposure to 30 mJ/cm2 UVB.  </td> </tr> <tr> <td> Zhang et al., 2012  </td> <td> In vivo. Adult mice were exposed to 20.6 kJ/m2 UV-irradiation (313 nm peak; 1.6 mW/cm2). GSH levels were measured in lens homogenates.  </td> <td> Decrease in GSH of about 1 and 2 µmol/g wet weight compared to control after 1 and 16 months respectively after 20.6 kJ/m2 UV (313 nm peak) at 1.6 mW/cm2.  </td> </tr> </tbody> </table>

Time Concordance  <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Tseng et al., 2014  </td> <td> In vitro. Neural stem/precursor cells isolated from mouse subventricular and hippocampal dentate subgranular zones were exposed to 1-200 cGy of 56Fe irradiation at dose rates ranging from 5-50 cGy/min. RONS were measured from 1 to 8 weeks post-irradiation.   </td> <td> Compared to sham-irradiated controls, a trend toward increasing oxidative stress was seen, particularly at 1- and 4-weeks post-irradiation where RONS levels showed dose-responsive increases. The greatest rise was also seen at 10 cGy where relative RONS levels increased ~2-fold from 1 to 4 weeks, ~3-fold from 4 to 6 weeks and ~2 fold from 6 to 8 weeks. RONS were also found increased at doses as low as 2 cGy at 12 and 24h post-irradiation.   </td> </tr> <tr> <td> Suman et al., 2013  </td> <td> In vivo. Female mice were exposed to either 1.3 Gy of 56Fe irradiation (1 GeV/nucleon; dose rate of 1 Gy/min) or 2 Gy of gamma irradiation (dose rate of 1 Gy/min). ROS were measured in cerebral cortical cells at 2 and 12 months.  </td> <td> ROS levels showed statistically significant increases after 56Fe irradiation at both 2 and 12 months, while gamma irradiation led to an increase at only 2 months. The percent fluorescence intensity of ROS levels for control, gamma irradiated and 56Fe-irradiated were approximately 100, 115 and 140 at 2 months, and 100, 90 and 125 at 12 months, respectively.    </td> </tr> <tr> <td> Limoli et al., 2004  </td> <td> In vitro. Neural stem/precursor cells isolated from mouse subventricular and hippocampal dentate subgranular zones were exposed to 1 or 5 Gy of 56Fe irradiation at dose rates ranging from 4.5 Gy/min. RONS were measured at various time points until 33 days post-exposure.   </td> <td> ROS levels exhibited statistically significant fluctuations, increasing over the first 12h before dropping at 18h and rising again at 24h. At 5 Gy, ROS levels fluctuated with a peak at 7 days, a decrease at 13 days, an increase at 25 days, and a decrease below control levels at 33 days. At 1 Gy, ROS levels peaked at 25 days and also decreased below control at 33 days.    </td> </tr> <tr> <td> Gledzinski et al., 2005  </td> <td> In vitro. Neural precursor cells derived from rats were irradiated with 1, 2, 5 and 10 Gy of proton (1.7-1.9 Gy/min). ROS levels were determined at 5-25h post-irradiation.   </td> <td> Proton irradiation led to a rapid rise in ROS levels, with the increase most marked at 6h (approximately 10-70% for 1 and 10 Gy, respectively). The increase in ROS persisted for 24h, mainly for 10 Gy where the ROS levels were around 30% above control at the 12, 18 and 24h mark.    </td> </tr> <tr> <td> Acharya et al., 2010  </td> <td> In vitro. Human neural stem cells were subjected to 1, 2 or 5 Gy of gamma irradiation at a dose rate of 2.2 Gy/min. RONS and superoxide levels were measured at various time points until 7 days.    </td> <td> Intracellular RONS and superoxide levels showed significant increase from 2- to 4-fold at 12h. At 7 days, levels of RONS increased and were dose-responsive, elevated by ~3- to 7-fold and 3- to 5-fold, respectively, over sham-irradiated controls.     </td> </tr> <tr> <td> Rugo and Schiestl, 2004  </td> <td> In vitro. Human lymphoblast cell lines (TK6 and TK6 E6) were irradiated with 2 Gy of X-irradiation at a dose rate of 0.72 Gy/min. ROS levels were measured at various time points until 29 days.   </td> <td> In the TK6 E6 clones, there was only a significant ROS increase at day 29 (45.7 DCF fluorescence units). In the TK6 clones, there were significant ROS increases at days 13 (26.0 DCF fluorescence units), 15 (26.3 DCF fluorescence units) and 20 (38.1 DCF fluorescence units), with a strong trend of increased ROS in the treated group at day 25. On day 18, ROS levels decreased in the irradiated group, and there was no significant difference at day 29.   </td> </tr> <tr> <td> Huang et al., 2018  </td> <td> In vitro. Murine RAW264.7 cells were irradiated with 2 Gy of gamma rays at a rate of 0.83 Gy/min. ROS levels were measured at 2 and 8 h post-irradiation.   </td> <td> ROS levels in irradiated RAW264.7 cells decreased by ~10% from 2 h post-exposure to 8 h post-exposure (from ~138% above control at 2 h to ~98% above control at 8).    </td> </tr> <tr> <td> Zhang et al., 2018  </td> <td> In vitro. hBMMSCs were irradiated with 2 Gy of X-rays at a rate of 0.6 Gy/min. Relative ROS concentration was measured at 0, 0.5, 2, 3, 6, 8, and 12 h post-irradiation.   </td> <td> ROS levels increased in time dependent manner until a peak of ~90% above control level at 3 h-post irradiation, and then steadily declined back to approximately control levels at 12 h post-irradiation.   </td> </tr> <tr> <td> Phillip et al., 2020  </td> <td> In vitro. Human TICAE cells were irradiated with 0.25, 0.5, 2, and 10 Gy of 60Co gamma rays at a rate of 400 mGy/min. Levels of the antioxidants, SOD1 and PRDX5 were measured at 4 h, 24 h, 48 h, and 1-week post-irradiation to assess oxidative stress from IR exposure.   </td> <td> SOD1 levels did not follow a time-dependent pattern. However, SOD1 decreased at 2 Gy for every timepoint post-irradiation. While PRDX5 levels stayed at approximately baseline levels for the first two days after exposure to 10 Gy of radiation, levels elevated by ~1.6-fold after 1 week.   </td> </tr> <tr> <td> Ramadan et al., 2020  </td> <td> In vitro. Human TICAE/TIME cells were irradiated with 0.1 and 5 Gy of X-rays at a rate of 0.5 Gy/min. Intracellular ROS production was measured at 45 mins, 2 h, and 3 h post-irradiation.  </td> <td> After irradiation, ROS production saw time-dependent decreases in both TICAE and TIME cells from 45 mins to 3 h post-exposure. ROS production was elevated at 45 mins but returned to approximately baseline levels at 2 and 3 h.   </td> </tr> <tr> <td> Shen et al., 2018  </td> <td> In vivo. 8-week-old, female, C57BL/6 mice were irradiated with 18 Gy of X-rays. Levels of the oxidative markers, 4-HNE and 3-NT, and the antioxidants, CAT and heme HO-1 were measured the aortas of the mice at 3, 7, 14, 28, and 84 days post-irradiation.  </td> <td> Significant changes were observed in 4-HNE, 3-NT, CAT, and HO-1 levels of irradiated mice after 3 days. 3-NT and HO-1 levels increased from days 3 to 7 and then progressively decreased, while 4-HNE levels followed the same pattern but with a peak at day 14. CAT levels were at their lowest at day 3 and followed a time dependent increase until day 84.   </td> </tr> <tr> <td> Ungvari et al., 2013  </td> <td> In vitro. The CMVECs of adult male rats were irradiated with 2, 4, 6, and 8 Gy of 137Cs gamma rays. Production of the reactive oxygen species, peroxide and superoxide, were measured at 1- and 14-days post-irradiation.   </td> <td> ROS production was generally higher at day 1 than day 14, with the difference becoming progressively more significant from 2-8 Gy. Peroxide production was reduced from a ~3.25-fold increase compared to controls at day 1 back to baseline levels at day 14. Superoxide production had a ~1.6-fold increase at day 1 recover to baseline levels at day 14.   </td> </tr> <tr> <td> Ahmadi et al., 2021  </td> <td> In vitro. HLEC and HLE-B3 cells were exposed to 0.1, 0.25 and 0.5 Gy of gamma irradiation at 0.3 and 0.065 Gy/min. ROS levels were measured.         </td> <td> In human LECs immediately exposed to 0.25 Gy gamma rays, the level of ROS positive cells increased by 5%, relative to control, 1 h post-irradiation.   </td> </tr> <tr> <td> Jiang et al., 2006  </td> <td> In vitro. HLECs were exposed to UV-irradiation at a wavelength over 290 nm (30 mJ/cm2). ROS levels were measured.   </td> <td> Approximately 10-fold increase in ROS generation 15 mins after exposure to 30 mJ/cm2 UV.   </td> </tr> <tr> <td> Pendergrass et al., 2010  </td> <td> In vivo. Female mice were irradiated with 11 Gy of X-irradiation at a dose rate of 2 Gy/min. ROS levels in the lenses were used to represent oxidative stress.   </td> <td> 9 months after irradiation with 11 Gy X-rays at 2 Gy/min there’s 2250% cortical ROS relative to the control.   3 months after there was no significant change.               </td> </tr> <tr> <td> Belkacemi et al., 2001  </td> <td> In vitro. Bovine lens cells were exposed to 10 Gy of X-irradiation at 2 Gy/min. GSH levels were measured.  </td> <td> The intracellular GSH pool was measured by a decrease of about 15% monobromobimane fluorescence relative to the control 24 h after exposure to 10 Gy X-rays at 2 Gy/min and there was a decrease of about 40% relative to the control by 120 h.   </td> </tr> <tr> <td> Weinreb and Dovrat, 1996  </td> <td> In vitro. Bovine lenses were irradiated with 22.4 J/cm2 (10 min) and 44.8 J/cm2 (100 min) of UVA-irradiation at 8.5 mW/cm2. CAT levels were determined.   </td> <td> CAT activity decreased from 1.75 (control) to 0.5 U/mg protein at 48-168 h after exposure to 44.8 J/cm2 UV-A.         </td> </tr> <tr> <td> Cencer et al., 2018  </td> <td> In vitro. HLECs were exposed to 0.014 and 0.14 J/cm2 of UVB-irradiation at 0.09, 0.9 mW/cm2 for 2 and 5 min. ROS levels (mainly H2O2) were measured.        </td> <td> About 5 min after exposure to both 0.09 and 0.9 mW/cm2 UVB for 2.5 mins there is an increase of about 4 average brightness minus control (densitometric fluorescence scanning for ROS, mostly indicating H2O2).    About 90 and 120 min after exposure to 0.9 mW/cm2 the average brightness minus control is about 35 and 20 respectively.               </td> </tr> <tr> <td> Yang et al., 2020  </td> <td> In vitro. HLECs were irradiated with 30 mJ/cm2 of UVB-irradiation. Intracellular ROS levels were measured.  </td> <td> The level of ROS production in HLECs increased approximately 5-fold as determined by 2’,7’-dichlorofluorescein diacetate 24 h after exposure to 30 mJ/cm2 UVB.  </td> </tr> <tr> <td> Zhang et al., 2012  </td> <td> In vivo. Adult mice were exposed to 20.6 kJ/m2 UV-irradiation (313 nm peak; 1.6 mW/cm2). GSH levels were measured in lens homogenates.   </td> <td> Decrease in GSH of about 1 and 2 µmol/g wet weight compared to control after 1 and 16 months respectively after 20.6 kJ/m2 UV (313 nm peak) at 1.6 mW/cm2.  </td> </tr> </tbody> </table>

The relationship between deposition of energy and increased oxidative stress leads to several feedforward loops. Firstly, ROS activates the transforming growth factor beta (TGF)-β, which increases the production of ROS. This process is modulated in normal cells containing PRDX-6, or cells with added MnTBAP, which will both prevent TGF-β from inducing ROS formation (Fatma et al., 2005). Secondly, ROS can damage human mitochondrial DNA (mtDNA), this can then cause changes to the cellular respiration mechanisms, leading to increased ROS production (Turner et al., 2002; Zhang et al., 2010; Tangvarasittichai & Tangvarasittichai, 2019, Ahmadi et al., 2021; Yves, 2000). Some other feedback loops through which deposition of energy causes oxidative stress are discussed by Soloviev & Kizub (2019).

High Male Moderate Female High Unspecific

High

Juvenile

Moderate

Adult

Moderate Moderate High Low

Most evidence is derived from in vitro studies, predominately using rabbit models. Evidence in humans and mice is moderate, while there is considerable available data using rat models. The relationship is applicable in both sexes, however, males are used more often in animal studies. No studies demonstrate the relationship in preadolescent animals, while adolescent animals were used very often, and adults were used occasionally in in vivo studies.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b42ea6495a8> AOPWiki 2022-09-28T12:01:48

2023-04-25T11:32:48

<upstream-id>8d6a4475-5817-4386-be53-eb911186774f</upstream-id> <downstream-id>0e490634-53e7-457f-8aa8-4395c8e932ae</downstream-id> Direct deposition of ionizing energy refers to imparted energy interacting directly with the DNA double helix and producing randomized damage. This can be in the form of double strand breaks (DSBs), single-strand breaks, base damage, or the crosslinking of DNA to other molecules (Smith et al., 2003; Joiner, 2009; Christensen, 2014; Sage and Shikazono, 2017). Among these, the most detrimental type of DNA damage to a cell is DSBs. They are caused by the breaking of the sugar-phosphate backbone on both strands of the DNA double helix molecule, either directly across from each other or several nucleotides apart (Ward, 1988; Iliakis et al., 2015). This occurs when high-energy subatomic particles interact with the orbital electrons of the DNA causing ionization (where electrons are ejected from atoms) and excitation (where electrons are raised to higher energy levels) (Joiner, 2009). The number of DSBs produced and the complexity of the breaks is highly dependent on the amount of energy deposited on and absorbed by the cell. This can vary as a function of the dose-rate (Brooks et al., 2016) and the radiation quality which is a function of its linear energy transfer (LET) (Sutherland et al., 2000; Nikjoo et al., 2001; Jorge et al., 2012). LET describes the amount of energy that an ionizing particle transfers to media per unit distance (Smith et al., 2003; Okayasu, 2012a; Christensen et al., 2014). High LET radiation, such as alpha particles, heavy ion particles, and neutrons can deposit larger quantities of energy within a single track than low LET radiation, such as γ-rays, X-rays, electrons, and protons (Kadhim et al., 2006; Franken et al., 2012; Frankenberg et al., 1999; Rydberg et al., 2002; Belli et al., 2000; Antonelli et al., 2015). As such, radiation with higher LETs tends to produce more complex, dense structural damage, particularly in the form of clustered damage, in comparison to lower LET radiation (Nikjoo et al., 2001; Terato and Ide, 2005; Hada and Georgakilas, 2008; Okayasu, 2012a; Lorat et al., 2015; Nikitaki et al., 2016). Thus, the complexity and yield of clustered DNA damage increases with ionizing density (Ward, 1988; Goodhead, 2006). However, clustered damage can also be induced even by a single radiation track through a cell. While the amount of DSBs produced depends on the radiation dose (see dose concordance), it also depends on several other factors. As the LET increases, the complexity of DNA damage increases, decreasing the repair rate, and increasing toxicity (Franken et al., 2012; Antonelli et al., 2015).

The strategy for collating the evidence on radiation stressors to support the relationship is described in Kozbenko et al 2022.  Briefly, a scoping review methodology was used to prioritize studies based on a population, exposure, outcome, endpoint statement.

Overall Weight of Evidence for this KER: High

The biological rationale linking the direct deposition of energy on DNA with an increase in DSB formation is strongly supported by numerous literature reviews that are available on this topic (J .F. Ward, 1988; Lipman, 1988; Hightower, 1995; Terato & Ide, 2005; Goodhead, 2006; Kim & Lee, 2007; Asaithamby et al., 2008; Hada & Georgakilas, 2008; Jeggo, 2009; Clement, 2012; Okayasu, 2012b; Stewart, 2012; M. E. Lomax et al., 2013; EPRI, 2014; Hamada, 2014; Moore et al., 2014; Desouky et al., 2015; Ainsbury, 2016; Foray et al., 2016; Hamada & Sato, 2016; Hamada, 2017a; Sage & Shikazono, 2017; Chadwick, 2017; Wang et al., 2021; Nagane et al., 2021; Sylvester et al., 2018; Baselet et al., 2019). Ionizing radiation can be in the form of high energy particles (such as alpha particles, beta particles, or charged ions) or high energy photons (such as gamma-rays or X-rays). Ionizing radiation can break the DNA within chromosomes both directly and indirectly, as shown through using velocity sedimentation of DNA through neutral and alkaline sucrose gradients. The most direct path entails a collision between a high-energy particle or photon and a strand of DNA. Additionally, excitation of secondary electrons in the DNA allows for a cascade of ionization events to occur, which can lead to the formation of multiple damage sites (Joiner, 2009). As an example, high-energy electrons will traverse a DNA molecule in a mammalian cell within 10-18 s and 10-14 s, resulting in 100,000 ionizing events per 1 Gy dose in a 10 μm cell (Joiner, 2009). The amount of damage can be influenced by factors such as the cell cycle stage and chromatin structure. It has been shown that in more condensed, packed chromatin structures such as those present in intact cells and heterochromatin, it is more difficult for the DNA to be damaged (Radulescu et al., 2006; Agrawala et al., 2008; Falk et al., 2008; Venkatesh et al., 2016). In contrast, DNA damage is more easily induced in lightly-packed chromatin such as euchromatin and nucleoids, (Radulescu et al., 2006; Falk et al., 2008; Venkatesh et al., 2016). Of the possible radiation-induced DNA damage types, DSB is considered to be the most harmful to the cell, as there may be severe consequences if this damage is not adequately repaired (Khanna & Jackson, 2001; Smith et al., 2003; Okayasu, 2012a; M. E. Lomax et al., 2013; Rothkamm et al., 2015). A considerable fraction of DSBs can also be formed in cells through indirect mechanisms.  In this case, deposited energy can split water molecules near DNA, which can generate a significant quantity of reactive oxygen species in the form of hydroxyl free radicals (Ward, 1988; Wolf, 2008; Desouky et al., 2015; Maier et al., 2016, Cencer et al., 2018; Bains, 2019; Ahmadi et al., 2021). Estimates using models and experimental results suggest that hydroxyl radicals may be present within nanoseconds of energy deposition by radiation (Yamaguchi et al., 2005). These short-lived but highly reactive hydroxyl radicals may react with nearby DNA. This will produce DNA damage, including single-strand breaks and DSBs (Ward, 1988; Sasaki, 1998; Desouky et al., 2015; Maier et al., 2016). DNA breaks are especially likely to be produced if the sugar moiety is damaged, and DSBs occur when two single-strand breaks are in close proximity to each other (Ward, 1988).

Empirical data strongly supports this KER. The evidence presented below is summarized in table 1. The types of DNA damage produced by ionizing radiation and the associated mechanisms, including the induction of DSBs, are reviewed by Lomax et al. (2013) and documents produced by international radiation governing frameworks (Valentin, 1998; UNSCEAR, 2000). Other reviews also highlight the relationship between the deposition of energy by radiation and DSB induction, and discuss the various methods available to detect these DSBs (Terato & Ide, 2005; Rothkamm et al., 2015; Sage & Shikazono, 2017). A visual representation of the time frames and dose ranges probed by the dedicated studies discussed here is shown in Figures 1 & 2 below.     <img alt="" src="https://aopwiki.org/system/dragonfly/production/2022/10/12/4zw4nw353c_ke1_mie_dsb_dose_v2.png" style="height:734px; width:1000px" /> Figure 1: Plot of example studies (y-axis) against equivalent dose (Sv) used to determine the empirical link between direct deposition of energy and DSBs. The z-axis denotes the equivalent dose rate used in each study. The y-axis is ordered from low LET to high LET from top to bottom.      <img alt="" src="https://aopwiki.org/system/dragonfly/production/2022/10/12/a85jspx5_ke1_mie_dsb_time_v2.png" style="height:706px; width:1000px" /> Figure 2: Plot of example studies (y-axis) against time scales used to determine the empirical link between direct deposition of energy and DSBs. The z-axis denotes the equivalent dose rate used in each study. The y-axis is ordered from low LET to high LET from top to bottom.      Dose Concordance There is evidence in the literature suggesting a dose concordance between the direct deposition of energy by ionizing radiation and the incidence (Grudzenski et al., 2010) of DNA DSBs. Results from in vitro (Aufderheide et al., 1987; Sidjanin, 1993; Bucolo, 1994; Frankenberg et al., 1999; Rogakou et al., 1999; Belli et al., 2000; Sutherland et al., 2000; Lara et al., 2001; Rydberg et al., 2002; Baumstark-Kham et al., 2003; Rothkamm and Lo, 2003; Long, 2004; Kuhne et al., 2005; Sudprasert et al., 2006; Beels et al., 2009; Grudzenski et al., 2010; Liao, 2011; Franken et al., 2012; Bannik et al., 2013; Shelke & Das, 2015; Antonelli et al., 2015; Markiewicz et al., 2015; Allen, 2018; Dalke, 2018; Bains, 2019; Ahmadi et al., 2021; Sabirzhanov et al., 2020; Ungvari et al., 2013; Rombouts et al., 2013; Baselet et al., 2017), in vivo (Reddy, 1998; Sutherland et al., 2000; Rube et al., 2008; Beels et al., 2009; Grudzenski et al., 2010; Markiewicz et al., 2015; Barnard, 2018; Barnard, 2019; Barnard, 2022; Schmal et al., 2019; Barazzuol et al., 2017; Geisel et al., 2012), ex vivo (Rube et al., 2008; Flegal et al., 2015)  and simulation studies (Charlton et al., 1989) suggest that there is a positive, linear, dose-dependent increase in DSBs with increasing deposition of energy across a wide range of radiation types (iron ions, X-rays, ultrasoft X-rays, gamma-rays, photons, UV light, and alpha particles) and radiation doses (1 mGy - 100 Gy) (Aufderheide et al., 1987; Sidjanin, 1993; Frankenberg et al., 1999; Sutherland et al., 2000; de Lara et al., 2001; Baumstark-Khan et al., 2003; Rothkamm & Lo, 2003; Kuhne et al., 2005; Rube et al., 2008; Grudzenski et al., 2010; Bannik et al., 2013; Shelke & Das, 2015; Antonelli et al., 2015; Dalke, 2018; Barazzuol et al., 2017; Ungvari et al., 2013; Rombouts et al., 2013; Baselet et al., 2017; Geisel et al., 2012). DSBs have been predicted to occur at energy deposition levels as low as 75 eV (Charlton et al., 1989).     Time Concordance There is evidence suggesting a time concordance between the direct deposition of energy and the incidence of DSBs. A number of different models and experiments have provided evidence of ionizing radiation-induced foci (IRIF), which can be used to infer DSB formation seconds (Mosconi et al., 2011) or minutes after radiation exposure (Rogakou et al., 1999; Rothkamm and Lo, 2003; Rube et al., 2008; Beels et al., 2009; Kuefner et al., 2009; Grudzenski et al., 2010; Antonelli et al., 2015; Acharya et al., 2010; Sabirzhanov et al., 2020; Rombouts et al., 2013; Nübel et al., 2006; Baselet et al., 2017; Zhang et al., 2017).       Essentiality Deposition of energy is essential for DNA strand breaks. They can also be caused through other routes, such as oxidative stress (Cadet et al., 2012), but under normal physiological conditions deposition of energy is necessary. This was tested through many studies using  various indicators such as 53BP1 foci/cell, γH2AX foci/cell, DNA migration, and the amount of DNA in tails for the comet assay. Various organisms such as humans, mice, rabbits, guinea pigs, and cattle were used. They showed that without the deposition of energy, there was only a negligible amount of DNA strand breaks (Aufderheide et al., 1987; Sidjanin, 1993; Bucolo, 1994; Reddy, 1998; Rogers, 2004; Bannik et al., 2013; Dalke, 2018; Bains, 2019; Barnard, 2019; Barnard, 2021).

Uncertainties and inconsistencies in this KER are as follows: <ul> <li>Studies have shown that dose-rates (Brooks et al., 2016) and radiation quality (Sutherland et al., 2000; Nikjoo et al., 2001; Jorge et al., 2012) are factors that can influence the dose-response relationship.  </li> <li>Low-dose radiation has been observed to have beneficial effects and may even invoke protection against spontaneous genomic damage (Feinendegen, 2005; Day et al., 2007; Feinendegen et al., 2007; Shah et al., 2012; Nenoi et al., 2015; Dalke, 2018). This protective effect has been documented in in vivo and in vitro, as reviewed by ICRP (2007) and UNSCEAR (2008) and can vary depending on the cell type, the tissue, the organ, or the entire organism (Brooks et al., 2016).</li> <li>Depositing ionizing energy is a stochastic event; as such this can influence the location, degree and type of DNA damage imparted on a cell. As an example, studies have shown that mitochondrial DNA may also be an important target for genotoxic effects of ionizing radiation (Wu et al., 1999).</li> </ul>

<table border="1"> <tbody> <tr> <td> Modulating Factor  </td> <td> Details   </td> <td> Effects on the KER   </td> <td> References   </td> </tr> <tr> <td> Nitroxides  </td> <td> Increased concentration  </td> <td> Decreased DNA strand breaks.  </td> <td> DeGraff et al., 1992; Citrin & Mitchel, 2014  </td> </tr> <tr> <td> 5-fluorouracil  </td> <td> Increased concentration  </td> <td> Increased DNA strand breaks.  </td> <td> De Angelis et al., 2006; Citrin & Mitchel, 2014  </td> </tr> <tr> <td> Thiols  </td> <td> Increased concentration  </td> <td> Decreased DNA strand breaks.  </td> <td> Milligan et al., 1995; Citrin & Mitchel, 2014  </td> </tr> <tr> <td> Cisplatin  </td> <td> Increased concentration  </td> <td> Decreased DNA break repair.  </td> <td> Sears & Turchi; Citrin & Mitchel, 2014  </td> </tr> </tbody> </table>

Quantitative understanding of this linkage suggests that DSBs can be predicted upon exposure to ionizing radiation. This is dependent on the biological model, the type of radiation and the radiation dose. In general, 1 Gy of radiation is thought to result in 3000 damaged bases (Maier et al., 2016), 1000 single-strand breaks, and 40 DSBs (Ward, 1988; Foray et al., 2016; Maier et al., 2016) . The table below provides representative examples of the calculated DNA damage rates across different model systems, most of which are examining DNA DSBs.     Dose Concordance  The following tables provide representative examples of the relationship, unless otherwise indicated, all data is significantly significant.  <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Ward, 1988  </td> <td> In vitro. Cells containing approximately 6 pg of DNA were exposed to 1 Gy.  </td> <td> Under the assumption of 6 pg of DNA per cell. 60 eV of energy deposited per event over a total of 1 Gy. Deoxyribose (2.3 pg/cell): 14,000 eV deposited, 235 events. Bases (2.4 pg/cell): 14.7 keV deposited, 245 events. Phosphate (1.2 pg/cell): 7,300 eV deposited, 120 events. Bound water (3.1 pg/cell): 19 keV deposited, 315 events. Inner hydration shell (4.2 pg/cell): 25,000 eV deposited 415 events.  </td> </tr> <tr> <td> Charlton, 1989  </td> <td> In-silico. A computer simulation/model was used to test various types of radiation with doses from 0 to 400 eV (energy deposited) on the amount of DNA damage produced.  </td> <td> Simulated dose-concordance prediction of increase in number of DSBs/54 nucleotide pairs as direct deposition of energy increases in the range 75-400 eV. In the range 100 - 150 eV: 0.38 DSBs/54 nucleotide pairs and at 400 eV: ~0.80 DSBs per 64 nucleotide pairs.  </td> </tr> <tr> <td> Sutherland, 2000  </td> <td> In vitro. Human cells were exposed to 137Cs γ-rays (0 – 100 Gy, 0.16 – 1.6 Gy/min). The frequency of DSBs was determined using gel electrophoresis.  </td> <td> Using isolated bacteriophage T7 DNA and 0-100 Gy of γ radiations, observed a response of 2.4 DSBs per megabase pair per Gy.  </td> </tr> <tr> <td> Rogakou et al., 1999  </td> <td> In vitro. Normal human fibroblasts (IMR90) and human breast cancer cells (MCF7 were exposed to 0.6 and 2 Gy 137Cs γ-rays delivered at 15.7 Gy/min. The number of DSBs were determined by immunoblotting for γ-H2AX.    </td> <td> Radiation doses of 0.6 Gy & 2 Gy to normal human fibroblasts (IMR90) and MCF7 cells resulted in 10.1 & 12.2 DSBs per nucleus on average (0.6 Gy), respectively; increasing to 24 & 27.1 DSBs per nucleus (2 Gy).  </td> </tr> <tr> <td> Kuhne et al., 2005  </td> <td> In vitro. Primary human skin fibroblasts (HSF2) were exposed to 0 – 70 Gy 60Co γ-rays (0.33 Gy/min), X-rays (29 kVp, 1.13 Gy/min), and CKX-rays (0.14 Gy/min). The number of DSBs were determined with pulsed-field gel electrophoresis.  </td> <td> γ-ray and X-ray irradiation of primary human skin fibroblasts (HSF2) at 0 - 70 Gy. γ-rays: (6.1 ± 0.2) x 10-9 DSBs per base pair per Gy, X-rays: (7.0 ± 0.2) x 10-9 DSBs per base pair per Gy. CKX -rays: (12.1 ± 1.9) x 10-9 DSBs per base pair per Gy.  </td> </tr> <tr> <td> Rothkamm, 2003  </td> <td> In vitro. Primary human fibroblast cell lines MRC-5 (lung), HSF1 and HSF2 (skin), and180BR (deficient in DNA ligase IV) were exposed to 1 mGy – 100 Gy X-rays (90 kV). Low doses were delivered at 6 – 60 mGy/min and high doses were delivered at 2 Gy/min. The number of DSBs were determined with pulsed-field gel electrophoresis.  </td> <td> X-ray irradiation of primary human fibroblasts (MRC-5) in the range 1 mGy - 100 Gy, 35 DSBs per cell per Gy.  </td> </tr> <tr> <td> Grudzenski et al, 2010  </td> <td> In vitro. Primary human fibroblasts (HSF1) and C57BL/6NCrl adult mice were exposed to X-rays (2.5 – 200 mGy, 70 mGy/min), and photons (10 mGy – 1 Gy, 2 Gy/min (100 mGy and 1 Gy), and 0.35 Gy/min (10 mGy)). γ-H2AX immunofluorescence was observed to determine DSBs.  </td> <td> X-rays irradiating primary human fibroblasts (HSF1) in the range 2.5 - 100 mGy yielded a response of 21 foci per Gy. When irradiating adult C57BL/6NCrl mice with photons a response of 0.07 foci per cell at 10 mGy was found. At 100 mGy the response was 0.6 foci per cell and finally, at 1 Gy; 8 foci per cell.  </td> </tr> <tr> <td> de Lara, 2001  </td> <td> In vitro. Chinese hamster cells (V79-4) were exposed to 0 – 20 Gy of  60Co γ-rays (2 Gy/min), and ultrasoft X-rays (0.7 – 35 Gy/min): carbon-K shell (0.28 keV), copper L-shell (0.96 keV), aluminum K-shell (1.49 keV), and titanium K-shell (4.55 keV). The number of DSBs were determined with pulsed-field gel electrophoresis.  </td> <td> V79-4 cells irradiated with γ-rays and ultrasoft X-rays (carbon K-shell, copper L-shell, aluminium K-shell and titanum K-shell) in the range 0 - 20 Gy. Response (DSBs per Gy per cell): γ-rays: 41, carbon K-shell: 112, copper L-shell: 94, aluminum K-shell: 77, titanium K-shell: 56.  </td> </tr> <tr> <td> Rübe et al., 2008  </td> <td> In vivo. Brain, lung, heart and small intestine tissue from adult SCID, A-T, BALB/c and C57BL/6NCrl mice; Whole blood and isolated lymphocytes from BALB/c and C57BL/6NCrl mice were exposed to 0.1 – 2 Gy of photons (whole body irradiation, 6 MV, 2 Gy/min) and X-rays (whole body irradiation, 90 kV, 2 Gy/min). γ-H2AX foci were determined with immunochemistry to measure DSBs.  </td> <td> Linear dose-dependent increase in DSBs in the brain, small intestine, lung and heart of C57BL/6CNrl mice after whole-body irradiation with 0.1 - 1.0 Gy of radiation. 0.8 foci per cell (0.1 Gy) and 8 foci per cell (1 Gy).  </td> </tr> <tr> <td> Antonelli et al., 2015  </td> <td> In vitro. Primary human foreskin fibroblasts (AG01522) were exposed to 0 – 1 Gy of 136Cs γ-rays (1 Gy/min), protons (0.84 MeV, 28.5 keV/um), carbon ions (58 MeV/u, 39.4 keV/um), and alpha particles (americium-241, 0.75 MeV/u, 0.08 Gy/min, 125.2 keV/um). γ-H2AX foci were determined with immunochemistry to measure DSBs.  </td> <td> Linear dose-dependent increase in the number of DSBs from 0 - 1 Gy for γ-rays and alpha particles as follows: γ-rays: 24.1 foci per Gy per cell nucleus, alpha particles: 8.8 foci per Gy per cell nucleus.  </td> </tr> <tr> <td> Barnard et al., 2019  </td> <td> In vivo. 10-week-old female C57BL/6 mice were whole-body exposed to 0.5, 1, and 2 Gy of 60Co γ-rays at 0.3, 0.063, and 0.014 Gy/min. p53 binding protein 1 (53BP1) foci were determined via immunofluorescence.  </td> <td> Central LECs showed a linear increase in mean 53BP1 foci/cell with the maximum dose and dose-rate displaying a 78x increase compared to control. Peripheral LECs and lower dose rates displayed similar results, with slightly fewer foci.  </td> </tr> <tr> <td> Ahmadi et al., 2021  </td> <td> In vitro. Human LEC cells were exposed to 137Cs γ-rays at doses of 0, 0.1, 0.25, and 0.5 Gy and dose rates of 0.065 and 0.3 Gy/min. DNA strand breaks were measured using the comet assay.  </td> <td> Human LECs showed a gradual increase in the tail from the comet assay with the maximum dose and dose-rate displaying a 3.7x increase compared to control. Lower dose-rates followed a similar pattern with a lower amount of strand breaks.  </td> </tr> <tr> <td> Hamada et al., 2006  </td> <td> In vitro. Primary normal human diploid fibroblast (HE49) cells were exposed to 0.1, 0.5, and 4 Gy X-rays at 240 kV with a dose rate of 0.5 Gy/min. The number of H2AX foci/cell, which represented DNA strand breaks, was determined 6 – 7 minutes after irradiation through fluorescence microscopy.  </td> <td> Cells displayed a linear increase in the number of H2AX foci/cell, with the maximum dose displaying a 125x increase compared to control.    </td> </tr> <tr> <td>Dubrova & Plumb, 2002</td> <td> </td> <td> At 1 Gy observe 70 DSBs, 1000 single-strange breaks and 2000 damaged DNA bases per cell per Gy. </td> </tr> <tr> <td>Sabirzhanov et al., 2020</td> <td>In vitro. Rat cortical neurons were exposed to 2, 8 or 32 Gy of X rays (320 kV) at a dose rate of 1.25 Gy/min. Western blot was used to measure γ-H2AX, p-ataxia telangiectasia mutated (ATM) and p- ATM/RAD3-related (ATR) levels.  </td> <td>In rat cortical neurons, p-ATM increased at 2, 8, and 32 Gy, with a 15-fold increase at 8 and 32 Gy. γ-H2AX levels increased at 8 and 32 Gy. </td> </tr> <tr> <td>Geisel et al., 2012 </td> <td>In vivo. Patients with suspected coronary artery disease receiving X-rays from computed tomography or conventional coronary angiography had levels of DSBs assessed in blood lymphocytes by γ-H2AX fluorescence. </td> <td>There was a correlation between effective dose (in mSv) and DSBs. For both conventional coronary angiography and computed tomography, a dose of 10 mSv produced about 2-fold more DNA DSBs than a dose of 5 mSv. </td> </tr> <tr> <td>Ungvari et al., 2013 </td> <td>In vitro. Rat cerebromicrovascular endothelial cells and hippocampal neurons were irradiated with 2-10 Gy of 137Cs gamma rays. DNA strand breaks were assessed with the comet assay. </td> <td>DNA damage increased at all doses (2-10 Gy). In the control, less than 5% of DNA was in the tail, while by 6 Gy, 35% of the DNA was in the tail in cerebromicrovascular endothelial cells and 25% was in the tail in neurons. </td> </tr> <tr> <td>Rombouts et al., 2013 </td> <td>In vitro. EA.hy926 cells and human umbilical vein endothelial cells were irradiated with various doses of X-rays (0.25 Gy/min). γ-H2AX foci were assessed with immunofluorescence. </td> <td>More γ-H2AX foci were observed at higher doses in both cell types. In human umbilical vein endothelial cells, few foci/nucleus were observed at 0.05 Gy, with about 23 at 2 Gy. In EA.hy926 cells, few foci/nucleus were observed at 0.05 Gy, with about 37 at 2 Gy. </td> </tr> <tr> <td>Baselet et al., 2017 </td> <td>In vitro. Human telomerase-immortalized coronary artery endothelial cells were irradiated with various doses of X-rays (0.5 Gy/min). Immunocytochemical staining was performed for γ-H2AX and 53BP1 foci. </td> <td>Doses of 0.05 and 0.1 Gy did not increase the number of γ-H2AX foci, but 0.5 Gy increased foci number by 5-fold and 2 Gy by 15-fold. A dose of 0.05 Gy did not increase the number of 53BP1 foci, but 0.1 Gy, 0.5 Gy and 2 Gy increased levels by 3-fold, 7-fold and 8-fold, respectively. </td> </tr> </tbody> </table>   Time Concordance  <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Rogakou et al., 1999  </td> <td> In vitro. Normal human fibroblasts (IMR90), human breast cancer cells (MCF7), human astrocytoma cells (SF268), Indian muntjac Muntiacus muntjak normal skin fibroblasts, Xenopus laevisA6 normal kidney cells, Drosophila melanogaster epithelial cells, and Saccharomyces cerevisiae were exposed to 0.6, 2, 20, 22, 100, and 200 Gy 137Cs γ-rays. Doses below 20 Gy were delivered at 15.7 Gy/min and other doses were delivered in 1 minute. DNA breaks were visualized using γ-H2AX antibodies and microscopy.  </td> <td> DSBs were present at 3 min and persisted from 15 - 60 min.  </td> </tr> <tr> <td> Hamada & Woloschak, 2017  </td> <td> In vitro. human LECs were exposed to 0.025 Gy X-rays at 0.42 – 0.45 Gy/min. 53BP1 foci were measured via indirect immunofluorescence.  </td> <td> In cells immediately exposed to 0.025 Gy, the level of 53BP1 foci/cell increased to 3.3x relative to control 0.5 h post-irradiation.  </td> </tr> <tr> <td> Hamada et al., 2006  </td> <td> In vitro. Primary normal human diploid fibroblast (HE49) cells were exposed to 0.1, 0.5, and 4 Gy (deposition of energy) at 240 kV with a dose rate of 0.5 Gy/min. The number of H2AX foci/cell, which represented DNA strand breaks, was determined through fluorescence microscopy.  </td> <td> In cells immediately exposed to 0.5 Gy, 11% of cells had 18 foci six min post-irradiation, compared to 90% of controls having 0 foci.   </td> </tr> <tr> <td> Acharya et al., 2010  </td> <td> In vitro. Human neural stem cells were exposed to 1, 2 and 5 Gy of γ-rays at a dose rate of 2.2 Gy/min. The levels of γ-H2AX phosphorylation post irradiation were assessed by immunocytochemistry, fluorescence-activated cell sorting (FACS) analysis and γ-H2AX foci enumeration.  </td> <td> The number of cells positive for nuclear γ-H2AX foci peaked at 20 min post-irradiation. After 1h, this level quickly declined.   </td> </tr> <tr> <td> Schmal et al., 2019  </td> <td> In vivo. Juvenile and adult C57BL/6 mice were exposed to whole body 6-MV photons at 2 Gy/min. Irradiations were done in 5x, 10x, 15x and 20x fractions of 0.1 Gy. Double staining for NeuN and 53BP1 was used to quantify DNA damage foci and the possible accumulation in the hippocampal dentate gyrus.  </td> <td> To assess possible accumulation of persisting 53BP1-foci during fractionated radiation, juvenile and adult mice were examined 72 h after exposure to 5×, 10×, 15×, or 20× fractions of 0.1 Gy, compared to controls. The number of persisting 53BP1-foci increased significantly in both juvenile and adult mice during fractionated irradiation (maximum at 1 m post-IR).  </td> </tr> <tr> <td> Dong et al., 2015  </td> <td> In vivo. C57BL/6J mice were exposed to 2 Gy of X-rays at 2 Gy/min using a 6 MV source. γ-H2AX foci were assessed with immunofluorescence in the brain.  </td> <td> At 0.5 h, about 14 γ-H2AX foci/cell were present. This decreased linearly to about 2 foci/cell at 24 h, with no foci/cell from 48 h to 6 weeks.  </td> </tr> <tr> <td> Barazzuol et al., 2017  </td> <td> In vivo. C57BL/6 mice were exposed to 0.1 or 2 Gy of X-rays (250 kV) at a rate of 0.5 Gy/min. 53BP1 foci were quantified with immunofluorescence in neural stem cells and neuron progenitors in the lateral ventricle.   </td> <td> At both 0.5 and 6 h post-irradiation, increased 53BP1 foci were observed, with the highest level at 0.5 h.  </td> </tr> <tr> <td> Sabirzhanov et al., 2020    </td> <td> In vitro. Rat cortical neurons were exposed to 2, 8 or 32 Gy of X rays (320 kV) at a dose rate of 1.25 Gy/min. Western blot was used to measure γ-H2AX, p-ATM and p-ATR levels.   </td> <td> In rat cortical neurons, γ-H2AX, p-ATM and p-ATR all increased at 30 minutes post-irradiation, with a sustained increase until 6 h.  </td> </tr> <tr> <td> Zhang et al., 2017  </td> <td> In vitro. HT22 hippocampal neuronal cellsT were irradiated with X-rays (320 kVp) at 8 or 12 Gy at a dose rate of 4 Gy/min. The comet assay was preformed to assess the DNA double strand breaks in HT22 cells. Western blot was used to measure γ-H2AX and p-ATM.  </td> <td> At 8 Gy, the comet assay showed an increased tail moment at both 30 minutes and 24 h post-irradiation. At 12 Gy, p-ATM was increased over 4-fold at both 30 minutes and 1 h post-irradiation. γ-H2AX was increased over 3-fold at 30 minutes post-irradiation and almost 2-fold at 1 and 24 h.  </td> </tr> <tr> <td> Geisel et al., 2012  </td> <td> In vivo. Patients with suspected coronary artery disease receiving X-rays from computed tomography or conventional coronary angiography had levels of DSBs assessed in blood lymphocytes by γ-H2AX fluorescence.  </td> <td> DSBs were increased at 1 h post-irradiation and returned to pre-irradiation levels by 24 h.  </td> </tr> <tr> <td> Park et al., 2022  </td> <td> In vitro. Human aortic endothelial cells were irradiated with 137Cs gamma rays at 4 Gy (3.5 Gy/min). γ-H2AX was measured with western blot. p-ATM and 53BP1 were determined with immunofluorescence.  </td> <td> γ-H2AX, p-ATM, and 53BP1 were shown increased at 1 h post-irradiation and slightly decreased for the rest of the 6 h but remained elevated above the control.  </td> </tr> <tr> <td> Kim et al., 2014  </td> <td> In vitro. Human umbilical vein endothelial cells were irradiated with 4 Gy of 137Cs gamma rays. γ-H2AX levels were determined with immunofluorescence.  </td> <td> γ-H2AX foci greatly increased at 1 and 6 h post-irradiation, with the greatest increase at 1 h.  </td> </tr> <tr> <td> Dong et al., 2014  </td> <td> In vitro. Human umbilical vein endothelial cells were irradiated with 2 Gy of 137Cs gamma rays. γ-H2AX levels were determined with immunofluorescence.  </td> <td> γ-H2AX foci increased 8-fold at 3 h, 7-fold at 6 h, and 2-fold at 12 and 24 h post-irradiation.  </td> </tr> <tr> <td> Rombouts et al., 2013  </td> <td> In vitro. EA.hy926 cells and human umbilical vein endothelial cells were irradiated with X-rays (0.25 Gy/min). γ-H2AX foci were assessed with immunofluorescence.  </td> <td> The greatest increase in γ-H2AX foci was observed 30 minutes post-irradiation, while levels were still slightly elevated at 24 h.  </td> </tr> <tr> <td> Nübel et al., 2006  </td> <td> In vitro. Human umbilical vein endothelial cells were irradiated with gamma rays at 20 Gy. DNA strand breaks were assessed with the comet assay and western blot for γ-H2AX.  </td> <td> The olive tail moment increased 5-fold immediately after irradiation and returned to control levels by 4 h. A large increase in γ-H2AX was observed at 0.5 h post-irradiation, with lower levels at 4 h but still above the control.  </td> </tr> <tr> <td> Baselet et al., 2017  </td> <td> In vitro. Human telomerase-immortalized coronary artery endothelial cells were irradiated with various doses of X-rays (0.5 Gy/min). Immunocytochemical staining was performed for γ-H2AX and 53BP1 foci.  </td> <td> Increased γ-H2AX and 53BP1 foci were observed at 0.5 h post-irradiation, remaining elevated at 4 h but returning to control levels at 24 h.  </td> </tr> <tr> <td> Gionchiglia et al., 2021  </td> <td> In vivo. Male CD1 and B6/129 mice were irradiated with X-rays at 10 Gy. Brain sections were single or double-stained with antibodies against γ-H2AX and p53BP1.   </td> <td> In the forebrain, cerebral cortex, hippocampus and subventricular zone (SVZ)/ rostral migratory stream (RMS)/ olfactory bulb (OB), γH2AX and p53BP1 positive cells increased at both 15 and 30 minutes post-irradiation, with the greatest increase at 30 minutes.  </td> </tr> </tbody> </table>

There is evidence of a response-response relationship between the deposition of energy and the frequency of DSBs. In studies encompassing a variety of biological models, radiation types and radiation doses, a positive, linear relationship was found between the radiation dose and the number of DSBs (Aufderheide et al., 1987; Sidjanin, 1993; Frankenberg et al., 1999; Sutherland et al., 2000; de Lara et al., 2001; Baumstark-Khan et al., 2003; Rothkamm & Lo, 2003; Kuhne et al., 2005; Rube et al., 2008; Grudzenski et al., 2010; Bannik et al., 2013; Shelke & Das, 2015; Antonelli et al., 2015; Hamada, 2017b; Dalke, 2018; Barazzuol et al., 2017; Geisel et al., 2012; Ungvari et al., 2013; Rombouts et al., 2013; Baselet et al., 2017). There were, however, at least four exceptions reported. When human blood lymphocytes were irradiated with X-rays in vitro, a linear relationship was only found for doses ranging from 6 - 500 mGy; at low doses from 0 - 6 mGy, there was a quadratic relationship reported (Beels et al., 2009). Secondly, simulation studies predicted that there would be a non-linear increase in DSBs as energy deposition increased, with a saturation point at higher LETs (Charlton et al., 1989). Furthermore, primary normal human fibroblasts exposed to 1.2 – 5 mGy X-rays at 5.67 mGy/min showed a supralinear relationship, indicating at low doses, the DSBs are mostly due to radiation-induced bystander effects. Doses above 10 mGy showed a positive linear relationship (Ojima et al., 2008). Finally, in the human lens epithelial cell line SRA01/04, DNA strand breaks appeared immediately after exposure to UVB (0.14 J/cm2) and were repaired after 30 minutes. They then reappeared after 60 and 90 minutes. Both were once again repaired within 30 minutes. However, the two subsequent stages of DNA strand breaks did not occur when exposed to a lower dose of UVB (0.014 J/cm2) (Cencer et al., 2018).

Data from temporal response studies suggests that DSBs likely occur within seconds to minutes of energy deposition by ionizing radiation. In a variety of biological models, the presence of DSBs has been well documented within 10 - 30 minutes of radiation exposure (Rogakou et al., 1999; Rube et al., 2008; Beels et al., 2009; Kuefner et al., 2009; Grudzenski et al., 2010; Antonelli et al., 2015; Acharya et al., 2010; Dong et al., 2015; Barazzuol et al., 2017; Sabirzhanov et al., 2020; Rombouts et al., 2013; Nübel et al., 2006; Baselet et al., 2017; Zhang et al., 2017; Gionchiglia et al., 2021); there is also evidence that DSBs may actually be present within 3 - 5 minutes of irradiation (Kleiman, 1990; Rogakou et al., 1999; Rothkamm & Lo, 2003; Rube et al., 2008; Grudzenski et al., 2010; Cencer et al., 2018). Interestingly, one study that focussed on monitoring the cells before, during and after irradiation by taking photos every 5, 10 or 15 seconds found that foci indicative of DSBs were present 25 and 40 seconds after collision of the alpha particles and protons with the cell, respectively. The number of foci were found to increase over time until plateauing at approximately 200 seconds after alpha particle exposure and 800 seconds after proton exposure (Mosconi et al., 2011). After the 30 minute mark, DSBs have been shown to rapidly decline in number. By 24 hours post-irradiation, DSB numbers had declined substantially in systems exposed to radiation doses between 40 mGy and 80 Gy (Aufderheide et al., 1987; Baumstark-Khan et al., 2003; Rothkamm & Lo, 2003; Rube et al., 2008; Grudzenski et al., 2010; Bannik et al., 2013; Markiewicz et al., 2015; Russo et al., 2015; Antonelli et al., 2015; Dalke, 2018; Bains, 2019; Barnard, 2019; Ahmadi et al., 2021; Dong et al., 2015; Dong et al., 2014; Sabirzhanov et al., 2020; Rombouts et al., 2013; Baselet et al., 2017; Gionchiglia et al., 2021), with the sharpest decrease documented within the first 5 h (Kleiman, 1990; Sidjanin, 1993; Rogakou et al., 1999; Rube et al., 2008; Kuefner et al., 2009; Grudzenski et al., 2010; Bannik, 2013; Markiewicz et al., 2015; Shelke and Das, 2015; Cencer et al., 2018; Acharya et al., 2010; Park et al., 2022; Kim et al., 2014; Nübel et al., 2006). Interestingly, DSBs were found to be more persistent when they were induced by higher LET radiation (Aufderheide et al., 1987, Baumstark-Khan et al., 2003; Antonelli et al., 2015).

Not identified.

High Unspecific

High

All life stages

High High High Low Low Low

This KER is plausible in all life stages, sexes, and organisms with DNA. The majority of the evidence is from In vivo adult mice and human In vitro models that do not specify the sex.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b42ea75e8d0> AOPWiki 2019-08-26T12:00:06

2023-05-15T13:35:01

<upstream-id>b813d3a8-c9cc-49b4-bd70-a0bb3e9e8efd</upstream-id> <downstream-id>0e490634-53e7-457f-8aa8-4395c8e932ae</downstream-id> Oxidative stress is an event that involves both a reduction in free radical scavengers and enzymes, and an increase in free radicals (Brennan et al., 2012). Oxidative stress needs to be maintained within an organism to avoid an excess of damage to biological structures, such as DNA. A redox homeostasis between the radicals and the scavengers is necessary. Between reactive oxygen species (ROS) and reactive nitrogen species (RNS), collectively known as RONS, ROS is particularly significant to oxidative damage and disease states.  Radicals such as singlet oxygen and hydroxyl radical are highly unstable and will react with molecules near their generation point, while radicals such as H2O2 are more stable and membrane permeable, meaning they can travel further to find electrons (Spector, 1990). Since DNA is mainly found in nucleus, ROS needs to reach the nucleus to induce breaks. Hydroxyl radicals, in addition to being highly reactive, are capable of causing DNA damage (Halliwell et al., 2021; Engwa et al., 2020). The regulation of these radicals is achieved by the antioxidant defense response (ADR), which includes enzymatic and non-enzymatic processes. The ADR is recruited to manage RONS levels, with antioxidants such as superoxide dismutase (SOD) functioning as the first line of defense (Engwa et al., 2020). These antioxidants act as scavengers to oxidants, reacting with them before reaching other structures within the cell such as DNA strands (Cabrera et al., 2011; Engwa et al., 2020). The backbone of DNA can fragment upon sustained exposure to ROS (Uwineza et al., 2019; Cannan et al., 2016). Due to low oxidation potentials, adenine and guanine are the DNA bases more prone to oxidation, with oxidation potentials (normal hydrogen electrode) at pH 7 of 1.3 eV and 1.42 eV compared to the 1.6 eV and 1.7 eV of cytosine and thymine (Fong, 2016; Halliwell et al., 2021; Poetsch, 2020). In fact, certain radicals even target guanine in a selective fashion, including carbonate anion radical (CO3•-) and singlet oxygen (1O2) (Halliwell et al., 2021).

The strategy for collating the evidence to support the relationship is described in Kozbenko et al 2022. Briefly, a scoping review methodology was used to prioritize studies based on a population, exposure, outcome, endpoint statement.

Overall Weight of Evidence: Moderate

The biological plausibility of the relationship between increased oxidative stress leading to increased DNA double strand breaks (DSBs) is highly supported by the literature. Evidence was collected from studies conducted using in vitro lens epithelial cell models and derived from humans, bovine and germ line cells (Spector, 1990; Stohs, 1995; Aitken et al., 2001; Spector, 1995).  As this evidence is derived from studies using a human cell model it limits the ability to compare between different taxonomies (Ahmadi et al., 2022; Cencer et al., 2018; Liu et al., 2013; Meng et al., 2021; Smith et al., 2015; Zhou et al., 2016). Other evidence comes from human-derived and rodent models of neuronal and endothelial cells (Cervelli et al., 2014; El-Missiry et al., 2018; Huang et al., 2021; Sakai et al., 2017; Ungvari et al., 2013; Zhang et al., 2017).  ROS that are generated specifically as a result of radiation are highly localized, increasing the likelihood of clustered regions of damage. Naturally generated ROS are more widespread and as a result less capable of generating clusters of damage. ROS will act on DNA bases to oxidize or delete them from the sequence, which create nicks on the strand (Cannan et al., 2016). This damage can occur to any DNA base but bases such as guanine and adenine are most vulnerable due to their low oxidation potentials (Fong, 2016). The mechanism through which the strand break occurs is a result of base excision repair (BER) happening at multiple sites that are too close together, resulting in the spontaneous conversion to DSBs prior to completion of repair. ROS damage to bases clustered together means that multiple sites of BER are happening very close together and while the strand may be able to support the damaged area for one repair, concurrent repairs make surrounding areas more fragile and the strand breaks at the nick sites are under added strain (Cannan et al., 2016). Endogenous damage to DNA as a result of radicals appears over time and mainly as isolated lesions, a pattern understood to be due to the diffusion of the radicals resulting in homogenous distribution patterns. This differs from the specific situations where radiation acts as the stressor to increase oxidative stress, as the radiation track will be highly localized and form radicals within that hit space. This leads to non-homologous lesions and clustered damage to the DNA (Ward et al., 1985).

This relationship is well supported through empirical evidence from studies using stressors such as H2O2, photons, γ- and X-ray, which cause an increase in markers of oxidative stress such as ROS-generating enzymes (lactate dehydrogenase, LDH), and a decrease in free radical scavengers, resulting in DNA strand fragmentation. These studies include both in vivo and in vitro human lens epithelial cells (LECs), mouse, rat and rabbit models, including neuronal cells lines and endothelial cells (Ahmadi et al., 2022; Cencer et al., 2018; Cervelli et al., 2014; El-Missiry et al., 2018; Huang et al., 2021; Liu et al., 2013; Meng et al., 2021; Spector et al., 1997; Ungvari et al., 2013; Zhang et al., 2017; Zhou et al., 2016; Sakai et al., 2017).    Dose/Incidence Concordance   There is high evidence to support a dose concordance between oxidative stress and DNA strand breaks. One in vitro study demonstrated that when ROS levels in LECs are 10% above control following 0.5 Gy gamma ray exposure, DNA strand breaks increased 15-20% above control (Ahmadi et al., 2021). Another study with ultraviolet (UV)B radiation demonstrated higher ROS levels after exposure to 0.14 J/cm2 on in vitro LECs as compared to a lower dose exposure (0.014 J/cm2) for the same time. This corresponded to DNA strand break levels also increasing following high dose rate exposure, but not with the low dose exposure (Cencer et al., 2018).  A 30 µM of H2O2 treatment of in vitro LECs is associated with a 1.4x increase in lactate dehydrogenase (LDH) and 55% more DNA strand breaks (Liu et al., 2013; Smith et al., 2015). Following exposure of in vitro LECs to 50 µM H2O2, increased ROS levels, 4x for LDH, and decreased antioxidant levels, 2x control for GSH-Px and SOD, are associated with a 3x increase in γ-H2AX, a marker of DNA strand breaks (Meng et al., 2021). SOD and GSH decreased by 2-fold following 100 µM H2O2 exposure on LECs with an in vitro model (Zhou et al., 2016). At 125 µM H2O2 intact DNA can be reduced to near 1% of pre-treatment levels for in vitro LECs (Spector et al., 1997). Following 400 µM H2O2 LDH increased to 1200% of control in neuroblastoma cells (Feng et al., 2016) and DNA strand breaks increased to over 150% of control in in vitro LECs (Li et al., 1998).  Exposure of in vitro mouse hippocampal neuronal cells (HT22 cell line) to 10 Gy of X-irradiation resulted in a 5x increase in ROS generation and 3x increase in γ-H2AX (Huang et al., 2021). Another study exposed the same cell line to 8 and 12 Gy of X-irradiation and found a ~2x increase in ROS at 8 Gy and a 4.4x and 3.2x increase in phosphorylation of ataxia telangiectasia mutated (ATM) and γ-H2AX, respectively, 30 minutes after 12 Gy (Zhang et al., 2017). A separate study exposed adult male rats to 4 Gy of γ-irradiation and found 2x increase in 4-hydroxy-2-nonenal (4-HNE) (lipid peroxidation marker) and 3x increase in protein carbonylation. Glutathione reductase decreased by approximately 5x, whereas glutathione and glutathione peroxidase levels decreased by approximately 3x each. Tail DNA %, tail length and tail moment (DNA strand break parameters) increased by approximately 2x, 3x and 6x, respectively (El-Missiry et al., 2018).  Endothelial cells exposed to irradiation also demonstrated the relation between oxidative stress and DNA strand breaks. Rat cerebromicrovascular endothelial cells (CMVECs) exposed to 8 Gy 137Cs gamma rays showed increased cellular peroxide production and mitochondrial oxidative stress. Tail DNA content indicating DNA damage was also increased from 0 to 45% (Ungvari et al., 2013). Human umbilical vein endothelial cells (HUVECs) were irradiated with single (0.125, 0.25, 0.5 Gy), or fractionated (2 × 0.125 Gy, 2 × 0.250 Gy) doses of X-rays. Intracellular ROS production increased in a dose-dependent manner following 0.125, 0.25, 0.5 Gy, and γ-H2AX foci positive cells were observed at all doses (Cervelli et al., 2014). Human aortic endothelial cells (HAECs) exposed to 100µM H2O2 showed 3.7-fold increase in intracellular ROS and a 3.4- and 4.7-fold increase in γ-H2AX and p-ATM, respectively (Sakai et al., 2017).   Time Concordance  There is low evidence to support a time concordance between oxidative stress to strand breaks on DNA. Non-protein-thiol levels, an antioxidant, in in vitro LECs decreased to near zero by 30 min post-exposure to 300 µM H2O2, before recovering to 70% of control by 120 min. At 60 min post-exposure to 125 µM H2O2 there was a start to a divergence from control level DNA fragmentation, one that increased logarithmically, with the treated group having a 14~18% reduction in intact DNA by 9 h post-exposure (Yang et al., 1998). Time response information is difficult to monitor for DNA strand breaks because repair will occur, reducing the number of breaks over time. At 0 minutes post in vitro exposure to 40 µM H2O2 LECs had ~145% of control level DNA strand breaks but that number dropped to ~105% by 30 minutes post-exposure (Li et al., 1998).    Essentiality   Oxidative stress has been found to increase levels of DNA strand breaks above background levels (Li et al., 1998; Liu et al., 2013; Cencer et al., 2018; Ahmadi et al., 2022; El-Missiry et al., 2018; Huang et al., 2021; Cervelli et al., 2017; Sakai et al., 2017). It has been shown that inhibition of oxidative stress leads to a reduction in DNA strand breaks. Sulforaphane (SFN) is an isothiocyanate, which provides chemical protection against ROS by activating the release of enzymatic scavengers. When SFN was added to in vitro LECs exposed to 30 µM H2O2, LDH decreased to near unexposed cell levels from the 1.4x control level without SFN. This LDH drop was associated with reducing the levels of DNA strand breaks induced by oxidative stress almost 3-fold as compared to cells without SFN (Liu et al., 2013). In another study, intact DNA levels were returned to control when treated with µPx-11 (peroxidase that breaks down H2O2), following exposure to 125 µM H2O2. This was a near 100% recovery compared to the drop seen in LECs that did not contain µPx-11 (Spector et al., 1997).  Within the brain of Wistar rats, epigallocatechin-3-gallate (EGCG) ameliorated radiation-induced increases in lipid peroxidation and protein carbonylation, as well as decreases in glutathione (GSH), glutathione peroxidase (GPx) and glutathione reductase (GR) and reverted the levels back to those similar to controls. DNA strand break parameters also returned to those similar to controls after treatment with EGCG (El-Missiry et al., 2018). Similar effects were also shown in another study using treatment mesenchymal stem cell-conditioned medium in mouse hippocampal cells exposed to 10 Gy of X-irradiation (Huang et al., 2021).  HUVECs pretreated with the antioxidant mixture RiduROS blunted ROS generation in a concentration-dependent manner by 65% ± 5.6% and 98% ± 2%, at 0.1 and 1 μg/mL, respectively, compared with cells irradiated without pretreatment. Low-dose irradiation also increased DSB-induced γ-H2AX foci compared with control cells and 24 h of RiduROS pretreatment reduced the γ-H2AX foci number by 41% (Cervelli et al., 2017). Additionally, HAECs treated with eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA) found significantly reduced intracellular ROS at 100µM, as well as reduced γ-H2AX foci formation by 47% and 48% following EPA and DHA treatment respectively. (Sakai et al., 2017).

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There is limited evidence demonstrating this relationship across different life stages/ages or sexes (Cencer et al., 2018; Li et al., 1998).   <table border="1"> <tbody> <tr> <td> Modulating Factors  </td> <td> MF Details  </td> <td> Effects on the KER  </td> <td> References  </td> </tr> <tr> <td> Age  </td> <td> Reduced antioxidant capacities have been linked to aged lenses (in humans >30 years old). The development of a chemical barrier between the cortex and the nucleus is partially responsible, as it prevents GSH from protecting aged lens cells from ROS.  </td> <td> Prevention of RONS-mediated damage is primarily achieved by antioxidants, so a lowered capacity would likely lead to reduced damage mitigation abilities. 78% of lens over 30 had a low level of GSH in the center compared to 14% of lens under 30. Lens epithelial cells have an associated 3-fold increase in γ-H2AX (marker of DNA damage) when GSH-PX decreases by 2-fold.  </td> <td> Taylor & Davies, 1987; Cabrera & Chihuailaf, 2011; Quinlan & Hogg, 2018; Sweeney & Truscott, 1998; Meng & Fang, 2021  </td> </tr> <tr> <td> Free radical scavengers  </td> <td> ROS-scavengers are essential components of the body’s natural defense against oxidative damage. Increased ROS production leads to increased incidence of electron donation by scavengers, thus reducing the overall level of free radical scavengers available to deal with ROS.  </td> <td> Isothiocyanates, such as sulforaphane (SFN), activate the release of more enzymatic scavengers. When SFN was added to in vitro LECs, LDH decreased to near unexposed cell levels and was associated with 3.3x less DNA strand breaks compared to the non-SFN cells following stressor exposure. Epigallocatechin-3-gallate (EGCG) also has antioxidant properties and was shown to alleviate radiation-induced increases in oxidative stress and DNA strand breaks within rat hippocampi.  </td> <td> Taylor et al., 1987; Cabrera et al., 2011; Liu et al., 2013; El-Missiry et al., 2018  </td> </tr> <tr> <td> Media  </td> <td> Mesenchymal stem cell-conditioned medium (MSC-CM), which has self-renewal, differential and proliferation capacities.  </td> <td> MSC-CM treatment has also been shown to improve ROS levels and decrease radiation-induced DNA strand breaks within mouse hippocampal neuronal cells.  </td> <td> Huang et al., 2021  </td> </tr> </tbody> </table>

The following tables provide representative examples of the relationship, unless otherwise indicated, all data is significantly significant.  Dose Concordance  <table border="1"> <tbody> <tr> <td> Reference   </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Cencer et al., 2018    </td> <td> In vitro, human LECs exposed to UVB and tested for 120 min post exposure with fluorescent probes to detect ROS production and mitochondrial superoxide, and tetramethylrhodamine-dUTP (TMR) red assay to detect strand breaks.  </td> <td> Both ROS and DNA strand breaks were increased by both 0.014 J/cm2 and 0.14 J/cm2 UVB radiation. At 0.014 J/cm2, cellular ROS increased a maximum of 15 fluorescence units above the control at 5 minutes post-UVB, while DNA strand breaks increased about 115 fluorescence units above the control at this time. At 0.14 J/cm2, cellular ROS increased a maximum of about 35 fluorescence units above the control at 90 minutes post-UVB, while mitochondrial superoxide increased about 30 fluorescence units above the control and DNA strand breaks increased about 125 fluorescence units above the control at this time.      </td> </tr> <tr> <td> Ahmadi et al., 2021   </td> <td> In vitro, human LECs exposed to 0.065-0.3 Gy/min gamma radiation, with dihydroethdium (DHE) fluorescent probes to measure ROS levels and comet assay to measure strand breaks.  </td> <td> Human LECs exposed in vitro to 0.1 - 0.5 Gy gamma rays showed a gradual increase in ROS levels and a corresponding gradual increase in DNA in the tail from the comet assay (indicative of increased DNA strand breaks) with the maximum dose displaying a 10% increase in ROS levels and a 17% increase in DNA strand damage.  </td> </tr> <tr> <td> Li et al., 1998   </td> <td> In vitro, bovine LECs were exposed to 40 and 400 µM H2O2 with an alkaline unwinding assay to determine strand break levels.  </td> <td> Immediately after LECs were exposed to 40 µM and 400 µM H2O2, there were ~145% and ~150% DNA strand breaks compared to the unexposed control level, respectively. The amounts of DNA strand breaks in cells exposed to both concentrations were reduced to ~105% of the unexposed control level after 30 mins. After 400 µM H2O2, oxidative stress as measured by LDH was 1200% of control in neuroblastoma cells.  </td> </tr> <tr> <td> Spector et al., 1997   </td> <td> In vitro, rat LECs exposed to 100 and 125 µM H2O2 with alkaline elution assay to determine single strand break level.  </td> <td> Exposure to 125 µM of H2O2 to lens epithelial cells resulted in reduction of intact DNA to near 1% by 9 hr post-exposure. Exposure to 100 µM H2O2 reduced SOD and GSH levels by 2-fold.  </td> </tr> <tr> <td> El-Missiry et al., 2018  </td> <td> In vivo, albino Wistar rats were exposed to 4 Gy of γ radiation (137Cs source) at 0.695 rad/s. Kits were used to measure 4-HNE (secondary product of lipid peroxidation) and protein  carbonyl group levels as markers of oxidative stress. Antioxidants including GSH, GPx  and GR were also assessed. The comet assay was used to analyze DNA strand breaks by visualizing DNA tail %, tail length and tail moment.  </td> <td> 4-HNE and protein carbonyl levels increased by approximately 2- and 3-fold after radiation exposure. GSH and GPx levels decreased by approximately 3-fold each, whereas GR levels decreased by approximately 5-fold. Tail DNA %, tail length and tail moment increased by approximately 2-, 3- and 6-fold after exposure to 4 Gy.  </td> </tr> <tr> <td> Ungvari et al., 2013  </td> <td> In vitro. CMVECs and rat hippocampal neurons were irradiated with 2-8 Gy 137Cs gamma rays. 5(and 6)-chloromethyl-2′,7′-dichlorodihydrofluorescein diacetate acetyl ester (CM-H2DCFDA) staining, and flow cytometry were used to measure ROS production. DNA damage was quantified by measuring the tail DNA content (as a percentage of total DNA) using the Comet Assay-IV software.  </td> <td> Day 1 post-irradiation showed increased cellular peroxide production and increased mitochondrial oxidative stress in CMVECs in a dose-dependent manner, increasing a maximum of ~3-fold at 8 Gy. Tail DNA content also increased in a dose-dependent manner with an approximate increase from 0 to 45% at 8 Gy.    </td> </tr> <tr> <td> Huang et al., 2021  </td> <td> In vitro, HT22 cells (mouse hippocampal neuronal cell line) were exposed to 10 Gy of X-irradiation at 6 Gy/min. ROS levels were measured using H2-DCFDA staining and fluorescence microscope analysis, whereas western blotting was used to detect γ-H2AX.  </td> <td> At 10 Gy, intracellular ROS generation increased by 5-fold and γ-H2AX increased by 3-fold.  </td> </tr> <tr> <td> Zhang et al., 2017  </td> <td> In vitro. HT22 cells were exposed to 8 and 12 Gy X-rays. Relative intracellular ROS levels were determined by DCFDA. p-ATM, γ-H2AX were measured with Western blot.  </td> <td> Following 8 Gy irradiation, intracellular ROS levels increased ~1.8-fold. Phosphorylation of ATM and γ-H2AX were increased 4.4-fold and 3.2-fold, respectively, 30 minutes after 12 Gy.    </td> </tr> <tr> <td> Cervelli et al., 2014  </td> <td> In vitro. HUVECs were irradiated with single doses (0.125, 0.25, 0.5 Gy), or fractionated doses (2 × 0.125 Gy, 2 × 0.250 Gy) of X-rays. Intracellular ROS generation was measured with a fluorescent dye, C-DCFDA, using a spectrofluorometer. Immunofluorescence microscopy was used to measure γ-H2AX foci.  </td> <td> Intracellular ROS production was significantly increased in a dose-dependent manner (1.6-, 2- and 2.8-fold at 0.125, 0.25, 0.5 Gy, respectively). When HUVECs were exposed to fractionated doses, no increase in ROS generation was observed, compared with respective single doses. 24h post-irradiation the percentage of foci-positive cells exposed to 0.125 Gy, 2 × 0.125 Gy, 0.250 Gy, 2 × 0.250 Gy and 0.5 Gy, was 1.68, 1.48, 3.53, 2.59, 8.74-fold over the control, respectively.  </td> </tr> <tr> <td> Sakai et al., 2017  </td> <td> In vitro. HAECs were exposed to 100uM H2O2. Intracellular ROS was measured by CM-H2DCFDA. DNA DSBs were detected by immunofluorescent analysis with γ-H2AX as a marker.  </td> <td> Intracellular ROS increased by ~3.7-fold  p-ATM increased by ~4.7-fold. γ-H2AX increased by ~3.4-fold.    </td> </tr> </tbody> </table>   Incidence Concordance  <table border="1"> <tbody> <tr> <td> Reference   </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Meng et al., 2021  </td> <td> In vitro, human LECs exposed to 50 µM H2O2 with DCFH-DA fluorescent probe to detect ROS levels and immunofluorescence and western blot assay to detect γ-H2AX.  </td> <td> 50 µM H2O2 exposure to lens epithelial cells increased oxidative stress, with ROS measured by LDH, by 4-fold and decreased the level of antioxidants by 2-fold as measured by SOD and GSH-PX. This resulted in 3-fold increase in γ-H2AX.  </td> </tr> <tr> <td> Smith et al., 2015  </td> <td> In vitro, human LECs exposed to 30 µM H2O2 with alkaline comet assay to determine amount of strand breaks.  </td> <td> Treatment of lens epithelial cells to 30 µM H2O2 induced DNA strand breaks by 55% at 0.5 hr after exposure and increased the level of LDH by ~1.4 fold at 24 hr post-exposure.  </td> </tr> <tr> <td> Liu et al. 2013  </td> <td> In vitro, human LECs exposed to 30 µM H2O2 with alkaline comet assay determination of strand breaks.  </td> <td> LDH increased by ~1.4 fold at 24 hr post-exposure, with a 5x increase from control levels in DNA strand breaks.  </td> </tr> </tbody> </table>   Time Concordance  <table border="1"> <tbody> <tr> <td> Reference   </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Yang et al., 1998   </td> <td> In vitro, rabbit LECs exposed to H2O2 with TCA addition and thiol assay to determine non-protein thiol (NP-SH) level and alkaline elusion assay to determine strand breaks.  </td> <td>  In rabbit LECs exposed in vitro to 125 µM H2O2, non-protein thiol levels decreased to <5% control (indicates oxidative stress) 30 min post-irradiation, and % DNA retained using alkaline elution decreased by 1.6 log (indicates increased DNA fragmentation) within the next 8.5 h.  </td> </tr> </tbody> </table>

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Low Unspecific Low Male

Low

Adult

Low

Not Otherwise Specified

Low Low Low Low Low

This KER is plausible in all life stages, sexes, and organisms with DNA. The evidence is from human, rodent, rabbit and bovine in vitro studies that do not specify the sex, as well as an adult rat in vivo study.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b42ea815be8> AOPWiki 2022-12-16T16:30:26

2023-03-22T09:48:24

<upstream-id>0e490634-53e7-457f-8aa8-4395c8e932ae</upstream-id> <downstream-id>1883f503-acf6-4e69-8704-91f462c360dc</downstream-id> DNA strand breaks can lead to altered signaling of various pathways through the DNA damage response. DNA strand breaks, which are a form of DNA damage, can induce ataxia telangiectasia mutated (ATM) and ATM/RAD3-related (ATR), two phosphoinositide 3-kinase (PI3K)-related serine/threonine kinases (PIKKs) (Abner and McKinnon, 2004; Lee and McKinnon, 2007; Nagane et al., 2021; Sylvester et al., 2018; Thadathil et al., 2019; Wang et al., 2020; Wang et al., 2017). Following DNA strand breaks, DNA damage response cellular signaling can phosphorylate downstream proteins and activate several transcription factors and pathways (Wang et al., 2017). Spontaneous DNA strand breaks from endogenous sources will induce signaling as a normal response to facilitate DNA repair. However, excessive DNA damage induced by a stressor will result in increased activation of these pathways and subsequent harmful downstream effects. Signaling pathways induced by DNA strand breaks include p53/p21 (Abner and McKinnon, 2004; Baselet et al., 2018; Lee and McKinnon, 2007; Nagane et al., 2021; Sylvester et al., 2018; Thadathil et al., 2019; Wang et al., 2020; Wang et al., 2017), caspase (Abner and McKinnon, 2004; Baselet et al., 2019; Wang et al., 2020; Wang et al., 2016) and mitogen-activated protein kinase (MAPK) family pathways (Ghahremani et al., 2002; Nagane et al., 2021).

The strategy for collating the evidence on radiation stressors to support the relationship is described in Kozbenko et al 2022. Briefly, a scoping review methodology was used to prioritize studies based on a population, exposure, outcome, endpoint statement.

Overall weight of evidence: Moderate

There is strong evidence supporting the link between DNA strand breaks leading to altered signaling pathways. Single strand breaks (SSBs) or double strand breaks (DSBs) in DNA from both endogenous and exogenous sources can induce the DNA damage response, which can result in the induction of various signaling pathways (Baselet et al., 2019). DNA strand breaks are well known to lead to the activation of ATM and ATR as part of the normal DNA damage response (Abner and McKinnon, 2004; Baselet et al., 2019; Lee and McKinnon, 2007; Nagane et al., 2021; Sylvester et al., 2018; Thadathil et al., 2019; Wang et al., 2020; Wang et al., 2017; Wang et al., 2016). While ATM tends to be recruited to DSBs, ATR is recruited by many types of DNA damage including both DSBs and SSBs (Maréchal and Zou, 2013; Wang et al., 2017). Following a DNA DSB, the Mre11-Rad50-Nbs1 (MRN) complex senses and directly binds to the DNA ends at the site of the break, which subsequently activates ATM (Lee and McKinnon, 2007; Maréchal and Zou, 2013). Following a DNA SSB, resection of the damaged strand by apurinic/apyrimidinic endonuclease (APE)1/APE2 is followed by coating the single-stranded DNA with replication protein A (RPA), where the recruitment of the ATR-ATR interacting protein (ATRIP) complex and the activation of ATR occurs (Caldecott, 2022; Maréchal and Zou, 2013).  ATM and ATR can phosphorylate over 700 proteins (Nagane et al., 2021), and phosphorylation of key signaling proteins by ATM/ATR will alter signaling in their respective pathways. High levels of DNA strand breaks induced by exogenous stressors will enhance ATM/ATR activation and subsequently further activate downstream signaling, leading to downstream consequences. The extracellular signal-regulated kinase (ERK), c-Jun N-terminal kinase (JNK) and p38 MAPK subfamily pathways can be phosphorylated and activated by ATM/ATR (Ghahremani et al., 2002; Nagane et al., 2021). Additionally, ATM/ATR can phosphorylate p53 on serine 15 to enhance the stability of p53, leading to activation of the p53 pathway and changes in the transcriptional activity of p53 (Abner and McKinnon, 2004; Baselet et al., 2019; Lee and McKinnon, 2007; Nagane et al., 2021; Sylvester et al., 2018; Thadathil et al., 2019; Wang et al., 2020; Wang et al., 2017; Wang et al., 2016). The apoptosis pathway downstream of p53 can also be activated by DNA strand breaks (Abner and McKinnon, 2004; Baselet et al., 2019; Lee and McKinnon, 2007; Thadathil et al., 2019; Wang et al., 2020).

Evidence for this relationship was collected from studies using in vivo mouse and rat models as well as in vitro mouse-derived, rat-derived and human-derived cell models. The stressors used to support this relationship include 137Cs gamma rays and X rays. Markers of DNA strand breaks in this KER include p53 binding protein 1 (53BP1), phosphorylation of H2AX (γ-H2AX), phosphorylation of ATR (p-ATR) and phosphorylation of ATM (p-ATM). Altered signaling was measured mostly by the protein expression of the p53/p21 and apoptosis pathways.    Dose Concordance  A few studies have indicated a dose concordance between the increase in DNA strand breaks and altered signaling pathways. X-ray irradiation of rat cortical neurons showed increased DNA damage markers, γ-H2AX, p-ATM and p-ATR and increased levels of signaling proteins, including p21, p-p53 and cleaved caspase 3 at both doses of 8 and 32 Gy (Sabirzhanov et al., 2020). 137Cs gamma irradiated cerebromicrovascular endothelial cells (CMVECs) and rat hippocampal neurons showed increased DNA strand breaks, measured by comet assay, at 2-10 Gy, and increased caspase 3/7 activity at 2, 4 and 6 Gy (Ungvari et al., 2013).    Time Concordance  Many studies demonstrate that DNA strand breaks occur before altered signaling in a time course. Although both KEs can occur quickly, Gionchiglia et al. (2021) showed in mice that γ-H2AX and p53BP1 foci were increased as early as 15 minutes after 10 Gy of X-ray irradiation while cleaved caspase 3 did not increase until 30 minutes after irradiation. In HT22 hippocampal neurons irradiated with 12 Gy of X-rays, γ-H2AX and p-ATM were increased at 30 minutes post-irradiation while p53 was increased after 1 h and caspase 3 was increased after 48 h (Zhang et al., 2017). Similarly, rat cortical neurons irradiated with 8 Gy of X-rays showed increased p-ATM, γ-H2AX and p-ATR after 30 minutes, while p-p53, p21 and cleaved caspase 3 did not increase until 3 or 6 h post-irradiation (Sabirzhanov et al., 2020). Multiple studies using human- and rat-derived endothelial cells irradiated with 4 Gy of 137Cs gamma rays show increased DNA strand breaks at 1 h post-irradiation, with altered signaling to p53 and p21 at 6 h and to caspase 3/7 at 18 h post-irradiation (Kim et al., 2014; Park et al., 2022; Ungvari et al., 2013). In a longer-term study irradiating human lung microvascular endothelial cells (HMVEC-L) with 15 Gy of X-rays, increased DNA strand breaks were observed at 14 days post-irradiation, while altered signaling in the p53 pathway was observed at 21 days post-irradiation (Lafargue et al., 2017).    Incidence concordance  A few studies have demonstrated an incidence concordance between DNA strand breaks and altered signaling at equivalent doses. Following X-ray irradiation of mice, DNA damage markers, γ-H2AX and p53BP1, increased by 10, 15 and 5-fold in different region of the brain, while cleaved caspase 3 signaling molecule increased by 1.4 and 2.6-fold (Gionchiglia et al., 2021). Gamma ray irradiation of Wistar rats showed a 6-fold increase in DNA damage marker compared to a 0.2-fold decrease in (B-cell lymphoma 2) Bcl-2 and a 2- to 4-fold increase in signaling proteins p53, Bcl-2-associated protein X (Bax) and caspase 3/8/9 (El-Missiry et al., 2018).    Essentiality  Some studies show that preventing an increase in DNA strand breaks will restore signaling. Treatment with mesenchymal stem cell-conditioned medium (MSC-CM) reduced γ-H2AX, decreased the levels of p53, Bax, cleaved caspase 3 and increased the levels of Bcl-2 in HT22 cells irradiated with 10 Gy of X-rays (Huang et al., 2021). The inhibition of microRNA (miR)-711 decreased levels of DNA damage markers, p-ATM, p-ATR and γ-H2AX, and decreased signaling molecules including p-p53, p21 and cleaved caspase 3 (Sabirzhanov et al., 2020).

None identified

<table border="1"> <tbody> <tr> <td> Modulating factor   </td> <td> Details   </td> <td> Effects on the KER   </td> <td> References   </td> </tr> <tr> <td> Media  </td> <td> MSC-CM  </td> <td> Treatment decreased γ-H2AX, p53, the Bax/Bcl-2 ratio and cleaved caspase 3 in irradiated neurons.  </td> <td> Huang et al., 2021  </td> </tr> <tr> <td> Genetic  </td> <td> miR-711  </td> <td> miR-711 inhibition reduced the DNA damage response including p-ATM, p-ATR, and γ-H2AX. It also decreased signaling molecules including p-p53, p21, and cleaved caspase 3.  </td> <td> Sabirzhanov et al., 2020    </td> </tr> <tr> <td> Drug  </td> <td> Minocycline  </td> <td> Treatment with minocycline in irradiated neurons reduced the DNA damage response through reduced γ-H2AX and p-ATM. Caspase 3 was also inhibited by minocycline, but p53 was not changed.  </td> <td> Zhang et al., 2017  </td> </tr> <tr> <td> Drug  </td> <td> Metformin  </td> <td> Treatment reduced p-ATM, p-p53 and p21 levels, but did not change the level of 53BP1 in irradiated HAECs.  </td> <td> Park et al., 2022  </td> </tr> </tbody> </table>

The tables below provide some representative examples of quantitative linkages between the two key events. All data that is represented is statistically significant unless otherwise indicated.

None identified

Moderate Male Low Female

Low

Juvenile

Moderate

Adult

Low Moderate Moderate

Evidence for this relationship is predominantly from studies using rat- and mouse-derived cells, with some in vivo evidence in mice and rats. There is in vivo evidence in male animals, but no in vivo studies specify the use of female animals. In vivo evidence is from adult models.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b42ea923418> AOPWiki 2023-03-10T15:07:20

2023-03-21T13:09:34

<upstream-id>b813d3a8-c9cc-49b4-bd70-a0bb3e9e8efd</upstream-id> <downstream-id>1883f503-acf6-4e69-8704-91f462c360dc</downstream-id> Oxidative stress occurs when the production of free radicals exceeds the capacity of cellular antioxidant defenses (Cabrera & Chihuailaf, 2011). Reactive oxygen species (ROS) and reactive nitrogen species (RNS) are both free radicals that can contribute to oxidative stress (Ping et al., 2020); however, ROS are more commonly studied than RNS (Nagane et al., 2021). ROS can mediate oxidative damage to biomacromolecules as they react with DNA, proteins and lipids, resulting in functional changes to these molecules (Ping et al., 2020). For example, ROS acting on lipids creates lipid peroxidation (Cabrera & Chihuailaf, 2011).    Many signaling pathways control and maintain physiological balance within a living organism, and these can be impacted by oxidative stress. Excessive reactive oxygen and nitrogen species (RONS) during oxidative stress can modify biological molecules and directly cause DNA damage, which can lead to altered signal transduction pathways (Hughson, Helm & Durante, 2018; Lehtinen & Bonni, 2006; Nagane et al., 2021; Ping et al., 2020; Ramadan et al., 2021; Schmidt-Ullrich et al., 2000; Soloviev & Kizub, 2019; Wang, Boerma & Zhou, 2016; Venkatesulu et al., 2018; Zhang et al., 2016). Different cell types can express distinct cellular pathways that can have varied response to an increase in oxidative stress. For example, oxidative stress in endothelial cells has been shown to inhibit the insulin-like growth factor 1 receptor (IGF-1R) and phosphatidylinositol-3-kinase/protein kinase B (PI3K/Akt) pathway and to activate the mitogen-activated protein kinase (MAPK) pathway, which can then have downstream detrimental effects (Ping et al., 2020). The MAPK family pathway is also activated in the central nervous system (CNS) in response to oxidative stress through calcium-induced phosphorylation of several kinases. These include phosphoinositide 3-kinase (PI3K), protein kinase A (PKA) and protein kinase C (PKC) and calcium/calmodulin-dependent protein kinase II (CaMKII) (Lehtinen & Bonni, 2006; Li et al., 2013; Ramalingam & Kim, 2012). Oxidative stress in bone cells can lead to increased expression of the receptor activator of nuclear factor kappa B ligand (RANKL) and Nrf2 activation (Tahimic & Globus, 2017; Tian et al., 2017). Following activation, Nrf2 then interferes with the activation of runt-related transcription factor 2 (Runx2), and depending on the level of oxidative stress, this may result in altered bone cell function (Kook et al., 2015).

The strategy for collating the evidence on radiation stressors to support the relationship is described in Kozbenko et al 2022. Briefly, a scoping review methodology was used to prioritize studies based on a population, exposure, outcome, endpoint statement.

Overall weight of evidence: High

Many reviews describe the role of oxidative stress in altered signaling. The mechanisms through which oxidative stress can contribute to changes in various signaling pathways are well-described. For example, oxidative stress can directly alter signaling pathways through protein oxidation (Ping et al., 2020; Schmidt-Ullrich et al., 2000; Valerie et al., 2007). Oxidation of cysteine and methionine residues, which are particularly sensitive to oxidation, can cause conformational change, protein expansion, and degradation, leading to changes in the protein levels of signaling pathways (Ping et al., 2020). Furthermore, oxidation of key residues in signaling proteins can alter their function, resulting in altered signaling. For example, oxidation of methionine 281 and 282 in the Ca2+/calmodulin binding domain of Ca2+/calmodulin-dependent protein kinase II (CaMKII) leads to constitutive activation of its kinase activity and subsequent downstream alterations in signaling pathways (Li et al., 2013; Ping et al., 2020). Similarly, during oxidative stress, tyrosine phosphatases can be inhibited by oxidation of a catalytic cysteine residue, resulting in increased phosphorylation of proteins in various signaling pathways (Schmidt-Ullrich et al., 2000; Valerie et al., 2007). Particularly relevant to this are the MAPK pathways. For example, the extracellular signal-regulated kinase (ERK) pathway is activated by upstream tyrosine kinases and relies on tyrosine phosphatases for deactivation (Lehtinen & Bonni, 2006; Valerie et al., 2007).    Furthermore, oxidative stress can indirectly influence signaling pathways through oxidative DNA damage which can lead to mutations or changes in the gene expression of proteins in signaling pathways (Ping et al., 2020; Schmidt-Ullrich et al., 2000; Valerie et al., 2007). DNA damage surveillance proteins like ataxia telangiectasia mutated (ATM) kinase and ATM/Rad3-related (ATR) protein kinase phosphorylate over 700 proteins, leading to changes in downstream signaling (Nagane et al., 2021; Schmidt-Ullrich et al., 2000; Valerie et al., 2007). For example, ATM, activated by oxidative DNA damage, phosphorylates many proteins in the ERK, p38, and Jun N-terminal kinase (JNK) MAPK pathways, leading to various downstream effects (Nagane et al., 2021; Schmidt-Ullrich et al., 2000).    The response of oxidative stress on signaling pathways has been studied extensively in various diseases. Herein presented are examples relevant to a few cell types related to vascular disease, impaired learning and memory, and bone loss. Many other pathways are plausible but available research has highlighted these to be critical to disease.    Endothelial cells: Endothelial cells can normally produce ROS. Antioxidant enzymes and the glutathione redox buffer control the redox state of vascular tissues. However, the dysregulation of signaling pathways can occur in the endothelium when oxidative stress is favored (Soloviev & Kizub, 2019). Oxidative stress can activate the acidic sphingomyelinase (ASMase)/ceramide pathway, the MAPK pathways, the p53/p21 pathway, and the signaling proteins p16 and p21, as well as inhibit the PI3K/Akt pathway (Hughson, Helm & Durante, 2018; Nagane et al., 2021; Ping et al., 2020; Ramadan et al., 2021; Soloviev & Kizub, 2019; Wang, Boerma & Zhou, 2016).    Bone cells: oxidative stress can induce signaling changes in the Wnt/β-catenin pathway, the RANK/RANKL pathway, the Nrf2/HO-1 pathway, and the MAPK pathways (Domazetovic et al., 2017; Manolagas & Almeida, 2007; Tian et al., 2017).     Brain cells: oxidative stress can induce alterations to various pathways such as the PI3K/Akt pathway, cAMP response element-binding protein (CREB) pathway, the p53/p21 pathway, as well as the MAPK family pathways, including JNK, ERK and p38 (Lehtinen & Bonni, 2006; Ramalingam & Kim, 2012).    Additionally, the electron transport chain in the mitochondria is an important source of ROS, which can damage mitochondria by inducing mutations in mitochondrial DNA. These mutations lead to mitochondrial dysfunction due to alterations in cellular respiration mechanisms that perpetuates oxidative stress and can then induce the release of signaling molecules related to apoptosis from the mitochondria. Pro-apoptotic markers (Bax, Bak and Bad) and anti-apoptotic markers (Bcl-2 and Bcl-xL) can regulate the caspase pathway that ultimately mediate apoptosis (Annunziato et al., 2003; Wang & Michaelis, 2010; Wu et al., 2019).    The mechanisms of oxidative stress leading to altered signaling may be different for each pathway. For example, although both the PI3K/Akt and MAPK pathways can be regulated by insulin-like growth factor (IGF)-1, ROS results in selective inhibition of the IGF-1R/PI3K/Akt pathway by inhibiting the IGF-1 receptor (IGF-1R) activation of IRS1 (Ping et al., 2020). Additionally, ROS-induced MAPK activation can be done through Ras-dependent signaling. Firstly, oxygen radicals mediate the phosphorylation of upstream epidermal growth factor receptors (EGFRs) on tyrosine residues, resulting in increased binding of growth factor receptor-bound protein 2 (Grb2) and subsequent activation of Ras signaling (Lehtinen & Bonni, 2006). Direct inhibition of MAPK phosphatases with hydroxyl radicals also activates this pathway (Li et al., 2013). In another mechanism, ROS competitively inhibit the Wnt/β-catenin pathway through the activation of forkhead box O (FoxO), which are involved in the antioxidant response and require binding of β-catenin for transcriptional activity (Tian et al., 2017).

Evidence for this relationship was collected from studies using in vivo mouse, rat, and pig models, as well as in vitro mouse-derived, rat-derived, bovine-derived, and human-derived models. The stressors used to support this relationship include gamma rays, X rays, microgravity, hydrogen peroxide, chronic cold stress, heavy ion radiation, simulated ischemic stroke and growth differentiation factor (GDF) 15 overexpression. These stressors were shown to increase levels of oxidative stress and induce changes within relevant signaling pathways (Azimzadeh et al., 2021; Azimzadeh et al., 2015; Fan et al., 2017; Xu et al., 2019; Suman et al., 2013; Limoli et al., 2004; Tian et al., 2020; Hladik et al., 2020; Diao et al., 2018; Hasan, Radwan & Galal, 2019; Xin et al., 2015; El-Missiry et al., 2018; Kenchegowda et al., 2018; Kook et al., 2015; Sun et al., 2013; Yoo, Han & Kim, 2016, Zhao et al., 2013; Bai et al., 2020; Chen et al., 2009; Carvour et al., 2008; Wortel et al., 2019; Azimzadeh et al., 2017; Park et al., 2016; Sakata et al., 2015; Ruffels et al., 2004; Crossthwaite et al., 2002).  Incidence concordance  A few studies demonstrate greater changes to oxidative stress than to altered signaling. Human umbilical vein endothelial cells (HUVECs) irradiated with 10 Gy of X-rays showed a 20-fold increase in ROS and a 0.5-fold decrease in the ratio of p-Akt/Akt (Sakata et al., 2015). Microgravity exposure to preosteoblast cells showed a 0.24-fold decrease to the antioxidant Cu/Zn-superoxide dismutase (SOD) and a 0.36-fold decrease to p-Akt (Yoo, Han & Kim, 2016). It was also shown in rats that MDA levels increased by 1.5-fold while angiotensin and aldosterone increased by 1.4-fold after 6 Gy of gamma rays (Hasan, Radwan & Galal, 2020). Bai et al. (2020) demonstrated with multiple endpoints that ROS levels increased, and antioxidant enzyme levels decreased more than signaling pathways were altered.    Dose Concordance  Many studies demonstrate dose concordance for this relationship, at the same doses. Low-dose (0.5 Gy) X-ray irradiation of human coronary artery endothelial cells (HCAECs) show increased protein carbonylation with decreased glutathione S-transferase omega-1 (GSTO1) antioxidant levels and a simultaneous alteration of signaling proteins Rho GDP-dissociation inhibitor (RhoGDI), p16, and p21 (Azimzadeh et al., 2017). A dose of about 2 Gy of gamma rays showed decreased antioxidants as well as decreased protein levels and activation of the PI3K/Akt pathway in pig cardiac tissue (Kenchegowda et al., 2018). Similarly, gamma irradiation at 6 Gy resulted in reduced levels of the antioxidant glutathione (GSH) and increased levels of the lipid peroxidation marker MDA as well as an increase in the renin angiotensin aldosterone system (RAAS) measured in rat heart tissue and blood serum, respectively (Hasan, Radwan & Galal, 2020). HUVECs irradiated with 10 Gy of X-rays demonstrated increased ROS while p-Akt decreased and p-ERK1/2 increased (Sakata et al., 2015). Gamma radiation at 15 Gy led to both increased ROS as well as attenuated p38 MAPK and Nrf2 signaling pathways in murine cardiac tissue (Fan et al., 2017). In contrast, 16 Gy X-ray exposure led to decreased levels of the antioxidant SOD, increased MDA as well as increased MAPK signaling in murine heart tissue (Azimzadeh et al., 2021). After simulated microgravity, changes to signaling pathways, increased ROS and MDA, and decreased antioxidants were found both in in vitro mouse-derived bone cells and in in vivo rat femurs. Increased ROS levels and decreased antioxidants were found with changes in the RANK/RANKL pathway, Wnt/β-catenin pathway, Runx2, PI3K/Akt pathway, and MAPK pathways (Diao et al., 2018; Sun et al., 2013; Xin et al., 2015; Yoo, Han & Kim, 2016).    A few studies also find that oxidative stress often occurs at lower doses than altered signaling pathways. Bai et al. (2020) measured oxidative stress, shown by increased ROS and decreased antioxidant expression, at 2, 5, and 10 Gy of gamma rays. They also found Runx2 increased at the same doses, but the p53/p21 pathway was only significantly altered at 5 and 10 Gy (Bai et al., 2020). At similar doses, X-ray irradiated mouse osteoblast-like cell line MC3T3-E1 cells showed increased ROS and decreased antioxidants both 4 and 8 Gy (Kook et al., 2015). While HO-1 also increased at both 4 and 8 Gy, Nrf2 and Runx2 were measured altered at 8 Gy (Kook et al., 2015). In another study, X-ray irradiation at 16 Gy resulted in decreased SOD and increased MDA and protein carbonylation, which were associated with decreased PI3K/Akt pathway activity and protein levels, decreased ERK activity and protein levels, increased p38 activity, and increased p16 and p21 protein levels in heart tissue (Azimzadeh et al., 2015). Azimzadeh et al. (2015) also showed that at 8 Gy oxidative stress was still observed, but fewer signaling molecule levels and activity were altered at this. Particularly, no changes to MAPK pathways were observed.    Within the rat hippocampus, El-Missiry et al. (2018) demonstrated that exposure to 4 Gy of X-irradiation results in increased 4-HNE (oxidative stress marker) levels, reduced antioxidant activity and an increase in p53 expression. In the cerebral cortex of mice, Suman et al. (2013) reported that 1.6 Gy of 56Fe and 2 Gy of gamma rays increased ROS levels, consequently increased p21 and p53 levels. Limoli et al. (2004) also reported increased ROS levels in mice and rat neural precursor cells after exposure to X-irradiation (1-10 Gy), accompanied by increased expression of p21 and p53. Hladik et al. (2020) exposed female mice to 0.063, 0.125 or 0.5 Gy of gamma-radiation, which resulted in increases of protein carbonylation, as well as increased phosphorylation of CREB, ERK1/2 and p38. Radiation-induced changes in apoptotic markers were also reported. More specifically, there was a significant rise in pro-apoptotic markers Bax and caspase 3, with significant reduction in anti-apoptotic marker Bcl-xL (Hladik et al., 2020). Furthermore, middle cerebral artery occlusion (MCAO) surgery known to simulate ischemic stroke in C57BL/6J mice was shown to increase ROS levels, as well as the phosphorylation of ERK1/2, p38 and JNK (Tian et al., 2020).    Other studies that have used hydrogen peroxide (H2O2) to induce oxidative stress within cell cultures, have also observed alterations in signaling pathways. Zhao et al. (2013) exposed mouse hippocampal-derived HT22 cells to varying concentrations of H2O2 and found a dose-dependent increase in ROS production from 250-1000 μM. Additionally, treating the cells to H2O2 resulted in a concentration-dependent increase of ERK1/2, JNK1/2 and p38 phosphorylation. Ruffels et al. (2004) incubated human neuroblastoma cells (SH-SY5Y) to varying concentrations of H2O2 that ranged from 0.5-1.25 mM and found a dose-dependent increase in JNK1/2, ERK1/2 and Akt phosphorylation. Another study exposed SH-SY5Y and rat pheochromocytoma (PC12) cells to 0.05-2 mM H2O2 and found a dose-dependent increase in ROS from 0-1 mM in SH-SY5Y cells, and from 0-2 mM in PC12 cells with a concentration-dependent increase in ERK1/2, p38 and JNK phosphorylation (Chen et al., 2009). Furthermore, Crossthwaite et al. (2002) incubated neuronal cultures from 15- to 16-day-old Swiss mice to 100, 300 and 1000 μm H2O2 and showed increased levels of ROS. A corresponding increase in ERK1/2 and Akt activation was observed at 100-300 μm, and for JNK1/2 the observation was observed at 1000 μm. Carvour et al. (2008) treated N27 cells (rat dopaminergic cell line) to 3-30 μM H2O2 and measured increased ROS levels, as well as increased apoptotic signaling molecules caspase 3 and proapoptotic kinase protein kinase C-δ (PKCδ) cleavage.     Time Concordance  Limited evidence shows that oxidative stress leads to altered signaling pathways in a time concordant manner. When irradiated with X-rays, HCAECs, BAECs and MCT3T3-E1 osteoblast-like cells show increase in ROS or levels of protein carbonylation, or a decrease in the levels of superoxide dismutase (SOD), catalase (CAT), GSTO1 or GSH at earlier timepoints than alterations in the signaling molecules p16, p21, Ceramide, Runx2, and HO-1 (Azimzadeh et al., 2017; Kook et al., 2015; Wortel et al., 2019). As the key events are both molecular-level changes, both can occur quickly after irradiation. Wortel et al. (2019) found that increased hydrogen peroxide levels could be observed in vitro as early as 2 minutes post-irradiation, while ASMase activity and ceramide levels were only increased 5 minutes post-irradiation.    When exposed to H2O2, PC12 cells show an increase production of ROS with a corresponding increase in phosphorylation of MAPK proteins in a time-dependent fashion. An increase in ERK1/2, JNK and p38 phosphorylation was observed within 5-15 minutes and sustained for over 2 hours (Chen et al., 2009). When exposed to cold stress for 1, 2 and 3 weeks, MDA levels increased in a time-dependent manner from 1-3 weeks within the brain tissue isolated from C57BL/6 mice. The expressions of JNK, ERK and p38 phosphorylation levels were all also significantly upregulated in chronic cold-stressed groups for all time-points (Xu et al., 2019). After gamma irradiation (2 Gy), ROS increased 2 months post-irradiation, while increased p21 and decreased Bcl-2 were only observed at 12 months (Suman et al., 2013). However, other signaling molecules were increased at both times.    Essentiality  Several studies have investigated the essentiality of the relationship, where the blocking or attenuation of the upstream KE causes a change in frequency of the downstream KE. The increase in oxidative stress can be modulated by certain drugs, antioxidants and media. L-carnitine injections decreased ROS and increased p-p38/p38 and p-Nrf2/Nrf2 signaling (Fan et al., 2017). Fenofibrate was found to return SOD, phosphorylated MAPK signaling proteins and increase Nrf2 levels (Azimzadeh et al., 2021). Antioxidants (N-acetyl cysteine, curcumin) were shown to restore or reduce ROS levels closer to control levels following radiation or microgravity exposure, respectively. Signaling proteins in the Nrf2/HO-1 pathway and the RANKL/osteoprotegerin (OPG) ratio were decreased and brought closer to control levels (Kook et al., 2015; Xin et al., 2015). Hydrogen rich medium showed reduced ROS with restoration of OPG and RANKL signaling levels to controls (Sun et al., 2013). Polyphenol S3 (60 mg/kg/d) treatment was found to reverse the effect of microgravity on CAT, SOD and MDA, returning the levels to near control values. Meanwhile, Runx2 mRNA levels and β-catenin/β-actin levels increased following treatment (Diao et al., 2018). Sildenafil is another drug that was found to reduce ROS generation by inhibiting O2- production and intracellular peroxynitrite levels in bovine aortic endothelial cells (BAECs) after gamma irradiation. As well, ASMase activity and ceramide levels were inhibited by sildenafil (Wortel et al., 2019).    Within brain cells, several antioxidants have been found to attenuate oxidative stress-induced alterations in signaling pathways. These antioxidants include Melandrii Herba extract, N-acetyl-L-cysteine (NAC), gallocatechin gallate/epigallocatechin-3-gallate, Cornus officinalis (CC) and fermented CC (FCC), L-165041, fucoxanthin, and edaravone. These antioxidants were shown to reduce ROS and subsequently decrease phosphorylation of MAPKs such as ERK1/2, JNK1/2 and p38 after exposure to radiation, H2O2 or LPS (Lee et al., 2017; Deng et al., 2012; Park et al., 2021; Tian et al., 2020; Schnegg et al., 2012; Zhao et al., 2017; Zhao et al., 2013; El-Missiry et al., 2018). Another documented modulator is mesenchymal stem-cell conditioned medium (MSC-CM), which was able to alleviate oxidative stress in HT22 cells and restore levels of p53 (Huang et al., 2021).

<ul> <li>MAPK pathways can exhibit varied responses after exposure to oxidative stress. The expected response is an increase in the activity of the ERK, JNK, and p38 pathways as protein phosphatases, involved in the inactivation of MAPK pathways, are deactivated by oxidative stress (Valerie et al., 2007). Although some studies observe this (Azimzadeh et al., 2021; Sakata et al., 2015), others show a decrease (Fan et al., 2017; Yoo, Han & Kim, 2016) or varying changes (Azimzadeh et al., 2015) in the MAPK pathways.</li> </ul>

<table border="1"> <tbody> <tr> <td> Modulating factor   </td> <td> Details   </td> <td> Effects on the KER   </td> <td> References   </td> </tr> <tr> <td> Drug  </td> <td> Fenofibrate (PPARα activator, PPARα is a transcription factor that can activate antioxidant response)  </td> <td> Treatment of mice with 100 mg/kg of body weight daily for 2 weeks before and 2 weeks after radiation restored SOD activity, returned the level of phosphorylated MAPK proteins and increased Nrf2 levels.  </td> <td> Azimzadeh et al., 2021  </td> </tr> <tr> <td> Drug  </td> <td> L-carnitine (antioxidant)  </td> <td> L-carnitine injections (100 mg/kg) following irradiation resulted in decreased DHE staining, indicating ROS, and increased p-p38/p38 and p-Nrf2/Nrf2.  </td> <td> Fan et al., 2017  </td> </tr> <tr> <td> Drug  </td> <td> N-acetyl cysteine (antioxidant)  </td> <td> Treatment of osteoblast-like cells with 5 mM restored ROS levels, SOD activity, and the level of proteins in the Nrf2/HO-1 pathway.  </td> <td> Kook et al., 2015  </td> </tr> <tr> <td> Drug  </td> <td> Curcumin (antioxidant)  </td> <td> Treatment of osteoblast-like cells with 4 µM reduced ROS levels and the RANKL/OPG ratio. Treatment of rats with 40 mg/kg of body weight reduced oxidative stress and the RANKL/OPG ratio.  </td> <td> Xin et al., 2015  </td> </tr> <tr> <td> Drug  </td> <td> Bradykinin potentiating factor (BFP)   (antioxidant)  </td> <td> Treatment with BFP (1ug/g) after irradiation showed decreased AngII and aldosterone levels compared to irradiation alone.   </td> <td> Hasan, Radwan & Galal, 2020  </td> </tr> <tr> <td> Media  </td> <td> Hydrogen-rich   (antioxidant)  </td> <td> Osteoblasts in a medium consisting of 75% H2, 20% O2, and 5% CO2 (vol/vol/vol) showed a reduction in ROS production and restoration of normal signaling.  </td> <td> Sun et al., 2013  </td> </tr> <tr> <td> Drug  </td> <td> Melatonin  (antioxidant)  </td> <td> Treatment with 200 nM melatonin reversed the effect of microgravity on Cu/Zn-SOD and Mn-SOD to control levels.   </td> <td> Yoo, Han & Kim, 2016  </td> </tr> <tr> <td> Drug  </td> <td> Polyphenol S3   </td> <td> Polyphenol S3 treatment reverses the effect of microgravity on CAT, SOD and MDA, returning the levels to near control values when S3 is used at high dose (60mg/kg/d). Runx2 mRNA levels and β-catenin/β-actin levels increased following treatment and simulated microgravity.  </td> <td> Diao et al., 2018  </td> </tr> <tr> <td> Drug  </td> <td> Sildenafil  </td> <td> Sildenafil (5 uM) inhibits O2- production and attenuates intracellular peroxynitrite in BAECs after 10 Gy irradiation. As well, ASMase activity and ceramide generation was inhibited.  </td> <td> Wortel et al., 2019  </td> </tr> <tr> <td> Drug  </td> <td> DPI   (NOX-inhibitor)  </td> <td> Inhibits O2-  production and intracellular H2O2 in BAECs after 10 Gy irradiation.  </td> <td> Wortel et al., 2019  </td> </tr> <tr> <td> Drug  </td> <td> Edaravone (EDA) which acts as a free radical scavenger  </td> <td> EDA treatment was able to reduce the levels of ROS and consequently decrease the expression levels of phosphorylated JNK, p38 and ERK1/2.  </td> <td> Zhao et al., 2013  </td> </tr> <tr> <td> Drug  </td> <td> Melandrii Herba extract (antioxidant)  </td> <td> The extract was able to reduce the H2O2-induced phosphorylation of ERK1/2, JNK1/2 and p38 in human neuroblastoma SH-SY5Y cells.  </td> <td> Lee et al., 2017  </td> </tr> <tr> <td> Drug  </td> <td> N-acetyl-L-cysteine (NAC) (antioxidant)  </td> <td> Attenuated the effects of H2O2 in BV-2 murine microglial cells as treatment with NAC reduced c-Jun and ERK1/2 phosphorylation.  </td> <td> Deng et al., 2012  </td> </tr> <tr> <td> Drug  </td> <td> Gallocatechin gallate (GCG) or epigallocatechin-3-gallate (EGCG), both of which have antioxidant properties  </td> <td> GCG and EGCG inhibits ROS accumulation in mouse hippocampal-derived HT22 cells and Wistar rats, respectively. This consequently reduced glutamate-induced phosphorylation of MAPKs (ERK and JNK) and returned p53 to control levels.  </td> <td> Park et al., 2021; El-Missiry et al., 2018  </td> </tr> <tr> <td> Drug   </td> <td> Cornus officinalis (CC) and fermented CC (FCC), both of which have antioxidant properties  </td> <td> Both CC and FCC were able to reduce intracellular ROS generation in H2O2-induced neurotoxicity in SH-SY5Y human neuroblastoma cells. This was accompanied with a decrease in ERK1/2, JNK and p38 phosphorylation.  </td> <td> Tian et al., 2020  </td> </tr> <tr> <td> Drug  </td> <td> L-165041, a PPARδ agonist (PPARα is a transcription factor that can activate antioxidant response).  </td> <td> 10 Gy of 137Cs irradiation resulted in an increase in intracellular ROS and c-Jun, MEK1/2 and ERK1/2 phosphorylation in BV-2 cells, all of which were attenuated with L-165041 treatment.  </td> <td> Schnegg et al., 2012  </td> </tr> <tr> <td> Drug  </td> <td> Fucoxanthin (antioxidant)  </td> <td> Fucoxanthin was able to inhibit the LPS-induced increase in intracellular ROS and phosphorylation of JNK, ERK and p38.  </td> <td> Zhao et al., 2017  </td> </tr> <tr> <td> Media  </td> <td> Mesenchymal stem-cell conditioned medium (MSC-CM)  </td> <td> MSC-CM was able to inhibit the X-ray-induced increase in ROS and MDA levels and decrease in SOD and GSH levels, resulting in activation of PI3/Akt.  </td> <td> Huang et al., 2021  </td> </tr> </tbody> </table>

The tables below provide representative examples of quantitative linkages between the two key events. It was difficult to identify a general trend across all the studies due to differences in experimental design and reporting of the data. All data that is represented is statistically significant unless otherwise indicated.

Dose/Incidence Concordance  <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Azimzadeh et al., 2017  </td> <td> In vitro. HCAECs were irradiated with 0.5 Gy of X-rays (0.5 Gy/min). Protein carbonylation and GSTO1 antioxidant levels were measured with a carbonylation assay and immunoblotting, respectively. Proteins from various signaling pathways including RhoGDI, p16, and p21 were measured with immunoblotting.  </td> <td> After 0.5 Gy, carbonyl content increased a maximum of 1.2-fold and GSTO1 decreased a maximum of 0.78-fold. After 0.5 Gy, p-RhoGDI decreased a maximum of 0.7-fold, p16 increased a maximum of 1.5-fold, and p21 increased a maximum of 1.2-fold.  </td> </tr> <tr> <td> Kenchegowda et al., 2018  </td> <td> In vivo. Male 3- to 5-month-old Gottingen minipigs and Sinclair minipigs were whole-body irradiated with 1.7-2.3 Gy of 60Co gamma rays (0.6 Gy/min). Both survivors (n=23) and euthanized moribund animals (n=17) had measurements taken for oxidative stress and altered signaling taken from the heart. SOD, CAT, and p67 (subunit of NADPH oxidase/NOX, involved in producing superoxide) levels were determined with western blot. ELISA and western blot were used to measure altered signaling in the IGF/PI3K/Akt pathway.  </td> <td> Compared to survivors, radiation induced a 2.1-fold increase in p67, 0.87-fold decrease in SOD, and a 0.83-fold decrease in CAT (non-significant, ns) in the deceased group. Compared to survivors, the ratio of activated (phosphorylated) to total IGF-1R and the ratio of activated (phosphorylated) to total Akt both decreased 0.5-fold in the deceased group.  </td> </tr> <tr> <td> Kook et al., 2015  </td> <td> In vitro. MC3T3-E1 osteoblast-like cells were irradiated with 2, 4, and 8 Gy of X-rays (1.5 Gy/min). ROS were measured with a fluorescent probe, and SOD, CAT, and GSH antioxidant activities were determined with assay kits. Protein levels in the Nrf2/HO-1 signaling pathway were determined by either western blot or RT-PCR.  </td> <td> ROS increased linearly at 2 and 4 Gy up to 1.4-fold at 8 Gy (significant changes at 4 Gy and 8 Gy). GSH and SOD were decreased 0.7-fold at 4 Gy and 0.5-fold at 8 Gy (insignificant increases at 2 Gy). CAT was also decreased but not significantly. HO-1 increased 3.3-fold after 4 Gy and 4.9-fold after 8 Gy (insignificant increase at 2 Gy). Nrf2 increased 2.3-fold after 8 Gy. Runx2 mRNA was decreased 0.5-fold after 8 Gy.  </td> </tr> <tr> <td> Bai et al., 2020  </td> <td> In vitro. Rat-derived bone marrow-derived mesenchymal stem cells (bmMSCs) were irradiated with 2, 5, and 10 Gy of 137Cs gamma rays. Mitochondrial and cellular ROS levels were determined with fluorescent probes. RT-qPCR was performed to measure antioxidant enzyme expression. Protein expression in various signaling pathways was measured by western blot.  </td> <td> Mitochondrial ROS increased 1.6-fold at 2 Gy (non-significant), 2-fold at 5 Gy, and 2.3-fold at 10 Gy. Cellular ROS increased 1.2-fold at 2 Gy, 1.5-fold at 5 Gy, and 2.1-fold at 10 Gy. Antioxidants SOD1, SOD2, and CAT all decreased about 0.9-fold (ns for SOD2) after 2 Gy, 0.8- to 0.7-fold at 5 Gy, and 0.7- to 0.4-fold at 10 Gy. Runx2 decreased 0.9-fold at 2 and 5 Gy and 0.6-fold at 10 Gy. p21 increased 1.6-fold at 5 Gy and 2.5-fold at 10 Gy. p53 increased 1.6-fold at 5 Gy and 1.7-fold at 10 Gy. p16 remained unchanged.  </td> </tr> <tr> <td> Fan et al., 2017    </td> <td> In vivo. 10-week-old male C57BL/6J mice were irradiated with 60Co gamma rays at 3 Gy/day for 5 days. Left ventricular cardiac tissue was harvested for analysis. ROS was detected by dihydroethidium (DHE) staining. MAPK and Nrf2 signaling molecules were measured by western blot.  </td> <td> Following irradiation, ROS production increased by 3.6-fold.  p-p38/p38 decreased by 0.36-fold and p-Nrf2/Nrf2 decreased by 0.14-fold.  </td> </tr> <tr> <td> Hasan, Radwan & Galal, 2020  </td> <td> In vivo. 6-week-old male Wistar rats were irradiated with 6 Gy 137Cs gamma rays. Oxidative stress was measured by MDA and GSH in heart tissue. Angiotensin II (AngII) and aldosterone, key molecules in the RAAS pathway, were measured with ELISA kits in serum.  </td> <td> Following irradiation, MDA levels increased by 1.5-fold and GSH levels decreased by 0.5-fold. AngII and aldosterone increased 1.4-fold compared to control.       </td> </tr> <tr> <td> Azimzadeh et al., 2015  </td> <td> In vivo. Male 10-week-old C57BL/6 mice were irradiated with 8 and 16 Gy of X-rays. SOD, MDA, and protein carbonylation levels were determined with immunoblotting, lipid peroxidation, and protein carbonylation assays, respectively, in heart tissue. Proteins in various signaling pathways were measured with immunoblotting in heart tissue.  </td> <td> SOD decreased 0.7-fold at both 8 and 16 Gy and MDA increased 1.4-fold after 8 Gy and 2.1-fold after 16 Gy. Protein carbonylation increased 1.4-fold after 16 Gy. Levels and activity of proteins in the PI3K/Akt pathway were decreased between 0.5- and 0.1-fold at both 8 and 16 Gy. The ERK/MAPK pathway was found decreased 0.5-fold at 16 Gy and the p38/MAPK pathway was found increased 1.3-fold at 16 Gy. p16 was increased 1.6-fold at both 8 and 16 Gy. p21 was increased 2.4-fold at both 8 and 16 Gy.  </td> </tr> <tr> <td> Sakata et al., 2015  </td> <td> In vitro. HUVECs were irradiated with 10 Gy X-rays at a dose rate of 5 Gy/min. Measurements were performed 0-72 h post-irradiation. ROS were detected by fluorescence microscopy. MAPK, Akt, p-p38, JNK and ERK1/2 signaling molecules were measured by western blot.  </td> <td> Following 10 Gy irradiation, the intensity representing ROS generation increased 20- and 30-fold at the 24 and 72 h timepoints, respectively.    MAPK, p38 and JNK remained unchanged for the 72 h measured following 10 Gy irradiation.     p-Akt/Akt in HUVECs after 10 Gy irradiation showed an initial decrease at 5 min and a delayed decrease of 0.5-fold at 6-24 h. p-ERK1/2 decreased at 5 min then increased to a maximum 1.75-fold change.  </td> </tr> <tr> <td> Wortel et al., 2019  </td> <td> In vitro. BAECs were irradiated with 10 Gy 137Cs gamma rays at a rate of 1.66 Gy/min. Extracellular H2O2 was measured by Amplex Red Assay, intracellular H2O2 levels were determined by HyPer sensor and peroxynitrite was quantified by chemiluminescence assay. Superoxide levels were quantified by luminescence after treatment with Diogenes Complete Enhancer Solution. The activation of the ASMase enzyme and the levels of ceramide were quantified by radioenzymatic assay to deter mine the changes on the ASMase/ceramide pathway.  </td> <td> Following 10 Gy irradiation, intracellular H2O2 increased to a maximum 1.35-fold. Extracellular H2O2 increased by 1.75-fold. Peroxynitrite increased by 2.86-fold after 10 Gy (Fig 5). Superoxide levels increased over 350% at 2 minutes after 10 Gy irradiation. ASMase activity increased to a maximum 5.6-fold at 5 min after irradiation, then decreased and remained unchanged until the 30 min time-point. Ceramide increased from -500 to over 3000 pmol/106 cells. The significance of these changes was not indicated against a control.  </td> </tr> <tr> <td> Azimzadeh et al., 2021  </td> <td> In vivo. Male C57BL/6J mice 8 weeks of age were irradiated with 16 Gy of X-rays to the heart. SOD antioxidant activity and MDA in heart tissue were determined with an assay kit and lipid peroxidation assay, respectively. The level of proteins in MAPK pathways were determined by ELISA in heart tissue.  </td> <td> After 16 Gy, SOD decreased 0.8-fold and MDA increased 1.3-fold. After 16 Gy, p-ERK increased 1.5-fold, p-p38 increased 1.3-fold, and p-JNK increased 1.3-fold.  </td> </tr> <tr> <td> Xin et al., 2015  </td> <td> In vitro and in vivo.  In vitro. MC3T3-E1 osteoblast-like cells were exposed to microgravity for 96 hours. ROS were determined with a fluorescent probe and the RANK/RANKL pathway was measured using RANKL and OPG assay kits.    In vivo. Male 8-week-old Sprague-Dawley rats were exposed to hind-limb suspension for 6 weeks. Femur and plasma MDA and femur sulfhydryl levels were measured with assay kits and the RANK/RANKL pathway was measured in the femur using RANKL and OPG assay kits.  </td> <td> In vitro. ROS increased 1.5-fold and the RANKL/OPG ratio increased 1.6-fold.    In vivo. Serum and femur MDA increased 1.4-fold and femur sulfhydryl decreased 0.6-fold. The RANKL/OPG ratio increased 3.5-fold.  </td> </tr> <tr> <td> Sun et al., 2013  </td> <td> In vitro. MC3T3-E1 osteoblast-like cells were exposed to microgravity (0.01G) for 96 hours. ROS production was measured by a fluorescent probe. The RANKL/OPG ratio was determined by assay kit and Runx2 mRNA expression was determined by RT-qPCR  </td> <td> ROS increased 1.5-fold. The RANKL/OPG ratio increased 1.6-fold. Runx2 expression decreased 0.4-fold.  </td> </tr> <tr> <td> Yoo, Han & Kim, 2016  </td> <td> In vitro. Preosteoblast MC3T3-E1 cells were exposed to microgravity conditions by a 3D clinostat at a speed from 1-10 rpm. Oxidative stress was measured by Cu/Zn-SOD, Mn-SOD and catalase activity. Signaling molecules, p-Akt, phosphorylation of the mechanistic target of rapamycin p-(mTOR), and p-ERK were measured by western blot.  </td> <td> Following microgravity exposure, Cu/Zn-SOD and Mn-SOD levels decreased by 0.24 and 0.65-fold, respectively. Signaling molecules p-Akt decreased by 0.36-fold. p-mTOR and p-ERK decreased by 0.58-fold.    </td> </tr> <tr> <td> Diao et al., 2018      </td> <td> In vivo. The left femur of rats was studied after exposure to simulated microgravity. Oxidative stress was measured by MDA, SOD, and CAT levels. RANK/RANKL signaling pathway was measured in rat femur by enzyme-linked immunoassay detection of OPG/RANKL molecules. Signaling molecule, Runx2, mRNA levels were measured by quantitative real time PCR. The Wnt/β-catenin pathway was measured by western blot for β-catenin protein levels.  </td> <td> MDA increased by 1.4-fold. SOD and CAT levels decreased by 0.4-fold. OPG/RANKL decreased by 0.6-fold. Runx2 mRNA levels decreased 0.04-fold (Fig 8d). β-catenin decreased 0.6-fold.  </td> </tr> <tr> <td> El-Missiry et al., 2018  </td> <td> In vivo. Male Wistar rats were irradiated with gamma rays (137Cs source, 4 Gy, 0.695 cGy/s) and measurements were taken from the hippocampus. Assay kits were used to assess levels of oxidative stress for marker 4-HNE (4-hydroxy-2-nonenal) and antioxidant markers GSH, glutathione peroxidase (GPx) and glutathione reductase (GR). Levels of p53 were determined using an assay kit.  </td> <td> After 4 Gy, 4-HNE increased 2.4-fold, protein carbonylation increased 3.2-fold, GSH decreased 0.4-fold, GPx decreased 0.3-fold, GR decreased 0.2-fold, and p53 increased 2.7-fold.  </td> </tr> <tr> <td> Suman et al., 2013  </td> <td> In vivo. Female adult C57BL/6J mice were irradiated with 1.6 Gy of 56Fe or 2 Gy of 137Cs gamma irradiation at 1 Gy/min, then measurements were taken from the cerebral cortex. ROS levels were determined with flow cytometry and 4-HNE levels were assessed with immunohistochemical staining. p21 and p53 levels were determined with immunoblotting.  </td> <td> ROS increased a maximum of 1.2-fold after gamma rays and 1.4-fold after 56Fe radiation. The number of 4-HNE+ cells increased a maximum of 4.4-fold after gamma radiation and 14-fold after 56Fe radiation. p21 increased a maximum of 1.5-fold after gamma rays and 3-fold after 56Fe radiation. p53 increased a maximum of 8.4-fold after gamma rays and 9-fold after 56Fe radiation.  </td> </tr> <tr> <td> Limoli et al., 2004  </td> <td> In vivo. Adult male C57BL/6J mice were irradiated with 1-10 Gy of X-ray at 1.75 Gy/min. MDA levels in the hippocampus were measured using an assay kit and western blot was used to determine p53 and p21 levels.    In vitro. Neural precursor cells from the rat hippocampus were irradiated with 1-10 Gy of X-ray at 4.5 Gy/min. ROS levels were measured using CM-H2DCFDA dye and Western blot was used to measure p53 and p21 levels.  </td> <td> MDA levels increased about 30% at 10 Gy. ROS increased a maximum of 31% at 1 Gy and 35% at 5 Gy, after 24 and 12 hours, respectively. At 5 Gy, p53 levels increased a maximum of 4-fold, while p- p21 also increased at this dose.  </td> </tr> <tr> <td> Tian et al., 2020  </td> <td> In vivo. C57BL/6J mice (including miR-137-/- and Src-/- models) underwent middle cerebral artery occlusion (MCAO) to simulate ischemic stroke and measurements were taken 7 days later in the cerebral cortex. ROS levels were measured with DCFH-DA fluorescent dye. Signaling molecules were measured with western blotting or RT-qPCR.  </td> <td> ROS increased 1.8-fold. ERK1/2, p38 and JNK mRNA increased 2- to 3- fold. The ratios of phosphorylated to total ERK1/2, p38 and JNK increased 2- to 3- fold as well.    </td> </tr> <tr> <td> Hladik et al., 2020  </td> <td> In vivo. Female B6C3F1 mice were exposed to total body 60Co gamma irradiation at 0.063, 0.125, or 0.5 Gy and at a dose rate of 0.063 Gy/min. Measurements from the hippocampus were taken up to 24 months post-irradiation. Protein levels in various signaling pathways (CREB, p38, ERK1/2, pro-apoptotic Bax and cleaved caspase 3, anti-apoptotic Bcl-xL) were determined with immunoblotting.  </td> <td> Carbonylated proteins (indicative of ROS levels) were elevated in the 0.125 and 0.5 Gy group by approximately 25% and 30%, respectively. CREB phosphorylation increased by approximately 20% and 25% at 0.063 and 0.125 Gy, respectively. Phosphorylated p38 increased by approximately 100% and 80% at 0.063 and 0.125 Gy, respectively. Phosphorylated ERK1/2 increased by approximately 100% and 90% at 0.063 and 0.125 Gy, respectively.     Anti-apoptotic BCL-xL decreased by 1.7-fold at 0.5 Gy, whereas pro-apoptotic Bax increased by approximately 2-fold at this dose. Caspase 3 also increased by approximately 2-fold at 0.5 Gy.  </td> </tr> <tr> <td> Carvour et al., 2008  </td> <td> In vitro. Mesencephalic dopaminergic neuronal cell line (N27) derived from rat mesencephalon were exposed to 3, 10, or 30 μM of H2O2. ROS levels were detected using dihydroethidine dye and flow cytometry. Western blot was used to detect cleaved PKCδ and Sytox fluorescence was used to measure caspase-3 enzyme activity.   </td> <td> Exposure to 10 and 30 μM of H2O2 resulted in 34 and 58% increases in ROS production, respectively, compared to untreated N27 cells. Exposure to 3, 10, and 30 μM hydrogen peroxide resulted in 2-, 10-, and 9-fold increases in caspase-3 enzyme activity. Lastly, exposure to 10 and 30 μM of H2O2 dose-dependently induced proteolytic cleavage of PKCδ.  </td> </tr> <tr> <td> Chen et al., 2009  </td> <td> In vitro. PC12 and SH-SY5Y human cells were incubated with H2O2. The production of ROS was measured by detecting the fluorescent intensity of oxidant-sensitive probe CM-H2DCFDA. Western blot analysis was used to assess activation of MAPKs.  </td> <td> Treatment with H2O2 for 24 h resulted in a concentration-dependent increase of ROS production at the concentrations of 0–1 mM in PC12 and SH-SY5Y cells. In comparison with PC12, SH-SY5Y cells appeared to be more sensitive to H2O2, thereby showing a decreased ROS production at 2 mM. Additionally, treatment of PC12 cells with H2O2 for 2 h increased phosphorylation of Erk1/2 and p38 in a concentration-dependent manner. Noticeably, H2O2-activation of JNK resulted in a robust (5–10-fold) increase of protein expression and phosphorylation of c-Jun at 0.3–1 mM. Similar results were also seen in SH-SY5Y cells (data not shown).  </td> </tr> </tbody> </table>

Time Concordance  <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Azimzadeh et al., 2017  </td> <td> In vitro. HCAECs were irradiated with 0.5 Gy of X-rays (0.5 Gy/min). Protein carbonylation and GSTO1 antioxidant level were measured with a carbonylation assay and immunoblotting, respectively. Proteins from various signaling pathways including RhoGDI, p16 and p21 were measured with immunoblotting. Measurements were taken at 1, 7, and 14 days after irradiation.  </td> <td> After 7 and 14 days, carbonyl content increased 1.2-fold (insignificant increase at 1 day post-irradiation). After 1-14 days, GSTO1 decreased 0.78-fold (significant decreases at all timepoints). After 1 and 7 days, p-RhoGDI decreased 0.7-fold (non-significant decrease at 14 days post-irradiation). p16 increased 1.2-fold after 7 days and 1.5-fold after 14 days (non-significant increase at 1 day post-irradiation). p21 increased 1.2-fold after 7 and 14 days (insignificant increase at 1 day post-irradiation).  </td> </tr> <tr> <td> Wortel et al., 2019  </td> <td> In vitro. BAECs were irradiated with 10 Gy 137Cs gamma rays at a rate of 1.66 Gy/min. Superoxide levels were quantified by luminescence after treatment with Diogenes Complete Enhancer Solution. The activation of the ASMase enzyme and the levels of ceramide were quantified by radioenzymatic assay to deter mine the changes on the ASMase/ceramide pathway.  </td> <td> Superoxide increased by over 350% at 2 minutes post-irradiation. ASMase activity increased to a maximum 5.6-fold at 5 min post-irradiation. Ceramide increased from -500 to over 3000 pmol/106 cells at 5 minutes post-irradiation. The significance of these changes was not indicated against a control.  </td> </tr> <tr> <td> Kook et al., 2015  </td> <td> In vitro. MC3T3-E1 osteoblast-like cells were irradiated with X-rays (1.5 Gy/min). ROS were measured with a fluorescent probe, and SOD, CAT, and GSH antioxidant activities were determined with assay kits. Protein levels in the Nrf2/HO-1 signaling pathway were determined by either western blot or RT-PCR.  </td> <td> After 1 day and 8 Gy, ROS increased 1.4-fold, GSH decreased 0.5-fold, and SOD decreased 0.5-fold. CAT was also decreased but not significantly. After 1 day and 8 Gy, Nrf2 increased 2.3-fold. After 2 days and 8 Gy, HO-1 increased 4.9-fold. After 3 days and 8 Gy, Runx2 mRNA was decreased 0.5-fold.  </td> </tr> <tr> <td> Suman et al., 2013  </td> <td> In vivo. Female adult C57BL/6J mice were irradiated with 1.6 Gy of 56Fe or 2 Gy of 137Cs gamma irradiation at 1 Gy/min, then measurements were taken from the cerebral cortex until up to 12 months. ROS levels were determined with flow cytometry and 4-HNE levels were determined with immunohistochemical staining. p21 and p53 levels were determined with immunoblotting.  </td> <td> All changes after 56Fe radiation were found after both 2 and 12 months post-irradiation. Most endpoints were also increased at both time points following gamma irradiation, however, p21 only increased at 12 months by 3-fold, but not 2 months, while oxidative stress was shown at 2 months (0.2-fold increase).    </td> </tr> <tr> <td> Xu et al., 2019  </td> <td> In vitro. Adult male C57BL/6 mice experienced chronic cold stress for various lengths (1, 2 and 3 weeks). Brain tissue was then collected, and Western blot was used to measure MDA and proteins of MAPK (JNK, ERK and p38).  </td> <td> MDA levels increased in a time-dependent manner. At 1 week, there was an approximate 3-fold increase, at 2 weeks was an approx. 4-fold increase and for 3 weeks, there was an approx. 5-fold increase in response to cold stress. Phosphorylated JNK increased by &sim;10% (1 week) and &sim;30% at 2 and 3 weeks compared to room temperature control. Phosphorylated ERK increased by &sim;60% at 1 week, &sim;150% at 2 weeks and &sim;140% at 3 weeks. Phosphorylated p38 increased by &sim;50% at 1 week, &sim;100% at 2 weeks and &sim;150% at 3 weeks.  </td> </tr> <tr> <td> Chen et al., 2009  </td> <td> In vitro. PC12 and SH-SY5Y human cells were incubated with hydrogen peroxide. The production of ROS was measured by detecting the fluorescent intensity of oxidant-sensitive probe CM-H2DCFDA. Western blot analysis was used to assess activation of MAPKs.  </td> <td> They observed that H2O2 induced phosphorylation of MAPKs in a time-dependent fashion. Within 5–15 min, H2O2 increased phosphorylation of Erk1/2, JNK and p38, and such phosphorylation was sustained for over 2 h. Consistently, high levels of c-Jun and phospho-c-Jun were induced. </td> </tr> </tbody> </table>

ROS can upregulate protein kinase C, which stimulates the production of ceramide from sphingomyelinase. Ceramide activates NADPH oxidase, which can then produce more ROS (Soloviev & Kizub, 2019). Another feedback loop exists between the Nrf2/HO-1 signaling pathway and oxidative stress. The Nrf2/HO-1 signaling pathway is involved in negative feedback of oxidative stress, activating transcription of anti-oxidative enzymes to regulate cellular ROS and maintain a redox balance (Tahimic & Globus, 2017; Tian et al., 2017). Lastly, the MAPK pathway also exhibits a feedback loop. ERK can regulate ROS levels indirectly through p22phox, which increases ROS and upregulates antioxidants by Nrf2 activation. JNK activation can lead to FoxO activation, thereby resulting in antioxidant production (Arfin et al., 2021; Essers et al., 2004).

High Male Low Female Low Unspecific

Moderate

Adult

Moderate

Juvenile

Low High High Moderate

Based on the prioritized studies presented here, the evidence of taxonomic applicability is low for humans despite there being strong plausibility as the evidence only includes in vitro human cell-derived models. The taxonomic applicability for mice and rats is considered high as there is much available data using in vivo rodent models that demonstrate the concordance of the relationship. The taxonomic applicability was determined to be moderate for pigs as only one in vivo study provided meaningful support to the relationship. In terms of sex applicability, all in vivo studies that indicated the sex of the animals used male animals, therefore, the evidence for males is high and females is considered to be low for this KER. The majority of studies used adolescent animals, with a few using adult animals. Preadolescent animals were not used to support the KER; however, the relationship in preadolescent animals is still plausible.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b42ea9e8560> AOPWiki 2022-09-28T12:40:29

2023-03-21T14:10:58

<upstream-id>8d6a4475-5817-4386-be53-eb911186774f</upstream-id> <downstream-id>1cc566f9-d2e5-4c9c-a06c-c40f7767c251</downstream-id> Deposition of energy from irradiation can affect nitric oxide (NO), a diffusible signaling molecule responsible for vasodilation (Dong et al., 2020; Mitchell et al., 2019; Soloviev & Kizub, 2019; Wang, Boerma & Zhou, 2016). NO activity is regulated by nitric oxide synthase (NOS) enzymes, which can be affected by NOS protein concentrations and cofactors tetrahydobiopterin (BH4), nicotinamide adenine dinucleotide phosphate (NADPH) and Ca2+ (Luiking, Engelen & Deutz, 2010). The deposition of energy can alter certain pathways involving NO and therefore indirectly alter NO levels. The phosphatidylinositol 3-kinase (PI3K)/Akt pathway, the RhoA/Rho kinase (ROCK) pathway, the renin-angiotensin-aldosterone system (RAAS), and the acidic sphingomyelinase/ceramide pathway can influence NO levels (Hemmings & Restuccia 2012; Millatt, Abdel-Rahman & Siragy, 1999; Nagane et al., 2021; Soloviev & Kizub, 2019; Yao et al., 2010).  Deposition of energy can alter NO levels through radiolysis and the direct formation of reactive oxygen species (ROS) (Azzam, Jay-Gerin & Pain, 2013). An increase in ROS and reactive nitrogen species (RNS) can influence NO levels; however, the involvement of RNS in NO production has not been strongly demonstrated in literature (Nagane et al., 2021). ROS acts as a modulator for the relationship between energy deposition leading to altered NO levels. Following ionizing radiation (IR) exposure, there are various enzymes and immune cells involved with indirectly increasing ROS, thereby influencing NO levels (Powers & Jackson, 2008; Soloviev & Kizub, 2019; Vargas-Mendoza et al., 2021).

Overall weight of evidence: High

The biological plausibility surrounding the connection between deposition of energy and altered NO levels is well-supported by reviews in the literature and mechanistic understanding. NO is a diffusible molecule produced by endothelial cells (Soloviev & Kizub, 2019; Wang, Boerma & Zhou, 2016). The enzyme NOS produces NO and can be used as a proxy to measure NO levels. eNOS and iNOS are common endpoints for assessing NO levels. Changes in eNOS phosphorylation (p-eNOS) can also indicate NO levels, with phosphorylation at serine 1177 increasing eNOS activity and phosphorylation at threonine 495 decreasing eNOS activity (Nagane et al., 2021). The deposition of energy from IR can indirectly lead to changes in NO levels through various pathways (Nagane et al., 2021; Soloviev & Kizub, 2019).  NO levels can be altered by deposition of energy through ROS. ROS can be produced directly through the radiolysis of water or indirectly through the mitochondrial electron transport chain (ETC) and various enzymes and immune cells (Azzam, Jay-Gerin & Pain, 2013; Powers & Jackson, 2008; Soloviev & Kizub, 2019; Vargas-Mendoza et al., 2021). NO bioavailability is reduced through the molecule’s reaction with ROS that produces peroxynitrite, or oxidation of the NOS cofactor BH4. This uncouples NOS causing it to produce ROS instead of NO, further driving down NO bioavailability (Forrester et al., 2019; Soloviev & Kizub, 2019). NO can also increase as a result of deposition of energy through activation of iNOS during oxidative stress (Nathan & Xie, 1994). However, this additional NO reacts with ROS to form peroxynitrite (Nagane et al., 2021; Soloviev & Kizub, 2019). The reaction of ROS with NO produces the RNS peroxynitrite, which can also further oxidize BH4 and uncouple NOS, resulting in further NO reduction (Soloviev & Kizub, 2019).  Deposition of energy can also alter NO levels by activating signaling pathways involved in NO regulation. Under normal conditions, the PI3K/Akt pathway can activate NOS through phosphorylation (Hemmings & Restuccia 2012; Nagane et al., 2021). The RhoA/ROCK pathway prevents both the expression and phosphorylation of NOS (Yao et al., 2010). Furthermore, the RAAS pathway can increase the production of ROS through NADPH oxidase (NOX), which causes decreased NO (Nguyen Dinh Cat et al., 2013). Additionally, the RAAS pathway can activate NOS as a countermeasure for vasoconstriction (Millatt, Abdel-Rahman & Siragy, 1999). Lastly, the acidic sphingomyelinase/ceramide pathway can also activate NOX, resulting in lower NO levels (Soloviev & Kizub, 2019). Deposition of energy from IR can change these pathways through oxidative stress and changes in protein expression, which results in altered NO levels (Schmidt-Ullrich et al., 2000).

The empirical evidence supporting this KER was collected from research using both in vivo and in vitro models. The use of X-rays and gamma rays as stressors with doses ranging from 0.05 Gy to 60 Gy demonstrates the relationship between deposition of energy and altered NO levels. The dose, dose rate, and radiation type will all influence the level of energy deposition and therefore of NO production, but insufficient evidence exists to quantify the relationship between these factors and NO production. Human coronary artery endothelial cells (HCAECs) (Azimzadeh et al. 2017), human aortic endothelial cells (HAoECs) (Azimzadeh et al., 2021), human umbilical vein endothelial cells (HUVECs) (Hong et al. 2013; Sakata et al., 2015; Sonveaux et al., 2003), bovine aortic endothelial cells (BAECs) (Hirakawa et al., 2002; Sonveaux et al., 2003), tumor models (Sonveaux et al., 2003), cardiac microvascular endothelial cells and serum (Azimzadeh et al. 2015), as well as in vivo rat pulmonary arterioles (Fuji et al., 2016), Wistar albino rat heart tissue and serum (Abdel-Mageid & Shedid, 2019), B6J mice aortic endothelium (Hamada et al., 2020; Hamada et al., 2021), WAG/RijCmcr rat heart tissue (Baker et al., 2009), and serum from the abdominal inferior vena cava (Ohta et al., 2007) were the models used in these studies. Both direct NO measures, such as ELISA assay, and indirect NO measures, such as eNOS, iNOS, citrulline (NOS product) and NOx (nitrite and nitrate, NO metabolites) levels and activity, were used to determine changes in NO levels following energy deposition.    Dose Concordance  Evidence shows concordant changes in NO levels and deposition of energy. The study by Dias et al. (2018) showed varying changes to eNOS after chronic gamma irradiation, with a general trend of decreasing eNOS expression with increasing energy deposition (i.e., at 0.5, 1, and 2 Gy). Similarly, HUVECs exposed to gamma irradiation showed slightly lower eNOS levels at 12.5 Gy than at 10 Gy (Sadhukhan et al., 2020). X-ray irradiation of mouse cardiac microvascular endothelial cells showed 0.6 and 0.2-fold decreases in p-eNOS at 8 and 16 Gy, respectively (Azimzadeh et al., 2015). X-ray irradiation of mouse blood serum showed NO decreased 0.3-fold and 0.2-fold following 8 Gy and 16 Gy X-ray irradiation, respectively (Azimzadeh et al., 2015).   In contrast, BAECs and HUVECs irradiated with X-rays showed a trend of increasing eNOS expression and activation from 2 to 20 Gy (Sonveaux et al., 2003). Using similar doses of X-rays also showed that NOx increased dose-dependently from 1 to 20 Gy and reached a maximum increase of 10-fold (Sakata et al., 2015). Other high X-ray doses from 19.6 to 31.5 Gy on mice showed about a 2-fold increase in serum nitrate concentration, which had a general increasing trend at larger doses (Ohta et al., 2017). In BAECs, eNOS expression did not significantly change from 1 to 60 Gy of X-rays, while iNOS expression increased to 6% of GAPDH expression after 1 Gy, 26% after 2 Gy and remained around 40% of GAPDH expression from 10 to 60 Gy (Hirakawa et al., 2002).  Many studies do not examine this relationship at multiple doses. After 0.5 Gy X-ray irradiation of HCAECs, p-eNOS and NO were both decreased (Azimzadeh et al., 2017). HUVECs irradiated with 4 Gy X-rays showed increases in iNOS and nitrotyrosine (peroxynitrite biomarker) levels (Hong et al., 2013). eNOS levels decreased in mouse aortic endothelial cells when exposed to 5 Gy gamma and X-ray irradiation at various acute and chronic regimens (Hamada et al., 2020; Hamada et al., 2021; Hamada et al., 2021). Rat cardiac and serum NOx increased as a response to 8 Gy gamma irradiation (Abdel-Mageid & Shedid, 2019). Total body irradiation (TBI) of rats with 10 Gy X-rays resulted in a decrease in eNOS, iNOS and NOx levels (Baker et al., 2009). Both cardiac eNOS and serum NO decreased to 73% and 63%, respectively, following 16 Gy X-ray irradiation (Azimzadeh et al., 2021).    Time Concordance  There is evidence that a change in NO levels occurs after energy deposition (i.e., irradiation) occurred. HUVECs irradiated with 4 Gy X-rays displayed an increase in nitrotyrosine (peroxynitrite biomarker) at 6 hours post-irradiation (Hong et al., 2013). They also showed increased iNOS after 1.5, 3, and 6 hours (Hong et al., 2013). Following a 26 Gy X-ray irradiation of mice, serum nitrate concentration showed a maximum increase after 3h, followed by a return to pre-irradiation levels at 12 h and 24 h post-irradiation (Ohta et al., 2007). As well, in HUVECs, 10 and 12 Gy gamma rays led to decreased eNOS levels after 4 and 24 h (Sadhukhan et al., 2020). BAECs and HUVECs irradiated with 6 Gy X-rays did not exhibit significant changes in eNOS 1-6 h post-irradiation, but eNOS increased after 12, 24, and 48 hours (Sonveaux et al., 2003)  HUVECs irradiated with 10 Gy X-rays also had no change in eNOS levels over 72 h, while p-eNOS (Ser1177) increased consistently over 72 h, p-eNOS (Thr495) decreased after 6 h and returned to initial levels after 72 h, iNOS did not change other than a non-significant increased after 24 h, citrulline increased after 6 h and remained the same for 72 h and NOx increased over 72 h (Sakata et al., 2015). After 2 Gy X-ray irradiation of BAECs, eNOS expression did not significantly change from 0-120 h, iNOS expression increased to a maximum 6 h post-irradiation and NO increased to a maximum 12 h (Hirakawa et al., 2002).  Following 0.5 Gy X-ray irradiation of HCAECs, NO was not significantly lower after 1 day, while the 7 and 14 day time-points showed significant decreases (Azimzadeh et al., 2017).  In human aortic endothelial cells, acute gamma radiation consistently showed decreased eNOS at each time-point up to 16 days after irradiation, while chronic irradiation decreased eNOS levels 1 and 4 days post-irradiation (Dias et al., 2018).  Finally, eNOS expression was lower 1, 3, and 6 months post-irradiation of mice aortic endothelial cells with 5 Gy gamma rays (Hamada et al., 2020).    Essentiality  Altered NO levels can occur due to a deposition of energy in the form of ionizing radiation. Since deposited energy initiates events immediately, the removal of deposited energy also supports the essentiality of the key event. Studies that do not deposit energy are observed to have no downstream effects. Deposition of energy cannot be blocked by chemicals, but radiation could be shielded. Currently, no studies show the effect of shielding on this relationship.

<ul> <li>Due to the high reactivity of NO, it can be difficult to directly measure it (Luiking, Engelen & Deutz, 2010). The inconsistencies in NO levels may be attributed to the challenges in measuring NO, such as its availability in cell (Azimzadeh et al., 2017; Hirakawa et al., 2002; Hong et al., 2013; Sakata et al., 2015; Sonveaux et al., 2003), serum (Abdel-Magied & Shedid,. 2019; Azimzadeh et al. 2015; Ohta et al., 2007) or tissue (Abdel-Magied & Shedid, 2019; Baker et al., 2009; Fuji et al., 2016; Hamada et al., 2020), its homeostasis with ROS, and its relationship to nitrosylation as accumulated damage and whether p-eNOS/eNOS is being measured.</li> </ul>

Examples of quantitative understanding of the relationship are provided. All data that is represented is statistically significant unless otherwise indicated.

Dose Concordance  <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Azimzadeh et al., 2015  </td> <td> In vivo. 10-week-old mice were exposed to X-ray irradiation at either 8 or 16 Gy. Cardiac levels of NO, eNOS and p-eNOS were determined. Protein levels were measured using immunoblotting, and NO was measured using an ELISA assay.  </td> <td> p-eNOS decreased 0.6-fold after 8 Gy and 0.2-fold after 16 Gy. NO decreased 0.3-fold after 8 Gy and 0.2-fold after 16 Gy. eNOS levels did not change.  </td> </tr> <tr> <td> Azimzadeh et al., 2017  </td> <td> In vitro. HCAECs were irradiated with 0.5 Gy X-ray irradiation over 1 minute. Phosphorylated and total eNOS levels were determined using immunoblotting, and NO levels were determined using ELISA assay. Measurements were taken after 1, 7, and 14 days.  </td> <td> p-eNOS/total eNOS was 0.7-fold lower than control and NO was 0.8-fold lower than control after 14 days.    </td> </tr> <tr> <td> Hong et al., 2013  </td> <td> In vitro. HUVECs were irradiated with 4 Gy X-rays (2.7 Gy/min) and amounts of eNOS, iNOS and nitrotyrosine (a biomarker for peroxynitrite) were determined by western blot. Measurements were taken after 1.5, 3, and 6 hours.  </td> <td> In 4 Gy irradiated HUVECs, iNOS was increased 6.6-fold and nitrotyrosine was increased 6.4-fold after 6 h. eNOS expression did not change.  </td> </tr> <tr> <td> Fuji et al., 2016  </td> <td> In vivo. eNOS levels in the smooth muscle layer of 4-week-old rat pulmonary arteries were determined using immunohistochemical staining after irradiation (dose not specified). A 6.5 GeV electron beam was converted into monochromatic X-rays. Measurements were taken after 2 weeks.  </td> <td> eNOS levels decreased 0.6-fold compared to control.  </td> </tr> <tr> <td> Abdel-Magied & Shedid, 2019  </td> <td> In vivo. Adult male rats were irradiated with 8 Gy 137Cs gamma irradiation (0.4092 Gy/min). Serum and heart total NOx was determined with a NOx assay kit. Measurements were taken after 14 days.  </td> <td> Cardiac and serum NOx increased 2 and 1.8-fold after exposure to 8 Gy, respectively.    </td> </tr> <tr> <td> Hamada et al., 2022  </td> <td> In vivo. 8-week-old mice were irradiated with 5 Gy 260 kVp X-rays at 0.5 Gy/min delivered in 25 or 100 daily fractions over 42 or 152 days, respectively, or delivered as an acute single dose. As well, chronic 137Cs gamma rays (<1.4 mGy/h for 153 days) were delivered in another regimen. eNOS levels were determined by immunofluorescence.    </td> <td> At 6 months after the start of irradiation, eNOS decreased 0.3-fold, 0.4-fold, and 0.4-fold following single dose irradiation, 25 fraction regimens, and 100 fraction regimens, respectively. eNOS also decreased 0.8-fold after chronic gamma ray irradiation. At 12 months after the start of irradiation, eNOS decreased 0.7-fold, 0.8-fold and 0.8-fold following acute, 25 fraction and 100 fraction regimens, respectively.   </td> </tr> <tr> <td> Hamada et al., 2020  </td> <td> In vivo. 8-week-old male mice were irradiated by 5 Gy 137Cs gamma rays (0.5 Gy/min). Immunofluorescence was used to determine eNOS levels. Measurements were taken after 1, 3, and 6 months.  </td> <td> 6 months post-irradiation of 5 Gy, eNOS decreased by 0.3- fold.  </td> </tr> <tr> <td> Baker et al., 2009  </td> <td> In vivo. Male rats were irradiated with 10 Gy X-rays (1.95 Gy/min). Western blotting was used to determine eNOS and iNOS. NOx content was determined using ozone chemiluminescence. Measurements were taken after 120 days.  </td> <td> Rats exposed to 10 Gy of TBI experienced a 27% decrease in eNOS, a 29% decrease in iNOS protein, and a 20% decrease in NOx (index of NO activity).  </td> </tr> <tr> <td> Sonveaux et al., 2003  </td> <td> In vitro. Levels of eNOS and p-eNOS after X-ray irradiation of BAECs and HUVECs were measured with immunoblotting at various doses (0.86 Gy/min). eNOS mRNA was also measured with RT-qPCR before and after 2 Gy-irradiated human tumour cells.  </td> <td> Compared to controls, 24 h after irradiation, eNOS increased 1.3-fold after 2 Gy (not significant), 2.1-fold after 4 Gy (not significant), 3-fold after 6 and 8 Gy, 4.3-fold after 10 Gy and 3.4-fold after 20 Gy. Compared to controls, 24 h after irradiation, p-eNOS increased 1.2-fold after 2 Gy and 1.7-fold after 6 Gy. eNOS increased in human tumor cells in every patient after 2 Gy irradiation.  </td> </tr> <tr> <td> Hirakawa et al., 2002  </td> <td> In vitro. BAECs were irradiated with X-rays at various doses. eNOS and iNOS were measured with semiquantitative RT-PCR and western blot. NO was measured through DAF-2, a NO-sensitive fluorescent dye after treatment with L-arginine.  </td> <td> eNOS expression did not significantly change from 1-60 Gy. iNOS expression did not occur in non-irradiated samples, but increased to 6% of GAPDH expression after 1 Gy, 26% after 2 Gy and remained around 40% of GAPDH expression from 10-60 Gy. NO increased a maximum of 5.2-fold 12h after 2 Gy irradiation.  </td> </tr> <tr> <td> Sakata et al., 2015  </td> <td> In vitro. HUVECs were irradiated with various doses of X-rays. eNOS, p-eNOS (Ser1177 & Thr495), iNOS, citrulline (produced by NOS with NO) and NOx levels were measured with western blots for proteins and various assay kits for molecules.  </td> <td> For maximum changes after 10 Gy, eNOS expression did not significantly change, p-eNOS (Ser1177) increased 1.8-fold, p-eNOS (Thr495) decreased 0.3-fold, iNOS showed a non-significant 1.4-fold increase and citrulline increased 1.3-fold. NOx increased consistently from 1-20 Gy and showed a maximum 10-fold increase after 10 Gy.  </td> </tr> <tr> <td> Ohta et al., 2007  </td> <td> In vivo. Levels of nitrate in mouse serum were measured by Griess assay after a whole-body X-ray irradiation of 6-week-old mice using various doses from 19.6 to 31.5 Gy.  </td> <td> Serum nitrate concentrations increased about 2-fold from 0-31.5 Gy.  </td> </tr> <tr> <td> Hamada et al., 2021  </td> <td> In vivo. Either acute or chronic doses of X-rays and 137Cs gamma rays were given to 8-week-old B6J mice, to a total 5 Gy dose. X-rays were given as a single acute dose, 25 fractions of 0.2 Gy/fraction spread over 42 days or 100 fractions of 0.05 Gy/fraction spread over 153 days all given at 0.5 Gy/min. Gamma rays were given as a single acute dose at 0.5 Gy/min or chronically at <1.4 mGy/h for 153 days. eNOS was measured through immunofluorescence.  </td> <td> All regimens of X-rays and acute gamma rays led to a 0.3- to 0.4-fold decrease in eNOS levels. Chronic gamma rays led to a 0.8-fold (nonsignificant) decrease in eNOS levels.  </td> </tr> <tr> <td> Azimzadeh et al., 2021  </td> <td> In vivo. 8-week-old male C57BL/6J were irradiated with an acute dose of 16 Gy X-rays. eNOS activity and NO were measured using fluorometric and Griess assays, respectively.  </td> <td> 16 Gy resulted in a decrease in eNOS activity to 73% in heart tissue. NO levels decreased to 63% in serum.  </td> </tr> <tr> <td> Sadhukhan et al., 2020  </td> <td> In vitro. HUVECs were irradiated with various regimens and doses of 137Cs gamma radiation. eNOS levels were determined by western blot. Fractionated doses were separated by 24 h.  </td> <td> At 5 doses of 2 Gy, eNOS decreased 0.2-fold. At 5 doses of 2.5 Gy, eNOS decreased 0.3-fold. At a single dose of 10 Gy, eNOS decreased 0.8-fold. At a single dose of 12 Gy, eNOS decreased 0.7-fold.  </td> </tr> <tr> <td> Dias et al., 2018  </td> <td> In vitro. HAoECs were irradiated with various regimens of 60Co gamma radiation. Acute radiation was given at 1 Gy/min, while chronic radiation was given at 6 mGy/h eNOS mRNA level was determined using qPCR. Each dose was measured at different times after irradiation.  </td> <td> An acute dose of 0.05 Gy led to a 0.6-fold decrease in eNOS while the chronic dose led to a 1.6-fold increase. At 0.5 Gy, the acute dose did not significantly change eNOS, while the chronic dose increased it 1.6-fold. Acute 1 Gy caused eNOS to decrease 0.5-fold without significant changes chronically. Acute 2 Gy caused a 0.3-fold decrease in eNOS without significant changes chronically.  </td> </tr> </tbody> </table>

Time Concordance  <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Azimzadeh et al., 2017  </td> <td> In vitro. HCAECs were irradiated with 0.5 Gy X-ray irradiation over 1 minute. Phosphorylated and total eNOS levels were determined using immunoblotting, and NO levels were determined using ELISA assay. Measurements were taken after 1, 7 or 14 days.  </td> <td> 1 day after 0.5 Gy irradiation, p-eNOS/total eNOS was 0.8-fold lower than control and NO did not change. After 7 days, p-eNOS/total eNOS was 0.6-fold lower than control and NO was 0.8-fold lower than control. After 14 days, p-eNOS/total eNOS was 0.7-fold lower than control and NO was 0.8-fold lower than control.  </td> </tr> <tr> <td> Hong et al., 2013  </td> <td> In vitro. HUVECs were irradiated with 4 Gy X-rays (2.7 Gy/min) and amounts of iNOS and nitrotyrosine (a biomarker for peroxynitrite) were determined by western blot at various times over 6 hours.  </td> <td> 1.5 h after 4 Gy irradiation, iNOS 1.9-fold higher than control and nitrotyrosine did not change. After 3 h, iNOS was 2.8-fold higher than control and nitrotyrosine did not change. After 6 h, iNOS was 6.6-fold higher than control and nitrotyrosine was 6.3-fold higher than control.  </td> </tr> <tr> <td> Hamada et al., 2020  </td> <td> In vivo. 8-week-old male mice were irradiated by 5 Gy 137Cs gamma rays (0.5 Gy/min). Immunofluorescence was used to determine eNOS levels. Measurements were taken after 1, 3 and 6 months.  </td> <td> eNOS levels decreased 0.5-fold at 1-month post-irradiation, 0.4-fold at 3 months post-irradiation and a maximal fold decrease of 0.3-fold 6 months post-irradiation.  </td> </tr> <tr> <td> Sonveaux et al., 2003  </td> <td> In vitro. Level of eNOS after X-ray irradiation of BAECs and HUVECs was measured with immunoblotting at various times.  </td> <td> At 6 Gy, eNOS did not significantly change 1-6 h post-irradiation, but increased 1.8-fold after 12 h, 2-fold after 24 h and 1.5-fold after 48 h.  </td> </tr> <tr> <td> Hirakawa et al., 2002  </td> <td> In vitro. BAECs were irradiated with 2 Gy X-rays at various times. eNOS and iNOS were measured with semiquantitative RT-PCR and western blotting. NO was measured through DAF-2, a NO-sensitive fluorescent dye after treatment with L-arginine.  </td> <td> eNOS expression did not significantly change from 0-120 h post-irradiation. iNOS expression did not occur in non-irradiated samples but increased to a maximum 26% of GAPDH expression 6 h post-irradiation, and slowly decreased to 16% after 120 h. Similarly, NO increased a maximum of 5.2-fold after 12 h, while NO levels slowly returned after 120 h.  </td> </tr> <tr> <td> Sakata et al., 2015  </td> <td> In vitro. HUVECs were irradiated with 10 Gy X-rays at various times. eNOS, p-eNOS (Ser1177 & Thr495), iNOS, citrulline (produced by NOS with NO) and NOx levels were measured with western blots for proteins and various assay kits for molecules.  </td> <td> eNOS expression did not significantly change over 72 h, p-eNOS (Ser1177) increased 1.8-fold over 72 h, p-eNOS (Thr495) decreased 0.3-fold after 6 h and returned to initial levels after 72 h, iNOS did not change other than a nonsignificant 1.4-fold increased after 24 h. Citrulline increased 1.3-fold after 6 h and remained the same for 72 h. NOx increased 3.7-fold after 6 h and 5-fold after 72 h.  </td> </tr> <tr> <td> Ohta et al., 2007  </td> <td> In vivo. Levels of nitrate in mouse serum were measured by Griess assay at various times after a whole-body 26 Gy X-ray irradiation of 6-week-old mice.  </td> <td> Serum nitrate concentration showed a maximum increase of 2.1-fold after 3 h, followed by a return to pre-irradiation levels at 12 h and 24 h post-irradiation.  </td> </tr> <tr> <td> Sadhukhan et al., 2020  </td> <td> In vitro. HUVECs were irradiated with various regimens and doses of 137Cs gamma radiation. eNOS levels were determined by western blot 4 and 24 h after irradiation. Fractionated doses were separated by 24 h.  </td> <td> After the 10 Gy doses, eNOS was lowest at 4 h, but after 12 Gy it was the same at both 4 and 24 h.  </td> </tr> <tr> <td> Dias et al., 2018  </td> <td> In vitro. HAoECs were irradiated with various regimens of 60Co gamma radiation. Acute radiation was given at 1 Gy/min, while chronic radiation was given at 6 mGy/h. eNOS mRNA level was determined using qPCR up to 16 days after irradiation. Each timepoint used a different dose. </td> <td> An acute dose of 0.05 Gy led to a 0.6-fold decrease in eNOS while the chronic dose led to a 1.6-fold increase 1 day post-irradiation. At 0.5 Gy, the acute dose did not significantly change eNOS, while the chronic dose increased it 1.6-fold 4 days post-irradiation. Acute 1 Gy caused eNOS to decrease 0.5-fold without significant changes chronically 8 days post-irradiation. Acute 2 Gy caused a 0.3-fold decrease in eNOS without significant changes chronically 16 days post-irradiation.  </td> </tr> </tbody> </table>

Not identified

Moderate Male Low Female Low Unspecific

Moderate

Adult

Moderate

Juvenile

Low High Moderate

The evidence for the taxonomic applicability to humans is low as evidence comes from in vitro human cell-derived models. The relationship has been shown in vivo in mice and rats, with considerable evidence in mice. The relationship has been shown in vivo in males and is likely in females. Most in vivo studies indicate adult or adolescent animal models used. In addition, the relationship is also likely in preadolescent animals.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b42eaa9ed88> AOPWiki 2022-09-28T12:44:42

2023-03-21T11:06:40

<upstream-id>b813d3a8-c9cc-49b4-bd70-a0bb3e9e8efd</upstream-id> <downstream-id>1329b7b0-4714-406a-a848-17ef92fd2b72</downstream-id> The increase in reactive oxygen species (ROS) and reactive nitrogen species (RNS) during a state of oxidative stress can stimulate an increase in pro-inflammatory mediators. Reactive oxygen and nitrogen species (RONS) cause cellular damage, which leads to the production of pro-inflammatory mediators (Slezak et al., 2015; Sylvester et al., 2018; Wang et al., 2019a). In addition, ROS can act as second messenger signalling molecules in activating pro-inflammatory transcription factor nuclear factor kappa B (NF-κB), resulting in increased production of pro-inflammatory cytokines and adhesion factors (Ping et al., 2020; Slezak et al., 2017; Slezak et al., 2015; Sylvester et al., 2018; Venkatesulu et al., 2018; Wang et al., 2019a). The inflammatory state induced by RONS will further increase RONS, leading to a cycle of chronic inflammation and oxidative stress (Venkatesulu et al., 2018; Wang et al., 2019a).

Overall weight of evidence: Moderate

The biological plausibility of the linkage between oxidative stress and pro-inflammatory mediators is strongly supported by review papers on the subject (Ping et al., 2020; Ramadan et al., 2021; Slezak et al., 2017; Slezak et al., 2015; Sylvester et al., 2018; Venkatesulu et al., 2018; Wang et al., 2019a). Pro-inflammatory mediators are released in instances of cell damage to recruit macrophages, monocytes, and other scavengers to ingest and degrade dead and damaged cells. As a major pathway of cell damage, oxidative stress causes upregulation of pro-inflammatory mediators including NF-κB, transforming growth factor-β (TGF-β), tumour necrosis factor-α (TNF-α), interleukin-1 (IL-1), and interleukin-6 (IL-6) (Ping et al., 2020; Slezak et al., 2017; Slezak et al., 2015; Sylvester et al., 2018; Venkatesulu et al., 2018; Wang et al., 2019a). Oxidative stress also stimulates a rise in pro-inflammatory adhesion factors, such as E-selectin, intercellular adhesion molecule-1 (ICAM1), and vascular cell adhesion molecule-1 (VCAM1), which facilitate inflammation by assisting the entrance of inflammatory cells into tissues and recruiting macrophages (Ping et al., 2020; Slezak et al., 2017; Slezak et al., 2015; Sylvester et al., 2018; Venkatesulu et al., 2018; Wang et al., 2019a).    Once antioxidant levels become exhausted in a state of oxidative stress, ROS are present in higher concentrations and can therefore act more effectively as second messenger signalling molecules in activating pro-inflammatory transcription factors, such as NF-κB, and stimulating production of pro-inflammatory cytokines, such as IL-1, IL-6 and TNF-α (Ping et al., 2020; Sylvester et al., 2018; Wang et al., 2019a). NF-κB is normally kept in an inactive state through formation of a complex with the IkB family of inhibitor proteins but is activated by oxidative stress through nuclear translocation of the complex to the promoter areas of inflammation regulatory genes (Ping et al., 2020; Slezak et al., 2017). The macrophages that are recruited in the resulting inflammatory response can also produce ROS and activate the pro-inflammatory mediator, TGF-β, forming a positive feedback loop (Venkatesulu et al., 2018). Another positive feedback loop is formed by ROS and NF-κB, as ROS activates NF-κB, resulting in the expression of the genes cyclooxygenase-2 (COX-2) and 5-lipoxygenase (5-LPO), which are responsible for ROS production (Ping et al., 2020). In addition, NF-κB is also involved in the production of the pro-inflammatory adhesion factors ICAM and VCAM (Ping et al., 2020; Slezak et al., 2017; Slezak et al., 2015).    Oxidative stress may also result in oxidation of low-density lipoproteins, allowing the lipoproteins to be ingested by macrophages. This could initiate the atherosclerotic process (plaque build-up in the arteries) and subsequently lead to lipid cells secreting pro-inflammatory cytokines, such as IL-1β and TGF-β (Ramadan et al., 2021; Slezak et al., 2017; Sylvester et al., 2018; Ping et al., 2020; Venkatesulu et al., 2018).

The empirical evidence relevant to this KER provides moderate support for the linkage between increases in oxidative stress and increases in pro-inflammatory mediators. The evidence to support this relationship comes from studies examining the effects of ionizing radiation, such as 137Cs gamma-rays, on the cardiovascular system. There is moderate evidence for a dose- and time-dependent relationship between oxidative stress and pro-inflammatory mediators (Abdel-Magied & Shedid, 2019; Chen et al., 2019; Cho et al., 2017; Ismail et al., 2015; Ismail et al., 2016; Karam et al., 2019; Philipp et al., 2020; Wang et al., 2019a).    Dose Concordance  Current literature on the dose-dependent relationship between oxidative stress and increases in pro-inflammatory mediators is moderate. Chen et al. (2019) exposed male Sprague Dawley rats to simulated microgravity for 7 and 21 days and measured levels of the oxidative stress marker H2O2 along with the pro-inflammatory mediators, IL-6, interferon-gamma (IFN-γ), and TNF-α. Their finding provided evidence of dose concordance between oxidative stress and pro-inflammatory mediators, as the study observed more significant changes to H2O2 levels at 7 days than 21 days, while IL-6, IFN-γ, and TNF-α levels were more significantly affected at day 21 than day 7 (Chen et al., 2019). Philipp et al. (2020) irradiated human telomerase-immortalized coronary artery endothelial cells (TICAE cells) with 0.25, 0.5, 2, and 10 Gy of 137Cs gamma rays. They found that superoxide dismutase (SOD) decreased consistently at 2 Gy, while pro-inflammatory mediators showed consistent increases at 2 or 10 Gy, but not at lower doses. Other studies using gamma rays show levels of oxidative stress markers increase with subsequent increases to pro-inflammatory mediators following exposure to high doses (>2 Gy), with some markers being significantly affected at low doses (<2 Gy) (Abdel-Magied & Shedid, 2019; Cho et al., 2017; Ismail et al., 2016; Ismail et al., 2015; Karam et al., 2019). However, not all studies demonstrated dose concordance. Wang et al. (2019b) irradiated human endothelial cells with various doses of gamma rays. They found pro-inflammatory mediators significantly increased at lower doses (0.2-5Gy) than oxidative stress (5 Gy only), although this could be due to the sensitivity of the assays.    Time Concordance  Current literature on the time-dependent relationship between oxidative stress and increases in pro-inflammatory mediators is low. Cho et al. (2017) irradiated male C57BL/6 mice and measured levels of superoxide and the pro-inflammatory mediators TNF-α and monocyte chemoattractant protein (MCP-1), at 4, 8, and 24 hours post-irradiation. Oxidative stress and pro-inflammatory mediators demonstrated a time concordant relationship, as ROS levels were significantly increased at 4 hours post-irradiation, while both pro-inflammatory mediators did not significantly change until 8 hours (Cho et al., 2017). Philipp et al. (2020) irradiated telomerase immortalized human coronary artery endothelial cells and measured levels of the antioxidant, SOD1, and the pro-inflammatory adhesion factor, ICAM1, at 4 hours, 24 hours, 48 hours, and 1 week post-irradiation. Neither SOD1 nor ICAM1 followed consistent changes over time across all doses. However, the earliest decrease in SOD1 and the earliest increase in ICAM1 were both found at 4 hours (Philipp et al., 2020).    Incidence concordance  Few studies demonstrated incidence concordance between oxidative stress and increased pro-inflammatory mediators. In human TICAE cells irradiated with 2 Gy of gamma rays, SOD1 was decreased 0.5-fold at 24h post-irradiation, while ICAM1 was increased 1.1-fold at this dose and time (Philipp et al., 2020).    Essentiality  Studies that treated models with countermeasures to suppress the increase in oxidative stress caused by ionizing radiation exposure found that subsequent increases in pro-inflammatory mediators were also significantly attenuated (Abdel-Magied & Shedid, 2019; Karam et al., 2019). Blocking ionizing radiation effect on oxidative stress (upstream KE) and analyzing the subsequent effect on pro-inflammatory mediators (downstream KE) provided evidence for essentiality between the KEs.    Treatment of irradiated albino rat heart tissue with the antidiabetic drug, metformin (50 mg/kg daily for 2 weeks), provided evidence for its efficacy as an antioxidant and anti-inflammatory drug. Metformin treatment following irradiation resulted in a recovery of SOD and catalase (CAT) activity to 90% and 44%, respectively, compared to irradiated groups. In addition to attenuating ionizing radiation effect on oxidative stress, metformin mitigated increases to NF-κB, TNF-α, and IL-6 levels, as well as reduced elevated E-selectin, ICAM, and VCAM levels by 0.5-, 0.55-, and 0.6-fold, respectively (Karam et al., 2019).    Irradiated Wistar albino rats were treated with the food and drug additive ZnO-NP (10 mg/kg daily for 2 weeks), which has antioxidant effects. Treatment with ZnO-NPs attenuated all IR-induced changes to oxidative and inflammatory markers. Compared to the irradiated group, ZnO-NP treatment resulted in restoration of CAT, SOD, glutathione (GSH), and glutathione peroxidase (GPx) levels by ~104%, ~73%, ~91%, and ~73%, respectively. Elevated levels of ICAM, TNF-α, IL-18, and C-reactive protein (CRP) were reduced by ~44%, ~46%, ~45%, and ~42%, respectively, compared to irradiated groups (Abdel-Magied & Shedid, 2019).    Irradiated Wistar rats were treated with flaxseed oil (FSO) (500 mg/kg-bw daily for 7 days), which has a uniquely high content of the antioxidant lignans. Treatment with FSO significantly reduced IR-induced changes to both antioxidants and pro-inflammatory mediators. IR-induced reductions to the activity of the antioxidants SOD, CAT, and GSH-Px were significantly alleviated back to levels statistically similar to the controls. Reduced levels of pro-inflammatory mediators, TNF-α, IL-1β, and IL-6, back to control levels was also observed (Ismail et al., 2016).

<ul> <li>Chen et al. (2019) found that levels of the pro-inflammatory mediators, IL-6, IFN-γ, and TNF-α, decreased following 7 and 21 days of microgravity exposure, contrary to the trend generally observed following ionizing radiation exposure (Chen et al., 2019).</li> </ul>

Several examples of studies that provide quantitative understanding of the relationship are summarized. All data represented is statistically significant unless otherwise indicated.

<ul> <li>Positive feedback loop: oxidative stress upregulates production of pro-inflammatory cytokines, which in turn upregulate ROS production. The macrophages that are recruited in an oxidative stress-induced inflammatory response can also produce ROS and activate the pro-inflammatory mediator, TGF-β (Venkatesulu et al., 2018). Another positive feedback loop is formed by ROS and NF-κB, as ROS activates NF-κB, resulting in expression of the genes, COX-2 and 5-LPO, which are responsible for ROS production (Ping et al., 2020).</li> </ul>

High Male Low Female Low Unspecific

Low

Juvenile

Low

Adult

Moderate

Not Otherwise Specified

Low High Moderate

Most evidence defining the relationship is derived from mice or rat models. A low number of in-vitro human studies were available. Males have been studied more often than females. The age of the models remained unspecified in several studies, while a few studies reported evidence from adult and adolescent models.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b42eab20bf8> AOPWiki 2022-09-28T12:41:20

2023-03-21T14:44:43

<upstream-id>8d6a4475-5817-4386-be53-eb911186774f</upstream-id> <downstream-id>870809d3-663d-4170-9ec7-b63bbcebbc41</downstream-id> Energy deposition can lead to ionization events that can directly interact with molecules within the cell and can subsequently lead to biological changes such as the formation of free radicals and the initiation of DNA damage repair mechanisms. Different radiation types have different physical properties and as a result the biological effects on cells may differ. Dose and dose rate of the deposited energy also play a role as these factors affect the amount and rate of energy deposited (Donaubauer et al., 2020). Repeated or prolonged exposure to radiation can exhaust the protective effect of the endothelium and lead to endothelial dysfunction (Baselet et al., 2019). Consequently, cells within the vascular endothelium may lose their integrity and become senescent or apoptotic via alterations to signaling pathways related to cell survival, leading to dysregulation of vasodilation and eventual endothelial dysfunction (Deanfield et al., 2007; Bonetti et al., 2003). Activation of the endothelium, consisting of inflammation, proliferation, thrombosis and low nitric oxide, occurs as a normal response to pathological conditions and oxidative stress from deposited energy (Krüger-Genge et al., 2019).

Overall weight of evidence: Moderate

The biological plausibility surrounding the connection between deposition of energy leading to endothelial dysfunction is well-supported by reviews in the literature and mechanistic understanding. The impact on endothelial dysfunction from deposited energy onto cells may vary with the radiation source and associated parameters of dose, dose rate, and type, which can influence the amount of energy absorbed, among other factors such as tissue type.    Radiation types such as gamma rays, X-rays, and charged particles at doses ranging from 0.05-18 Gy and dose rates as low as 2.4 mGy/h induce endothelial dysfunction through an increase in cellular markers of apoptosis and cellular senescence in human cell and animal models as well as diminished relaxation response of vessels in animal models (Yentrepalli et al., 2013a; Yentrepalli et al., 2013b; Soucy et al., 2011; On et al., 2001; Hatoum et al., 2006; Soucy et al., 2010; Soloviev et al., 2003; Baselet et al., 2017; Shen et al., 2017). Following irradiation, endothelial cells may lose their integrity and become senescent or apoptotic via alterations to signaling pathways related to cell survival, leading to endothelial dysfunction (Deanfield et al., 2007; Bonetti et al., 2003). Senescent endothelial cells show changes in cell morphology, cell-cycle arrest, and increased senescence-associated β-galactosidase (SA-β-gal) staining. They also have a pro-inflammatory secretory phenotype, which further contributes to negative effects. These changes lead to endothelial dysfunction, which results in dysregulation of vasodilation (Wang et al., 2016; Hughson et al., 2018; Ramadan et al., 2021). Prolonged chronic inflammation following irradiation causes an ineffective healing process, further worsened by a decrease in endothelium-dependent relaxation. This leads to endothelial dysfunction, making the vasculature more vulnerable to damage from non-laminar flow (Sylvester et al., 2018). Since the endothelium is largely responsible for controlling fluid flow, dysfunctions in the endothelium can lead to fluid imbalance, blood pressure changes, and blood clot formation (Konukoglu & Uzun, 2017; Korpela & Liu, 2014; Verma et al., 2003).

The empirical evidence supporting this KER is gathered from research utilizing both in vivo and in vitro models. Many in vitro studies have examined this relationship using human endothelial cell cultures, such as telomerase-immortalized coronary artery endothelial cells (TICAE) and human umbilical vein endothelial cells (HUVECs) (Baselet et al., 2017; Yentrapalli et al., 2013b). In vivo studies analyzed changes in murine aorta, white rabbit thoracic aorta, and rat aorta and microvessels (Hatoum et al., 2006; Shen et al., 2018; Soloviev et al., 2003; Soucy et al., 2010; Soucy et al., 2011). The evidence includes use of gamma, X-ray, and heavy ion radiation in the dose range of 0.05-18 Gy. SA-β-gal, a marker for cellular senescence, and therefore endothelial dysfunction, relaxation in response to acetylcholine (ACh) and apoptosis were examined in these studies (Baselet et al., 2017; Hatoum et al., 2006; Shen et al., 2018; Soloviev et al., 2003; Soucy et al., 2010; Soucy et al., 2011; Yentrapalli et al., 2013).    Dose Concordance  Studies using in vitro and in vivo models have shown that deposition of energy as produced by acute and chronic doses of ionizing radiation administered from 0.05-18 Gy, with dose rates ranging from 2.4 mGy/h to 2.43 Gy/min, cause endothelial dysfunction as indicated by cellular markers such as cellular senescence and apoptosis as well as decreased maximum relaxation response of vessels in response to ACh.   For example, chronic gamma irradiation of human endothelial cells at a dose rate as low as 2.4 mGy/h led to an increase of 2-fold in SA-β-gal staining (Yentrapalli et al., 2013a). A similar study showed that 4.1 mGy/h chronic gamma irradiation caused no significant changes after 0.69 Gy, but there was a 2-fold increase in SA-β-gal at 2.07 Gy and a 3-fold increase after 4.13 Gy (Yentrapalli et al., 2013b). Another study also looking at SA-β-gal found a 1.7-fold maximum increase in SA-β-gal after irradiating human endothelial cells with 0.05-2 Gy X-rays (Baselet et al., 2017). A study measuring radiation-induced apoptosis in mouse aorta found a 5-fold increase in the number of apoptotic cells after 18 Gy X-ray irradiation (Shen et al., 2018).     Many studies have measured endothelial dysfunction through the relaxation response of vessels in response to ACh. After rabbit thoracic aortas were irradiated with 6 Gy gamma rays, there was a 0.5-fold decrease in maximum relaxation response to ACh, with a linear decrease in relaxation from 0, 1, 2, and 4 Gy gamma rays (Soloviev et al., 2003). A study that irradiated rat aorta with 0.5 and 1 Gy 56Fe ions found a 0.8-fold decrease in maximum relaxation response to ACh (Soucy et al., 2011). Microvessels from rat intestines irradiated with 2250 cGy of fractionated X-rays showed an ACh-induced maximum dilation of 3%, while controls showed a maximum dilation of 87%. A significant decrease was seen after only three doses of 250 cGy (Hatoum et al., 2006). Gamma ray irradiation of rat aorta at 5 Gy showed a 0.6-fold decrease in relaxation response to ACh (Soucy et al., 2010). Similar results were found in a study using 10 Gy gamma rays on rat aortas, which showed a 9% maximum relaxation response to ACh compared to the 48% maximum relaxation in the control group (On et al., 2001).    Time Concordance  There is moderate evidence to suggest a time concordance between the deposition of energy and endothelial dysfunction. A chronic gamma irradiation study at a dose rate of 2.4 mGy/h examined human endothelial cells in vitro after 1, 6, 10 and 12 weeks and showed an increase in SA-β-gal as early as 10 weeks, with levels remaining significantly increased at 12 weeks (Yentrapalli et al., 2013a). A study by the same group also looked at the effects of 4.1 mGy/h gamma rays on human endothelial cells in vitro, but at 1, 3 and 6 weeks of chronic irradiation, revealing an increase in cellular senescence, indicated by increased SA-β-gal, as soon as 3 weeks, with levels remaining significantly increased at 6 weeks (Yentrepalli et al., 2013b). After mouse aorta were irradiated with 18 Gy X-rays, the number of apoptotic cells were determined over 84 days. The number of apoptotic cells was significantly higher than the controls after 3, 7, 14, 28, and 84 days, but was highest (5-fold higher than control) after 7 days followed by a linear decrease (Shen et al., 2018).     Similarly, rabbit thoracic aortas irradiated with 6 Gy gamma rays and exposed to ACh showed a 0.5-fold decrease in maximum relaxation after both 9 and 30 days (Soloviev et al., 2003). When rats were irradiated with 1 Gy 56Fe ions, ACh-induced relaxation in the aorta decreased 0.25-fold compared to controls 4 months after irradiation, and went from a 67.5% relaxation response in the control group to 59% in the irradiated group after 6 months (although this change was not statistically significant). Relaxation response to ACh remained non-significant at 8 months post-irradiation (Soucy et al., 2011).    Essentiality  Endothelial dysfunction can be triggered in response to an injury or a stressor. Therefore, with a reduction in stressor severity, there should be less endothelial dysfunction. As deposition of energy is a physical stressor, it cannot be blocked/decreased using chemicals; however, it can be shielded, though currently no available data used shielding of radiation and measured the impact on endothelial dysfunction. Since deposited energy initiates events immediately, the removal of deposited energy also supports the essentiality of the key event. Studies that do not deposit energy are observed to have no downstream effects.

Much evidence for this relationship comes from high dose studies (>2 Gy); further work is needed at varying doses and dose rates to better understand the relationship.

Examples of quantitative understanding of the relationship are shown in the table below. All data represented is statistically significant unless otherwise indicated.

Dose Concordance   <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Baselet et al., 2017    </td> <td> In vitro. X-ray radiation was delivered to human endothelial cells at a dose rate of 0.5 Gy/min for total doses of 0.05, 0.1, 0.5 and 2 Gy. SA-β-gal activity was used as a marker for senescence and endothelial dysfunction and was measured 14 days post exposure.   </td> <td> SA-β-gal activity for all radiation doses was significantly elevated above the non-irradiated control and increased with an increase in radiation dose. At the highest dose of 2 Gy, there was a 1.7-fold increase compared to control.   </td> </tr> <tr> <td> Soucy et al., 2011    </td> <td> In vivo. 3-4 months-old rats were whole body irradiated with 0.5 or 1 Gy of 56Fe ions before their aortas were harvested and the endothelium dependent vasodilation response to ACh was evaluated at 4 months post-irradiation.     </td> <td> A 0.5 Gy dose did not show significant changes to maximum relaxation response to ACh compared to non-irradiated control.   Following a 1 Gy dose there was a 0.8-fold decrease in maximum relaxation response to ACh compared to non-irradiated control.   </td> </tr> <tr> <td> Soucy et al., 2010     </td> <td> In vivo. 4-months-old rats were irradiated with 5 Gy of 137Cs, and the endothelium dependent vasodilation response to ACh of harvested aortas was evaluated.   </td> <td> There was a 0.6-fold decrease in maximum relaxation response to ACh in the aorta compared to the non-irradiated control.   </td> </tr> <tr> <td> Soloviev et al., 2003   </td> <td> In vivo. The endothelium dependent vasorelaxation response to ACh of aortas from rabbits exposed to 6 Gy of 60Co whole body irradiation was evaluated 9 days post exposure. Furthermore, endothelium dependent relaxation response following exposure to 1, 2, and 4 Gy on the 7th day post exposure were also evaluated.     </td> <td> 9 days after exposure to 6 Gy, there was a 0.5-fold decrease in maximum relaxation response to ACh.     At 7 days post irradiation, maximum relaxation response to ACh decreased with an increase in radiation dose, with 60% maximum relaxation at 0 Gy dropping down to 30% after 4 Gy.   </td> </tr> <tr> <td> Shen et al., 2018   </td> <td> In vivo. 18 Gy of X-ray radiation was delivered to 8-week-old mice. Apoptosis was evaluated using TUNEL assays at 3-, 7-, 14-, 28- and 84-days post irradiation.   </td> <td> Apoptosis levels in 18 Gy irradiated groups were significantly elevated above sham irradiated control at all time points tested. The difference peaked 7-days post irradiation at a 5-fold increase compared to control.   </td> </tr> <tr> <td> Hatoum et al., 2006     </td> <td> In vivo. Rats were whole body irradiated with up to 2250 cGy via 9 fractions of 250 cGy X-rays at a dose rate of 243 cGy/min. Endothelium dependent vasodilation response to ACh of harvested submucosal vessels was evaluated at various radiation doses. </td> <td> After the final dose (total 22.5 Gy) there was a 0.03-fold decrease in maximum relaxation response to ACh in irradiated rat microvessels compared to non-irradiated controls.   Following dose 1 and 2 (250 cGy and 500 cGy total dose), maximum dilation remained similar to non-irradiated control (~90% maximum dilation). However, following the third dose (750 cGy total), maximum dilation dropped below 10% and remained significantly below non-irradiated control levels for all remaining doses tested.   </td> </tr> </tbody> </table>

Time Concordance <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Yentrapalli et al., 2013a    </td> <td> In vitro. Chronic gamma irradiation (137Cs) was delivered to human umbilical vein endothelial cells (HUVECs) at a dose rate of 1.4 mGy/h or 2.4 mGy/h. SA-β-gal activity was used as a marker for premature endothelial cell senescence and was evaluated at 1-, 3-, 6-, 10- and 12-weeks post irradiation.   </td> <td> Between 1 to 6 weeks post irradiation no significant differences were observed between either of the irradiated groups and the sham irradiated control.   At the 10- and 12-week time points, the 1.4 mGy/h exposure continued to show no significant changes from control, while the 2.4 mGy/h group showed a 1.7-fold increase at 10-weeks and 1.9-fold increase at 12-weeks.    </td> </tr> <tr> <td> Yentrapalli et al., 2013b    </td> <td> In vitro. Chronic gamma (137Cs) of HUVECs at a dose rate of 4.1 mGy/h for up to 6 weeks for final doses of 0.69, 2.07 and 4.13 Gy. SA-β-gal activity was used as a marker for premature endothelial cell senescence and was evaluated at 1-, 3- and 6-weeks post exposure.   </td> <td> No significant changes in SA-β-gal activity were observed between irradiated and sham irradiated groups in the first week.   SA-β-gal activity was significantly elevated in irradiated HUVECs at the 3- and 6-week timepoints, showing a 2-fold and 3-fold elevation above control respectively.    </td> </tr> <tr> <td> Soucy et al., 2011    </td> <td> In vivo. 3-4 months-old rats were whole body irradiated with 0.5 or 1 Gy of 56Fe ions before their aortas were harvested and the endothelium dependent vasodilation response to ACh was evaluated.   </td> <td> At 4 months post radiation there was a 0.8-fold decrease in maximum relaxation response to ACh with a return to control levels by 6 months.  </td> </tr> <tr> <td> Soloviev et al., 2003    </td> <td> In vivo. The maximum endothelium dependent vasorelaxation response to ACh of aortas from rabbits having been whole body irradiated to 6 Gy 60Co gamma-rays was evaluated 9- and 30-days post exposure.  </td> <td> At both 9- and 30-days post-irradiation there was a ~0.5-fold decrease in maximum relaxation response to ACh compared to non-irradiated control. There was no significant difference in maximum relaxation between the 9- and 30-day timepoints.   </td> </tr> <tr> <td> Shen et al., 2018   </td> <td> In vivo. 18 Gy of X-ray radiation was delivered to 8-week-old mice with apoptosis levels being measured for up to 84 days post-irradiation in the aorta.  </td> <td> There was a significant increase of 3-fold in apoptosis as soon as 3 days post-irradiation with a peak of 7-fold after 7 days. There was a gradual return to a 3-fold increase by 84 days.  </td> </tr> </tbody> </table>

Moderate Male Moderate Female Low Unspecific

Moderate

Adult

Low

Juvenile

Moderate

Not Otherwise Specified

Low Moderate Moderate Moderate

The evidence for the taxonomic applicability to humans is low as the majority of the evidence is from in vitro human-derived cells. The relationship is supported by both sexes of mouse, rat, and rabbit models. The in vivo studies were mostly undertaken in adolescent or adult rats and mice. In addition, the relationship is likely at any life stage.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b42eaca8750> AOPWiki 2022-09-28T12:44:53

2023-03-21T10:37:26

<upstream-id>1883f503-acf6-4e69-8704-91f462c360dc</upstream-id> <downstream-id>1cc566f9-d2e5-4c9c-a06c-c40f7767c251</downstream-id> Multiple signaling pathways can regulate nitric oxide (NO) levels. The phosphatidylinositol 3-kinase (PI3K)/Akt pathway can activate nitric oxide synthase (NOS), an enzyme that produces NO, through phosphorylation (Hemmings & Restuccia 2012; Nagane et al., 2021). The RhoA/Rho kinase (ROCK) pathway inhibits both the expression and phosphorylation of NOS (Yao et al., 2010). Furthermore, the renin-angiotensin-aldosterone system (RAAS) can both inhibit NOS to reduce vasodilation and activate NOS as a countermeasure for vasoconstriction (Millatt, Abdel-Rahman & Siragy, 1999). The extracellular signal-regulated protein kinase 5 (ERK5)/kruppel-like factor 2 (KLF2) pathway can increase transcription of endothelial NOS (eNOS), which results in increased NO levels (Paudel, Fusi & Schmidt, 2021). Lastly, the acidic sphingomyelinase/ceramide pathway can activate NADPH oxidase (NOX) production of reactive oxygen species (ROS) that react with NO, resulting in lower NO levels (Soloviev & Kizub, 2019). Alterations in these pathways will result in altered NO levels.

Overall weight of evidence: Moderate

The biological plausibility surrounding the connection between altered signaling pathways and altered NO levels is well-supported by the literature. Studies have shown that altered signaling pathways lead to altered NO levels (Azimzadeh et al., 2015; Azimzadeh et al., 2017; Hasan, Radwan & Galal, 2019; Shi et al., 2012; Siamwala et al., 2010).  Multiple signaling pathways can influence NO levels. Under normal physiological conditions, the PI3K/Akt pathway is regulated by various growth factors and other signaling molecules to modulate eNOS phosphorylation and therefore NO production (Hemmings & Restuccia, 2012). Phosphorylation of eNOS can affect its function. Phosphorylation at Ser1177 activates the enzyme, while phosphorylation at Thr495 acts in reverse and decreases enzymatic activity instead (Forstermann, 2010; Nagane et al., 2021). In endothelial cells, Thr495 is constitutively phosphorylated by kinases like protein kinase C (PKC) (Forstermann, 2010). Following Akt activation, eNOS is phosphorylated on Ser1177 to activate the enzyme and thus NO production is upregulated (Karar & Maity, 2011). Although peroxisome proliferator-activated receptor α (PPARα) can increase eNOS directly through transcription, it can also increase vascular endothelial growth factor (VEGF) (Du, Wagner & Wagner, 2020), which can activate the PI3K pathway to activate eNOS (Hicklin & Ellis, 2005). Alternatively, activation of the RhoA/ROCK pathway leads to a decrease in NO bioavailability (Yao et al., 2010). The RhoA/ROCK pathway causes eNOS mRNA destabilization and prevents Ser1177 eNOS phosphorylation by Akt (Yao et al., 2010). However, phosphorylation of RhoGDI, a regulator in the RhoA/ROCK pathway, causes increased affinity to GTP-RhoA, sequestering the active form of RhoA and preventing eNOS inhibition (Dovas & Couchman, 2005). The RAAS pathway results in the production of angiotensin II (AngII) which can cause various downstream effects on vascular homeostasis, including inducing vasoconstriction (Millatt, Abdel-Rahman & Siragy, 1999). AngII can downregulate eNOS Ser1177 phosphorylation to prevent vasodilation (Ding et al., 2020; Millatt, Abdel-Rahman & Siragy, 1999), or activate eNOS as a corrective measure (Millatt, Abdel-Rahman & Siragy, 1999). As a result, depending on the mechanism, NO levels can either increase or decrease after AngII stimulation. The ERK5/KLF2 pathway results in activation of the KLF2 transcription factor, which increases transcription of eNOS (Paudel, Fusi & Schmidt, 2021).

The empirical evidence to support this KER is provided by in vivo mouse and rat models and in vitro models of human coronary artery endothelial cell (HCAECs), human umbilical vein endothelial cells (HUVECs), and EA.hy96 cells (a hybrid of HUVECs and A549 cells). The effects of altered signaling pathways on the levels of NO as well as inducible NOS (iNOS) and eNOS have been investigated. These studies examined levels of signaling molecules in insulin-dependent PI3K/Akt pathway such as IGFR1, PPARα, PI3K, Akt, and the phosphorylated versions of each (Azimzadeh et al., 2015; Azimzadeh et al., 2021; Shi et al., 2012), RAAS pathway indicator, AngII (Hasan, Radwan & Galal, 2019), phosphorylated Rho GDP-dissociation inhibitor (p-RhoGDI) in the RhoA/ROCK pathway (Azimzadeh et al., 2017), ERK5 and KLF2 (Sadhukhan et al., 2020), and the effect they have on NO. The studies used stressors such as X-rays (Azimzadeh et al., 2015; Azimzadeh et al., 2017; Azimzadeh et al., 2021), gamma rays (Hasan, Radwan & Galal, 2019; Sadhukhan et al., 2020) and altered gravity (Shi et al., 2012; Siamwala et al., 2010).    Dose Concordance  There is moderate evidence to demonstrate dose concordance between altered signaling pathways leading to altered NO levels. Activated RhoA reduces the activity and abundance of eNOS (Yao et al., 2010). HCAECs irradiated with 0.5 Gy demonstrated a 0.7-fold decrease in p-RhoGDI, a 0.6-fold decrease in p-eNOS and a 0.8-fold decrease in cellular NO, 7 days after exposure (Azimzadeh et al., 2017). The decrease in p-RhoGDI, involved in the RhoA/ROCK pathway, was correlated with a decrease in p-eNOS and NO (Azimzadeh et al., 2017). HSP90, a protein that binds to and activates eNOS (Karar & Maity, 2011), decreased 0.8-fold following 0.5 Gy X-ray irradiation (Azimzadeh et al., 2017).  High doses (>2 Gy) also show dose concordance between altered signaling pathways and altered NO levels. X-ray irradiation of BAECs and HUVECs caused a 5.6-fold increase in p-Akt/Akt after 6 Gy (Sonveaux et al., 2003). eNOS increased 3-fold after 6 Gy, while p-eNOS increased 1.2-fold after 2 Gy and 1.7-fold after 6 Gy (Sonveaux et al., 2003). Although the p-Akt/Akt ratio was not significantly increased after 2 Gy, p-Akt alone had increased, indicating changes in the PI3K/Akt pathway had occurred at 2 Gy. Therefore, a decrease in PI3K/Akt pathway proteins, such as p-IGFR1, p-Akt and p-PI3K are correlated with decreases in p-eNOS and NO (Azimzadeh et al., 2015), while an increase in p-Akt correlates with increases in eNOS and p-eNOS (Shi et al., 2012; Sonveaux et al., 2003). In rat blood serum, 6 Gy gamma ray irradiation led to a 1.4-fold increase in AngII and aldosterone levels and a 3.3-fold increase in iNOS expression indicating that a change in the RAAS pathway can lead to altered NO levels (Hasan, Radwan & Galal, 2019).  Mice that received high dose (8 or 16 Gy) X-ray irradiation on the heart showed alterations in the levels of signalling proteins involved in the PI3K/Akt pathway, including a 0.4 and 0.3-fold decrease of p-IGFR1, a 0.5-fold decrease of PI3K, a 0.2 and 0.1-fold decrease of p-Akt and a 0.6 and 0.2-fold decrease of p-eNOS at 8 Gy and 16 Gy, respectively (Azimzadeh et al., 2015). Concurrently to changes to PI3K/Akt pathway signaling molecules, the level of serum NO decreased 0.3-fold and 0.3-fold following 8 Gy and 16 Gy X-ray irradiation, respectively (Azimzadeh et al., 2015). As well, following X-ray irradiation, the ERK/MAPK pathway decreased 0.5-fold at 16 Gy and the p38/MAPK pathway increased 1.3-fold at 16 Gy (Azimzadeh et al., 2015). X-ray irradiation in mice at 16 Gy also inhibited PPARα resulting in decreased activity and phosphorylation of eNOS through the PI3K/Akt pathway and decreased NO levels (Azimzadeh et al., 2021). This study also showed an increase in p-ERK, and p-p38 at 16 Gy (Azimzadeh et al., 2021). After irradiation of HUVECs with 10 or 12 Gy, p-ERK5, KLF2 and eNOS were all found to decrease after fractionated doses, while KLF2 and eNOS both decreased after acute doses (Sadhukhan et al., 2020).  Microgravity in HUVEC-C was simulated by a clinostat using rotation at a slow speed to negate centrifugal force. Following clinorotation, levels of eNOS and p-eNOS expression were significantly increased by 5.2-fold and 5.5-fold, respectively. As well, p-Akt increased by 2.9-fold after exposure to simulated microgravity in HUVEC-C (Shi et al., 2012).    Time Concordance  There is some evidence to demonstrate time concordance between altered signaling pathways leading to altered NO levels. After 0.5 Gy X-ray of HCAECs, p-Rho-GDI significantly decreased after 1 day, while NO was not significantly lower after 1 day (Azimzadeh et al., 2017). After 24 h, p-Akt was increased, while eNOS was increased 24 and 48 h after X-ray irradiation of BAECs and HUVECs (Sonveaux et al., 2003). After 7 days, p-Rho-GDI decreased further and NO showed a significant decrease. HUVECs irradiated with 10 or 12 Gy acute or fractionated X-rays showed decreased levels of p-ERK5, KLF2 and eNOS at 4 h, and decreased KLF2 and eNOS at 24 h after irradiation (Sadhukhan et al., 2020).    Incidence Concordance  The evidence of incidence concordance for this relationship is moderate, as a few studies demonstrated incidence concordance. In mice hearts irradiated with X-rays, p-Akt, the direct upstream activator of eNOS, decreased 0.2-fold after 8 Gy and 0.1-fold after 16 Gy, while NO decreased 0.3-fold after 8 Gy and 0.2-fold after 16 Gy (Azimzadeh et al., 2015). Similarly, after 0.5 Gy X-ray irradiation of HCAECs, p-RhoGDI decreased 0.7-fold while NO decreased 0.8-fold (Azimzadeh et al., 2017). X-ray irradiation of BAECs and HUVECs at 6 Gy resulted in a 5.6-fold increase in the ratio of p-Akt/Akt and a 3-fold increase to p-eNOS (Sonveaux et al., 2003).     Essentiality  Several studies have investigated the essentiality of various signalling pathways in altering NO levels. Under normal conditions, the phosphorylation of eNOS by the PI3K/Akt pathway activates NOS, resulting in NO production. LY294002 treatment, a PI3K inhibitor, led to significant decreases in eNOS and p-eNOS levels in HUVEC-C samples (Shi et al., 2012). After exposure to simulated microgravity by clinorotation, LY294002 treatment led to decreased p-Akt levels to below control levels (Shi et al., 2012). The inhibition of the PI3K/Akt pathway with wortmanin, another PI3K inhibitor, also led to a 0.3-fold decrease in NO production (Siamwala et al., 2010). Irradiation inhibited PPARα and eNOS, while treatment with fenofibrate, a PPARα activator, kept both p-PPARα and NO at control levels (Azimzadeh et al., 2021).  Bradykinin-potentiating factor (BPF) is implicated in muscle contraction, inflammatory responses and angiotensin-converting enzyme (ACE) inhibition. ACE in endothelial cells converts AngI to AngII. Irradiation at 6 Gy led to increased iNOS, AngII, and aldosterone, while the subsequent treatment with BPF decreased all endpoints measured following irradiation, including the levels of iNOS, AngII, and aldosterone. The recovery of iNOS serum levels in 6 Gy gamma irradiated rats following BPF treatment indicates that the signaling pathways play a role in NO levels (Hasan, Radwan & Galal, 2019). In addition, bradykinin signaling through bradykinin receptor 2 (B2R) activates eNOS (Ancion et al., 2019). Nitrite concentration was found to increase after both microgravity and treatment with bradykinin (Siamwala et al., 2010).  Various inhibitors of the mevalonate pathway were used to recover KLF2 levels after irradiation, which resulted in increased KLF2 and eNOS levels (Sadhukhan et al., 2020).

<ul> <li>Due to the high reactivity of NO, it can be difficult to obtain its direct measures (Luiking, Engelen & Deutz, 2010). The inconsistencies in NO levels may be attributed to the challenges in measuring NO. Directionality of NO changes cannot be compared between studies due to a variety of experimental conditions like stressor type, dose, dose rate, model and time course of the experiment.</li> </ul>

<div> <table cellspacing="0" class="MsoTableGrid" style="border-collapse:collapse; border:none"> <tbody> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:1px solid black; height:26px; vertical-align:top; width:156px"> Modulating factor </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; height:26px; vertical-align:top; width:156px"> Details </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; height:26px; vertical-align:top; width:156px"> Effects on the KER </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; height:26px; vertical-align:top; width:156px"> References </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Drug </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> BPF (ACE inhibitor) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> BPF treatment led to decreased iNOS, angiotensin II and aldosterone following irradiation. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Hasan, Radwan & Galal, 2019 </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Drug </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> LY294002 (PI3K inhibitor) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> LY294002 treatment led to reduced phosphorylation of both Akt and eNOS. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Shi et al., 2012; </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Drug </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Wortmanin (PI3K inhibitor) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Wortmanin decreased NO production. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Siamwala et al., 2010 </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Drug </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Fenofibrate (PPARα activator) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Treatment with PPARα prevented the decrease in eNOS levels after irradiation. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Azimzadeh et al., 2021 </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Drug </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Atorvastatin (HMG-CoA reductase inhibitor) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Treatment with atorvastatin increased KLF2 and eNOS levels slightly. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Sadhukhan et al., 2020  </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Drug </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Gamma tocotrienol (HMG-CoA reductase inhibitor) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Treatment with gamma tocotrienol prevented the radiation-induced decrease in KLF2 and eNOS levels. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Sadhukhan et al., 2020  </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Drug </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Geranylgeranyltransferase I inhibitor 298 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Treatment with Geranylgeranyltransferase I inhibitor 298 prevented the radiation-induced decrease in KLF2 and eNOS levels. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Sadhukhan et al., 2020  </td> </tr> </tbody> </table> </div>

The following are a few examples of quantitative understanding of the relationship. All data that is represented is statistically significant unless otherwise indicated.

Not identified

Low Male Low Female Moderate Unspecific

Low

Adult

Moderate

Not Otherwise Specified

Low Moderate Moderate

The majority of the evidence for this KER is from rat and mouse models. Most evidence regarding sex and lifestage is unspecified with a small amount of evidence from adult models.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b42eae207b8> AOPWiki 2022-09-28T12:41:39

2023-03-21T11:44:55

<upstream-id>8d6a4475-5817-4386-be53-eb911186774f</upstream-id> <downstream-id>c79fec82-3db8-4eaf-b5ef-0fdcffb5dfe8</downstream-id> Deposition of energy can trigger vascular remodeling through many pathways (Tapio, 2016) including changes to vessel structure and blood flow (Patel, 2020; Sylvester et al., 2018). Pro-inflammatory mediators can be increased, which can result in a low level of inflammation causing intimal thickening (Sylvester et al., 2018). Deposition of energy can generate reactive oxygen species (ROS) and highly reactive radicals sparsely from low- linear energy transfer (LET) radiation and densely from high-LET radiation, which can cause endothelial dysfunction and subsequent vascular remodeling (Boerma et al., 2015; Hughson, Helm & Durante, 2017; Slezak et al., 2017; Soloviev & Kizub, 2019; Sylvester et al., 2018). Increased production of ROS changes the bioavailability of nitric oxide (NO), a diffusible molecule responsible for vasodilation, which leads to inhibited vasomotion and cellular senescence as components of endothelial dysfunction (Patel, 2020; Soloviev & Kizub, 2019). Changes in the expression or activity of proteins in many signaling pathways can lead to endothelial dysfunction (Schmidt-Ullrich et al., 2000; Tapio, 2016). In addition, the increased pro-inflammatory mediators can lead to endothelial dysfunction and therefore, vascular remodeling (Tapio, 2016). Another possible vascular remodeling change is age accelerated atherosclerosis (EPRI, 2020; Hamada et al., 2014). Studies using varying LET, delivered at acute and chronic dose-rates, have shown remodeling of the vasculature (reviewed in Tapio, 2016).

Overall weight of evidence: High

The biological plausibility suggesting that deposition of energy leads to vascular remodeling is well-supported by reviews and mechanistic studies published in the literature. Vascular remodeling may occur due to aging and diet (Zieman, Melenovsky & Kass, 2005). However, the deposition of energy from ionizing radiation (IR) can accelerate vascular remodeling in the form of accelerated atherosclerosis (Boerma et al., 2015; Boerma et al., 2016; EPRI, 2020; Hamada et al., 2014; Hughson, Helm & Durante, 2017; Mitchell et al., 2019; Sylvester et al., 2018), which can be demonstrated by arterial thickening or the amount of oxidized low-density lipoprotein (oxLDL) (Poznyak et al., 2021). Remodeling normally allows adaptation to long-term hemodynamic changes but can also contribute to vascular diseases (Gibbons & Dzau, 1994). Various short-term post-spaceflight studies have shown vascular remodeling after deposition of energy from space IR (Patel, 2020).     Under physiological conditions, the body maintains a balance of ROS and NO levels. IR generates ROS that can react with NO and reduce its bioavailability, causing endothelial dysfunction and vascular stiffness (Patel, 2020). Similarly, signaling pathways can cause vascular remodeling through endothelial dysfunction and altered NO (Tapio, 2016). Increased ROS or altered signaling can cause a prolonged inflammatory response; this has been observed in animal models exposed to high-LET radiation (Hughson, Helm & Durante, 2017; Sylvester et al., 2018; Tapio, 2016). The low level of inflammation results in intimal thickening and inhibits tissue and vessel recovery (Sylvester et al., 2018). Microvascular injury and inflammation may cause angiogenesis, which prevents vascular resistance (Slezak et al., 2017). However, depending on the source, radiation may have different effects on angiogenesis (Grabham & Sharma, 2013). An increase in pro-angiogenic factors, such as vascular endothelial growth factor (VEGF), secreted as a consequence of photon irradiation can promote angiogenesis. Exposure to low-LET protons and high-LET heavy ion radiation can disturb angiogenesis due to decreased VEGF secretion and tubule formation (Grabham & Sharma, 2013; Sylvester et al., 2018). Matrix metalloproteinases (MPPs) are involved in remodeling of the extracellular matrix (ECM) and can affect various pathological processes after irradiation (Slezak et al., 2017). Following radiation, existing collagen in the heart may be remodeled, which indicates ECM remodeling (Shen et al., 2018; Sridharan et al., 2020; Zieman, Melenovsky & Kass, 2005). Thus, many mechanisms exist via which the deposition of energy can lead to vascular remodeling; these are generally well understood and described in the literature, leading to a strong weight of evidence for the biological plausibility of this KER.

There is moderate empirical evidence supporting the connection between deposition of energy leading to vascular remodeling. The evidence was gathered from studies using in vivo rat and mouse models, as well as in vitro models of human vessels and ex vivo human vessel biopsies (Grabham et al., 2011; Hamada et al., 2020; Hamada et al., 2021; Hamada et al., 2022; Russel et al., 2009; Sarkozy et al., 2019; Shen et al., 2018; Soucy et al., 2007; Soucy et al., 2010; Soucy et al., 2011; Sridharan et al., 2020; Yu et al., 2011). The models used stressors such as iron ions (56Fe) (Soucy et al., 2011; Yu et al., 2011), gamma rays (Hamada et al., 2020; Hamada et al., 2021; Hamada et al., 2022; Soucy et al., 2007; Soucy et al., 2010), X-rays (Hamada et al., 2021; Hamada et al., 2022; Shen et al., 2018), electrons (Sarkozy et al., 2019), protons and oxygen ions (16O) (Sridharan et al., 2020). Vascular stiffness measured by pulse wave velocity (PWV) and vascular composition (Hamada et al., 2020; Hamada et al., 2021; Hamada et al., 2022; Shen et al., 2018; Soucy et al., 2007; Soucy et al., 2010; Soucy et al., 2011; Sridharan et al., 2020), vessel thickness/diameter measured directly (Sarkozy et al., 2019; Shen et al., 2018; Soucy et al., 2011; Sridharan et al., 2020; Yu et al., 2011) and other morphological changes (Hamada et al., 2020; Hamada et al., 2021; Hamada et al., 2022) were used as endpoints for vascular remodeling.     Dose Concordance  PWV is often measured to determine vascular stiffness and therefore remodeling, where PWV is quadratically proportional to Young's modulus (measure of a material’s stiffness) (Pereira, Correia & Cardoso, 2015; Soucy et al., 2011). The PWV of irradiated rats increased with an increase in radiation dose. Across several studies, 0.5, 1, 1.6, and 5 Gy doses were tested, with responses of 1.1 to 1.2-fold changes to PWV (Soucy et al., 2007, 2010, 2011). PWV is also quadratically proportional to the vessel diameter and inversely quadratically proportional to the vessel wall thickness. However, Soucy et al. (2011) found no significant change in the aortic wall thickness:lumen diameter ratio, indicating the change in PWV was due to changes in vessel composition and not geometric remodeling. Collagen composition in the membrane was also measured to determine vascular composition and therefore remodeling, with increased collagen indicating stiffness (Zieman, Melenovsky & Kass, 2005). Collagen accumulation increased 1.4-fold after 18 Gy X-ray irradiation (Shen et al., 2018) and 1.5-fold after various regiments of 5 Gy X-rays (Hamada et al., 2021; Hamada et al., 2022) in mice. Following 12 months of 16O exposure, the tissue content of collagen type III peptide increased 2.3-fold and 2-fold at 0.05 Gy and 0.25 Gy, respectively (Sridharan et al, 2020). VE-cadherin, a marker for adherens junctions, decreased 0.2- to 0.3-fold in mice after various regimens of 5 Gy gamma and X-rays, with a high decrease in the acute and fractionated doses and no change after chronic gamma rays (Hamada et al., 2022; Hamada et al., 2021; Hamada et al., 2020). oxLDL increased in mice after X-ray irradiation at both 8 and 16 Gy; however, the increase was not greater at 16 Gy relative to 8 Gy (Azimzadeh et al., 2015).    Vessel thickness was also measured in various studies. Aortic thickness increased 1.4-fold after 18 Gy X-ray irradiation of mice (Shen et al., 2018). Iron ion irradiation on apolipoprotein E (apoE)-deficient mice, a model for atherosclerosis, at both 2 and 5 Gy showed a 1.4-fold increase in carotid artery intima thickness compared to sham-irradiated apoE-deficient mice (Yu et al., 2011). Markers of vascular remodeling, anterior and inferior wall thicknesses in systole (AWTs and IWTs) and diastole (AWTd and IWTd) were measured after a high dose of 50 Gy electrons. AWTs increased 1.3-fold, IWTs increased 1.1-fold, AWTd increased 1.5-fold and IWTd increased 1.2-fold (Sarkozy et al., 2019). In turn, patients having undergone radiation treatment for head and neck or breast cancer with radiation doses totaling 66 Gy and 49 Gy, respectively, observed increased intima-media ratios (IMR) of 1.5- and 1.4-fold, respectively (Russel et al., 2009). Mice that did not show endothelial detachments before irradiation showed high frequencies of detachments after acute doses of 5 Gy gamma and X-rays (Hamada et al., 2020; Hamada et al., 2021). Vascular permeability was also significantly increased after 5 Gy gamma rays in mice (Hamada et al., 2020).     Radiation type can affect the nature of vascular remodeling following exposure. A study using 3D vessel models of endothelial cells in a gel matrix showed that high-LET iron-ions reduces vessel length in both mature and developing vessels after only 0.8 Gy. In contrast, low-LET protons only inhibited growth in developing vessels at 0.8 Gy and required a dose of 3.2 Gy to affect mature vessels. Gamma radiation required a dose of 0.8 Gy to inhibit vessel growth and 6.4 Gy for any significant breakdown of mature vessels (Grabham et al., 2011).   Time Concordance  In rats irradiated with 5 Gy gamma rays, PWV as a measure of arterial stiffness increased from 3.9 m/s before irradiation to around 4.5 m/s after 1 day, 1 week and 2 weeks (Soucy et al., 2007). Under the same conditions, PWV increased from 4.1 m/s before irradiation to 4.6 m/s after 1 day, 4.9 m/s after 1 week, and 4.8 after 2 weeks in a separate study by the same authors (Soucy et al., 2010). Using 1 Gy iron ions instead, PWV increased by 0.5 m/s both 4 months and 8 months post-irradiation (Soucy et al., 2011). Mice irradiated with 18 Gy X-rays showed significantly increased aortic thickness 1.4-fold after 7 days and 1.3-fold after 14 and 28 days, while collagen accumulation increased 1.4-fold after 14, 28 and 84 days (Shen et al., 2018). ApoE-deficient mice that were irradiated with both 2 and 5 Gy iron ions showed a 1.4-fold increase in carotid artery intima thickness after 13 weeks compared to controls (Yu et al., 2011). Mice irradiated with 8 and 16 Gy of X-rays showed increased oxLDL levels at 16 weeks post-irradiation (Azimzadeh et al., 2015). Ventricular posterior wall thickness decreased at 3- and 5-months post-irradiation, and PWV increased at 12 months post-irradiation with 16O irradiation (Sridharan et al., 2020). The results suggest that 16O irradiation may lead to long-term vascular dysfunction in rats similar to iron ions in the studies by Soucy et al. (2011) and Yu et al. (2011). Collagen type III increased 2.4-fold after 12 months of 0.5 Gy proton irradiation (Sridharan et al., 2020). CD2, CD4 and CD8 markers for the T-protein lymphocytes that are thought to be involved with promoting hypertension and microvascular remodeling increased in rat hearts 6 months after a 0.1 Gy dose of 16O, and 12 months after a single 0.5 Gy dose of 16O (Sridharan et al., 2020).  Mice irradiated with 5 Gy gamma rays showed increased endothelial detachments, increased vascular permeability and VE-cadherin expression at 1, 3 and 6 months after irradiation (Hamada et al., 2020; Hamada et al., 2021). VE-cadherin was also increased 12 months after irradiation, but endothelial detachments were no longer present (Hamada et al., 2022).    Essentiality   Vascular remodeling such as arterial stiffness occurs naturally with aging, but deposition of energy can accelerate this process (Zieman, Melenovsky & Kass, 2005). Since deposited energy initiates events immediately, the removal of deposited energy also supports the essentiality of the key event. Studies that do not deposit energy are observed to have no downstream effects.

<ul> <li> Not all results show the expected dose-response. For example, total collagen and collagen type III peptide levels studied in Sridharan et al. (2020) did not consistently increase with increasing dose. Similarly, oxLDL levels were higher at the 8 Gy dose compared to the 16 Gy dose (Azimzadeh et al., 2015).  </li> <li> Yu et al. (2011) showed that intimal thickness increased at 13 weeks after iron ion irradiation of apoE-deficient mice. At 40 weeks post-irradiation, intimal thickness remained at similar levels, but the level was no longer statistically significant because the sham-irradiated group showed higher intimal thickness.  </li> </ul>

Dose Concordance  <table border="1"> <tbody> <tr> <td> Reference </td> <td> Experimental Description </td> <td> Result </td> </tr> <tr> <td> Sridharan et al., 2020  </td> <td> In vivo. In study B, male rats were exposed to oxygen ions (0.01-0.25 Gy). Heart tissue analysis was performed 6-7 and 12 months after radiation using histology and western blots. </td> <td> Study B:  At 12 months after 16O exposure, the tissue content of the 75 kDa collagen type III peptide increased 2.3-fold and 2-fold at 0.05 Gy and 0.25 Gy respectively.     </td> </tr> <tr> <td> Soucy et al., 2007  </td> <td> In vivo. Sprague-Dawley rats were irradiated with 137Cs gamma rays (0.5, 1.6, 5 Gy). Vascular stiffness was calculated using PWV with an ECG and doppler probe.  </td> <td> At 0.5 and 1.6 Gy, PWV increased from 3.9 m/s (before irradiation) to 4.2 m/s. At 5 Gy PWV increased to 4.6 m/s.  </td> </tr> <tr> <td> Grabham et al., 2011      </td> <td> In vitro. 3D models of human vessels were created using human endothelial cells in gel matrix. Mature and developing vessel models were exposed to iron-ion (1 GeV/nucleon; LET 151 keV/um) and proton (1 GeV/nucleon; LET 0.22 keV/um) radiation at a 0.1-1 Gy/min dose rate or 137Cs gamma radiation at 85 cGy/min dose rate.      Vessel length as determined by length of capillary with lumen per cell was evaluated by DTAF staining for proteins and propidium iodide for nuclei.   </td> <td> In mature vessels, iron-ion exposure reduced vessel length significant starting after 0.8 Gy with a 44% decrease in length after 1.6 Gy. Proton exposure produced no significant change, while gamma exposure required a dose of 6.4 Gy for significant vessel length breakdown.   In developing vessels iron-ion exposure of 0.8 Gy decreased length by 50%, proton exposure of 0.4 Gy inhibited development and 0.8 Gy decreased length by 60%, while gamma exposure of 0.8 Gy inhibited vessel growth, and 6.4 Gy was required to reduce vessel length.  </td> </tr> <tr> <td> Soucy et al., 2011  </td> <td> In vivo. 56Fe ions were used to irradiate rats at 0.5 or 1 Gy (0.5 Gy/min). PWV measured with Doppler probe and ECG and aortic wall thickness:lumen diameter ratio measured with histological analysis were used to determine vascular remodeling.  </td> <td> After 0.5 Gy, there was no significant change in PWV, but at 1 Gy there was a significant 1.1-fold increase in PWV. No change in aortic wall thickness:lumen diameter was observed after either dose.  </td> </tr> <tr> <td> Yu et al., 2011  </td> <td> In vivo. 56Fe ions were irradiated onto apoE-/- mice, and intima thickness of the carotid artery was measured using hematoxylin and eosin staining at 0, 2 and 5 Gy doses and a 13- and 40-weeks post-irradiation.   </td> <td> 13-weeks post irradiation, both 2 and 5 Gy doses showed a maximum 1.5-fold elevation in thickness compared to controls.   40-weeks post irradiation there was no significant difference between the control, 2 Gy, and 5 Gy groups.  </td> </tr> <tr> <td> Soucy et al., 2010  </td> <td> In vivo. Rats were irradiated with 5 Gy 137Cs gamma radiation and PWV, measured with Doppler probe and ECG was used as a measure of aortic stiffness and vascular remodeling.  </td> <td> After 5 Gy, maximum PWV increased 1.2-fold compared to PWV pre-irradiation.  </td> </tr> <tr> <td> Hamada et al., 2020    </td> <td> In vivo. Male mice were irradiated by 5 Gy 137Cs gamma rays (0.5 Gy/min) and compared to 0 Gy control. Vascular remodeling was measured through a Miles assay to show vascular permeability, vascular endothelial cadherin (VE-cadherin, a marker for adherens junctions) levels and number of endothelial detachments.  </td> <td> Vascular permeability increased to a maximum of 16-fold as shown by staining intensity; VE-cadherin decreased to a maximum of 0.2-fold and percent of mice with total endothelial detachments increased from 0 to 90% at the maximum response.  </td> </tr> <tr> <td> Hamada et al., 2021      </td> <td> In vivo. Either acute or chronic doses of X-rays and 137Cs gamma rays were given to B6J mice, all resulting in a total 5 Gy dose. X-rays were given as a single acute dose, 25 fractions of 0.2 Gy/fraction spread over 42 days or 100 fractions of 0.05 Gy/fraction spread over 153 days all given at 0.5 Gy/min. Gamma rays were given as a single acute dose at 0.5 Gy/min or chronically at <1.4 mGy/h for 153 days. Vascular remodeling was measured by IMT, collagen content (aniline blue staining), VE-cadherin levels and number of mice with detachments 6 months post-irradiation.   </td> <td> IMT in the aorta increased about 2-fold after all X-ray treatments, but chronic gamma rays did not cause a change and acute gamma rays were not measured. Stained intensity to show collagen content increased about 1.5-fold for both the acute and 25 fractions X-ray regimens, but not in others. VE-cadherin decreased after acute gamma rays (0.2-fold) and all X-ray doses (0.2- to 0.3-fold). After acute gamma and X-rays and 25 fraction X-rays the percent of mice with a detachment went from 0 to a maximum of 100%.  </td> </tr> <tr> <td> Hamada et al., 2022  </td> <td> In vivo. Either acute or chronic doses of X-rays and 137Cs gamma rays were given to B6J mice, all resulting in a total 5 Gy dose. X-rays were given as a single acute dose, 25 fractions of 0.2 Gy/fraction spread over 42 days or 100 fractions of 0.05 Gy/fraction spread over 153 days all given at 0.5 Gy/min. Gamma rays were given chronically at <1.4 mGy/h for 153 days. Vascular remodeling was measured by IMT, collagen content (aniline blue staining), VE-cadherin levels and number of mice with detachments 12 months post-irradiation.  </td> <td> IMT in the aorta increased about 1.3-fold after X-rays in acute or 25 fraction regimens, but not after other regimens. Collagen content increased about 1.1-fold for all regimens except X-rays at 100 fractions. VE-cadherin decreased after all X-ray doses (maximum 0.5-fold). No mice were observed to have a detachment.  </td> </tr> <tr> <td> Azimzadeh et al., 2015        </td> <td> In vivo. 10-week-old male C57Bl/6 mice received cardiac irradiation at 8 or 16 Gy with X-rays. Serum oxLDL was measured using ELISA.  </td> <td> oxLDL increased 1.2-fold after 8 Gy and 1.1-fold after 16 Gy compared to 0 Gy control.  </td> </tr> <tr> <td> Shen et al., 2018  </td> <td> In vivo. Male mice irradiated with 18 Gy X-rays had measurements of aortic thickness determined at various times using hematoxylin and eosin staining. The accumulation of collagen was measured using Sirius red staining. Endpoints were evaluated at 3-, 7-, 14-, 28- and 84-days post exposure and compared to a sham-irradiated control group.   </td> <td> Sham-irradiated control:   Thickness and collagen accumulation remained unchanged throughout the timepoints tested.   18 Gy irradiated:   Thickness and collagen accumulation were elevated above control at all time points tested. Thickness peaked 7-days post exposure with a 1.4-fold increase compared to controls.   Collagen accumulation peaked at 14-days post exposure with a 1.4-fold increase above controls.    </td> </tr> <tr> <td> Sarkozy et al., 2019  </td> <td> In vivo. Rats were irradiated with 50 Gy electrons (5 Gy/min). Various markers of vascular remodeling, anterior and inferior wall thicknesses in systole (AWTs and IWTs) and diastole (AWTd and IWTd) were measured with an echocardiograph. Endpoints were evaluated at 1, 3 and 19-weeks post-irradiation.  </td> <td> All three timepoints tested showed significant differences in cardiac structure measures between 50 Gy irradiated and 0 Gy control groups. By week 19, AWTs increased 1.3-fold, IWTs increased 1.1-fold, AWTd increased 1.5-fold and IWTd increased 1.2-fold.  </td> </tr> <tr> <td> Russel et al., 2009       </td> <td> Ex vivo. Recipient arteries from 147 patients receiving reconstructive surgery following treatment for head and neck cancer (H&N) or breast cancer (BC) were studied. H&N treatment group received 66 Gy (standard deviation 7 Gy) dose of radiation, while BC group received 49 Gy (standard deviation 3 Gy).     Intimal-medial thickness in the form of intima-media ratio (IMR) was assessed through histology and proteoglycan and collagen content were scored. Analysis compared irradiated vessels, unirradiated vessels from the same patient as well as donor controls.   </td> <td> In the H&N group the IMR was 1.5-fold greater without correction for the control artery.     In the BC group the IMR increased 1.4-fold after correction for the control artery at a mean of 4 years following irradiation.     There was an increase in the proteoglycan content of the intima of the irradiated IMA vessels, from 65% to 73%.   </td> </tr> </tbody> </table>

Time concordance <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Sridharan et al., 2020  </td> <td> In vivo. Male rats were exposed to oxygen ions and whole-body protons (0.5 Gy) to measure cardiac function and blood flow in study A. Measurements were taken at 3, 5, 9 and 12 months after radiation. Ultrasound, histology and Western blots were used as measurement methods.  In study B, male rats were exposed to oxygen ions (0.01-0.25 Gy). Heart tissue analysis was performed 6-7 and 12 months after radiation using histology and western blots.  </td> <td> Study A:  At 3 and 5 months after proton and oxygen ion radiation, there was a significant decrease in left ventricular posterior wall thickness.   12 months after oxygen ion exposures, velocity measures of pulsed wave Doppler of abdominal aorta increased.   Cardiac volume increased at all time points in proton exposed rats, with a significance at 3 and 9 months.  Collagen type III increased 2.4-fold after 12 months of 0.5 Gy protons radiation.   Protein T lymphocyte markers CD2, CD4 and CD8 content in the rat hearts increased 6 months after a dose of 16O at 0.1 Gy.   CD2, CD4 and CD8 increased 1.4-fold 12 months after 16O at 0.5 Gy.  </td> </tr> <tr> <td> Soucy et al., 2007  </td> <td> In vivo. Sprague-Dawley rat aorta was irradiated with 0.5, 1.6 and 5 Gy 137Cs gamma rays. A measure of vascular stiffness and vascular remodeling was calculated using PWV and measurements taken with an ECG and doppler probe.  </td> <td> At 1 day post irradiation in each group PWV significantly increased. At 1- and 2-weeks levels remained similar to the 1-day results, but slightly increased in the 500 cGy group and slightly decreased in the 50 and 160 cGy groups.  </td> </tr> <tr> <td> Soucy et al., 2011  </td> <td> In vivo. 56Fe ions were used to irradiate rats (0.5 Gy/min). PWV measured with Doppler probe and ECG was used to determine vascular remodeling.  </td> <td> At 4 months after irradiation with 1 Gy, PWV increased by ~0.5 m/s.   At 8 months, PWV remained at a 0.5 m/s increase over control.  </td> </tr> <tr> <td> Yu et al., 2011  </td> <td> In vivo. 2 and 5 Gy 56Fe ions were irradiated onto apoE-/- mice, and intima thickness of the carotid artery was measured using hematoxylin and eosin staining after 13 and 40 weeks.  </td> <td> At both doses, after 13 weeks, intima thickness increased 1.4-fold. Intima thickness was the same as control after 40 weeks.  </td> </tr> <tr> <td> Soucy et al., 2010  </td> <td> In vivo. Rats were irradiated with 5 Gy 137Cs gamma radiation and PWV, measured with Doppler probe and electrocardiogram (ECG) over 2 weeks, was used as a measure of aortic stiffness and vascular remodeling.  </td> <td> PWV increased 1.1-fold (not significant) after 1 day, 1.2-fold after 1 week and 1.2-fold after 2 weeks.  </td> </tr> <tr> <td> Hamada et al., 2020   </td> <td> In vivo. Male mice were irradiated by 5 Gy 137Cs gamma rays (0.5 Gy/min). Vascular remodeling was measured after 1, 3 and 6 months through a Miles assay to show vascular permeability, vascular endothelial cadherin (VE-cadherin, a marker for adherens junctions) levels and number of endothelial detachments.  </td> <td> Vascular permeability increased 8-fold at 1 month, 16-fold at 3 months and 5-fold at 6 months shown by staining intensity. VE-cadherin decreased about 0.2-fold after 1, 3, and 6 months. Percent of mice with total endothelial detachments increased from 0 to 20% at 1 month, 34% at 3 months and 90% at 6 months post-irradiation.  </td> </tr> <tr> <td> Shen et al., 2018  </td> <td> In vivo. Male mice irradiated with 18 Gy X-rays had measurements of aortic thickness determined from 3 to 84 days post-irradiation using hematoxylin and eosin staining. The accumulation of collagen was measured using Sirius red staining from 3 to 84 days.  </td> <td> Aortic thickness increased 1.1-fold after 3 days (ns), 1.4-fold after 7 days, 1.3-fold after 14 and 28 days and 1.2-fold (ns) after 84 days. Collagen increased 1.4-fold 14-, 28- and 84- days post-irradiation.  </td> </tr> <tr> <td> Azimzadeh et al., 2015      </td> <td> In vivo. 10-week-old male C57Bl/6 mice received cardiac irradiation at 8 or 16 Gy with X-rays. Serum oxLDL was measured using ELISA 16 weeks post-irradiation.  </td> <td> After 16 weeks, oxLDL increased 1.2-fold after 8 Gy and 1.1-fold after 16 Gy compared to 0 Gy control.  </td> </tr> </tbody> </table>

None exist

High Male High Female

Moderate

Juvenile

High

Adult

High Moderate High

The relationship has been shown in vivo in mice and rats and ex vivo in human models. Majority of studies used males. Evidence came from either adult or adolescent animals. However, the relationship is plausible at any life stage.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b42eafd3c40> AOPWiki 2022-10-17T16:54:29

2023-03-21T10:32:53

<upstream-id>b813d3a8-c9cc-49b4-bd70-a0bb3e9e8efd</upstream-id> <downstream-id>1cc566f9-d2e5-4c9c-a06c-c40f7767c251</downstream-id> The increased production of reactive oxygen species (ROS) and reactive nitrogen species (RNS) during oxidative stress can lead to altered nitric oxide (NO) levels, specifically a reduction in its bioavailability.     Although RNS can also interfere with NO levels, most studies focus on ROS and not RNS (Nagane et al., 2021). Oxidative stress influences the production and activity of endothelial nitric oxide synthase (eNOS), thereby altering and reducing NO levels and its bioavailability. eNOS, otherwise known as NOS3, is an enzyme that catalyzes NO production from the amino acid L-arginine in vascular endothelial cells. NO mediates vascular tone and blood flow via the activation of soluble guanylate cyclase (sGC) within the vascular smooth muscle (Chen, Pittman and & Popel, 2008). A form of ROS known as superoxide anion (O2-) causes increased NO degradation (Incalza et al., 2018), converting NO into the RNS peroxynitrite. In addition, ROS can uncouple eNOS by oxidation of the enzyme’s cofactor, BH4 (Matsubara et al., 2015; Forstermann, 2010). Uncoupled eNOS produces ROS instead of NO, which can further convert existing NO into peroxynitrite (Forstermann, 2010).

Overall weight of evidence: Moderate

The biological rationale for the relationship between increased oxidative stress and altered NO levels is well-supported by the literature, validated by many studies presented in this area of research. The biological mechanism of this relationship is well-known and widely accepted. The reaction between NO and free radicals leads to the creation of peroxynitrite, which results in reduced NO bioavailability (Incalza et al., 2018; Mitchell et al., 2019; Nagane et al., 2021; Soloviev & Kizub, 2019; Wang, Boerma & Zhou, 2016). In addition, in vascular tissues, increased levels of O2- or peroxynitrite can oxidize BH4 and lead to uncoupled eNOS (Forstermann, 2010; Matsubara et al., 2015; Soloviev & Kizub, 2019). When uncoupled, eNOS transfers electrons to O2- rather than L-arginine, causing O2- production instead of NO production, further reducing NO bioavailability. Although much of the biological plausibility indicates that NO decreases with oxidative stress, ROS have been associated with increased NO as well (Nagane et al., 2021; Soloviev & Kizub, 2019). This is likely due to the complexity of NO regulation in signaling pathways, upregulation of inducible nitric oxide synthase (iNOS), as well as variations in stressors, doses, dose rates, models, duration of study and diseases present in studies (Nagane et al., 2021).

The empirical data for this KER somewhat supports the relationship of increased oxidative stress eventuating in altered NO levels. The evidence was gathered from both in vivo and in vitro models. Gamma rays (Abdel-Magied & Shedid, 2020; Hasan, Radwan & Galal, 2019; Soucy et al., 2010), X-rays (Cervelli et al., 2017; Yan et al., 2020), altered gravity (Zhang et al., 2009) and heavy ions (Soucy et al., 2011) were used as stressors with dose levels ranging from 0.25 to 10 Gy. Direct and indirect measures of NO, including iNOS, eNOS and nitrite levels, were used as endpoints to investigate the effect of increased oxidative stress on NO levels. Evidence from radiation sources that induce oxidative stress in human umbilical vein endothelial cells (HUVECs) or rat aorta have demonstrated a change in NO levels (Abdel-Magied & Shedid, 2020; Cervelli et al., 2017; Hasan, Radwan & Galal, 2019; Soucy et al., 2010; Soucy et al., 2011; Yan et al., 2020; Zhang et al., 2009).    Dose Concordance Moderate evidence is available within current literature that demonstrates dose-concordance between increased oxidative stress and altered NO levels. Elevations in the production of ROS have been reported with exposure to altered gravity and different doses (0.25 Gy, 1 Gy, 4 Gy, 5 Gy, and 10 Gy) of radiation in HUVECs and rat aorta/tissue models, and provide supporting evidence that an increase in oxidative stress can lead to subsequent alterations in NO levels (Abdel-Magied & Shedeed, 2020; Cervelli et al., 2017; Hasan, Radwan & Galal, 2019; Sakata et al., 2015; Soucy et al., 2010; Soucy et al., 2011; Yan et al., 2020; Zhang et al., 2009). For example, a study involving HUVEC X-ray irradiation at 0.25 Gy found increases in both ROS and nitrite/nitrate levels (Cervelli et al., 2017). As well, a study involving 1 Gy of iron ions demonstrated elevations in ROS production with accompanying reductions in NO levels using the DAF-FM DA fluorescent probe (Soucy et al., 2011).   At high doses (>2 Gy), dose concordance between increased oxidative stress and altered nitric oxide can be observed. For example, 5 Gy gamma irradiation leads to increased ROS production with decreased NO levels measured using the DAF-FM DA fluorescent probe (Soucy et al., 2010) Gamma irradiation of rats with 6 Gy led to changes in oxidative stress indices that showed an increase in ROS, while iNOS levels were also increased (Hasan, Radwan & Galal, 2019). Rat cardiac tissue exposed to 8 Gy gamma irradiation also showed an increase in ROS through oxidative stress indices and an increase in nitrite/nitrate content (Abdel-Magied & Shedid, 2020). A study investigating HUVECs and rat aorta/tissue responses to 10 Gy and 4 Gy X-ray radiation, respectively, showed reduced eNOS dimerization and nitrite levels with elevations in ROS production in both models (Yan et al., 2020). Also in HUVECs, 10 Gy X-rays resulted in increased ROS, increased p-eNOS (Ser1177), decreased p-eNOS (Thr495), increased citrulline and increased NOx (nitrite and nitrate, NO proxies) (Sakata et al., 2015). NOx was also significantly increased after 20 Gy (Sakata et al., 2015).  Simulated microgravity showed increased eNOS and iNOS in rats, while superoxide levels, although not quantitatively shown, increased following altered gravity (Zhang et al., 2009). Although, the authors state that the increase in NOS would likely cause decreased NO production because of NOS uncoupling and peroxynitrite production (Zhang et al., 2009).    Time Concordance  Very few studies demonstrate time concordance of this relationship. In HUVECs irradiated with 0.25 Gy of X-rays, ROS levels increased after 45 minutes, while nitrite and nitrate levels increased after 24 hours. In rats exposed to 6 Gy of gamma rays, both oxidative stress indices and iNOS increased 4 weeks post-irradiation (Hasan, Radwan & Galal, 2019).    Incidence concordance  There is moderate evidence of incidence concordance for this relationship. Yan et al. (2020) showed both in vivo in a rat model irradiated with 4 Gy of X-rays and in vitro in HUVECs irradiated with 10 Gy of X-rays that superoxide levels increased more than NO and eNOS were decreased following irradiation. Rats irradiated with 1 Gy of 56Fe ions showed a 1.8-fold increase in ROS and just a 0.8-fold decrease in NO levels (Soucy et al., 2011). Similarly, gamma irradiation of rats at 5 Gy resulted in a 1.7-fold increase in ROS and a 0.7-fold decrease in NO levels (Soucy et al., 2010). In HUVECs irradiated with 10 Gy of X-rays, ROS increased 15.5-fold while NOx levels increased just 10-fold (Sakata et al., 2015).     Essentiality     Studies scrutinizing the usefulness of antioxidants in inhibiting oxidative stress show a moderately supported relationship between elevated oxidative stress and altered NO levels. Antioxidants have been implicated in the activation of the NOS family of enzymes by preventing BH4 oxidation, thereby increasing NO bioavailability (Kojsova et al., 2006). One study observed that treatment with bradykinin potentiating factor (BPF), which can affect eNOS regulation through the renin aldosterone angiotensin system, contributed to elevations in the antioxidant glutathione (GSH) and a marker of antioxidant potential, ferric reducing antioxidant power (FRAP) (Hasan, Radwan & Galal, 2019). BPF contributed to reductions in malondialdehyde (MDA), a biomarker for oxidative stress, while also increasing iNOS levels (Hasan, Radwan & Galal, 2019).  Another study demonstrated that a composition of antioxidants (resveratrol, extramel, seleno-L-methionine, Curcuma longa, reduced glutathione, and vitamin C (RiduROS) inhibited increases in ROS while restoring NO levels (Cervelli et al., 2017). Furthermore, rat mesenteric arteries and HUVECs treated with 2,4-diamino-6-hydroxypyrimidine (DAHP), an inhibitor of Gch1 that is important for BH4 synthesis, lowered both the dimer:monomer eNOS ratio and the nitrite (measure of NO) concentration, and increased superoxide following irradiation (Yan et al., 2020). The notable elevation in O2- demonstrates that inhibiting Gch1 and limiting BH4 activity can prevent eNOS activity and elevate ROS Levels (Yan et al., 2020).  In addition, two studies demonstrated that treatment with oxypurinol (OXP), a xanthine oxidase (XO) inhibitor where XO is responsible for generating cardiac ROS, led to reductions in ROS well under the control level whereas NO increased to control levels demonstrating that oxidative damage contributes to a decrease in NO bioavailability (Soucy et al., 2010; Soucy et al., 2011).  Zinc oxide nanoparticles (ZnO-NPs) can act as antioxidants; Abdel-Magied & Shedid (2020) found that low concentrations (10 mg/kg ZnO-NPs) decreased oxidation and NO levels in rats after gamma irradiation. Angiotensin II type 1 (AT1) receptors are able to regulate NOS levels. Treatment with losartan, an AT1 receptor antagonist, led to a decrease in O2- , iNOS and eNOS levels, demonstrating that the increase in NOS and ROS can be prevented by blocking AT1 receptors (Zhang et al., 2009). Biotin was found to increase GSH content, superoxide dismutase (SOD), catalase (CAT) activity and return NOx levels to near control values following irradiation in hippocampus (Abdel-Magied & Shedid, 2020).

<ul> <li>The directionality of changes to NO is inconsistent between studies, as some studies show increased NO levels and other studies show decreased NO levels. Improved methods are needed to assess NO levels directly, which may facilitate an understanding of the relationship (Cervelli et al., 2017, Hasan, Radwan & Galal, 2019). This, along with variation in experimental conditions, can account for the inconsistencies in NO changes between studies.</li> </ul>

The following are a few examples of quantitative understanding of the relationship. All data that is represented is statistically significant unless otherwise indicated.

Moderate Male Low Female Low Unspecific

Moderate

Adult

Low

Juvenile

Low High

The evidence for the taxonomic applicability to humans is low as evidence comes from in vitro human cell-derived models. Many studies use in vivo rat models, predominately males. Occasionally, animal age is not specified in studies; most studies indicate the animals are adult or adolescent. In addition, the relationship is also plausible in preadolescent animals.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b42eb067bc0> AOPWiki 2022-09-28T12:41:55

2023-03-21T11:35:56

<upstream-id>b813d3a8-c9cc-49b4-bd70-a0bb3e9e8efd</upstream-id> <downstream-id>870809d3-663d-4170-9ec7-b63bbcebbc41</downstream-id> Oxidative stress describes the imbalances in reactive oxygen and reactive nitrogen species (RONS) radical formation as well as antioxidants and reactive oxygen species (ROS) scavengers (Beckhauser et al., 2016; Elahi et al., 2009; Ray et al., 2012). Oxidative stress can lead to endothelial dysfunction. Within the cardiovascular system, every vessel is lined with a single layer of endothelial cells (Augustin et al., 1994; Fishman, 1982). This endothelial layer plays a crucial role in the regulation of vascular homeostasis through controlling various factors such as vascular permeability, vasomotion, and immune response (Baran et al., 2021; Bonetti et al., 2003; Hughson et al., 2018; Slezak et al., 2017; Sylvester et al., 2018). Of the vascular wall components, the endothelium is also the most vulnerable to damage from ROS (Soloviev & Kizub, 2018). Endothelial cells normally exist in a quiescent state characterized by high nitric oxide (NO) bioavailability (Carmeliet & Jain, 2011); however, cells can become activated as part of a normal host-defence response following tissue injury or oxidative stress (Deanfield et al., 2007; Krüger-Genge et al., 2019). Sustained activation leads to the pathological state of endothelial dysfunction which is defined by decreased NO bioavailability, increased vessel permeability, altered vasomotion, and a pro-thrombotic and inflammatory environment (Baran et al., 2021; Bonetti et al., 2003; Deanfield et al., 2007; Schiffrin, 2008). Shifting redox balance towards oxidation is known to indirectly lead to endothelial dysfunction through various mechanisms (Hughson et al., 2018; Ramadan et al., 2020; Soloviev & Kizub, 2018). There are several ways through which imbalanced ROS can affect endothelium function, including decreasing NO bioavailability through direct scavenging, which forms the RNS peroxynitrite (ONOO-) (Hatoum et al., 2006; Li et al., 2002; Schiffrin, 2008; Soloviev & Kizub, 2018; Venkatesulu et al., 2018), as well as impeding NO production and diffusion (Hatoum et al., 2006; Li et al., 2002; Schiffrin, 2008; Soloviev & Kizub, 2018; Venkatesulu et al., 2018; Schiffrin, 2008; Soloviev & Kizub, 2018). Additionally, elevated ROS contribute to introducing a pro-inflammatory and pro-thrombotic milieu characteristic of dysfunction (Hughson et al., 2018; Schiffrin, 2008; Slezak et al., 2017; Tapio, 2016; Venkatesulu et al., 2018). It is also linked to decreased vasomotion (Schiffrin, 2008; Soloviev & Kizub, 2018; Venkatesulu et al., 2018) and finally the onset of endothelial cell apoptosis and premature senescence (Borghini et al., 2013; Hughson et al., 2018; Tapio, 2016; Wang et al., 2016).

Overall weight of evidence: Moderate

Mechanisms for oxidative stress leading to endothelial dysfunction are outlined in various reviews on the topic (Hughson et al., 2018; Nagane et al., 2021; Slezak et al., 2017; Soloviev & Kizub, 2018; Venkatesulu et al., 2018; Wang et al., 2016). Elevated ROS can indirectly lead to endothelial dysfunction by causing an imbalance of NO, specifically the decrease in NO bioavailability. Firstly, ROS can react with NO directly; if quenching outpaces NO production, it will cause reduced NO bioavailability underlying endothelial dysfunction (Hatoum et al., 2006; Li et al., 2002; Soloviev & Kizub, 2018). In particular, the superoxide anion (•O-2) reacts with NO to form peroxynitrite, both reducing available NO and further accelerating NO degradation (Li et al., 2002; Soloviev & Kizub, 2018). In addition, superoxide and peroxynitrite can uncouple eNOS which produces more ROS instead of NO (Soloviev & Kizub, 2018). Peroxynitrite can cause cellular senescence as a part of endothelial dysfunction (Nagane et al., 2021). eNOS downregulation and subsequent drop in NO levels are caused in part by increased endothelin-1 (ET-1), a vasoconstrictor with enhanced secretion during an oxidative stress state (Marasciulo, Montagnani & Potenza, 2006; Ramadan et al., 2020). ROS is also involved in perturbing NO diffusion from the endothelial cells (Soloviev & Kizub, 2018). Overall, the decreased NO bioavailability causes reduced vasodilation and endothelial dysfunction (Soloviev & Kizub, 2018). Oxidative stress also affects endothelial function through inhibition of endothelium-dependent vasodilation (Soloviev & Kizub, 2018; Venkatesulu et al., 2018). ROS in both endothelial cells and surrounding vascular smooth muscle cells (VSMCs) act as second messengers to many cellular pathways that mediate VSMC contractility and endothelial permeability and function, causing disruption to these endothelial functions (Hughson et al., 2018; Li et al., 2002; Ramadan et al., 2020; Soloviev & Kizub, 2018; Ungvari et al., 2013; Venkatesulu et al., 2018). Specifically, impaired endothelium-dependent vasomotion following radiation (Venkatesulu et al., 2018) was suggested to be due to the loss of PGF2α inhibition and therefore, vasoconstriction (Li et al., 2002). Oxidative stress is also involved in inducing the pro-thrombotic and inflammatory environment of endothelial dysfunction. In the case of radiation induced endothelial injury, radiation type, fraction size used, and endothelial cell model used all influence the resulting downstream endpoints (Venkatesulu et al., 2018). Possible changes to the endothelial milieu include alterations of cell adhesion molecule levels, creation of pro-thrombotic environment, endothelial cell apoptosis and inflammation (Hughson et al., 2018; Nagane et al., 2021; Slezak et al., 2017; Tapio, 2016; Venkatesulu et al., 2018). When induced by oxidative stress, nuclear factor kappa B (NF-кB) can target genes involved with the upregulation of prothrombotic markers associated with endothelial dysfunction (Slezak et al., 2017). Free radicals produced by macrophages have also been shown to stimulate TGF-β, thus accelerating the creation of a profibrotic milieu (Venkatesulu et al., 2018). ROS can also oxidize low-density lipoproteins (LDL) resulting in structural complications as oxidized LDL accumulates in blood circulation due to decreased cell uptake (Nagane et al., 2021; Slezak et al., 2017). Furthermore, endothelial cells can undergo morphological changes following oxidative injury, as the cells become enlarged, and form fibrin networks, showing increased levels of activated platelets and leukocytes with membrane protrusions and pseudopodial extensions, which are all indicative of an inflammatory and pro-thrombotic state (Li et al., 2002). Furthermore, ROS can induce premature endothelial cell senescence, which in turn contributes to overall endothelial dysfunction (Hughson et al., 2018; Nagane et al., 2021; Tapio, 2016). In contrast to replicative senescence attributed to telomere dysfunction, oxidative stress is one of several injuries causing stress-induced premature senescence (Nagane et al., 2021). This is thought to occur through oxidative stress causing the induction of the p53/p21 pathway which regulates cell senescence (Borghini et al., 2013; Wang et al., 2016). Once senescent, the endothelial cells contribute to dysfunction in multiple ways. Firstly, senescence can stimulate a pro-inflammatory response and trigger apoptosis through decreased cell repair (Nagane et al., 2021; Ramadan et al., 2020). Additionally, senescent cells themselves are sources of ROS, furthering both genomic instability causing additional senescence in neighbouring cells and endothelial dysfunction itself (Tapio, 2016; Wang et al., 2016). Senescent cells also lack proper endothelial cell function, contributing to changing the environment to a dysfunctional one (Hughson et al., 2018; Tapio, 2016). This lack of function includes a decrease in NO production, increased monocyte adhesion, and loss of cell barrier integrity paired with increased levels of ET-1 (Hughson et al., 2018; Nagane et al., 2021; Tapio, 2016). Finally, oxidative stress has been shown to lead to mitochondrial dysfunction and dysregulation, which is thought to play an important role in the development of endothelial dysfunction (Borghini et al., 2013; Hughson et al., 2018; Nagane et al., 2021; Slezak et al., 2017).

Empirical evidence provides a moderate level of support to this KER. Examples of this evidence are summarized here and further in attached tables.The evidence to support the relationship between oxidative stress leading to endothelial dysfunction was gathered from studies using in vitro and in vivo rat and mice models (Delp et al., 2016; Hatoum et al., 2006; Shen et al., 2018; Soucy et al., 2007; Soucy et al., 2010; Soucy et al., 2011; Ungvari et al., 2013), in vivo pig models (Li et al., 2002) and human in vitro cells (Ramadan et al., 2020). Various stressors were applied, including X-rays, hindlimb unloading (HU), heavy ions (56Fe ions and 32P) and gamma rays with a dose range of 0.1 to 22.5 Gy. To determine the effect of oxidative stress on endothelial dysfunction, various assays and end-points were measured. Senescence-associated β-galactosidase (SA ꞵ-gal), insulin-like growth factor-binding protein-7 (IGFBP-7) and growth differentiation factor 15 (GDF-15) as senescence markers, caspase 3/7 activity as an apoptosis marker, endothelin-1 levels for the ratio of apoptotic to normal cells, 4-hydroxynonenal (4-HNE) and 3-nitrotyrosine (3-NT) as aortic oxidative damage markers, superoxide production, vascular tension, and ROS detection via xanthine oxidase (XO) activity, and fluorescent dyes, such as dihydroethidium fluorescence were all used as end-point measures. Dose Concordance Dose concordance between the two key events is supported by numerous studies. Hatoum et al. (2006) explored the effect of 3 to 9 cumulative X-ray doses of 0.25 Gy on murine intestinal arterioles. The study found that superoxide and hydrogen peroxide generation was increased, while at the same doses vasodilation in response to acetylcholine (ACh) was decreased (Hatoum et al., 2006). Another study using various doses on cerebral microvascular endothelial cells found a significant change in cellular peroxide and mitochondrial oxidative stress following the 4 Gy dose, while SA β-gal showed the first large increase at 4 Gy (Ungvari et al., 2013). Ramadan et al. (2020) used X-ray irradiation in multiple types of human endothelial cells has shown that ROS production is significantly higher in both the 0.1 Gy and 5 Gy compared to the control. These changes were correlated to endothelial dysfunction as SA β-gal activity and endothelial apoptosis were also affected, showing a response of greater magnitude following the 5 Gy dose compared to 0.1 Gy (Ramadan et al., 2020). Exposure of mice to 18 Gy X-rays led to a ~1.8-fold and ~2.2-fold decrease in 4-HNE and 3-NT respectively, both being markers of aortic oxidative damage. Simultaneously, the 18 Gy dose caused a ~5-fold increase in aortic apoptosis, a marker of endothelial dysfunction (Shen et al., 2018). Two studies (Soucy et al., 2007; Soucy et al., 2010) found that varying doses (0.5 and 5 Gy) of gamma radiation led to significant increases in ROS levels in rat aorta. Subsequently, endothelial function was affected with a 0.5 Gy dose resulting in a ~30% decrease in ACh-induced vasodilation response (Soucy et al., 2007), and a 5 Gy dose leading to a ~13-15% decrease in ACh-induced vasodilation (Soucy et al., 2010). Iron ion irradiation resulted in oxidative stress and endothelial dysfunction both occurring after 1 Gy (Soucy et al., 2011). Similarly, Delp et al. (2016) showed total body 56Fe irradiation at 1 Gy led to a ~2-fold increase in XO activity and a ~10% decrease in ACh response in mice (Delp et al., 2016). Delp et al. (2016) also explored the effects of HU on mice and found no significant changes to XO activity or ACh response following 2-week HU, while HU in combination with 1 Gy 56Fe radiation led to a ~2.2-fold increase in XO activity and the same ~10% decrease in ACh response. Li et al. (2002) showed ~3.5-fold increase in superoxide anion production and a ~25-80% decrease in vasoconstriction and vasodilation response to various vasomotive substances following 20 Gy 32P radiation in pig coronary arteries. Time Concordance There is some evidence of time concordance between oxidative stress and endothelial dysfunction. Ramadan et al. (2020) used human endothelial cells and showed ROS production first increased 45 minutes after 5 Gy X-ray exposure before returning to baseline levels after 2 hours. Apoptosis markers Annexin V and Caspase 3/7 were first increased after 4 hours. Cellular senescence evaluated with the SA-β-gal activity was first measured only after 7 days (Ramadan et al., 2020). ROS production and dilation response to ACh after irradiation were measured at various times as cumulative doses were given, which showed increased ROS and decreased dilation of rat intestinal microvessels both after 5 days of cumulative 0.25 Gy X-ray doses (Hatoum et al., 2006). Work using 4-HNE and 3-NT as biomarkers of oxidative stress in mice aorta following 18 Gy 6MV X-ray at 3, 7, 14, 28, and 84 days after irradiation showed both markers to be significantly elevated starting after 3 days. Exposure also led to significantly increased apoptosis, indicating endothelial dysfunction after 3 days (Shen et al., 2018).   Incidence Concordance There is moderate support in current literature for an incidence concordance relationship between oxidative stress and endothelial dysfunction. Three of the primary research studies used to support this AOP demonstrated an average change to endpoints of oxidative stress that was greater or equal to that of endothelial dysfunction (Soucy et al., 2011; Soucy et al., 2010; Soucy et al., 2007). Essentiality   Essentiality in the relationship was demonstrated in the following studies. Studies by Soucy et al. (2007, 2010, 2011) explored the relationship between radiation exposure, ROS levels and endothelial function, all focusing on the role of XO. Soucy et al. (2007) incubated aortic rings from irradiated rats in the XO inhibitor oxypurinol (Oxp) and saw this treatment result in recovery of ACh vasodilation response. Soucy et al. (2010) showed that administration of allopurinol (a superoxide scavenger) following irradiation led to significantly decreased ROS levels. Additionally, the latter two studies showed that XO inhibition by dietary administration of Oxp significantly decreased XO activity and ROS levels while simultaneously recovering ACh response (Soucy et al., 2010, 2011). Similar results were observed when treatment with manganese tetrakis (4-benzoic acid) porphyrin chloride (MnTBAP), a SOD mimetic, returned peroxide and superoxide levels and significantly improved ACh response irradiated rats (Hatoum et al., 2006). Dietary treatment with Tempol, a water-soluble SOD-mimetic likewise increased vasomotion and decreased superoxide levels (Hatoum et al., 2006). Human bone marrow mesenchymal stem cells (hBMSCs) have also been studied for their ability to prevent radiation-induced aortic injury. Both high and low doses of hBMSCs were shown to increase catalase and heme oxygenase 1 (HO-1) antioxidant activity, and decrease levels of the aortic oxidative damage markers 4-HNE and 3-NT. Subsequently, this treatment also significantly decreased levels of apoptosis in the aorta (Shen et al., 2018). Finally, blocking Connexin43 hemichannels using TAT-Gap19 peptide also significantly reduced oxidative stress and resultant cell senescence and death, suggesting the role of intracellular communication in mediating radiation response (Ramadan et al., 2020). Studies have also shown that transgenic mice overexpressing superoxide dismutase (SOD) have a twofold reduction in aortic lesions following X-ray exposure compared to control (Hughson et al., 2018). Overexpression of SOD also mitigates atherosclerotic plaque formation, further outlining the relationship between oxidative stress and the pathological environment of endothelial dysfunction (Tapio, 2016).

Work by Ramadan et al. (2020) explored the use of TAT-Gap19 to block endothelial intracellular communication in order to modulate radiation response of intercellular connexin proteins. Overall, TAT-Gap19 was shown to reduce ROS production and subsequent senescence (SA β-gal activity) and apoptosis (Annexin V and Caspase 3/7) markers. However, treatment with TAT-Gap19 led to an increase in SA β-gal in non-irradiated control at the 9-day point. Additionally, the 0.1 Gy irradiated group showed persistent SA β-gal activity at all time points studied, while the 5 Gy group demonstrated an unexpected decrease before day 14.

<div> <table cellspacing="0" class="MsoTableGrid" style="border-collapse:collapse; border:none"> <tbody> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:1px solid black; height:26px; vertical-align:top; width:156px"> Modulating factor </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; height:26px; vertical-align:top; width:156px"> Details </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; height:26px; vertical-align:top; width:156px"> Effects on the KER </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; height:26px; vertical-align:top; width:156px"> References </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Drug </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> MnTBAP (a superoxide dismutase mimetic) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Treatment with MnTBAP after irradiation was able to reduce superoxide and peroxide levels and restore vasodilation ability </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> (Hatoum et al., 2006) </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Drug </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Tempol (a superoxide dismutase mimetic) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Treatment with tempol after irradiation was able to restore vasodilation ability </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> (Hatoum et al., 2006) </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Drug </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> TAT-Gap19 (inhibitor of connexin 43 which is associated with atherogenesis and endothelial stiffness) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Treatment with TAT-Gap19 led to a decrease in ROS and SA β-gal levels after irradiation </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> (Ramadan et al., 2020) </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Drug </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> hBMSCs (protect against vascular damage through antioxidant properties) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Treatment with hBMSCs after irradiation caused increased catalase and HO-1, as well as decreased oxidative damage and apoptosis </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> (Shen et al., 2018) </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Drug </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Oxp (can inhibit XO, a source of ROS) </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> Treatment with Oxp showed decreased XO activity and ROS production along with increased vasodilation after irradiation </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:26px; vertical-align:top; width:156px"> (Soucy et al., 2007; Soucy et al., 2010; Soucy et al., 2011) </td> </tr> </tbody> </table> </div>

The following are a few examples of quantitative understanding of the relationship. All data that is represented is statistically significant unless otherwise indicated.

Dose/incidence concordance <table cellspacing="0" class="Table" style="border-collapse:collapse; border:none; width:624px"> <tbody> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:1px solid black; vertical-align:top; width:162px"> Reference </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; vertical-align:top; width:200px"> Experiment Description </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:1px solid black; vertical-align:top; width:262px"> Result </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:25px; vertical-align:top; width:162px"> Soucy et al. 2007 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:25px; vertical-align:top; width:200px"> In vivo. Sprague-Dawley rats were whole-body irradiated with 137Cs gamma radiation at 50, 160 and 500 cGy. XO is a primary source of cardiac ROS and was used as a measure of oxidative stress. Vasodilation response to ACh was used to evaluate endothelial function. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:25px; vertical-align:top; width:262px"> At 500 cGy, XO activity was found to be 2-fold elevated compared to control, and there was also an increase in XO quantity. Simultaneously, there was endothelial dysfunction as seen with a ~30 percentage point decrease in vasodilation response to ACh. </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:104px; vertical-align:top; width:162px"> Soucy et al. 2010 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:104px; vertical-align:top; width:200px"> In vivo. In Sprague-Dawley rats were whole-body irradiated with 137Cs gamma radiation at 5 Gy. ROS were measured using dihydroethidium fluorescence. Aortic relaxation response to ACh was also measured. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:104px; vertical-align:top; width:262px"> After 5 Gy, ROS increased 1.7-fold and relaxation decreased 0.7-fold. </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:51px; vertical-align:top; width:162px"> Soucy et al. 2011 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:51px; vertical-align:top; width:200px"> In vivo. Wistar rats were exposed to 0.5 and 1 Gy doses of 56Fe-ion radiation. ROS production rates were evaluated using dihydroethidium fluorescence. ACh-induced vasodilation responses were measured. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:51px; vertical-align:top; width:262px"> 56Fe irradiation at 1 Gy produced a 1.8-fold increase in ROS levels. At 10-5 M ACh, aorta without irradiation relaxed by 87%, while aorta with 1 Gy irradiation had significantly lower relaxation of 76%. </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:195px; vertical-align:top; width:162px"> Li et al. 2002 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:195px; vertical-align:top; width:200px"> In vivo. Surgically exposed coronary arteries of yucatan pigs were irradiated with 20 Gy 32P β-irradiation. Oxidative stress was evaluated through superoxide production. Endothelial function was evaluated through endothelial-dependent vasomotor response and morphological changes. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:195px; vertical-align:top; width:262px"> Superoxide production increased 3.5-fold between the control and 20 Gy irradiated groups. Contractile response to KCl dropped over 50% in the irradiated group. Morphological changes were also observed, with irradiated arteries seeing enlarged endothelial cells, formation of fibrin networks, activated platelets, leukocytes exhibiting membrane protrusions and pseudopodial extensions all indicative of an inflammatory and pro-thrombotic state of endothelial dysfunction. </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:125px; vertical-align:top; width:162px"> Shen et al. 2018 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:125px; vertical-align:top; width:200px"> In vivo. Male mice were irradiated with 18 Gy X-rays. Oxidative stress was measured with 4-HNE and 3-NE oxidative damage markers and antioxidant enzymes catalase and HO-1, measured by immunohistological staining. Endothelial dysfunction was determined through apoptosis. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:125px; vertical-align:top; width:262px"> 4-HNE showed a maximum increase of ~1.8-fold, and 3-NT showed a maximum increase of ~2.2-fold. Apoptosis levels peaked at a ~5-fold increase above control levels. </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:83px; vertical-align:top; width:162px"> Ramadan et al. 2020 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:83px; vertical-align:top; width:200px"> In vitro. Telomerase-immortalized human Coronary Artery and Microvascular Endothelial cells (TICAE) and Telomerase Immortalized human Microvascular Endothelial cells (TIME) were exposed to X-rays (0.1 and 5 Gy). ROS production was measured using CM-H2DCFDA combined with Incucyte live cell imaging.   Endothelial dysfunction was evaluated through:   endothelial apoptosis <ul> <li>Annexin V and Caspase 3/7 marker levels</li> <li>Dextran fluorescein dye uptake level by necrotic cells  </li> </ul> cell senescence <ul> <li>SA β-gal activity</li> <li>IGFBP-7 and GDF-15 senescence marker levels</li> </ul> </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:83px; vertical-align:top; width:262px"> ROS production was increased in TIME cells after 0.1 and 5 Gy dose.   In both coronary and endothelial cells, apoptosis mainly occurred after 5 Gy radiation. With coronary cells demonstrating an increase in Annexin V and Caspase 3/7 markers and endothelial cells showing elevated Annexin V and membrane leakage.    SA β-gal activity significantly increased for both 0.1 and 5 Gy doses. IGFBP-7 and GDF-15 levels were also elevated in both cell types; GDF-15 increasing at both 0.1 and 5 Gy doses, while IGFBP-7 only showed significant elevation at the 5 Gy dose. With the 0.1 Gy dose, there was a significant increase in SA β-gal activity of ~3-fold, while at 5 Gy the activity increased ~5-fold.   Endothelin-1 was found to be significantly elevated following 5 Gy irradiation in both cell types. </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:59px; vertical-align:top; width:162px"> Ungvari et al. 2013 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:59px; vertical-align:top; width:200px"> In vitro. Cerebral microvascular endothelial cells (CMVECs) from F344×BN rats were harvested and cultured. Following culture, cells were irradiated with 137Cs gamma radiation in doses between 2-8 Gy.   Oxidative stress was evaluated through cellular peroxide and superoxide production.   Endothelial dysfunction was evaluated through cell senescence via SA-β-gal presence, and apoptosis via caspase 3/7 maker and ratio of apoptotic:viable cells. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:59px; vertical-align:top; width:262px"> Oxidative stress increased in a dose-dependent manner following irradiation. Change of ROS became significant after 4 Gy at a ~1.5-fold increase and reached ~3-fold increase at the highest studied dose of 8 Gy. Mitochondrial oxidative stress also became significant after 4 Gy and increased linearly for a peak of a ~1.5-fold increase at 8 Gy.   Endothelial cell senescence and apoptosis were similarly found to increase in a dose dependent manner. With ~30% of cells being SA-β-gal positive after 8 Gy irradiation, signalling premature senescence. Ratio of dead cells peaked at 10% and Caspase 3/7 peaking at a ~5.5-fold change following 18h post irradiation. </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:305px; vertical-align:top; width:162px"> Hatoum et al. 2006 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:305px; vertical-align:top; width:200px"> In vivo. Effect of cumulative radiation doses on rat gut microvessels was studied. Rats were exposed to 1 to 9 fractions of 250 cGy for a total dose of up to 2250 cGy. Following exposure, the animals were euthanized, and submucosal vessels isolated.   Oxidative stress was measured through superoxide and peroxide levels.   Endothelial function was assessed through ACh vasodilation response. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:305px; vertical-align:top; width:262px"> After the final cumulative dose of 2250 cGy, superoxide was ~1.6-fold elevated and peroxides were ~1.7-fold elevated compared to non-irradiated controls. ROS levels increased sharply after the second dose, immediately preceding drop in ACh vasodilation response.   Max dilation dropped from 87% to 3% between pre-irradiation and post-final radiation dose. ACh response remained within control levels following fractions 1 and 2, however following fraction 3, response dropped below 30% for all remaining doses. </td> </tr> <tr> <td style="border-bottom:1px solid black; border-left:1px solid black; border-right:1px solid black; border-top:none; height:305px; vertical-align:top; width:162px"> Delp et al. 2016 </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:305px; vertical-align:top; width:200px"> In vivo. The effects of HU and 1 Gy dose of 56Fe radiation of the gastrocnemius muscle feed arteries and coronary arteries of C57BL/6 mice was studied.   Xanthine oxidase (XO) levels were used as a measure of ROS production and therefore oxidative stress.   Endothelial function was evaluated through vasodilation response to ACh. </td> <td style="border-bottom:1px solid black; border-left:none; border-right:1px solid black; border-top:none; height:305px; vertical-align:top; width:262px"> Following 2-week HU, there were no significant changes to XO levels or vasomotor response.   Following total body irradiation with 1 Gy, there was a ~2-fold increase in XO activity in both gastrocnemius muscle feed and coronary arteries. Vasodilation response subsequently decreased ~10 percentage points.   Combined HU and total body irradiation led to a ~2.2-fold increase in XO activity in both artery types and vasodilation response decrease of ~10 percentage points. </td> </tr> </tbody> </table>

Time concordance  <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Soucy et al. 2007  </td> <td> In vivo. Sprague-Dawley rats were whole-body irradiated with 137Cs gamma radiation at various doses. 2 weeks after irradiation, the animals were euthanized, and aortas were harvested. XO was used as a measure of oxidative stress. Dose-dependent vasodilation response to ACh was used to evaluate endothelial function.  </td> <td> XO activity was elevated 2-fold compared to control, and there was also an increase in XO quantity. Simultaneously, endothelial dysfunction was seen with a ~30% decrease in vasodilation response to ACh.     </td> </tr> <tr> <td> Soucy et al. 2010   </td> <td> In vitro. Sprague-Dawley rats were whole-body irradiated with 137Cs gamma radiation. 2 weeks after receiving radiation dose, the animals were euthanized and aortas were harvested. ROS were measured using dihydroethidium fluorescence. Aortic relaxation response to ACh was also measured.  </td> <td> ROS increased 1.7-fold and relaxation simultaneously decreased 0.7-fold.  </td> </tr> <tr> <td> Soucy et al. 2011   </td> <td> In vivo. Wistar rats were exposed to 56Fe-ion radiation. Rats were euthanized at 4 months post-irradiation and aorta was harvested. ROS production rates evaluated using dihydroethidium along the ACh-induced vasodilation response were measured.  </td> <td> ROS levels increased by 75% 4 months post-irradiation.   ACh vasodilation response decreased by 13%.  </td> </tr> <tr> <td> Shen et al. 2018   </td> <td> In vivo. Male mice were irradiated with 18 Gy X-rays. Immunohistochemical staining assessed oxidative stress using 4-HNE and 3-NE as markers for oxidative damage. Endothelial dysfunction was determined through apoptosis.  </td> <td> Exposure to 18 Gy caused increased 4-HNE and 3-NT levels. 4-HNE showed a maximum ~1.8-fold increase at 14-days post radiation, and 3-NT showed a maximum ~2.3-fold increase 7 days post radiation.   Apoptosis levels peaked at 7 days post-irradiation with a ~5-fold increase above control levels.  </td> </tr> <tr> <td> Ramadan et al. 2020   </td> <td> In vitro. TICAE and TIME cells were exposed to X-rays (0.1 and 5 Gy).     Oxidative stress was evaluated through intracellular ROS production.     Endothelial dysfunction was evaluated through:   endothelial apoptosis   <ul> <li> Annexin V and Caspase 3/7 marker levels  </li> <li> Dextran fluorescein dye uptake level by necrotic cells    </li> </ul> cell senescence   <ul> <li> SA β-gal activity  </li> <li> IGFBP-7 and GDF-15 senescence marker levels  </li> </ul> Endothelin-1 levels  </td> <td> Highest response was observed for both doses at 45 minutes after irradiation followed by a decline at the 2- and 3-hour time points but remaining elevated above non-irradiated control levels. Endothelial cells studied produced more ROS than the coronary cells.     Caspase 3/7 and annexin V increased linearly until 100h.    SA β-gal activity significantly increased at 7 and 9 days. GDF-15 and IGFBP-7 were increased after 7 days.    Endothelin-1 was found to be significantly elevated after 7 days.  </td> </tr> <tr> <td> Ungvari et al. 2013   </td> <td> In vitro. Cerebral microvascular endothelial cells (CMVECs) from F344×BN rats were harvested and cultured. Following culture, cells were irradiated with 137Cs gamma radiation.    Oxidative stress was evaluated through cellular peroxide and superoxide production.    Endothelial dysfunction was evaluated through cell senescence via SA-β-gal presence, and apoptosis via caspase 3/7 maker and ratio of apoptotic:viable cells.   </td> <td> Superoxide and peroxide increased 1 day but not 14 days post-irradiation    ~30% of cells were SA-β-gal positive after 8 Gy irradiation, measured 7 days post-irradiation. 24 h after irradiation, 10% of cells were dead. Caspase 3/7 increased from 2 to 18 h, peaking at a ~5.5-fold change following 18 h post-irradiation but decreased at 24 h.  </td> </tr> <tr> <td> Hatoum et al. 2006  </td> <td> In vivo. Effect of cumulative radiation doses on rat gut microvessels was studied. Rats were exposed to 1 to 9 cGy in 3 fractions per week on alternate days for 3 successive weeks for a total dose of up to 2250 cGy over a total time of 19 days.    Oxidative stress was measured through superoxide and peroxide levels from various fluorescent markers.    Endothelial function was assessed through ACh vasodilation response.  </td> <td> After 19 days, superoxide was ~1.6-fold elevated and peroxides were ~1.7-fold elevated compared to non-irradiated controls. ROS levels increased at day 5, at the same time as a drop in ACh vasodilation response.     Max dilation dropped from 87% to 3% between day 1 and day 19.  </td> </tr> </tbody> </table>

High Male Low Female Low Unspecific

Moderate

Adult

Low

Not Otherwise Specified

Low Moderate High Low

The evidence is derived from rat in vivo and in vitro models. Mice cell-derived studies were also available but less in-vivo evidence was available from this species. There was a low number of studies containing human or pig models to support this KER. Males have been studied more often than females. There are a few studies with unspecified lifestage of models, while the studies with a defined age typically used adult models.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b42eb168178> AOPWiki 2022-09-28T12:43:32

2023-03-21T09:23:09

<upstream-id>1883f503-acf6-4e69-8704-91f462c360dc</upstream-id> <downstream-id>870809d3-663d-4170-9ec7-b63bbcebbc41</downstream-id> Altered signaling pathways can disrupt cellular homeostasis and induce endothelial dysfunction, characterized by a prolonged state of endothelial activation (Deanfield et al., 2007). Signaling pathways involved in triggering endothelial dysfunction include the p53-p21 pathway, the Akt/phosphtidylinositol-3-kinase (PI3K)/mechanistic target of rapamycin (mTOR) pathway, the RhoA-Rho-kinase pathway, and the acid sphingomyelinase (ASM)/ceramide (Cer) pathway (Venkatsulu et al., 2018; Soloviev et al., 2019; Wang et al., 2016). Activation of the signaling molecule p53 by phosphorylation enhances its stability, leading to cell cycle arrest and premature senescence in endothelial cells and can alternatively lead to a caspase cascade resulting in cellular apoptosis. Activation of the sphingomyelinase ceramide pathway can also contribute to endothelial apoptosis through production of ceramide that activates mitogen-activated protein kinase (MAPK) and extracellular-signal-regulated kinase (ERK). Signaling molecules MAPK and ERK can also be activated as a direct response to a stressor and prompt a cascade of events resulting in endothelial cell apoptosis. Impairment of the Akt/PI3K/mTOR pathway can lead to apoptosis by preventing cell survival signaling and can also lead to downregulation of Rho cytoskeletal proteins for senescence of endothelial cells (Venkatesulu et al., 2018; Soloviev et al., 2019; Nagane et al., 2021; Ramadan et al., 2021; Hughson et al., 2018).

Overall weight of evidence: Moderate

The biological plausibility of the connection between altered signaling pathways leading to an increase in endothelial dysfunction is well-supported by literature and the mechanisms are generally understood. Multiple signaling pathways influence endothelial function.   Modulation of the Akt/PI3K/mTOR pathway downregulates downstream Rho cytoskeletal proteins, which leads to partial-to-full senescence of endothelial cells, resulting in increased vascular permeability and endothelial dysfunction (Venkatsulu et al., 2018). Modulation of the Akt/PI3K/mTOR pathway also acts upstream of the p53-p21 pathway to mediate endothelial cell senescence (Wang et al., 2016). Unlike in other cell types, in endothelial cells the p53-p21 pathway is more important than the p16-Rb pathway for induction of cell senescence (Wang et al., 2016). Senescent endothelial cells show changes in cell morphology, cell-cycle arrest, and increased senescence-associated β-galactosidase (SA-β-gal) staining. These changes lead to endothelial dysfunction, which results in dysregulation of vasodilation (Wang et al., 2016; Hughson et al., 2018; Ramadan et al., 2021). Phosphorylation of p53 is another important moderator of apoptosis in endothelial cells, as well as the ASM/Cer pathway, where the production of ceramide mediates endothelial apoptosis through sphingomyelinase, activating MAPK and ERK, which prompt a cascade of events culminating in endothelial cell apoptosis, another cellular marker for endothelial dysfunction (Venkatsulu et al., 2018; Soloviev et al., 2019).

Empirical Evidence  The empirical evidence supporting this KER was gathered from research utilizing both in vivo and in vitro models. Many in vitro studies have examined the relationship using endothelial cell cultures. Endothelial dysfunction due to altered signaling can be measured by premature endothelial cell senescence, apoptosis and impaired contractile response (Korpela & Liu, 2014). Levels of signaling molecules in pathways such as the Akt/PI3K/mTOR, RhoA-Rho-kinase, and ASM/Cer pathways, and the effect they have on endothelial dysfunction as characterized by endothelial cellular senescence, apoptosis and contractile response, was examined in these studies (Chang et al., 2017; Cheng et al., 2017; Su et al., 2020; Summers et al., 2008; Yentrapalli et al., 2013a; Yentrapalli et al., 2013b).    Dose Concordance  Studies have used in vitro models with acute and chronic doses of radiation administered from 0.23 to 10 Gy to show that the dose at which significant alterations in signaling pathways occurs is concordant with doses at which endothelial dysfunction occurs. Alterations in various signaling pathways, including decreases in the Akt/PI3K/mTOR pathway and increases in the p53-p21 pathway, both occurred at 2.4 and 4 Gy; endothelial cell senescence was found only at 4 Gy (Yentrapalli et al., 2013a). Another study using 4.1 mGy/h chronic gamma irradiation found significant changes in the p53-p21 and ERK signaling pathways after both 2.1 and 4.1 Gy (Yentrapalli et al., 2013b). These changes were associated with an increase in endothelial dysfunction at both doses as well, with only a slight increase in cell senescence after 2.1 Gy (Yentrapalli et al., 2013b). Endothelial cells exposed to a single dose of 10 Gy of X-rays showed an increase in Akt and p-Akt that was correlated with an increase in endothelial dysfunction indicated by a 5-fold increase in apoptosis (Chang et al., 2017).  Various studies also showed that altered signaling and endothelial dysfunction can occur at the same stressor severity during hindlimb unloading (HU). Rats exposed to microgravity for 20 days showed both decreased RhoA signaling molecule and impaired vasodilation (Summers et al., 2008). Two studies observing the effects of microgravity for 4 weeks in rats found that a decrease in the ASM/Cer pathway resulted in better endothelial function, while an increase in the pathway resulted in endothelial dysfunction (Cheng et al., 2017; Su et al., 2020).    Time Concordance  There is limited evidence to suggest a time concordance between altered signaling pathways and endothelial dysfunction. In a study using gamma irradiation in vitro, the p53-p21 and ERK signaling pathways were altered after 3 weeks of chronic gamma irradiation, while cell senescence was increased only slightly after 3 weeks, and increased more after 6 weeks (Yentrapalli et al., 2013b). In a similar study, alterations in various signaling pathways including decreases in the Akt/PI3K/mTOR pathway were found as early as 1 week during chronic gamma irradiation (Yentrapalli et al., 2013a). The earliest significant increase in cellular senescence occurred after 10 weeks (Yentrapalli et al., 2013a).    Incidence Concordance  There is limited support in current literature for an incidence concordance relationship between altered signaling and endothelial dysfunction. One out of the 6 primary research studies used to support this KER demonstrated an average change to endpoints of altered signaling that was greater or equal to that of endothelial dysfunction (Cheng et al., 2017; Yentrapalli et al., 2013b). After 6 weeks of gamma irradiation of HUVECs, altered signaling marker, p21, was increased by 3.5-fold, compared to the 3-fold increase in SA-β-gal staining, a marker for endothelial cell senescence (Yentrapalli et al., 2013b).    Essentiality  Changes in endothelial signaling pathways can trigger endothelial dysfunction. Therefore, in the absence of altered signaling endothelial dysfunction is not expected. Through the use of signaling molecule inhibitors such as Y27632 and dpm, altered signaling can be suppressed which results in reduced endothelial dysfunction (Venkatsulu et al., 2018; Soloviev et al., 2019; Wang et al., 2016). A study observing the effects of Y27632, a Rho kinase inhibitor, found that when rat abdominal aortas were incubated with Y27632 the contractile response, which was hindered by HU, was returned to control levels (Summers et al., 2008). Similar results have been shown by other groups that have looked at endothelial cells incubated in mesenchymal stem cell conditioned media (MSC-CM), which is thought to exhibit therapeutic potential for microvascular injury through angiogenic cytokines. MSC-CM prevented an increase in cleaved capsase-3 and increased both Akt and p-Akt, which was associated with a substantial decrease in apoptosis (Chang et al., 2017). It has also been found that enhancing the ASM/Cer pathway with C6-ceramide increases downstream caspase-3 and endothelial dysfunction, measured by apoptosis, while ASM inhibitors dpm and DOX partially decreased both caspase-3 and apoptosis (Su et al., 2020; Cheng et al., 2017).

<ul> <li>Much of the evidence for this relationship comes from in vitro studies; further work is needed to determine the certainty of the relationship at the tissue level.</li> </ul>

<table border="1"> <tbody> <tr> <td> Modulating factor  </td> <td> Details  </td> <td> Effects on the KER  </td> <td> References  </td> </tr> <tr> <td> Drug  </td> <td> Ceramide-6 (Cer-6)  </td> <td> Increases downstream caspase-3 and apoptosis  </td> <td> (Cheng et al., 2017; Su et al., 2020)  </td> </tr> <tr> <td> Media  </td> <td> Mesenchymal stem cell conditioned media  </td> <td> Prevented an increase in cleaved capsase-3, increased both Akt and p-Akt, and decreased apoptosis  </td> <td> (Chang et al., 2017)  </td> </tr> <tr> <td> Drug  </td> <td> Y27632 (Rho kinase inhibitor)  </td> <td> Recovered contractile response that was attenuated by HU  </td> <td> (Summers et al., 2008)  </td> </tr> <tr> <td> Drug  </td> <td> Desipramine (dpm) (ASM inhibitor)  </td> <td> Partially decreased caspase-3 and apoptosis  </td> <td> (Cheng et al., 2017; Su et al., 2020)  </td> </tr> <tr> <td> Drug  </td> <td> Dopexin hydrochloride (DOX) (ASM inhibitor)  </td> <td> Partially decreased caspase-3 and apoptosis  </td> <td> (Cheng et al., 2017; Su et al., 2020)  </td> </tr> </tbody> </table>

The following are a few examples of quantitative understanding of the relationship. All data that is represented is statistically significant unless otherwise indicated.

Dose/incidence concordance  <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Yentrapalli et al., 2013a    </td> <td> In vitro. Human umbilical vein endothelial cells (HUVECs) were irradiated with 137Cs gamma radiation at 1.4 mGy/h or 2.4 mGy/h dose rates over a period of 12 weeks. Signaling molecules were used as measures of altered signaling. Cell senescence was used as a measure of endothelial dysfunction.  </td> <td> 1.4 mGy/h dose rate: This dose rate led to alterations in the Akt/PI3K/mTOR pathway including a 3-fold increase in p21, a 0.7-fold decrease in p-Akt and PI3K, and a 0.8-fold decrease in mTOR. However, this dose rate did not induce endothelial dysfunction (no significant changes in SA-β-gal staining).  2.4 mGy/h dose rate: This dose rate led to alterations in the Akt/PI3K/mTOR pathway including a 3-fold increase in p21, a 1.5-fold increase in Akt, a 0.7-fold decrease in p-Akt and PI3K, a 0.8-fold decrease in mTOR and a 0.5-fold decrease in p-ERK. This dose rate induced endothelial dysfunction as indicated by a 2-fold increase in SA-β-gal staining, a marker for endothelial cell senescence.  </td> </tr> <tr> <td> Yentrapalli et al., 2013b    </td> <td> In vitro. HUVECs were irradiated with 137Cs gamma radiation at a 4.1 mGy/h dose rate over the course of 6 weeks. Signaling molecules were used as measures of altered signaling. Cell senescence was used as a measure of endothelial dysfunction.  </td> <td> Chronic irradiation led to a 2.5-fold increase in p-p53, a key signaling molecule in initiating endothelial cell senescence, after week 6 (4.13 Gy). There was no significant change in total p53 or p-ERK2 at any time point, while total ERK2 showed a 0.5-fold decrease after 3 weeks (2.07 Gy). p21, which acts downstream of p53, increased 3.5-fold by week 6 (4.13 Gy). Alterations in the above signaling molecules correlated with an increase in endothelial dysfunction as indicated by a 1.5-fold increase in SA-β-gal staining, a marker for endothelial cell senescence, after 3 weeks and 3-fold after 6 weeks (4.13 Gy).  </td> </tr> <tr> <td> Chang et al., 2017    </td> <td> In vitro. HUVECs were irradiated with 10 Gy of X-rays. Signaling molecules were used as measures of altered signaling. Apoptosis was used as a measure of endothelial dysfunction.  </td> <td> A 2-fold increase in cleaved caspase-3, part of the apoptosis signaling cascade, in the irradiated endothelial cells was correlated with an increase in endothelial dysfunction indicated by a 4-fold increase in the percentage of apoptotic cells.  </td> </tr> <tr> <td> Summers et al., 2008   </td> <td> In vivo. Adult male Wistar rats underwent 20-day HU after which abdominal aortas were harvested. Signaling molecules were used as measures of altered signaling. Contractile response was used as a measure of endothelial dysfunction.  </td> <td> Following HU, signaling molecule RhoA decreased by 0.5-fold and endothelial dysfunction was induced in the form of reduced contractile response to phenylephrine by 0.5-fold.  </td> </tr> <tr> <td> Su et al., 2020  </td> <td> In vivo. Sprague-Dawley male rats underwent 4 weeks of HU, following which cerebral and mesenteric arteries were harvested. Signaling molecules were used as measures of altered signaling. Apoptosis was used as a measure of endothelial dysfunction.  </td> <td> Simulated microgravity revealed a decrease in signaling molecules ASM and Cer in cerebral arteries but an increase in mesenteric arteries of rats. Caspase-3, a signaling molecule in the apoptosis cascade, decreased 0.5-fold in cerebral arteries and increased 2-fold in mesenteric arteries. This was associated with an increase in endothelial function in cerebral arteries by decreasing apoptosis 0.3-fold and an increase in endothelial dysfunction in mesenteric arteries by increasing apoptosis 2-fold.  </td> </tr> <tr> <td> Cheng et al., 2017    </td> <td> In vivo. Sprague-Dawley male rats underwent 4 weeks of hindlimb unloading following which carotid arteries were harvested. Signaling molecules were used as measures of altered signaling. Apoptosis was used as a measure of endothelial dysfunction.    </td> <td> 2 weeks of simulated microgravity resulted in a 0.6-fold decrease in signaling molecules ASM and caspase-3, which led to a decrease of endothelial dysfunction indicated by 0.5-fold reduced apoptosis in the carotid arteries of rats.  </td> </tr> </tbody> </table>

Time concordance  <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Yentrapalli et al., 2013a  </td> <td> In vitro. HUVECs were irradiated with 137Cs gamma radiation at 1.4 mGy/h or 2.4 mGy/h dose rates over a period of 10 weeks. Signaling molecules were used as measures of altered signaling. Cell senescence was used as a measure of endothelial dysfunction.  </td> <td> 1.4 mGy/h dose rate: This dose rate led to alterations in the Akt/PI3K/mTOR pathway including a 3-fold increase in p21 after 10 weeks, a 0.7-fold decrease in p-Akt and PI3K after 6 and 10 weeks, and a 0.8-fold decrease in mTOR after 10 weeks. However, this dose rate did not induce endothelial dysfunction (no significant changes in SA-β-gal staining at any timepoint).  2.4 mGy/h dose rate: This dose rate caused alterations in the Akt/PI3K/mTOR pathway including a 3-fold increase in p21 after 10 weeks, a 1.5-fold increase in Akt after 1 week, a 0.7-fold decrease in p-Akt and PI3K after 6 and 10 weeks, a 0.8-fold decrease in mTOR and a 0.5-fold decrease in p-ERK after 10 weeks. This dose rate induced endothelial dysfunction indicated by a 2-fold increase in SA-β-gal staining, after 12 weeks.  </td> </tr> <tr> <td> Yentrapalli et al., 2013b  </td> <td> In vitro. HUVECs were irradiated with 137Cs gamma radiation at a 4.1 mGy/h dose rate over the course of 6 weeks. Signaling molecules were used as measures of altered signaling. Cell senescence was used as a measure of endothelial dysfunction.  </td> <td> Chronic irradiation led to a 2.5-fold increase in p-p53, a key signaling molecule in initiating endothelial cell senescence, after week 6 (4.13 Gy). There was no significant change in total p53 or p-ERK2 at any time point, while total ERK2 showed a 0.5-fold decrease after 3 weeks (2.07 Gy). p21, which acts downstream of p53, increased 3.5-fold by week 6 (4.13 Gy). Alterations in the above signaling molecules correlated with an increase in endothelial dysfunction as indicated by a 1.5-fold increase in SA-β-gal staining after 3 weeks and 3-fold after 6 weeks (4.13 Gy).  </td> </tr> </tbody> </table>

Moderate Male Low Female Low Unspecific

Moderate

Adult

Low

Juvenile

Low Moderate

Evidence for this KER is supported through in vivo rat and in vitro human studies. The in vivo studies were conducted in male animals, although the relationship is still plausible in females. The in vivo studies were undertaken in adolescent and adult rats.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b42eb2c3360> AOPWiki 2022-09-28T12:42:18

2023-03-21T09:20:47

<upstream-id>1329b7b0-4714-406a-a848-17ef92fd2b72</upstream-id> <downstream-id>870809d3-663d-4170-9ec7-b63bbcebbc41</downstream-id> An increase in pro-inflammatory mediators including the cytokines tumor necrosis factor-α (TNF-α), interleukin 1 beta and 6 (IL-1β, IL-6), chemokines monocyte chemoattractant protein 1 (MCP-1) and intercellular adhesion molecule 1 (ICAM-1) can lead to inflammatory response which can disrupt cellular homeostasis and if persistent can lead to eventual endothelial dysfunction (Venkatesulu et al., 2018; Korpela & Liu, 2014). Normally, an inflammatory response provides a protective effect to the endothelium but if prolonged (over months) it can exhaust this protective inflammatory effect, as a result, endothelial cells may become senescent or apoptotic, leading to endothelial dysfunction (Deanfield et al., 2007; Bonetti et al., 2003, Wang et al., 2016; Hughson et al., 2018; Ramadan et al., 2021).

Overall weight of evidence: Moderate

The biological plausibility connecting increased pro-inflammatory mediators to increased endothelial dysfunction is well-supported by literature (Bonetti et al., 2003; Deanfield et al., 2007; Hughson et al., 2018; Ramadan et al., 2021; Wang et al., 2019; Wang et al., 2016), and has been demonstrated in animal studies and human cell models (Shen et al., 2018; Chang et al., 2017; Baselet et al., 2017; Ramadan et al., 2020; Ungvari et al., 2013).   Inflammation can initially provide a protective effect to the endothelium, but chronic inflammation can exhaust this protective inflammatory effect resulting in loss of endothelial integrity and resident cells becoming senescent or apoptotic, leading to endothelial dysfunction (Deanfield et al., 2007; Bonetti et al., 2003). Senescent endothelial cells show changes in cell morphology, cell-cycle arrest, and increased senescence-associated β-galactosidase (SA-β-gal) staining. These changes lead to endothelial dysfunction, which also leads to dysregulation of vasodilation (Wang et al., 2016; Hughson et al., 2018; Ramadan et al., 2021). The inflammatory response is regulated by a balance between pro-inflammatory and anti-inflammatory mediators, and specific cytokine profiles are dependent on parameters of the stressor/exposure/insult (Wang et al., 2019). The pro-inflammatory cytokines TNF-α and IL-1 play a critical role by triggering a cytokine cascade, which initiates an inflammatory response to promote healing and restore tissue function. TNF-α is able to induce apoptotic cell death, which is implicated in endothelial dysfunction. Nuclear factor kappa B (NF-кB) is also activated, which targets multiple genes coding for vascular cell adhesion proteins (VCAM), intercellular adhesion molecule (ICAM), and IL-1, as well as prothrombotic markers (Slezak et al., 2017). NF-кB mediates a pro-survival and pro-inflammatory state. Inflammation persisting for months leads to prolonged chronic inflammation, which causes an ineffective healing process that is further worsened by a decrease in endothelium-dependent relaxation. This causes endothelial dysfunction, making vasculature more vulnerable to damage from non-laminar flow (Sylvester et al., 2018). Senescent cells also have a pro-inflammatory secretory phenotype, which further contributes to negative effects on the endothelium. Increased pro-inflammatory mediators may be due to increased expression but may also be attributed to increased permeability of the endothelium as seen after irradiation in animal models, which results in increased transmigration of inflammatory cells into the endothelium and can lead to eventual dysfunction (Hughson et al., 2018).

The empirical evidence supporting this KER is gathered from research utilizing both in vivo and in vitro models. Many in vitro studies have examined the relationship using endothelial cell cultures. Levels of pro-inflammatory mediators such as TNF-α, IL-1β, IL-6, IL-8, MCP-1 and ICAM-1, and the effect they have on endothelial dysfunction, as characterized by endothelial cellular senescence and apoptosis, have been examined in these studies. The evidence is derived from stressors of gamma and X-ray radiation in the range of 0.05-18 Gy (Shen et al., 2018; Baselet et al., 2017; Ramadan et al., 2020; Ungvari et al., 2013; Chang et al., 2017).  Dose Concordance  There is moderate evidence to demonstrate dose concordance between an increase in pro-inflammatory mediators and endothelial dysfunction. Most studies do not show statistically significant effects across all doses; however, biological trends across multiple doses have been included to support this relationship.   Studies examining a range of doses from 0.05 Gy to 18 Gy using predominantly X-rays as the source of stressor support the relationship between increased pro-inflammatory mediators and endothelial dysfunction. An in vitro study using X-ray irradiation on human endothelial cells showed an increase in pro-inflammatory cytokines IL-6 and CCL2 at a dose as low as 0.05 Gy. Though the increase was not statistically significant, there was a biological trend showing significant increases at higher doses. All doses demonstrated a correlation to endothelial dysfunction (Baselet et al., 2017). Another study using human endothelial cells exposed to X-rays showed similar results with increases in pro-inflammatory mediators, including IL-6, VCAM-1, and IL-8, as well as SA-β-gal activity at 5 Gy (Ramadan et al., 2020).  A gamma ray study that exposed rat endothelial cells to a 6 Gy dose showed an increase in pro-inflammatory mediators IL-6, IL-1α, IL-1β, and MCP-1 associated with an increase in endothelial cell senescence (Ungvari et al., 2013). Another single dose X-ray study at 10 Gy also revealed increases in pro-inflammatory mediators with a 1.2-fold increase in IL-1α, and a 6-fold increase in IL-6 and TNF-α. This was associated with an increase in endothelial dysfunction, indicated by a 5-fold increase in apoptotic cells (Chang et al., 2017). A study using X-rays on mouse aortas found that there was a 2-fold increase in the pro-inflammatory mediators TNF-α and ICAM-1 after 18 Gy, and a 5-fold increase in endothelial apoptosis, which is a defined marker for endothelial dysfunction (Shen et al., 2018).  Time Concordance  There is limited evidence to suggest a time concordance between increased pro-inflammatory mediators and endothelial dysfunction. An in vitro study using 5 Gy of X-rays found that pro-inflammatory mediators, including IL-6, VCAM-1, TNF-α, ICAM-1, IL-1β and MCP-1, increased as soon as 1 day post-irradiation. SA-β-gal, a marker for cellular senescence, showed the first increase 7 days post-irradiation (Ramadan et al., 2020). A study using 18 Gy of X-rays examined mouse aortas after 3-84 days post-irradiation and found an increase in both pro-inflammatory mediators and endothelial dysfunction as early as 3 days post-irradiation (Shen et al., 2018).  Incidence concordance  Few studies demonstrated incidence concordance. In an in vitro study using human endothelial cells irradiated with X-rays, incidence concordance was demonstrated at both 0.5 and 2 Gy as pro-inflammatory mediators IL-6 and CCL2 were increased 2-fold or greater while SA-β-gal increased a maximum of 1.5-fold (Baselet et al., 2017). Similarly, 5 Gy of X-rays resulted in increases to multiple pro-inflammatory mediators (IL-1β, IL-6, IL-8, MCP-1, and VCAM) between 1.5- and 4-fold, while SA-β-gal activity increased 1.5-fold in human endothelial cells (Ramadan et al., 2020).  Essentiality  An increase in inflammation can trigger endothelial dysfunction. Therefore, in the absence of an increase in pro-inflammatory mediators endothelial dysfunction is not expected. Through the use of certain treatments, such as TAT-Gap19 and mesenchymal stem cells, the increase in pro-inflammatory mediators can be greatly supressed, but not fully blocked, which results in reduced but not completely prevented endothelial dysfunction such as apoptosis and cellular senescence (Venkatsulu et al., 2018; Soloviev et al., 2019; Wang et al., 2016). These treatments demonstrate the essentiality of the relationship and are described below; however, the available empirical data supporting essentiality for this KER is limited.   A study observing the effects of TAT-Gap19, a connexin43 hemichannel blocker, found the increase in pro-inflammatory mediators seen following the stressor was largely, though not fully in all mediators, prevented. This was also associated with a decrease in SA-β-gal, a marker of endothelial cell senescence and dysfunction, compared to the irradiated group (Ramadan et al., 2020). Similar results have been shown by other groups that have examined human endothelial cells incubated in mesenchymal stem cell conditioned media (MSC-CM), which is thought to exhibit therapeutic potential for microvascular injury through angiogenic cytokines. This study revealed a significant but not complete prevention of pro-inflammatory mediators IL-1α, IL-6 and TNF-α. The same pattern was seen in endothelial dysfunction, where apoptosis was significantly prevented but was still slightly above control levels (Chang et al., 2017).

<ul> <li> Much of the evidence for this relationship comes from in vitro studies; further work is needed to determine the certainty of the relationship at the tissue level.  </li> <li> Although studies often measure pro-inflammatory mediators at a few specific time points, chronic inflammation is what contributes to endothelial dysfunction. More human studies should examine the temporal concordance of this relationship to identify whether the inflammation is chronic.  </li> </ul>

The following are a few examples of quantitative understanding of the relationship. All data that is represented is statistically significant unless otherwise indicated.

Pro-inflammatory mediators can induce endothelial cell senescence and subsequent endothelial dysfunction. Senescent endothelial cells can secrete many pro-inflammatory cytokines and chemokines, contributing to further senescence of other endothelial cells and further endothelial dysfunction (Hughson et al., 2018; Wang et al., 2016).

Moderate Male Low Female Low Unspecific

Low

Adult

Moderate

Juvenile

Low Moderate Low

The majority of the evidence is derived from in vitro studies, and a single in vivo study in male pre-adolescent mice.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b42eb3eeac8> AOPWiki 2022-09-28T12:43:54

2023-03-21T10:39:07

<upstream-id>870809d3-663d-4170-9ec7-b63bbcebbc41</upstream-id> <downstream-id>c79fec82-3db8-4eaf-b5ef-0fdcffb5dfe8</downstream-id> Proper endothelial activation is a key step in the growth of new vessels through the process of angiogenesis, a process also affected by a dysfunctional endothelium (Rajashekhar et al., 2006). Regional responses to stressors are also possible, with mechanical stressors differentially affecting pressure in vessels above (superior to) and below (inferior to) the heart (Hargens & Watenpaugh, 1996; Zhang, 2001). The endothelial layer is responsive to these variations in mechanical stresses and can adapt through altering the balance between hypertrophic and hypotrophic remodeling in smooth vessel cells lining vasculature (Baeyens et al., 2016) in part through altering the progression of the acid sphingomyelinase (ASM)/ceramide (Cer) pathway (Cheng et al., 2017).

Overall weight of evidence: Moderate

The relationship between endothelial function and vascular remodeling is well supported through a number of in-depth reviews about the mechanisms behind the connection. In a functional endothelial layer, the endothelial cells both contribute and react to the high levels of bioavailable nitric oxide (NO). The result is a vasomotive balance primed for vasodilation and an elevated ratio of antioxidant to pro-oxidant species (Deanfield et al., 2007). Endothelial cells can become activated through various signals, including vascular endothelial growth factor (VEGF), that subsequently induce angiogenesis (Carmeliet & Jain, 2011). Lastly, the endothelial cells form tight junctions and together with pericytes form a basement membrane in tight control of vessel and cell permeability (Carmeliet & Jain, 2011).     Initial endothelial tissue injury can lead to premature cellular senescence resulting in endothelial dysfunction (Hughson et al., 2018). Cell death in the vessels can lead to overall cell loss and reduced vascular density. Although recovery in the form of revascularization following the decrease in vascular density occurs, it is complicated in cases of continuous exposure as it negatively impacts the angiogenic process (Hughson et al., 2018). Stressors such as radiation are thought to disturb angiogenesis through decreasing VEGF secretion causing a decrease in tubule formation (Sylvester et al., 2018). Following the initial tissue injury, the body’s ability to heal is also compromised, in part due to the prolonged state of oxidative stress and simultaneous decrease in endothelium-dependent vasorelaxation leading to increase in non-laminar blood flow. To compensate for the damage caused by this turbulent flow, there is intima-media thickening and potential for eventual atherosclerosis (Bonetti et al., 2003; Hughson et al., 2018; Slezak et al., 2017; Sylvester et al., 2018). Continued injury also leaves the vessels vulnerable to maladaptive repair and ensuing fibrosis (Hsu et al., 2019).    When activated within healthy limits, endothelial cells loosen at their junction and the presence of VEGF induces increased vessel permeability allowing for the vessels to expand and undergo angiogenesis (Carmeliet & Jain, 2011). However, prolonged increase in permeability is also a marker of dysfunction, where increases in adhesion proteins, and elevated levels of cell senescence accompany this change (Demontis et al., 2017; Hughson et al., 2018). Disruption of endothelial integrity can also lead to cell detachment from the basement membrane; with cardiovascular deconditioning following bedrest leading to significantly elevated levels of circulating endothelial cells in microcirculation (Zhang, 2013). In addition to vessel permeability, there are changes to permeability of endothelial cells themselves, which is shown to increase and not recover following removal of the stressor (Baran et al., 2021).     The elevated bioavailability of NO associated with proper endothelial function correlates with a decrease in prothrombotic factors, while a dysfunction in the endothelium creates a pro-thrombotic environment (Krüger-Genge et al., 2019). This is also true in the case of certain kinds of vascular damage, where a dysfunctional endothelial layer following stressor exposure has been shown to lead to lymphocyte adhesion and thrombus formation. This pro-thrombotic environment causes vessel occlusion and a decrease in capillary and vascular density, which in turn results in increased vascular resistance requiring further vascular remodeling as compensation (Slezak et al., 2017). Cell senescence following damage also induces monocyte adhesion and can further contribute to creating a pro-atherosclerotic environment (Hughson et al., 2018).    There is also evidence that the relationship between endothelial function and vascular remodeling is regionally affected following the exposure to localized varied mechanical stressors such as microgravity. Under microgravity conditions (and in simulated microgravity models such as hindlimb unloading (HU)), there is a cephalic shift in blood and fluid resulting in a change of transmural pressure, increasing pressure in vessels above the heart and decreasing in those below (Hargens & Watenpaugh, 1996; Zhang, 2001). The heart continues to pump as usual; however, above the heart the arterial flow is no longer pushing against gravity resulting in increased arterial vascular pressure, while venous return is simultaneously slowed without gravitational assistance. This results in changes such distended veins and arteries in the upper body, increased carotid intima-media thickness and vascular stiffness (Garrett-Bakelman et al., 2019). In the lower limb, the opposite is true as arterial perfusion is decreased and venous return is increased resulting in muscle atrophy. Blood pressure and related fluid shear stress act as important mechanical input for the mechanosensing endothelial cells, which translate these forces into biochemical signals that guide vascular remodeling through affecting the balance between vascular smooth muscle cell proliferation and apoptosis (Baeyens et al., 2016). Above the heart, remodeling presents as hypertrophy and increase in vasoreactivity, while below the heart there is hypotrophy and decrease in myogenic tone and vasoreactivity (Zhang, 2013). This trend has been observed by various reviews and studies; a review examining studies using HU rats summarized that the models studied showed both a decrease in response to drugs inducing vasodilation and constriction and a subsequent increased stiffness in the aorta and carotid arteries (Platts et al., 2014). In humans, bedrest study participants showed both a decrease in endothelium-dependent vasodilation and an increase in circulating endothelial cells – both markers of endothelial dysfunction. Additional bedrest studies also show a decrease in vessel diameter and intimal-medial thickness in arteries below the heart while those above the heart remain unaffected (Zhang, 2013). Ultrasound measurements of cosmonauts having travelled aboard Mir and Salyut-7 showed that after spaceflight, blood supply to the brain remained stable while below the heart vascular tone and arterial resistance was severely compromised (Zhang, 2013).     Research comparing the changes in vascular structure and vasodilation response between the various muscle resistance arteries following HU also showed changes to regionally vary between the vessels studied (Delp et al., 2000; Zhang, 2013). Additionally, HU models showed differences in the arterial response to vasoconstrictors and changes to artery diameter in cutaneous versus skeletal muscle arteries (Tarasova et al., 2020). The ASM and Cer pathway has been investigated for its role in remodeling. The work of Cheng et al. (2017) and Su et al. (2020) both found that a decrease in ASM activity and subsequent Cer production was linked to a decrease in apoptosis levels and a resulting thickening of vessel structure (Cheng et al., 2017; Su et al., 2020). It is important to note that some of the vascular changes following microgravity are protective adaptations that serve to safeguard the cardiovascular system in altered gravity conditions. Under continued microgravity conditions, these changes maintain their protective purpose and are thought not to contribute to adverse outcome progression. The problem arises upon return to earth when the vessels that have adapted to microgravity blood distribution are faced with earth conditions and issues like a decreased orthostatic tolerance surface.

Dose Concordance  There is some evidence in the literature supporting dose concordance between endothelial dysfunction and vascular remodeling. For example, gamma irradiation at 0.5 Gy led to a 9% decrease in endothelium-dependent vasodilation and no significant changes to vascular stiffness, while a 1 Gy dose led to 13% decrease in vasodilation corresponding to a 16% increase in vascular stiffness (Soucy et al., 2011). A dose of 5 Gy gamma rays significantly attenuated endothelium-dependent vasodilation, while simultaneously increasing vascular stiffness compared to a non-irradiated control (Soucy et al., 2010). Work exploring apoptosis as a measure of endothelial dysfunction demonstrated that an 18 Gy dose of X-rays increased the number of cells with apoptotic DNA fragmentation ~4.5-fold and increased aortic thickness ~1.5 fold compared to control (Shen et al., 2018).    Studies in rat models of HU by Su et al. (2020), Delp et al. (2000), and Cheng et al. (2017) all demonstrate the regional effects of changes in pressure on resulting vascular adaptation and remodeling. Su et al. (2020) compared the effects of 4-week unloading on the cerebral versus mesenteric artery, showing the balance moving towards cell proliferation above the heart (cerebral artery) with a decrease in apoptosis and increase in intima-media thickness and cross-sectional area. Meanwhile, below the heart (mesenteric artery) apoptosis increased and intima-media thickness and cross-sectional area decreased (Su et al., 2020). This agrees with a similar study that found apoptosis decreased and intima-media thickness of the carotid artery increased following HU (Cheng et al., 2017). Delp et al. (2000) showed that regional adaptations to changes in pressure following HU are also affected by how this change in pressure manifests. Vascular structure changes and endothelium-dependent vasodilation were observed in the gastrocnemius and soleus primary arterioles. Both vessels are in the hindlimbs of mice and therefore are subject to a decrease in pressure following cephalic shift in fluid in 2-week HU. In the gastrocnemius muscle, which saw a decrease in transmural pressure, the decrease in vessel cross sectional area (CSA) was due to muscle atrophy shown by a drop in media thickness but no change in outer media perimeter. In contrast, the soleus muscle experienced a drop in wall shear stress and saw a drop in vessel perimeter with no change in media thickness. Simultaneously, arterioles in the gastrocnemius muscle saw no change in acetylcholine (ACh) response, while those in soleus muscles saw a 50% decrease following the unloading and recovery to control levels at the 4-week time point (Delp et al., 2000).      Time Concordance  There is limited evidence supporting the time concordance of endothelial dysfunction and vascular remodeling. Aortic relaxation response to ACh in Sprague-Dawley rats was found to decrease 20-30% 2 weeks after gamma irradiation with an increase in vascular stiffness measured by pulse-wave velocity (PWV) from around 3.9 m/s to 4.9 m/s (Soucy et al., 2010; Soucy et al., 2007). Shen et al. (2018) demonstrated that endothelial dysfunction assessed via apoptosis became significant 3 days after 18 Gy X-rays in a mouse model, then had a linear decrease which tapered off by day 84. Aortic wall thickness, in turn, showed no significant increase on day 3 post-irradiation, only reaching maximal increase on day 7 before also decreasing linearly to day 84 (Shen et al., 2018). At 4 and 8 months post- 56Fe-ion irradiation, Wistar rats had 13% and 16% decreased endothelial relaxation response respectively. As well, PWV increased from 4.03 m/s to 4.45 m/s at 4 months and 4.53 m/s to 5.06 m/s at 8 months post-irradiation (Soucy et al., 2011).    Incidence concordance  Incidence concordance is moderate in this KER, as multiple studies demonstrate greater changes to endothelial dysfunction than to vascular remodeling. Three studies by Soucy et al. (2011, 2010, 2007) in rats demonstrate greater changes to vasodilation than to vascular stiffness after 5 Gy of gamma rays or 1 Gy of iron ions. In addition, 18 Gy X-ray irradiation of mice showed a 4.5-fold increase in apoptosis and a 1.4-fold increase in aortic thickness (Shen et al., 2018). Following 1 or 4 weeks of HU, greater changes to apoptosis were observed compared to changes in vascular remodeling in the cerebral and small mesenteric arteries of rats (Su et al., 2020).    Essentiality  Human bone marrow-derived mesenchymal stem cells (hBMSCs) can prevent endothelial dysfunction and vascular remodeling via their antioxidant and anti-inflammatory properties. While a radiation dose of 18 Gy X-rays in a mouse model increased both the amount of apoptosis in the aorta (indicating endothelial dysfunction) and aortic wall thickness, treatment with hBMSCs reversed these changes. TUNEL positive cells decreased but remained elevated above the control, while aortic wall thickness returned to control levels (Shen et al., 2018).    In work exploring the role of the ASM/Cer pathway, a decrease in the ASM activity and resulting Cer production corresponded to a decrease in apoptosis and increase in cell-proliferation in rat models of simulated microgravity (Cheng et al., 2017; Su et al., 2020). Incubation with permeable Cer (C6-Cer) returned apoptosis to control levels. Treatment with the ASM inhibitor desipramine (dpm) led to an overall decrease in apoptosis in all arteries tested, while treatment with doxepin hydrochloride (DOX) led to significant increases in cell proliferation and subsequent intima medial thickness (IMT) and cross-sectional areas (CSA) (Su et al., 2020).

<ul> <li> Lower doses (0.5 Gy and 1.6 Gy) did not show changes in vasomotion compared to control, but vascular stiffness increased at these doses (Soucy et al., 2007).   </li> <li> Tarasova et al. (2020) showed differences in the vascular remodeling and vasoconstriction responses between skeletal and cutaneous arteries. While the two groups demonstrated differences, all vessels followed different trends showing no clear relationship between KEs.   </li> <li> Studies exploring vasoreactivity, vascular structure and vessel stiffness endpoints in humans (Lee et al., 2020) and mice (Sofronova et al., 2015) flown in space, found changes in these endpoints to be inconsistent and/or changes were not statistically significant.   </li> <li> C6-Cer incubation in cerebral arteries showed increased apoptosis with HU in the study by Cheng et al. (2017); however, in Su et al. (2020), there was a slight decrease in apoptosis, measured by TUNEL.   </li> </ul>

<table border="1"> <tbody> <tr> <td> Modulating factor   </td> <td> Details   </td> <td> Effects on the KER   </td> <td> References   </td> </tr> <tr> <td> Drug  </td> <td> Oxp (xanthine oxidase inhibitor)  </td> <td> Treatment with Oxp after irradiation led to increased vasodilation and decreased PWV  </td> <td> Soucy et al., 2007; Soucy et al., 2010; Soucy et al., 2011  </td> </tr> <tr> <td> Drug  </td> <td> hBMSCs (protect against vascular damage)  </td> <td> Treatment with hBMSCs after irradiation led to decreased apoptosis and aortic thickness  </td> <td> Shen et al., 2018  </td> </tr> <tr> <td> Drug  </td> <td> C6-Cer (activates the ASM/Cer pathway)  </td> <td> Treatment with C6-Cer after microgravity caused increased apoptosis along with a return of proliferation to control levels  </td> <td> Cheng et al., 2017; Su et al., 2020  </td> </tr> <tr> <td> Drug  </td> <td> dpm (ASM inhibitor)  </td> <td> dpm treatment showed apoptosis and proliferation levels returned to control after microgravity  </td> <td> Su et al., 2020  </td> </tr> <tr> <td> Drug  </td> <td> DOX (ASM inhibitor)  </td> <td> Treatment with DOX after microgravity showed decreased apoptosis and increased IMT in rat carotid arteries  </td> <td> Su et al., 2020  </td> </tr> </tbody> </table>

The following are a few examples of quantitative understanding of the relationship. All data that is represented is statistically significant unless otherwise indicated.

Dose/Incidence Concordance  <table border="1"> <tbody> <tr> <td> Reference  </td> <td> Experiment Description  </td> <td> Result  </td> </tr> <tr> <td> Soucy et al., 2007   </td> <td> In vivo. Sprague-Dawley rats were whole-body irradiated with 137Cs gamma radiation at 0.5 Gy, 1.6 Gy and 5 Gy. Vasodilation response to ACh was used to evaluate endothelial function. Vascular stiffness was measured by PWV.  </td> <td> No changes in endothelial relaxation were observed after 0.5 or 1.6 Gy, but relaxation decreased about 20 percentage points at 5 Gy. At 0.5 and 1.6 Gy, PWV increased from 3.9 m/s (before irradiation) to 4.2 m/s. At 5 Gy PWV increased to 4.6 m/s.  </td> </tr> <tr> <td> Soucy et al., 2010   </td> <td> In vivo. Sprague-Dawley rats were whole-body irradiated with 137Cs gamma radiation at 5 Gy. Aortic relaxation response to ACh and PWV were measured.  </td> <td> Relaxation decreased about 30 percentage points after 5 Gy. PWV increased to 4.93 m/s from 4.06 m/s (control) after 5 Gy.  </td> </tr> <tr> <td> Soucy et al., 2011  </td> <td> In vivo. Wistar rats were exposed to 0.5 and 1 Gy doses of 56Fe-ion radiation. ACh-induced vasodilation response was measured. PWV was measured with Doppler probe and electrocardiogram (ECG) while aortic wall thickness:lumen diameter ratio was measured by histological analysis.  </td> <td> 0.5 Gy dose – 9% (non-significant) decrease in endothelium-dependent vasodilation and no significant change in PWV.  1 Gy dose - 13% decrease in endothelium-dependent vasodilation and PWV increased by 0.42 m/s.  Neither dose showed changes to aortic wall thickness:lumen diameter.  </td> </tr> <tr> <td> Shen et al., 2018   </td> <td> In vivo. Male mice were irradiated with 18 Gy X-rays. Endothelial dysfunction was determined through a TUNEL apoptosis assay. To measure vascular remodeling, aortic thickness was determined at various times using hematoxylin and eosin (HE) staining, and the accumulation of collagen was measured using Sirius red staining.  </td> <td> Irradiation with 18 Gy caused a ~4.5-fold maximum increase in TUNEL positive cells and a maximum increase of 1.4-fold in aortic thickness and collagen content compared to controls.  </td> </tr> <tr> <td> Su et al., 2020   </td> <td> In vivo. The effects of simulated microgravity by 0 day, 3 day, 1 week, 2 week or 4 week HU on cerebral and small mesenteric rat arteries were studied. Apoptosis was used as a measure of endothelial dysfunction by TUNEL assay and IMT and media CSA were used as measures of vascular remodeling.  </td> <td> Significant changes occurred in small mesenteric artery following 1 week of HU with a ~2-fold increase in apoptosis, ~0.8-fold decrease in IMT and ~0.6-fold decrease in CSA.     Rat cerebral artery exhibited significant changes following 4 weeks of HU at which point there was a 0.3-fold decrease in apoptosis, a ~2-fold increase in IMT, and a ~2.1-fold increase in CSA.      </td> </tr> <tr> <td> Delp et al., 2000   </td> <td> In vivo. Male Sprague-Dawley rats had 2 arteries and 2 arterioles analyzed after 2-week HU. Endothelial dysfunction was measured through the relaxation response to ACh and remodeling was determined through media cross-sectional area, wall thickness and perimeter.  </td> <td> 2-week HU led to the following changes:   Soleus muscle feed artery –   Cross sectional area decreased 0.5-fold, media thickness did not change, outer-media perimeter decreased 0.7-fold.  Response to ACh (i.e. endothelium-dependent vasodilation) in arterioles decreased 50% after 2-week HU, but no change was observed after 4-week HU.    Gastrocnemius muscle feed artery –   Cross sectional area decreased 0.5-fold, media thickness decreased 0.6-fold and outer-media perimeter did not significantly change.  Response to ACh (i.e. endothelium-dependent vasodilation) in arterioles did not significantly change.  </td> </tr> <tr> <td> Tarasova et al., 2020  </td> <td> In vivo. Male Wistar rats’ skin and skeletal muscle arteries were analyzed after 2-week HU. Endothelial dysfunction was determined through contractile response to noradrenaline and serotonin vasoconstrictors, and remodeling was determined through vessel inner diameter.  </td> <td> Following 2-week HU, the following changes occurred:   Forelimb arteries -   Brachial artery: Inner artery diameter increased 22.5%, active tension response to noradrenaline increased ~2-fold while response to serotonin increased ~1.5-fold.    Median artery: Inner artery diameter increased 10%, no significant changes in noradrenaline or serotonin responses.   Hindlimb arteries -   Sural artery: Inner artery diameter decreased 16.8%, ~0.7-fold decrease in active tension response for both noradrenaline and serotonin.   Saphenous artery: Inner artery diameter showed no significant change, and active tension response following noradrenaline and serotonin was elevated above the control but this increase was not significant across all doses studied.   </td> </tr> <tr> <td> Cheng et al., 2017   </td> <td> In vivo. Male rat carotid arteries were studied with HU. Apoptosis, measured by a TUNEL assay, was used as a measure of endothelial dysfunction. IMT and CSA were markers of vascular remodeling using HE staining.   </td> <td> Following HU, IMT in the carotid artery increased 1.8-fold and intima-media area increased 2.1-fold. Apoptosis decreased by 0.5-fold after HU.  </td> </tr> <tr> <td> Lee et al., 2020  </td> <td> In vivo. Ten male and three female astronauts who participated in various durations of spaceflight (189 ± 61 days). Irradiation measurements showed the crew experienced an average of 0.048 ± 0.018 Gy from 0.031 to 0.077 Gy. Endothelial dysfunction was measured through flow-mediated vasodilation of the brachial artery. Vascular remodeling was measured through intima-media area and vascular stiffness.  </td> <td> No changes in endothelium-dependent or -independent vasodilation were observed from preflight to postflight. From preflight to postflight, intima-media area increased by 1.04 mm2 and stiffness increased by 4.4 arbitrary units. However, none of the changes were significant.  </td> </tr> <tr> <td> Sofronova et al., 2015  </td> <td> In vivo. Male mice were placed under microgravity environment to study the properties of cerebral arteries, including endothelial dysfunction measured by vascular tension, vessels response to ACh (vasodilator) and vascular remodeling determined by elastin-collagen content using staining tissue with Verhoeff-van Gieson.  </td> <td> Spaceflight mice (SF) showed a 30% decrease in relaxation response of basilar arteries to ACh compared to habitat control (HC) group.    No significant changes in elastin or elastin-collagen ratio were observed between HC, vivarium control (VC) and SF mice. There was a 5% collagen content increase in VC mice compared to HC and SF groups.  </td> </tr> </tbody> </table>

High Male Moderate Female

Moderate

Adult

Low

Juvenile

Moderate

Not Otherwise Specified

Moderate Moderate High

There is a substantial amount of evidence for this KER from in vivo rodent models and from human studies. The sex applicability is high for males and moderate for females as many studies were done only using male animals. Most studies indicated that the animals used were adult.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b42ebe779b8> AOPWiki 2022-10-17T16:54:17

2023-03-21T09:43:16

<upstream-id>1cc566f9-d2e5-4c9c-a06c-c40f7767c251</upstream-id> <downstream-id>870809d3-663d-4170-9ec7-b63bbcebbc41</downstream-id> Altered nitric oxide (NO) levels can lead to endothelial dysfunction (Soloviev & Kizub, 2019).  In a functional endothelium, NO is bioavailable and is involved in preventing inflammation, proliferation and thrombosis (Deanfield, Halcox & Rabelink, 2007; Kruger-Genge et al., 2019). An increase in reactive oxygen species (ROS) along with increased NO can drive cellular senescence in endothelial cells (ECs) and catalyze endothelial dysfunction (Nagane et al., 2021; Wang, Boerma & Zhou, 2016). Another driver of endothelial dysfunction is reduced vasomotion. In a functional state, the endothelium requires a balance of vasoconstrictors and vasodilators (like NO); an interruption of this balance can lead to dysfunction (Deanfield, Halcox & Rabelink, 2007; Marti et al., 2012; Nagane et al., 2021; Schulz, Gori & Münzel, 2011; Soloviev & Kizub, 2019). Decreased NO due to direct reactions with ROS or uncoupling of NOS enzymes will lead to a reduced ability of smooth muscle cells (SMCs) to relax (Soloviev & Kizub, 2019).

Overall weight of evidence: Moderate

The biological plausibility surrounding the connection between altered NO levels leading to endothelial dysfunction is well-supported by literature. NO is synthesized from L-arginine and oxygen with the aid of enzymes and cofactors (Nagane et al., 2021). NO regulates ECs by binding to soluble guanylyl cyclase (sGC) to create cGMP and activate protein kinase G (PKG), leading to activation of Ca2+-dependent vasodilation and smooth muscle relaxation (Nagane et al., 2021; Soloviev & Kizub 2019). NOS isoforms, such as inducible NO synthase (iNOS) and endothelial NO synthase (eNOS) that synthesize NO are indirect measures of NO. Lower NO reduces the ability of SMCs relaxation and dilates the blood vessel leading to an inability to control vasodilation, a component of endothelial dysfunction (Soloviev & Kizub 2019).  Increased expression of NOS enzymes can result in reduced NO levels in the case of insufficient L-arginine substrate or BH4 cofactor leading to ROS production instead of NO (Zhang et al., 2009). ROS can decrease NO bioavailability by uncoupling/downregulating eNOS or converting NO to peroxynitrite (Mitchel et al 2019; Schulz, Gori & Münzel, 2011; Soloviev & Kizub 2019; Wang, Boerma & Zhou, 2016). A further decrease in NO occurs as peroxynitrite oxidizes BH4 to BH2 and induces eNOS to produce ROS, continuing the uncoupling of more NOS enzymes (Hong et al., 2013; Soloviev & Kizub, 2019; Zhang et al., 2009). A reduction in NO bioavailability due to ROS is an important mediator of endothelial dysfunction (Schulz, Gori & Münzel, 2011).  Another component of endothelial dysfunction influenced by NO levels is cellular senescence (Nagane et al., 2021). iNOS expression increases following an increase in oxidative stress (Nathan & Xie, 1994). In the presence of oxidative stress, NO is converted to peroxynitrite, which is a reactive nitrogen species (RNS) that can modify proteins and lead to cellular senescence (Hong et al., 2013; Nagane et al., 2021; Soloviev & Kizub, 2019). Although NO can increase at first and cause cell senescence, senescent ECs show downregulation and/or uncoupling of eNOS that contributes to a decrease of NO in the endothelium (Wang, Boerma & Zhou, 2016). These changes in senescent ECs lead to endothelial dysfunction.

The empirical evidence to support this KER was gathered from research that utilized in vivo rat and rabbit models (Soucy et al., 2010; Soucy et al., 2011; Hong et al., 2013; Yan et al., 2020; Zhang et al., 2009), ex vivo rabbit models and in vitro HUVEC models (Hong et al., 2013). Stressors used to alter NO levels and increase endothelial dysfunction include 56Fe ions (Soucy et al., 2011), X-rays (Hong et al., 2013; Yan et al., 2020), gamma rays (Soucy et al., 2010) and altered gravity by hindlimb unweighting (HU) (Zhang et al., 2009). Irradiation dose levels ranged from 0.5 Gy to 16 Gy (Hong et al., 2013; Soucy et al., 2010; Soucy et al., 2011; Yan et al., 2020). Studies used various endpoints to measure NO levels while endothelial function was consistently determined through relaxation response to acetylcholine (ACh). Methods to measure NO included DAF-FM DA fluorescent probe (Soucy et al., 2010; Soucy et al., 2011), Griess reagent NO assay kit (Yan et al., 2020), eNOS dimer to monomer ratio (Yan et al., 2020), and NOS protein and mRNA levels (Hong et al., 2013; Zhang et al., 2009).   Dose Concordance  There is moderate evidence to demonstrate dose concordance between a decrease in nitric oxide and endothelial dysfunction. 56Fe ion irradiation in rat aorta showed a decrease in both NO levels and endothelial relaxation at 1 Gy. However, endothelial relaxation did not significantly decrease at a lower dose of 0.5 Gy (Soucy et al., 2011). After 5 Gy gamma irradiation, NO production and endothelial relaxation both decreased 0.7-fold in rat aorta (Soucy et al., 2010). Increased NOS levels can cause further NO decreases due to uncoupling and production of more peroxynitrite, which is the result of NO reacting with ROS and can therefore, be used to determine NO levels (Hong et al., 2013; Zhang et al., 2009). In rat mesenteric arteries, NO levels decreased 0.6-fold after 4 Gy X-ray irradiation, while endothelial relaxation decreased 0.1-fold compared to non-irradiated arteries (Yan et al., 2020). X-ray irradiated human umbilical vein endothelial cells (HUVECs) showed significantly increased iNOS and peroxynitrite after 4 Gy, and X-ray irradiated rabbit carotid arteries had decreased relaxation after 8 and 16 Gy (Hong et al., 2013). Hong et al (2013) also showed that endothelial relaxation was lower after 16 Gy than 8 Gy ex vivo.   Altered gravity resulted in an increase of eNOS and increase of iNOS in carotid arteries (Zhang et al., 2009). Increases in eNOS and iNOS corresponded to a decrease in carotid artery relaxation from 64% to 33% at the same level of HU (Zhang et al., 2009). These studies showed increases in NOS isoforms and corresponding decreases in endothelial function, such as increased vasoconstriction and impaired vasodilation following altered gravity.    Time Concordance  Evidence of time concordance between altered NO levels and endothelial dysfunction is limited from the studies cited. HUVECs irradiated with 4 Gy X-rays displayed an increase in nitrotyrosine (peroxynitrite biomarker indicating reduced NO) after 6 hours post-irradiation (Hong et al., 2013). Rabbit carotid arteries in vivo and ex vivo irradiated with 8 or 16 Gy X-rays showed decreased ACh-induced endothelial relaxation only after 20 hours post-irradiation (Hong et al., 2013).   Incidence Concordance   There is limited support in current literature for an incidence concordance relationship between altered NO and endothelial dysfunction. A primary research study that supports this AOP demonstrated an average change to endpoints of altered NO that was greater or equal to that of endothelial dysfunction (Zhang et al., 2009).   Essentiality  Many studies show the essentiality of decreased NO levels in endothelial dysfunction. After stressors like irradiation, xanthine oxidase (XO) can produce cardiac ROS that can react with NO and decrease its concentration or oxidize BH4 and uncouple eNOS. Oxypurinol (Oxp), an inhibitor of XO, has been shown to reverse these effects and reduce ROS levels, restore NO levels, and increase endothelial relaxation following irradiation (Soucy et al., 2010; Soucy et al., 2011). L-nitroarginine (L-NA, general NOS inhibitor) and aminoguanidine (AG, specific iNOS inhibitor) together were able to reduce relaxation of rabbit carotid arteries, suggesting that reduced NO levels are a key cause to endothelial dysfunction (Hong et al., 2013). In addition, L-NA and AG were able to reduce iNOS and nitrotyrosine (peroxynitrite biomarker) levels after irradiation (Hong et al., 2013). Gch1 is an enzyme involved in the synthesis of BH4, a cofactor for eNOS coupling. DAHP, a Gch1 inhibitor, caused the ratio of coupled-to-uncoupled eNOS to decrease and endothelial relaxation to also decrease, showing how coupled eNOS is necessary for endothelial function (Yan et al., 2020). Angiotensin II (AngII) type 1 (AT1) receptor activation can activate NOS. HU rats treated with losartan, an AT1 receptor antagonist, show reduced NOS levels and increased endothelial relaxation (Zhang et al., 2009).

<ul> <li> Directionality of NO changes cannot be compared between studies due to a variety of experimental conditions like stressor type, dose, dose rate, model and time course of the experiment. </li> <li> Irradiating in vivo rabbit carotid arteries with X-rays showed that endothelial dysfunction was higher after 8 Gy than 16 Gy (Hong et al., 2013). This was inconsistent with the ex vivo model, where endothelial dysfunction was highest after 16 Gy (Hong et al., 2013). Endothelial dysfunction was shown through a relaxation response to ACh. </li> </ul>

Not identified

Moderate Male Low Female Low Unspecific

Low

Adult

Moderate

Not Otherwise Specified

Low Moderate Low

The majority of the evidence is derived from in vivo rat models. A limited number of studies were in human and rabbit models. The relationship has been more commonly shown in vivo in male animals, specifically in adult male rodents.

#<Reference::ActiveRecord_Associations_CollectionProxy:0x00007b42ebc84070> AOPWiki 2022-10-20T15:07:53

2023-03-21T11:58:08

Deposition of energy leads to vascular remodeling

Cataia Ives

Tatiana Kozbenko1,2, Nadine Adam, Veronica Grybas1, Benjamin Smith1, Dalya Alomar1, Robyn Hocking1, Janna Abdelaziz3, Amanda Pace3, Carole Yauk2, Ruth Wilkins1, Vinita Chauhan1 (1) Health Canada, Ottawa, Ontario, K1A 0K9, Canada   (2) University of Ottawa, Ottawa, Ontario K1N 6N5, Canada  (3) Carelton University, Ottawa, Ontario K1S 5B6, Canada  Consultants Marjan Boerma1, Omid Azimzadeh2, Steve Blattnig3, Nobuyuki Hamada4 (1) University of Arkansas for Medical Sciences, Little Rock, AR 72205, USA (2) Federal Office for Radiation Protection (BfS), Section Radiation Biology, 85764 Neuherberg, Germany (3) NASA Langley Research Center Hampton, VA  23681, USA (4) Biology and Environmental Chemistry Division, Sustainable System Research Laboratory, Central Research Institute of Electric Power Industry (CRIEPI), Tokyo, Japan

2.5

The present qualitative AOP (AOP#470) summarizes the evidence for a progression beginning with the deposition of energy  to vascular remodeling. The pathway is initiated by ionization/excitation events from the deposition of energy (MIE: Event #1686) leading to an enviorment  of reactive oxygen species (ROS), if this occurs at a rate that outpaces the antioxidant defense system, oxidative stress ensues (KE: Event #1392). Deposition of energy can concurrently induce DNA strand breaks (KE: Event #1635) either directly or through damage from ROS. Excessive ROS damages cellular compartments, thereby altering signaling pathways (KE: Event #2066), and increasing levels of pro-inflammatory mediators (KE: Event #1493). Within the vascular wall, activation of certain signaling molecules can alter nitric oxide (NO) levels (KE: Event #2067). All the upstream KEs of the pathway then converge to cause endothelial dysfunction (KE: Event #2068). Modified levels of NO can alter the blood flow within the endothelium resulting in subsequent compensatory vascular remodeling (AO: Event #2069). Vascular remodeling is an important precursor for many diverse cardiovascular pathologies and serves as an important marker for cardiovascular disease. Vascular remodeling includes many structural changes such as increased vessel stiffness, vessel wall thickening, and decreased capillary density. Studies informing this AOP include clinical follow-up studies of radiotherapy patients, epidemiological cohort studies of atomic bomb survivors and nuclear plant workers as well as biological studies using mouse and rat models. Knowledge gaps in the weight of evidence include inconsistencies in NO evaluation, and relatively few studies exploring chronic and low dose exposures including the lack of studies focusing on female biology.  Cardiovascular disease (CVD) includes any health condition affecting the heart and blood vessels. CVD is one of the leading causes of death worldwide, accounting for millions of deaths yearly and is surpassed in some countries, by only cancer (Bray et al., 2021; Tsao et al., 2022). This class of diseases includes congenital defects, as well as CVDs that can develop throughout life such as peripheral artery disease, atherosclerosis, coronary artery disease and myocardial infarction. While the progression to a CVD outcome is slow, many CVDs are often preceded by much earlier changes to vascular structure. Vascular remodeling entails various structural changes of existing vasculature arising from cell death, cell migration and changes to the endothelial cell membrane. It is important to note that changes to vascular structure are not inherently detrimental, and the cardiovascular system undergoes continuous adaptation to protect vascular health (Pries et al., 2001; Santamaría et al., 2020; Zakrzewicz et al., 2002). However, certain remodeling can also serve as an important marker and risk factor for future adverse cardiovascular events (Cohn et al., 2004; Van Varik et al., 2012). Changes to vascular structure can be triggered through perturbations such oxidative stress, inflammation, and alterations to cellular signaling pathways. Adverse remodeling of the vasculature encompasses structural and functional changes to vessel wall intima-media, elevated stiffness, and decreased lumen diameter which are all predictive of the development of and mortality and morbidity from CVD (Heald et al., 2006; Hodis et al., 1998; Polak et al., 2011; Zieman et al., 2005).   The risk of CVD increases with several factors such as age and available evidence suggests that environmental factors such as radiation can also contribute to increased risk (Belzile-Dugas & Eisenberg, 2021; Boerma et al., 2016; Francula-Zaninovic & Nola, 2018; Wang et al., 2019). The deposition of energy from radiation is a stochastic event, with adverse effects emerging years or decades after the exposure (Boerma et al., 2016; Dörr, 2015; EPRI, 2020; Menezes et al., 2018). The effects of high-dose radiation on the cardiovascular system have been well-characterized while the effects of low-dose exposure are more contended. However, growing evidence suggests that lower doses than previously thought are linked to cardiovascular outcomes (Boerma et al., 2016; EPRI, 2020; Little et al., 2021; UNSCEAR, 2008). Much of the high-dose data is from follow up studies in radiotherapy patient cohorts who have elevated risk for adverse cardiovascular events (Zou et al., 2019). In addition to clinical exposure scenarios, epidemiological studies of occupational exposures and Japanese atomic bomb survivors provide supporting evidence. Cohort studies of atomic bomb survivors show CVD risk can be modulated by factors such as age at exposure and estimated dose received (Ozasa et al., 2012; Preston et al., 2003; Shimizu et al., 2010; Takahashi et al., 2017). Long-term follow up of individuals exposed in the Chernobyl disaster also identified statistically significant elevation in CVD risk (Ivanov et al., 2006; Kashcheev et al., 2017). Occupational exposure studies have also been conducted in various countries in an effort to understand the relationship between low-dose chronic exposure and cardiovascular health of nuclear workers (Azizova et al., 2018; Gillies et al., 2017; Zielinski et al., 2009). Occupational exposure studies suggest positive associations between received dose and excessive relative risk of circulatory diseases (Zielinski et al., 2009), CVD mortality (Gillies et al., 2017) and occurrence of ischemic and cerebrovascular disease (Azizova et al., 2018).   Beyond earth, space travel presents an additional radiation exposure scenario. With future missions planned beyond low Earth orbit and the protective shield of the magnetosphere, understanding the unique challenges of space radiation is crucial for protection of travelers. In space, radiation is present in the form of high linear energy transfer (LET) particles and high mass, high energy ions (HZE) which indiscriminately impacts the whole body at a low fluence rate (Baker et al., 2011; Durante & Cucinotta, 2008; Norbury et al., 2016). While the present AOP includes an MIE focused on deposition of energy following radiation exposure, it is important to note that the space exposome contains multiple stressors to which space travelers will be exposed simultaneously. Particularly important, in the case of the cardiovascular system, is microgravity. The cardiovascular system is gravity sensitive, with the endothelial layer being responsive to changes in shear stress and blood pressure (Hughson et al., 2018; Maier et al., 2015; Versari et al., 2013). Variation to the pressure gradient throughout the body can also trigger regional adaptations to vascular structure (L. F. Zhang, 2013). While determining a mechanism for an MIE of microgravity has proven challenging, microgravity exposure has been shown to contribute to KEs in the pathway and therefore, evidence from microgravity studies has been included in the weight of evidence (WOE).

The present AOP (#470) is a part of a broader network that links the deposition of energy with four AOs: vascular remodeling, bone loss, learning and memory impairment, and cataract formation. The AOP development process began with the creation of a preliminary version of the network through narrative review of key literature as well as extensive subject matter expert consultation (Chauhan et al., 2021).   Many KEs were proposed for the preliminary network; however, attention was focused on those deemed most essential for disease progression and for which  biological plausibility  was strong for connectivity to the rest of the pathway. The preliminary network served as the basis for the next stage in which a scoping review methodology was used to collect a WOE and refine KEs and KERs. A risk-of-bias evaluation was not undertaken,  details of the methodology are described by Kozbenko et al. and are summarized below (Kozbenko et al. 2022).    In short, literature collection consisted of structured database searches, followed by prioritization and screening stages. Focused searches were completed for each of the KERs in the pathway using combinations of key words related to involved KEs, as well as an overarching search, collecting all references broadly related to the MIE and AO. Following the literature searches, the results were processed in two stages to determine their inclusion in the pathways WOE. Both stages determined reference relevance according to inclusion and exclusion criteria that had been outlined prior to screening in a PEOE (Population, Exposure, Outcome, and Endpoint) statement.   In the first stage, search results were prioritized using the SWIFT Review software. This software was used to identify the references most closely related to the PEOE statement as determined by the software created tags. The most relevant references identified by SWIFT were then screened by human screeners using screening forms created in Distiller SR. Screening in Distiller was split into title and abstract, full text and data extraction levels and human screeners were instructed to include references that met the  PEOE criteria. Human screeners additionally evaluated references for their demonstration of Bradford Hill criteria. At the end of the process, collected studies were used for the WOE.   The final pathway includes several KEs previously existing in the AOP-Wiki, these include the deposition of energy (KE #1686), oxidative stress (KE #1392), increased DNA strand breaks (KE #1635) and increased pro-inflammatory mediators (KE #1493). Newly created KEs for this pathway include altered signaling pathways (KE #2066), altered NO levels (KE #2067), increased endothelial dysfunction (KE #2068) and occurrence of vascular remodeling (AO: KE #2069).

It is well documented that ionizing radiation( (eg. X-rays, gamma, photons, alpha, beta, neutrons, heavy ions) leads to energy deposition on the atoms and molecules of the substrate. Many studies, have demonstrated that the type of radiation and distance from source has an impact on the pattern of energy deposition (Alloni, et al. 2014). High linear energy transfer (LET) radiation has been associated with higher-energy deposits (Liamsuwan et al., 2014) that are more densely-packed and cause more complex effects within the particle track (Hada and Georgakilas, 2008; Okayasu, 2012ab; Lorat et al., 2015; Nikitaki et al., 2016) in comparison to low LET radiation. Parameters such as mean lineal energy, dose mean lineal energy, frequency mean specific energy and dose mean specific energy can impact track structure of the traversed energy into a medium (Friedland et al., 2017). The detection of energy deposition by ionizing radiation can be demonstrated with the use of fluorescent nuclear track detectors (FNTDs). FNTDs used in conjunction with fluorescent microscopy, are able to visualize radiation tracks produced by ionizing radiation (Niklas et al., 2013; Kodaira et al., 2015; Sawakuchi and Akselrod, 2016). In addition, these FNTD chips can quantify the LET of primary and secondary radiation tracks up to 0.47 keV/um (Sawakuchi and Akselrod, 2016). This co-visualization of the radiation tracks and the cell markers enable the mapping of the radiation trajectory to specific cellular compartments, and the identification of accrued damage (Niklas et al., 2013; Kodaira et al., 2015). There are no known chemical initiators or prototypes that can mimic the MIE.

adjacent

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Moderate non-adjacent

Low

High non-adjacent

Low

Moderate non-adjacent

Low

High non-adjacent

Low

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High Unspecific

High

All life stages

High High High Low

Summary of evidence (KE & KER Relationships and evidence)   The AOP is supported by high biological plausibility and moderate empirical evidence. Research, primarily from laboratory studies, has supported  dose- and temporal-concordance for each KER.   Biological Plausibility   Described below is the well-established understanding of the mechanisms underlying this AOP with supporting literature. More detailed examples of the empirical data can be found in the individual entries for each KER.  It is well accepted that when energy is deposited in the cell from ionizing radiation (IR), direct damage to cellular structures can occur (Desouky et al., 2015). When traveling through a cell, IR can induce the radiolysis of water forming reactive oxygen species (ROS). Deposition of energy can also induce feedback loops of ROS production where structures and molecules damaged by ROS including the mitochondria and NADPH oxidase (NOX) further produce ROS (Mittal et al., 2014; Soloviev & Kizub, 2019). Additionally, deposited energy can directly upregulate enzymes involved in ROS and reactive nitrogen species (RNS) (collectively RONS) production (de Jager, Cockrell and Du Plessis, 2017). If reactive nitrogen species RONS production outpaces the antioxidant defense, a state of oxidative stress occurs (Fletcher et al., 2010; Slezak et al., 2017; Tahimic & Globus, 2017; Wang et al., 2019). Damage to macromolecules can occur due to oxidative stress, including strand breaks to DNA, oxidation of amino acid residues in proteins and peroxidation of lipids (Ping et al., 2020). Lipid peroxidation can induce further damage to cellular structures as a chain reaction is created by the lipid peroxidation radicals, attacking other lipids, proteins and nucleic acids (Ping et al., 2020). Consequently, oxidative stress can directly lead to multiple downstream KEs including altered signaling pathways, increased DNA strand breaks, increased pro-inflammatory mediators and altered nitric oxide (NO) levels.  DNA strand breaks in endothelial cells can be induced either directly through energy deposition or indirectly through oxidative stress. DNA strand breaks can recruit and activate the protein kinases ataxia telangiectasia mutated (ATM) and ATM/RAD3-related (ATR) (Nagane et al., 2021). Downstream signaling pathways involved in cell death and senescence like the p53/p21 pathway can be activated by ATM/ATR. Furthermore, DNA strand breaks induced by radiation directly or through oxidative stress can cause mutations or changes in transcription of proteins in signaling pathways (Ping et al., 2020; Schmidt-Ullrich et al., 2000). Therefore, DNA strand breaks will induce death and senescence of endothelial cells through altered signaling, resulting in endothelial dysfunction.  Oxidative stress can also induce altered signaling pathways. The effects of oxidative stress on signaling pathways occur through protein oxidation of signaling components (Ping et al., 2020; Schmidt-Ullrich et al., 2000; Valerie et al., 2007). Oxidation of cysteine and methionine residues, which are particularly sensitive to oxidation, can result in structural and functional detriments to the protein (Ping et al., 2020). RONS can influence various pathways including the Akt/PI3K/mTOR pathway, where impaired cell survival signaling can induce cellular senescence (Hassan et al., 2013; Ping et al., 2020). Additionally, inhibition of tyrosine phosphatases by ROS can increase the phosphorylation of mitogen-activated protein kinase (MAPK) pathways, resulting in various downstream effects (Schmidt-Ullrich et al., 2000; Valerie et al., 2007). A phosphorylated p53 induced by oxidative DNA damage can also activate MAPK signaling and initiates a cascade ending in apoptosis (Ashcroft et al., 1999; Gen, 2004). Through affecting cell signaling pathways, damage caused by elevated RONS affects cells beyond those that have been directly irradiated (Ramadan et al., 2021).  Excessive RONS produced by IR disrupt cellular balance and can increase pro-inflammatory mediators (Lumniczky et al., 2021; Schaue et al., 2015). Similar to activation of the immune system by damage from a pathogen, activation by oxidative stress promotes many repair mechanisms, some of which involve rapid release of pro-inflammatory cytokines (Stanojković et al., 2020). The cytokines released vary based on tissue type and radiation parameters (Di Maggio et al., 2015), but tumor necrosis factor (TNF)-α and interleukin (IL)-1 can trigger a cytokine cascade that initiates an inflammatory response (Slezak et al., 2017; Srinivasan et al., 2017). A prolonged state of inflammation in endothelial cells can lead to endothelial dysfunction (Baran et al., 2021).  Both oxidative stress and altered signaling can directly result in altered NO levels. NO is synthesized from L-arginine by the three nitric oxide synthase (NOS) enzymes, endothelial NOS (eNOS), inducible NOS (iNOS) and neuronal NOS (nNOS). ROS can directly reduce NO levels by reacting with NO to produce the RNS peroxynitrite (Deanfield et al., 2007). Furthermore, the cofactor of NOS enzymes, tetrahydrobiopterin (BH4), can be oxidized by RONS leading to inhibition of NOS dimerization, also called NOS uncoupling (Deanfield et al., 2007). Uncoupled NOS will produce superoxide instead of NO, leading to a positive feedback loop of ROS production and reduced NO (Förstermann, 2010; Förstermann & Münzel, 2006; Mitchell et al., 2019; Nagane et al., 2021; Soloviev & Kizub, 2019).  Modulation of NO through altered signaling pathways occurs through changing the activity of NOS enzymes. Phosphorylation of eNOS at Ser1177 will activate the enzyme while phosphorylation at Thr495 inhibits it (Förstermann, 2010; Nagane et al., 2021). Protein kinase B (Akt), part of the phosphoinositide 3-kinase (PI3K)/Akt pathway, can activate eNOS through phosphorylation at Ser1177 to increase NO production (Karar & Maity, 2011). In contrast, activation of the RhoA/Rho kinase (ROCK) pathway will inhibit NO production by destabilizing eNOS mRNA and preventing Ser1177 phosphorylation by Akt (Yao et al., 2010). Angiotensin II (AngII), the end product of the renin-angiotensin-aldosterone system (RAAS), is involved in both downregulating Ser1177 phosphorylation to prevent NO creation (Ding et al., 2020) and activating eNOS as a corrective measure (Millatt et al., 1999). Alterations to these pathways due to IR will result in changes in NO levels.  Each of the components of the pathway described above converge at endothelial dysfunction. Endothelial cells lining the blood vessels throughout the body are an important component for maintaining vascular homeostasis (Bonetti et al., 2003; Deanfield et al., 2007). Endothelial cells are quiescent with high levels of NO most of the time (Carmeliet & Jain, 2011). Endothelial dysfunction can occur due to prolonged activation of the endothelium, characterized by the prolonged lack of bioavailable NO, lack of endothelium-dependent vasodilation and chronic pro-thrombotic and inflammatory state (Baran et al., 2021; Bonetti et al., 2003; Deanfield et al., 2007; Krüger-Genge et al., 2019). A prolonged reduction in NO will decrease vasodilation, increase leukocyte adhesion and increase fibrous plaque formation contributing to the pro-thrombotic dysfunctional environment (Schiffrin, 2008; Senoner & Dichtl, 2019; Venkatesulu et al., 2018). Furthermore, signaling in pathways like p53/p21 or PI3K/Akt/mammalian target of rapamycin (mTOR) can induce apoptosis or premature senescence of endothelial cells as part of endothelial dysfunction due to DNA damage or oxidative stress (Borghini et al., 2013; Hughson et al., 2018; Schiffrin, 2008; Senoner & Dichtl, 2019; Soloviev & Kizub, 2019). Senescent cells have decreased levels of NO production and a pro-inflammatory secretory phenotype, which feed back to further promote endothelial dysfunction (Ungvari et al., 2013; Wang et al., 2016).  Endothelial dysfunction subsequently leads to vascular remodeling, which encompasses multiple structural changes to the vasculature. Chronic inflammation combined with impaired healing and lack of endothelium-dependent vasodilation during endothelial dysfunction increases vulnerability to damage from non-laminar flow and maladaptive repair (Sylvester et al., 2018). As compensation, vessel walls can thicken and atherosclerotic risk can increase (Hughson et al., 2018; Slezak et al., 2017; Sylvester et al., 2018). In cases of maladaptive repair of vessels, vascular remodeling can be exhibited through an increase in fibrosis (Hsu et al., 2019). The pro-thrombotic environment with increased lymphocyte adhesion induced by endothelial cell senescence can increase the likelihood of vessel occlusion, decreasing vascular density such that the corresponding increase in vascular resistance will induce remodeling as a compensatory measure (Slezak et al., 2017). Thus, increased leukocyte adhesion during endothelial dysfunction occurs early in the development of atherosclerosis (Senoner & Dichtl, 2019). Increased arterial stiffness can also occur in response to endothelial dysfunction (Boerma et al., 2015, 2016; Patel et al., 2020), with increased collagen and smooth muscle content paired with decreased elastin and degradation of the extracellular matrix (Zieman et al., 2005). The changes to the vascular structure in response to the deposition of energy are similar to a form of accelerated age-related atherosclerosis (Boerma et al., 2016; Sylvester et al., 2018; Vernice et al., 2020).  Temporal, Dose, and Incidence Concordance   Evidence for time, dose, and incidence concordance in this AOP is moderate. It has been repeatedly shown using many study designs and systems that deposition of energy occurs immediately following irradiation, and downstream events occur at a later timepoint. Endpoints indicating oxidative stress have been observed within minutes following irradiation (Wortel et al., 2019). Studies show that oxidative stress, increased DNA strand breaks, increased pro-inflammatory mediators, and altered signaling may occur over a similar time period; however, alteration in signaling pathways, increased DNA strand breaks, and increased pro-inflammatory mediators can be observed following oxidative stress (Ramadan et al., 2020; Baselet et al., 2017; Sakata et al., 2015; Yang et al., 1998). Increases in NO levels occur in hours to weeks after irradiation (Azimzadeh et al., 2017; Sonveaux et al., 2003; Sakata et al., 2015). Then, from weeks to months following irradiation both endothelial dysfunction and vascular remodeling occur, though concordance between these events is difficult to determine, possibly due to inter-study differences in experimental design and markers (Yentrepalli et al., 2017; Soucy et al., 2007; Yu et al., 2011; Shen et al., 2018).  Overall, the majority of studies demonstrate that upstream KEs occur at the same or lower doses and earlier or the same time as downstream KEs. For example, endothelial cells show a dose-dependent increase in oxidative stress to X-ray irradiation at 0.1 and 5 Gy, while 0.1 Gy induced few changes in pro-inflammatory mediators with significant increases only observed at 5 Gy (Ramadan et al., 2020). Some studies also show that the upstream and downstream KEs can be observed at the same doses of radiation. For example, X-ray irradiation of mice resulted in oxidative stress, altered signaling and reduced NO levels at both 8 and 16 Gy (Azimzadeh et al., 2015). Dose concordance is not consistent across studies, but this may be due to differences in models, timepoints, and radiation types used.   A limited number of studies support incidence concordance. In these, the upstream KE demonstrates a greater change than the downstream KE following exposure to a stressor. For example, mice exposed to 18 Gy of X-rays showed a roughly 2-fold increases in both oxidative stress and pro-inflammatory markers. A 1.3-fold increase in markers for endothelial dysfunction was observed (Shen et al., 2018). Uncertainties and inconsistencies   The collection of WOE identifies several important uncertainties in the literature. These include lack of quantitative understanding, low-dose or chronic-exposure studies, data from female models and consistency in measurement of NO levels.   The WOE contained data from a wide variety of interdisciplinary fields; consequently, experimental design was equally varied. Overall, studies did not use consistent doses, radiation types, time-points, or evaluation of endpoints. Since dose and type of radiation can affect biological responses, quantitative understanding of relationships could not be determined and was low overall. Additionally, most studies used single or select doses, with limited studies exploring relatively low doses (<0.5 Gy (EPRI, 2020)). Harmonized experiments evaluating changes to adjacent endpoints across a wide range of doses or time-points with consistency of radiation type would greatly benefit quantitative understanding for this AOP.   Similarly, the WOE is lacking in evidence using female models. Sex is an important modulating factor in cardiovascular changes and studies suggest vascular remodeling responses of astronauts can vary by sex (Hughson et al., 2016). The consequence of the general bias in clinical research (Rios et al., 2020; Yakerson, 2019) from which the current WOE draws, is the very large knowledge gaps in mechanistic data for the female body. Filling these knowledge gaps at all levels of biological organization will be an important step in solidifying the AOP.    Evaluation of NO levels was inconsistent between studies. According to the biological plausibility, deposition of energy and subsequent oxidative stress would lead to a decrease in NO that then contributes to impaired vascular relaxation as part of endothelial dysfunction. However, primary research concludes that NO can either increase (Abdel-Magied & Shedid, 2020; Hirakawa et al., 2002; Sakata et al., 2015; Sonveaux et al., 2003) or decrease (Baker et al., 2009; Fuji et al., 2016) following irradiation. Proxy measures used to detect NO, like NOS enzyme activities or nitrite/nitrate levels, may not directly correspond to changes in NO levels. Further standardization in NO measurement and interpretation could refine this KE to become the depletion of NO.  The empirical evidence supports that this AOP is relevant to human (Hong et al., 2013; Siamwala et al., 2010; Jiang et al., 2020; Lee, et al., 2020; Ramadan et al., 2020), rat (Hatoum et al., 2006; Soucy et al., 2010; Hong et al., 2013; Abdel-Magied & Shedid, 2019: Hasan et al., 2020), mouse (Yu et al., 2011; Coleman et al., 2015; Sofronova et al., 2015; Shen et al., 2018; Hamada et al., 2020), and rabbit (Soloviev et al., 2003, Hong et al., 2013) models. Biological plausibility suggests that events in this AOP are not sex specific; however, more studies used male models. Similarly, while biological plausibility suggests the pathway is not age-specific, most studies used adult models.

The essentiality of the MIE to a downstream KE is supported by a non-irradiated control. The comparison of irradiated and non-irradiated groups has shown that the effects of downstream events are enhanced or accelerated by the deposition of energy.   The essentiality of other KEs can be determined by the impact of the manipulation of the upstream KE on the resulting downstream effects. For example, the essentiality of oxidative stress is frequently assessed through antioxidant treatments, which can decrease oxidative stress markers through decreased ROS production or strengthened antioxidant defense activity. SOD administration decreased free radicals, superoxide and peroxide, and improved endothelium-dependent vasodilation, a downstream KE, which had been previously decreased due to radiation exposure (Hatoum et al., 2006). Additionally, oxypurinol treatment inhibited xanthine oxidase (XO) enzyme, which limited the enzyme’s contribution to cardiac ROS and improved endothelium-dependent vasodilation and the recovery of vascular stiffness to control levels (Soucy et al., 2007, 2010, 2011).  The essentiality of DNA strand breaks was not assessed often. One study used mesenchymal stem cell conditioned media (MSC-CM) to reduce the level of ROS-mediated DNA double-stranded breaks and found decreases in signaling molecules including p53, Bax and cleaved caspase 3 (Huang et al., 2021).  The essentiality for altered signaling pathways KE was evaluated by studies using pathway inhibitors or conditioned media. Signaling pathways were shown to be suppressed by inhibitors such as ROCK inhibitor Y27632 and acid sphingomyelinase (ASM) inhibitor desipramine (dpm), which have demonstrated decreased apoptosis and recovered endothelium-dependent vasodilation (Soloviev & Kizub, 2019; Venkatesulu et al., 2018; Wang et al., 2016). Incubation of endothelial cells in MSC-CM was shown to increase cell signaling components, Akt and p-Akt, and decrease apoptosis (Chang et al., 2017). PI3K inhibitors, such as LY294002 and wortmannin, and angiotensin-converting enzyme inhibitor bradykinin-potentiating factor (BPF) were studied for their impact on NO levels. The increase in p-Akt and subsequently eNOS, p-eNOS and NO levels were reversed following PI3K inhibition (Shi et al., 2012; Siamwala et al., 2010). AngII and iNOS levels were returned to control following BPF treatment of irradiated groups (Hasan et al., 2020). Further studies are required for a better understanding of the changes in NO levels and endothelial dysfunction due to altered signaling pathways. Overall, the flexibility of signaling pathways makes it difficult to assess essentiality.  The essentiality for pro-inflammatory mediators was assessed by studies that suppress their expression. The decrease in pro-inflammatory mediators was observed following the use of TAT-Gap19 to block connexin43 hemichannels. This decrease was associated with a decrease in radiation-induced endothelial cell senescence (Ramadan et al., 2020). Additionally, MSC-CM incubation resulted in decreased pro-inflammatory cytokines, IL-1α, IL-6 and TNF-α and decreased endothelial apoptosis (Chang et al., 2017).  Changes in vascular remodeling were evaluated through vascular structure, among other endpoints. Following hindlimb unloading, ASM inhibition in the small mesenteric artery was found to reverse the changes in apoptosis and intima-media thickness (IMT) (Su et al., 2020). Comparisons between irradiated and sham or non-irradiated control groups of various studies using animal and human models have demonstrated differences in vascular structures (Hamada et al., 2020, 2021; Sárközy et al., 2019; Shen et al., 2018; Sridharan et al., 2020; Yu et al., 2011). Essentiality of the key events  <table border="1"> <tbody> <tr> <td>   </td> <td> Defining Question  </td> <td> High  </td> <td> Moderate  </td> <td> Low  </td> </tr> <tr> <td> Support for Essentiality of KEs  </td> <td> Are downstream KEs and/or the AO prevented if an upstream KE is blocked?  </td> <td> Direct evidence from specifically designed experimental studies illustrating essentiality for at least one of the important KEs  </td> <td> Indirect evidence that sufficient modification of an expected modulating factor attenuates or augments a KE  </td> <td> No or contradictory experimental evidence of the essentiality of any of the KEs  </td> </tr> <tr> <td> MIE: KE #1689 Deposition of energy  </td> <td colspan="4"> Evidence for Essentiality of KE: High  This event is difficult to test for essentiality as deposition of energy is a physical stressor and cannot be blocked/decreased using chemicals. However, studies show that control or sham-irradiated groups do not show the occurrence of downstream KEs.  </td> </tr> <tr> <td> KE #1392 Oxidative stress  </td> <td colspan="4"> Evidence for Essentiality of KE: High  Essentiality was well supported within the literature. Antioxidant treatments led to recovery of antioxidant enzyme activity, decreases in DNA strand breaks, and decreases in pro-inflammatory mediators, while ZNO-NP also restored NO levels. Oxypurinol (Oxp) treatment was found to aid in the acetylcholine (ACh) vasodilation response and restore NO levels as it decreased xanthine oxidase (XO) activity and reactive oxygen species (ROS).   </td> </tr> <tr> <td> KE #1635  Increase, DNA strand breaks  </td> <td colspan="4"> Evidence for Essentiality of KE: Low  Few studies use countermeasures to reduce the number of DNA strand breaks in cells. A few studies show that reducing DNA strand breaks induced by radiation restores signaling pathways and reduces endothelial dysfunction.  </td> </tr> <tr> <td> KE #1493 Increase, pro-inflammatory mediators  </td> <td colspan="4"> Evidence for Essentiality of KE: Low  Essentiality of this event can be determined with countermeasures that limit the increase of pro-inflammatory mediators. Limited research does show essentiality, evidenced by a decrease in apoptosis of endothelial cells following treatment with MSC-CM, which contains angiogenic cytokines that have therapeutic potential for microvascular injury, and a decrease in endothelial cell senescence following treatment with TAT-Gap19, a connexin hemichannel blocker.  </td> </tr> <tr> <td> KE #2066  Altered signaling pathways  </td> <td colspan="4"> Evidence for Essentiality of KE: Moderate  Essentiality of this relationship can be determined with the use of signaling molecule inhibitors. Signaling molecule inhibitors reduced downstream changes in eNOS, NO, p-Akt, angiotensin II (AngII) and aldosterone following stressors such as irradiation and altered gravity.  Inhibitors also prevented impaired contractile response and decreased apoptosis in the arterial endothelium.  </td> </tr> <tr> <td> KE #2067 Altered, NO levels  </td> <td colspan="4"> Evidence for Essentiality of KE: Moderate  The evidence for essentiality of this KE can be determined by using countermeasures that limit changes in NO levels, such as Oxp, L-NA (NOS inhibitor), AG (iNOS inhibitor), DAHP (Gch1 inhibitor) and losartan (AT1 receptor antagonist). Use of these countermeasures reduced NOS levels and decreased the ratio of couple-to-uncoupled eNOS. Endothelial relaxation increased after Oxp and losartan treatment after microgravity exposure, while relaxation decreased in the presence of DAHP, L-NA and AG. When treated with these countermeasures following radiation or microgravity, changes to NO were limited or restored and as a result, endothelial dysfunction was limited.  </td> </tr> <tr> <td> KE #2068  Increase, endothelial dysfunction  </td> <td colspan="4"> Evidence for Essentiality of KE: Moderate  The essentiality of endothelial dysfunction leading to vascular remodeling is moderately supported within literature. Oxp treatment, an XO inhibitor, restored vasodilator response and reduced vascular stiffness following irradiation. Both dpm and DOX decreased apoptosis and reduced Caspase-3 protein expression. Ceramide treatment following microgravity was found to return proliferation to control levels and increase apoptosis. </td> </tr> </tbody> </table>

<table border="1"> <tbody> <tr> <td>   </td> <td> Defining Question  </td> <td> High  </td> <td> Moderate  </td> <td> Low  </td> </tr> <tr> <td> Review of Biological Plausibility for the KER    </td> <td> Is there a mechanistic (structural or functional) relationship between the upstream KE and downstream KE consistent with established biological knowledge  </td> <td> The relationship is well understood based on extensive previous documentation and has an established mechanistic basis and broad acceptance  </td> <td> The KER is plausible based on an analogy to accepted biological relationships but scientific understanding is not completely established  </td> <td> There is empirical support for a statistical association between KEs but structural or functional relationship between them is not understood  </td> </tr> <tr> <td> Deposition of energy (MIE: KE #1686) leads to oxidative stress (KE #1392)  </td> <td colspan="4"> Evidence for Biological Plausibility of KER: High   Deposition of energy onto the water and biological components of a cell creates ROS, and as ROS production outpaces the cell’s antioxidant defense system, oxidative stress is induced. Both ROS production and subsequent oxidative stress have been extensively studied and the mechanisms are well described in numerous review articles across many biological systems.   </td> </tr> <tr> <td> Deposition of energy (MIE: KE #1686) leads to increase, DNA strand breaks (KE #1635)  </td> <td colspan="4"> Evidence for Biological Plausibility of KER: High  The deposition of energy onto the DNA molecule will directly cause single- or double-strand breaks in the DNA. Deposited energy can induce chemical modifications to the phosphodiester backbone of both strands of the DNA, possibly resulting in breaks in one or both strands.  </td> </tr> <tr> <td> Oxidative stress (KE #1392) leads to increase, DNA strand breaks (KE #1635)  </td> <td colspan="4"> Evidence for Biological Plausibility of KER: High  Increased ROS during oxidative stress can result in the oxidation of bases on the DNA strand, triggering base excision repair, which removes the oxidized bases. When multiple bases in close proximity are removed, the repair efforts cause strain which can lead to strand breaks.  </td> </tr> <tr> <td> Increase, DNA strand breaks (KE #1635) leads to altered signaling pathways (KE #2066)  </td> <td colspan="4"> Evidence for Biological Plausibility of KER: High  Strand breaks induce the recruitment of the kinases ataxia-telangiectasia mutated (ATM) and ATM/RAD3-related (ATR). ATM and ATR can subsequently phosphorylate multiple downstream signaling molecules. High levels of DNA strand breaks can increase the recruitment of ATM and ATR, leading to greater activation of pathways like the p53/p21 pathway and subsequently greater downstream effects.  </td> </tr> <tr> <td> Oxidative stress (KE #1392) leads to increase, pro-inflammatory mediators (KE #1493)  </td> <td colspan="4"> Evidence for Biological Plausibility of KER: High  Excess ROS during oxidative stress damages cellular structures and thus activates the immune system and repair mechanisms, many of which involve release of pro-inflammatory mediators. Cells involved with host-defense can themselves also produce ROS, further exacerbating the state of oxidative stress. The biological plausibility of the linkage between oxidative stress and increases in pro-inflammatory mediators is highly supported in literature.    </td> </tr> <tr> <td> Oxidative stress (KE #1392) leads to altered signaling pathways (KE #2066)  </td> <td colspan="4"> Evidence for Biological Plausibility of KER: High  Oxidative stress can alter signaling pathways both directly and indirectly. Directly, oxidative stress conditions can lead to oxidation of amino acid residues. This can cause conformational changes, protein expansion, and protein degradation, leading to changes in the activity and level of signaling proteins. Oxidation of key functional amino acids can also alter the activity of signaling proteins, resulting in downstream alterations in signaling pathways. Indirectly, oxidative stress can damage DNA causing changes in the expression of signaling proteins as well as the activation of DNA damage response signaling. The mechanisms of this relationship are widely accepted.   </td> </tr> <tr> <td> Oxidative stress (KE #1392) leads to increase, endothelial dysfunction (KE #2068)   </td> <td colspan="4"> Evidence for Biological Plausibility of KER: High  ROS can interact with NO, taking a vasodilator crucial for endothelial function and turning it into peroxynitrite, a RNS that further contributes to oxidative stress. Furthermore, cellular senescence, inhibition of vasodilation, induced inflammatory environments and cellular apoptosis are all part of endothelial dysfunction that can be indirectly caused by oxidative stress.   </td> </tr> <tr> <td> Increase, pro-inflammatory mediators (KE #1493) leads to increase,  endothelial dysfunction (KE #2068)  </td> <td colspan="4"> Evidence for Biological Plausibility of KER: High  Inflammation provides a protective effect to the endothelium but prolonged or repeated exposure to a stressor can exhaust this, leading to senescence or apoptosis in endothelial cells and subsequent leading to endothelial dysfunction. This endothelial dysfunction can also manifest as a dysregulation of vasodilation. Prolonged inflammation is a widely accepted component in the development of endothelial dysfunction.   </td> </tr> <tr> <td> Altered signaling pathways (KE #2066) leads to increase, endothelial dysfunction (KE #2068)  </td> <td colspan="4"> Evidence for Biological Plausibility of KER: High  Signaling pathways including the PI3K/Akt/mTOR, RhoA-Rho-kinase, ASM/cer pathway, and the p53-p21 pathway have downstream effects on endothelial apoptosis, premature endothelial cell senescence and cytoskeletal proteins to impair contraction, indicators of endothelial dysfunction.  </td> </tr> <tr> <td> Increase, endothelial dysfunction (KE #2068) leads to occurrence, vascular remodeling (AO: KE #2069)  </td> <td colspan="4"> Evidence for Biological Plausibility of KER: High  Key components of endothelial dysfunction include deficiency in bioavailable NO, impaired vasodilation, inflamed endothelium and prothrombotic environment. These events can ultimately lead to vascular remodeling to compensate for decreased capillary and vascular density and increased vascular resistance. Regional pressure changes in vessels due to microgravity can also result in regional changes to vascular structure.  </td> </tr> <tr> <td> Deposition of energy (MIE: KE #1686) leads to increase, endothelial dysfunction (KE #2068)  </td> <td colspan="4"> Evidence for Biological Plausibility of KER: High  Irradiation can cause cellular and tissue level markers of endothelial dysfunction. Following prolonged exposure to radiation, the protective effect of the endothelium can become exhausted and lead to endothelial dysfunction. Consequently, endothelial cells may lose their integrity and become senescent or apoptotic via alterations to signaling pathways, leading to endothelial dysfunction evidenced by dysregulation of vasodilation. Endothelial dysfunction is commonly considered a hallmark for the development of various cardiovascular pathologies.   </td> </tr> <tr> <td> Deposition of energy (MIE: KE #1686) leads to occurrence, vascular remodeling (AO: KE #2069)  </td> <td colspan="4"> Evidence for Biological Plausibility of KER: High  Radiation can accelerate the natural processes of vascular remodeling related to aging. An increase in ROS, produced by IR, can reduce NO bioavailability, leading to endothelial dysfunction and vascular stiffness. In addition, the low level of inflammation during early stages of radiation leads to inhibition of tissue and vessel recovery, and later results in intimal thickening and vascular remodeling. Changes in vessel composition, such as collagen content, may also occur from energy deposition and affect vascular remodeling.  </td> </tr> <tr> <td> Deposition of energy (MIE: KE#1686) leads to altered, NO levels (KE #2067)  </td> <td colspan="4"> Evidence for Biological Plausibility of KER: High  NO is produced by NOS enzymes or by the reduction of nitrite to NO. Deposition of energy can interfere with this process in several ways. Radiolysis of water forms ROS that interacts with NO to produce peroxynitrite which reduces NO bioavailability. ROS can also cause NOS uncoupling, which can reduce NO levels. In contrast, NO can also increase as a result of IR through activation of iNOS during oxidative stress. IR can also influence various signaling pathways that control NO levels, causing radiation to indirectly affect NO levels.   </td> </tr> <tr> <td> Oxidative stress (KE #1392) leads to altered, NO levels (KE #2067)  </td> <td colspan="4"> Evidence for Biological Plausibility of KER: High  It is thought that excessive ROS production can lead to altered NO bioavailability both through direct interaction and indirectly through decreasing its production. Elevated O2- can interact with NO converting it to peroxynitrite leading to decreased bioavailability. ROS can also oxidize the eNOS cofactor BH4, causing eNOS uncoupling inhibiting NO production. Electron leakage in uncoupled eNOS produces additional ROS, exacerbating the state of oxidative stress.   </td> </tr> <tr> <td> Altered signaling pathways (KE #2066) leads to altered, NO levels (KE #2067)  </td> <td colspan="4"> Evidence for Biological Plausibility of KER: High  Various pathways are well known to influence NO levels. Some well-studied examples include the PI3K/Akt pathway, the RhoA/ROCK pathway, the RAAS pathway and the acidic sphingomyelinase/ceramide pathway. The PI3K/Akt, RhoA/ROCK and RAAS pathways and their components are involved in the phosphorylation of various eNOS residues affecting the enzymes activation. The activation or deactivation of eNOS affects the levels of NO production. In contrast, the acidic sphingomyelinase/ceramide pathway can activate NADPH oxidase (NOX), leading to the production of ROS that goes on to scavenge NO decreasing its bioavailability.   </td> </tr> <tr> <td> Altered, NO levels (KE #2067) leads to increase, endothelial dysfunction (KE #2068)  </td> <td colspan="4"> Evidence for Biological Plausibility of KER: High  Lack of bioavailable NO is considered one of the key drivers of endothelial dysfunction. Under normal conditions NO binds with soluble guanylyl cyclase (sGC) creating cGMP and cAMP to activate cellular kinase cascades and Ca2+-dependent vasodilation through smooth-muscle relaxation. Lack of bioavailable NO interrupts this process, reducing the relaxation of smooth muscle cells and dilation of the blood vessels. In contrast, an increase in NO combined with simultaneous excessive ROS can drive cellular senescence through increased peroxynitrite formation. Prolonged impaired vasodilation and elevated premature endothelial cell senescence are important characteristics of endothelial dysfunction.  </td> </tr> </tbody> </table>

In addition to drugs and treatments that modulate individual relationships, there are a variety of factors that modulate the overall progression of CVD and related vascular remodeling.   Genetics and heritable traits are the basis for individual predisposition and CVD risk; indeed, traits associated with heart disease are associated with genetics and are heritable. For example, baseline vascular stiffness, carotid intima-medial thickness (CIMT) and coronary calcification are in part hereditary and vary between ethnicities (Berk & Korshunov, 2006; Winham et al., 2014). Furthermore, how CVD manifests also differs between biological sexes. For example, men have higher age-adjusted mortality rates for coronary heart disease while the women living with CVD and dying of stroke outnumber men (Mosca et al., 2011). The genetic component of sex also plays a role, with polymorphisms of genes related to adverse cardiac remodeling found on the Y chromosome and X chromosome inactivation being linked to remodeling (Berk & Korshunov, 2006). Overall, sex plays a role in both predisposition for CVD risk and the form in which adverse cardiovascular effects progress.   Age is an additional well-established risk factor. Natural aging of the cardiovascular system leads to structural changes like increased arterial stiffness, wall thickening and decreased endothelial function (North & Sinclair, 2012; Zieman et al., 2005). With aging also comes changes to sex-hormone levels further modulating CVD risk and progression (Kessler et al., 2019; Mosca et al., 2011; Rodgers et al., 2019; Winham et al., 2014). Some radiation induced vascular changes have been compared to accelerated versions of the natural aging process (Boerma et al., 2016; EPRI, 2020). The age at which radiation exposure occurs also plays a role in the eventual outcomes, with younger age of exposure leading to elevated risk of radiation induced CVD (Belzile-Dugas & Eisenberg, 2021; Yang et al., 2021).   Finally, lifestyle factors contribute to the risk for CVD. In fact, positive modifications to lifestyle are sometimes considered the most impactful prevention strategy for CVD (Chiuve et al., 2006). Poor diet, inadequate physical activity, smoking and adiposity are linked with downstream risk factors like high blood pressure, elevated cholesterol and perturbations of glucose-insulin homeostasis (Mozaffarian et al., 2008). Several measures such as income level, educational attainment, employment status and neighborhood socioeconomic factors can be predictive of CVD risk (Schultz et al., 2018). Importantly, health and fitness requirements for astronauts leads to a cohort subject to the healthy worker effect (Chowdhury et al., 2017) and can serve to reduce the occurrence of CVD potentially masking adverse effects of workplace hazards. <table border="1"> <tbody> <tr> <td> Modulating Factor </td> <td> Influence or Outcome </td> <td> KER(s) involved </td> </tr> <tr> <td> Genetic </td> <td> Baseline vascular stiffness and intima thickness can vary by individual. This can affect how radiation influences these parameters.  </td> <td> Deposition of energy leading to vascular remodeling Endothelial dysfunction leading to vascular remodeling </td> </tr> <tr> <td> Age  </td> <td> Older age can increase baseline vascular stiffness and intima thickness and decrease endothelial function.  Younger age at radiation exposure leads to an elevated risk of CVD.  </td> <td> Deposition of energy leading to oxidative stress Deposition of energy leading to endothelial dysfunction Deposition of energy leading to vascular remodeling Endothelial dysfunction leading to vascular remodeling </td> </tr> <tr> <td> Sex  </td> <td> Sex can have various influences on CVD. For example, men have higher age-adjusted mortality rates for coronary heart disease while the women dying from a stroke outnumber men.  </td> <td> Deposition of energy leading to vascular remodeling Endothelial dysfunction leading to vascular remodeling </td> </tr> <tr> <td> Diet and smoking </td> <td> Linked with high blood pressure, elevated cholesterol and perturbations of glucose-insulin homeostasis.  </td> <td> Deposition of energy to vascular remodeling </td> </tr> <tr> <td> Physical activity </td> <td> Can be associated with various risk factors for CVD. Certain groups (e.g., astronauts) may have greater levels of physical activity which could influence changes to CVD outcomes.  </td> <td> Deposition of energy to vascular remodeling </td> </tr> </tbody> </table>

Despite biological plausibility and empirical evidence demonstrating the qualitative linkages within the AOP, quantitative understanding is low. As described above, the lack of quantitative understanding of the KERs is due to the diversity in experimental design, including doses tested and radiation types used. The evidence is primarily from laboratory studies that show dose and time response relationships for KEs; however, the strength of the response can vary with factors such as dose-rate, type of radiation, and cell type. Particularly relevant are the relative lack of low-dose studies and exposure scenarios relevant to space radiation. Future work could use the present qualitative AOP to guide experimental design and strengthen quantitative understanding. Standardized studies simultaneously measuring endpoints across several KEs, and across a range of doses and timepoints would be beneficial in filling important gaps in the quantitative understanding. <table border="1"> <tbody> <tr> <td> Deposition of energy (MIE: KE #1686) leads oxidative stress (KE #1392)  </td> <td> Evidence for Quantitative Understanding of KER: High  There is a large amount of evidence supporting how much of a change in the deposition of energy is needed to produce a change in the level of oxidative stress. Several different endpoints representing oxidative stress have been used, including changes in the levels or activity of catalase, GSH, superoxide dismutase, GSH-Px, MDA, and ROS. Measurements have also been made over a large range of doses and dose rates, and changes to oxidative stress levels have been shown to depend on the nature, dose and dose rate of energy deposition.   </td> </tr> <tr> <td> Deposition of energy (MIE: KE #1686) leads to increase, DNA strand breaks (KE #1635)  </td> <td> Evidence for Quantitative Understanding of KER: High  Studies examining energy deposition leading to strand breaks suggest a positive, linear relationship between these two events. The exact number of strand breaks is difficult to predict from the deposition of energy. The relationship depends on the biological model, the type of radiation, and the dose.  </td> </tr> <tr> <td> Oxidative stress (KE #1392) leads to increase, DNA strand breaks (KE #1635)  </td> <td> Evidence for Quantitative Understanding of KER: Moderate  There is a considerable amount of evidence showing increased DNA strand breaks following exposure to oxidative stress. However, no model has emerged that predicts the number of DNA strand breaks following oxidative stress. Measurements of oxidative stress vary across studies.  </td> </tr> <tr> <td> Increase, DNA strand breaks (KE #1635) leads to altered signaling pathways (KE #2066)  </td> <td> Evidence for Quantitative Understanding of KER: Moderate  There is much evidence showing changes in the expression or activity of signaling pathways following increased DNA strand breaks. However, no model has been developed to accurately predict the changes to signaling pathways due to increased DNA strand breaks. Furthermore, the changes to signaling pathways are very context- and cell type-dependent.  </td> </tr> <tr> <td>Oxidative Stress (KE #1392) leads to altered signaling pathways (KE #2066)</td> <td> Evidence for Quantitative Understanding of KER: Low  The quantitative understanding of oxidative stress leading to altered signaling pathways is low as a precise quantitative relationship between the key events is difficult to determine due to differences in experimental design. The exact changes to signaling pathways due to oxidative stress will depend on the cell type and species. </td> </tr> <tr> <td> Oxidative stress (KE #1392) leads to increase, endothelial dysfunction (KE #2068)  </td> <td> Evidence for Quantitative Understanding of KER: Low  Although studies quantitatively measure both oxidative stress and endothelial dysfunction following a stressor, it is difficult to compare results and identify a quantitative relationship as studies use different models, stressors, doses and time scales. In addition, many factors and pathways can contribute to endothelial dysfunction. Thus, no model has been established to predict the extent of changes in endothelial dysfunction after oxidative stress.   </td> </tr> <tr> <td> Oxidative stress (KE #1392) leads to increase, pro-inflammatory mediators (KE #1493)  </td> <td> Evidence for Quantitative Understanding of KER: Moderate  Current primary research shows that an increase in oxidative stress will be followed by a more significant increase in pro-inflammatory mediators. A quantitative association between the two KEs is difficult to determine, as multiple positive feedback mechanisms exist between oxidative stress and inflammation </td> </tr> <tr> <td> Increase, pro-inflammatory mediators (KE #1493) leads to increase, endothelial dysfunction (KE #2068)  </td> <td> Evidence for Quantitative Understanding of KER: Low  Although studies reveal increases in markers for endothelial dysfunction in response to increased pro-inflammatory mediators, no quantitative understanding has been established to predict the changes in endothelial dysfunction markers. There are various pro-inflammatory mediators that may contribute to various markers of endothelial dysfunction such as apoptosis and cellular senescence. Studies investigate changes in the levels of different pro-inflammatory mediators and different measures of endothelial dysfunction; therefore, it is difficult to compare the results and identify trends.   </td> </tr> <tr> <td> Altered signaling pathways (KE #2066) leads to increase, endothelial dysfunction (KE #2068)  </td> <td> Evidence for Quantitative Understanding of KER: Low  Although studies show increases in markers for endothelial dysfunction in response to altered signaling pathways, no quantitative understanding has been established to predict the changes in endothelial dysfunction markers. There are various signaling pathways that may contribute to endothelial dysfunction, including the Akt/PI3K/mTOR pathway, the RhoA-Rho-kinase pathway, and the ASM/cer pathway. Studies investigate changes to the levels of different signaling pathway molecules; therefore, it is difficult to compare the results and identify trends.   </td> </tr> <tr> <td> Increase, endothelial dysfunction (KE #2068) leads to occurrence, vascular remodeling (AO: KE #2069)  </td> <td> Evidence for Quantitative Understanding of KER: Low  Vascular remodeling is consistently shown with endothelial dysfunction. However, it is difficult to compare results and identify a quantitative relationship as various models, stressors, doses and endpoint measures were used. Thus, no model has been established to accurately predict the changes in vascular remodeling.   </td> </tr> <tr> <td> Deposition of energy (MIE: KE #1686) leads to increase, endothelial dysfunction (KE #2068)  </td> <td> Evidence for Quantitative Understanding of KER: Low  Studies revealed consistent increases in levels of indicators of endothelial dysfunction such as apoptosis, premature endothelial cell senescence and diminished relaxation response. There is consistent evidence that shows that as the dose increases, the maximum relaxation response decreases. However, more studies are required to quantify this association to show how this relates to levels of cellular markers of apoptosis and senescence.   </td> </tr> <tr> <td> Deposition of energy (MIE: KE #1686) leads to occurrence, vascular remodeling (AO: KE #2069)  </td> <td> Evidence for Quantitative Understanding of KER: Low  Deposition of energy from IR is consistently demonstrated to drive vascular remodeling. However, it is difficult to compare results and quantify relationships as each study uses different models, stressors, doses and time scales. In addition, many factors and pathways contribute to the components of vascular remodeling. Thus, no model has been established to predict the changes in vascular remodeling after deposition of energy.   </td> </tr> <tr> <td> Deposition of energy (MIE: KE #1686) leads to altered, NO levels (KE #2067)  </td> <td> Evidence for Quantitative Understanding of KER: Low  Altered nitric oxide levels occur consistently with deposition of energy. However, it is difficult to compare results and determine a quantitative relationship as each study uses different models, stressors, doses and endpoint measures of NO. As well, cancerous cells and normal cells can show different production of NO. Thus, no model has been established to predict the changes in nitric oxide levels at a given dose of IR.  </td> </tr> <tr> <td> Oxidative stress (KE #1392) leads to altered, NO levels (KE #2067)  </td> <td> Evidence for Quantitative Understanding of KER: Low  Alterations in NO levels cannot be predicted from relevant measures of oxidative stress changes, such as increased ROS production and antioxidant enzyme activity. Nevertheless, a general decrease in NO is observed following ROS production.  </td> </tr> <tr> <td> Altered signaling pathways (KE #2066) leads to altered, NO levels (KE #2067)  </td> <td> Evidence for Quantitative Understanding of KER: Low  Altered NO, iNOS and eNOS levels occur in response to altered signaling pathways; however, a model has not been established to predict the changes in NO levels. Different models, stressors, time scales, doses and dose rates make trends difficult to identify. The studies investigated the levels of different altered signaling pathway molecules and their effects on NO levels, making it difficult to compare and identify quantitative relationships across the results.   </td> </tr> <tr> <td> Altered, NO levels (KE #2067) leads to increase, endothelial dysfunction (KE #2068)  </td> <td> Evidence for Quantitative Understanding of KER: Low  Increased vascular tension occurs consistently with decreased NO levels. Although many studies quantitatively measure a change in endothelial function after changes in NO levels, no model has been established. Each study cited used different models, stressors, time scales, doses and dose rates, which makes it difficult to determine if response levels are consistent between studies. </td> </tr> </tbody> </table>

The present AOP serves as a platform to promote broader collaborative efforts to understand non-cancer health risks from radiation exposures. It will be a foundational AOP of regulatory interest to researchers seeking areas of knowledge gaps to prioritize research in understanding mechanisms of CVD. The AOP is also relevant to space agencies and clinicians working to improve the guidance on health risks from long-term spaceflight and radiotherapy treatments, respectively. The WOE for this AOP may also inform parameters in biologically based risk models and can serve to develop countermeasures to protect the cardiovascular systems of future space travelers on deep space missions. The present qualitative AOP can be used to guide the design of experiments that will provide quantitative understanding for the KERs to support risk-model development and inform additional guidelines for radiation protection; additionally, the identified research gaps could help prioritize research needs for funding strategies.

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Abdel-Magied, N., & Shedid, S. M. (2020). Impact of zinc oxide nanoparticles on thioredoxin-interacting protein and asymmetric dimethylarginine as biochemical indicators of cardiovascular disorders in gamma-irradiated rats. Environmental Toxicology, 35(4), 430–442. https://doi.org/10.1002/tox.22879  Ashcroft, M., Kubbutat, M. H. G., & Vousden, K. H. (1999). Regulation of p53 Function and Stability by Phosphorylation. Molecular and Cellular Biology, 19(3), 1751. https://doi.org/10.1128/MCB.19.3.1751  Azimzadeh, O., Subramanian, V., Sievert, W., Merl-Pham, J., Oleksenko, K., Rosemann, M., Multhoff, G., Atkinson, M. J., & Tapio, S. (2021). Activation of pparα by fenofibrate attenuates the effect of local heart high dose irradiation on the mouse cardiac proteome. Biomedicines, 9(12). https://doi.org/10.3390/biomedicines9121845  Azimzadeh, O. et al. (2017), “Proteome analysis of irradiated endothelial cells reveals persistent alteration in protein degradation and the RhoGDI and NO signalling pathways”, International Journal of Radiation Biology, Vol. 93/9, Informa, London, https://doi.org/10.1080/09553002.2017.1339332.  Azimzadeh, O. et al. (2015), “Integrative Proteomics and Targeted Transcriptomics Analyses in Cardiac Endothelial Cells Unravel Mechanisms of Long-Term Radiation-Induced Vascular Dysfunction”, Journal of Proteome Research, Vol. 14/2, American Chemical Society, Washington, https://doi.org/10.1021/pr501141b.  Azizova, T. V., Batistatou, E., Grigorieva, E. S., McNamee, R., Wakeford, R., Liu, H., De Vocht, F., & Agius, R. M. (2018). An Assessment of Radiation-Associated Risks of Mortality from Circulatory Disease in the Cohorts of Mayak and Sellafield Nuclear Workers. Radiation Research, 189(4), 371–388. https://doi.org/10.1667/RR14468.1  Baeyens, N., Bandyopadhyay, C., Coon, B. G., Yun, S., & Schwartz, M. A. (2016). Endothelial fluid shear stress sensing in vascular health and disease. The Journal of Clinical Investigation, 126(3), 821–828. https://doi.org/10.1172/JCI83083  Baker, J. E., Fish, B. L., Su, J., Haworth, S. T., Strande, J. L., Komorowski, R. A., Migrino, R. Q., Doppalapudi, A., Harmann, L., Allen Li, X., Hopewell, J. W., & Moulder, J. E. (2009). 10 Gy total body irradiation increases risk of coronary sclerosis, degeneration of heart structure and function in a rat model. International Journal of Radiation Biology, 85(12), 1089–1100. https://doi.org/10.3109/09553000903264473  Baker, J. E., Moulder, J. E., & Hopewell, J. W. (2011). Radiation as a Risk Factor for Cardiovascular Disease. Antioxidant and Redox Signaling, 15(7). https://doi.org/10.1089/ars.2010.3742  Balasubramanian, D. (2000). Ultraviolet radiation and cataract. Journal of Ocular Pharmacology and Therapeutics, 16(3), 285–297. https://doi.org/10.1089/jop.2000.16.285  Baran, R., Marchal, S., Campos, S. G., Rehnberg, E., Tabury, K., Baselet, B., Wehland, M., Grimm, D., & Baatout, S. (2021). The Cardiovascular System in Space: Focus on In Vivo and In Vitro Studies. Biomedicines 2022, Vol. 10, Page 59, 10(1), 59. https://doi.org/10.3390/BIOMEDICINES10010059  Baselet, B., Belmans, N., Coninx, E., Lowe, D., Janssen, A., Michaux, A., Tabury, K., Raj, K., Quintens, R., Benotmane, M. A., Baatout, S., Sonveaux, P., & Aerts, A. (2017). Functional gene analysis reveals cell cycle changes and inflammation in endothelial cells irradiated with a single X-ray dose. Frontiers in Pharmacology, 8. https://doi.org/10.3389/fphar.2017.00213  Belzile-Dugas, E., & Eisenberg, M. J. (2021). Radiation‐Induced Cardiovascular Disease: Review of an Underrecognized Pathology. Journal of the American Heart Association: Cardiovascular and Cerebrovascular Disease, 10(18), 21686. https://doi.org/10.1161/JAHA.121.021686  Berk, B. C., & Korshunov, V. A. (2006). Genetic determinants of vascular remodelling. Canadian Journal of Cardiology, 22(SUPPL. B). https://doi.org/10.1016/s0828-282x(06)70980-1  Boerma, M., Nelson, G. A., Sridharan, V., Mao, X.-W., Koturbash, I., & Hauer-Jensen, M. (2015). Space radiation and cardiovascular disease risk. World Journal of Cardiology, 7(12), 882. https://doi.org/10.4330/wjc.v7.i12.882  Boerma, M., Sridharan, V., Mao, X.-W., Nelson, G. A., Cheema, A. K., Koturbash, I., Singh, S. P., Tackett, A. J., & Hauer-Jensen, M. (2016). Effects of Ionizing Radiation on the Heart. Mutation Research - Reviews in Mutation Research, 770, 319–327. https://doi.org/10.1016/j.mrrev.2016.07.003  Bonetti, P. O., Lerman, L. O., & Lerman, A. (2003). Endothelial dysfunction: A marker of atherosclerotic risk. Arteriosclerosis, Thrombosis, and Vascular Biology, 23(2), 168–175. https://doi.org/10.1161/01.ATV.0000051384.43104.FC  Borghini, A., Luca Gianicolo, E. A., Picano, E., & Andreassi, M. G. (2013). Ionizing radiation and atherosclerosis: Current knowledge and future challenges. Atherosclerosis, 230(1), 40–47. https://doi.org/10.1016/j.atherosclerosis.2013.06.010  Bray, F., Laversanne, M., Weiderpass, E., & Soerjomataram, I. (2021). The ever-increasing importance of cancer as a leading cause of premature death worldwide. Cancer, 127(16), 3029–3030. https://doi.org/10.1002/cncr.33587  Carmeliet, P., & Jain, R. K. (2011). Molecular mechanisms and clinical applications of angiogenesis. https://doi.org/10.1038/nature10144  Cervelli, T., Panetta, D., Navarra, T., Gadhiri, S., Salvadori, P., Galli, A., Caramella, D., Basta, G., Picano, E., & Del Turco, S. (2017). A New Natural Antioxidant Mixture Protects against Oxidative and DNA Damage in Endothelial Cell Exposed to Low-Dose Irradiation. Oxidative Medicine and Cellular Longevity, 2017. https://doi.org/10.1155/2017/9085947  Chang, P. Y., Zhang, B. Y., Cui, S., Qu, C., Shao, L. H., Xu, T. K., Qu, Y. Q., Dong, L. H., & Wang, J. (2017). MSC-derived cytokines repair radiation-induced intra-villi microvascular injury. Oncotarget, 8(50). https://doi.org/10.18632/oncotarget.21236  Chauhan, V., Hamada, N., Monceau, V., Ebrahimian, T., Adam, N., Wilkins, R. C., Sebastian, S., Patel, Z. S., Huff, J. L., Simonetto, C., Iwasaki, T., Kaiser, J. C., Salomaa, S., Moertl, S., & Azimzadeh, O. (2021). Expert consultation is vital for adverse outcome pathway development: a case example of cardiovascular effects of ionizing radiation. International Journal of Radiation Biology, 97(11). https://doi.org/10.1080/09553002.2021.1969466  Cheng, Y. P., Zhang, H. J., Su, Y. T., Meng, X. X., Xie, X. P., Chang, Y. M., & Bao, J. X. (2017). Acid sphingomyelinase/ceramide regulates carotid intima-media thickness in simulated weightless rats. Pflugers Archiv European Journal of Physiology, 469(5–6). https://doi.org/10.1007/s00424-017-1969-z  Chiuve, S. E., McCullough, M. L., Sacks, F. M., & Rimm, E. B. (2006). Healthy lifestyle factors in the primary prevention of coronary heart disease among men: Benefits among users and nonusers of lipid-lowering and antihypertensive medications. Circulation, 114(2), 160–167. https://doi.org/10.1161/CIRCULATIONAHA.106.621417  Chowdhury, R., Shah, D., & Payal, A. (2017). Healthy worker effect phenomenon: Revisited with emphasis on statistical methods-A review. Indian Journal of Occupational and Environmental Medicine, 21(1), 2–8. https://doi.org/10.4103/ijoem.IJOEM_53_16  Cohn, J. N., Quyyumi, A. A., Hollenberg, N. K., & Jamerson, K. A. (2004). Surrogate markers for cardiovascular disease: Functional markers. Circulation, 109(25).  de Jager, T. L., A. E. Cockrell, S. S. Du Plessis (2017), “Ultraviolet Light Induced Generation of Reactive Oxygen Species”, in: Ultraviolet Light in Human Health, Diseases and Environment, vol 996. Springer Cham, https://doi.org/10.1007/978-3-319-56017-5_2. Deanfield, J. E., Halcox, J. P., & Rabelink, T. J. (2007). Endothelial function and dysfunction: Testing and clinical relevance. Circulation, 115(10), 1285–1295. https://doi.org/10.1161/CIRCULATIONAHA.106.652859  Desouky, O., Ding, N., & Zhou, G. (2015). Targeted and non-targeted effects of ionizing radiation. Journal of Radiation Research and Applied Sciences, 8(2), 247–254. https://doi.org/10.1016/J.JRRAS.2015.03.003  Di Maggio, F. M., Minafra, L., Forte, G. I., Cammarata, F. P., Lio, D., Messa, C., Gilardi, M. C., & Bravatà, V. (2015). Portrait of inflammatory response to ionizing radiation treatment. In Journal of Inflammation (Vol. 12, Issue 1, pp. 1–11). BioMed Central Ltd. https://doi.org/10.1186/s12950-015-0058-3  Ding, J., Yu, M., Jiang, J., Luo, Y., Zhang, Q., Wang, S., Yang, F., Wang, A., Wang, L., Zhuang, M., Wu, S., Zhang, Q., Xia, Y., & Lu, D. (2020). Angiotensin II Decreases Endothelial Nitric Oxide Synthase Phosphorylation via AT1R Nox/ROS/PP2A Pathway. Frontiers in Physiology, 11, 1245. https://doi.org/10.3389/fphys.2020.566410  Dörr, W. (2015). Radiobiology of tissue reactions. Annals of the ICRP, 44, 58–68. https://doi.org/10.1177/0146645314560686  Durante, M., & Cucinotta, F. A. (2008). Heavy ion carcinogenesis and human space exploration. Nature Reviews Cancer, 8(6), 465–472. https://doi.org/10.1038/nrc2391  Eaton, J. W. (1994). UV-mediated cataractogenesis: A radical perspective. Documenta Ophthalmologica, 88(3–4), 233–242. https://doi.org/10.1007/BF01203677  EPRI. (2020). Cardiovascular Risks from Low Dose Radiation Exposure: Review and Scientific Appraisal of the Literature.  Fletcher, A. E. (2010). Free radicals, antioxidants and eye diseases: Evidence from epidemiological studies on cataract and age-related macular degeneration. Ophthalmic Research, 44(3), 191–198. https://doi.org/10.1159/000316476  Forrester, S. J., Kikuchi, D. S., Hernandes, M. S., Xu, Q., & Griendling, K. K. (2018). Reactive oxygen species in metabolic and inflammatory signaling. Circulation Research, 122(6), 877–902. https://doi.org/10.1161/CIRCRESAHA.117.311401  Förstermann, U. (2010). Nitric oxide and oxidative stress in vascular disease. Pflugers Archiv European Journal of Physiology, 459(6), 923–939. https://doi.org/10.1007/s00424-010-0808-2  Förstermann, U., & Münzel, T. (2006). Endothelial Nitric Oxide Synthase in Vascular Disease. Circulation, 113(13), 1708–1714. https://doi.org/10.1161/CIRCULATIONAHA.105.602532  Francula-Zaninovic, S., & Nola, I. A. (2018). Management of Measurable Variable Cardiovascular Disease’ Risk Factors. Current Cardiology Reviews, 14(3), 153–163. https://doi.org/10.2174/1573403x14666180222102312  Fuji, S., Matsushita, S., Hyodo, K., Osaka, M., Sakamoto, H., Tanioka, K., Miyakawa, K., Kubota, M., Hiramatsu, Y., & Tokunaga, C. (2016). Association between endothelial function and micro-vascular remodeling measured by synchrotron radiation pulmonary micro-angiography in pulmonary arterial hypertension. General Thoracic and Cardiovascular Surgery, 64(10), 597–603. https://doi.org/10.1007/s11748-016-0684-6  Ganea, E., & Harding, J. J. (2006). Glutathione-related enzymes and the eye. In Current Eye Research (Vol. 31, Issue 1, pp. 1–11). Taylor & Francis. https://doi.org/10.1080/02713680500477347  Gen, S. W. (2004). The functional interactions between the p53 and MAPK signaling pathways. Cancer Biology and Therapy, 3(2), 156–161. https://doi.org/10.4161/cbt.3.2.614  Gillies, M., Richardson, D. B., Cardis, E., Daniels, R. D., O’Hagan, J. A., Haylock, R., Laurier, D., Leuraud, K., Moissonnier, M., Schubauer-Berigan, M. K., Thierry-Chef, I., & Kesminiene, A. (2017). Mortality from circulatory diseases and other non-cancer outcomes among nuclear workers in France, the United Kingdom and the United States (inworks). Radiation Research, 188(3), 276–290. https://doi.org/10.1667/RR14608.1  Grabham, P., & Sharma, P. (2013). The effects of radiation on angiogenesis. Vascular Cell, 5(1), 19. https://doi.org/10.1186/2045-824X-5-19  Hamada, N., Kawano, K. I., Nomura, T., Furukawa, K., Yusoff, F. M., Maruhashi, T., Maeda, M., Nakashima, A., & Higashi, Y. (2021). Vascular damage in the aorta of wild-type mice exposed to ionizing radiation: Sparing and enhancing effects of dose protraction. Cancers, 13(21). https://doi.org/10.3390/cancers13215344  Hamada, N., Kawano, K. I., Yusoff, F. M., Furukawa, K., Nakashima, A., Maeda, M., Yasuda, H., Maruhashi, T., & Higashi, Y. (2020). Ionizing irradiation induces vascular damage in the aorta of wild-type mice. Cancers, 12(10), 1–11. https://doi.org/10.3390/cancers12103030  Hasan, H. F., Radwan, R. R., & Galal, S. M. (2020). Bradykinin-potentiating factor isolated from Leiurus quinquestriatus scorpion venom alleviates cardiomyopathy in irradiated rats via remodelling of the RAAS pathway. Clinical and Experimental Pharmacology and Physiology, 47(2), 263–273. https://doi.org/10.1111/1440-1681.13202  Hassan, B., Akcakanat, A., Holder, A. M., & Meric-Bernstam, F. (2013). Targeting the PI3-Kinase/Akt/mTOR Signaling Pathway. Surgical Oncology Clinics of North America, 22(4), 641–664. https://doi.org/10.1016/j.soc.2013.06.008  Hatoum, O. A., Otterson, M. F., Kopelman, D., Miura, H., Sukhotnik, I., Larsen, B. T., Selle, R. M., Moulder, J. E., & Gutterman, D. D. (2006). Radiation induces endothelial dysfunction in murine intestinal arterioles via enhanced production of reactive oxygen species. Arteriosclerosis, Thrombosis, and Vascular Biology, 26(2), 287–294. https://doi.org/10.1161/01.ATV.0000198399.40584.8c  Heald, C. L., Fowkes, F. G. R., Murray, G. D., & Price, J. F. (2006). Risk of mortality and cardiovascular disease associated with the ankle-brachial index: Systematic review. Atherosclerosis, 189(1), 61–69. https://doi.org/10.1016/J.ATHEROSCLEROSIS.2006.03.011  Hemmings, B. A., & Restuccia, D. F. (2012). PI3K-PKB/Akt pathway. Cold Spring Harbor Perspectives in Biology, 4(9). https://doi.org/10.1101/cshperspect.a011189  Hirakawa, M., Oike, M., Masuda, K., & Ito, Y. (2002). Tumor cell apoptosis by irradiation-induced nitric oxide production in vascular endothelium. Cancer Research, 62(5), 1450–1457.  Hladik, D., & Tapio, S. (2016). Effects of ionizing radiation on the mammalian brain. Mutation Research - Reviews in Mutation Research, 770, 219–230. https://doi.org/10.1016/j.mrrev.2016.08.003  Hodis, H. N., Mack, W. J., LaBree, L., Selzer, R. H., Liu, C. R., Liu, C. H., & Azen, S. P. (1998). The role of carotid arterial intima - Media thickness in predicting clinical coronary events. Annals of Internal Medicine, 128(4), 262–269. https://doi.org/10.7326/0003-4819-128-4-199802150-00002  Hong, C. W., Kim, Y. M., Pyo, H., Lee, J. H., Kim, S., Lee, S., & Noh, J. M. (2013). Involvement of inducible nitric oxide synthase in radiation-Induced vascular endothelial damage. Journal of Radiation Research, 54(6), 1036–1042. https://doi.org/10.1093/jrr/rrt066  Hsu, T., Nguyen-Tran, H. H., & Trojanowska, M. (2019). Active roles of dysfunctional vascular endothelium in fibrosis and cancer. Journal of Biomedical Science, 26(1), 1–12. https://doi.org/10.1186/s12929-019-0580-3  Huang, Y. et al. (2021), “Mesenchymal Stem Cell-Conditioned Medium Protects Hippocampal Neurons From Radiation Damage by Suppressing Oxidative Stress and Apoptosis”, Dose-response, Vol. 19/1, Sage Publishing, https://doi.org/10.1177/1559325820984944  Hughson, R. L., Helm, A., & Durante, M. (2018). Heart in space: Effect of the extraterrestrial environment on the cardiovascular system. In Nature Reviews Cardiology (Vol. 15, Issue 3). https://doi.org/10.1038/nrcardio.2017.157  Hughson, R. L., Robertson, A. D., Arbeille, P., Shoemaker, J. K., Rush, J. W. E., Fraser, K. S., & Greaves, D. K. (2016). Increased postflight carotid artery stiffness and inflight insulin resistance resulting from 6-mo spaceflight in male and female astronauts. American Journal of Physiology - Heart and Circulatory Physiology, 310(5), H628–H638. https://doi.org/10.1152/ajpheart.00802.2015  Ismail, A. F. M., & El-Sonbaty, S. M. (2016). Fermentation enhances Ginkgo biloba protective role on gamma-irradiation induced neuroinflammatory gene expression and stress hormones in rat brain. Journal of Photochemistry and Photobiology B: Biology, 158, 154–163. https://doi.org/10.1016/j.jphotobiol.2016.02.039  Ivanov, V. K., Maksioutov, M. A., Chekin, S. Y., Petrov, A. V., Biryukov, A. P., Kruglova, Z. G., Matyash, V. A., Tsyb, A. F., Manton, K. G., & Kravchenko, J. S. (2006). The risk of radiation-induced cerebrovascular disease in chernobyl emergency workers. Health Physics, 90(3), 199–207. https://doi.org/10.1097/01.HP.0000175835.31663.EA  Karam, H. M., & Radwan, R. R. (2019). Metformin modulates cardiac endothelial dysfunction, oxidative stress and inflammation in irradiated rats: A new perspective of an antidiabetic drug. Clinical and Experimental Pharmacology and Physiology, 46(12), 1124–1132. https://doi.org/10.1111/1440-1681.13148  Karar, J., & Maity, A. (2011). PI3K/AKT/mTOR Pathway in Angiogenesis. Frontiers in Molecular Neuroscience, 4, 51. https://doi.org/10.3389/fnmol.2011.00051  Karimi, N., Monfared, A., Haddadi, G., Soleymani, A., Mohammadi, E., Hajian-Tilaki, K., & Borzoueisileh, S. (2017). Radioprotective effect of hesperidin on reducing oxidative stress in the lens tissue of rats. International Journal of Pharmaceutical Investigation, 7(3), 149. https://doi.org/10.4103/jphi.jphi_60_17  Kashcheev, V. V., Chekin, S. Y., Karpenko, S. V., Maksioutov, M. A., Menyaylo, A. N., Tumanov, K. A., Kochergina, E. V., Kashcheeva, P. V., Gorsky, A. I., Shchukina, N. V., Lovachev, S. S., Vlasov, O. K., & Ivanov, V. K. (2017). Radiation Risk of Cardiovascular Diseases in the Cohort of Russian Emergency Workers of the Chernobyl Accident. Health Physics, 113(1), 23–29. https://doi.org/10.1097/HP.0000000000000670  Kessler, E. L., Rivaud, M. R., Vos, M. A., & Van Veen, T. A. B. (2019). Sex-specific influence on cardiac structural remodeling and therapy in cardiovascular disease. Biology of Sex Differences, 10(1). https://doi.org/10.1186/s13293-019-0223-0  Konukoglu, D., & Uzun, H. (2016). Endothelial dysfunction and hypertension. In Advances in Experimental Medicine and Biology (Vol. 956, pp. 511–540). Adv Exp Med Biol. https://doi.org/10.1007/5584_2016_90  Korpela, E., & Liu, S. K. (2014). Endothelial perturbations and therapeutic strategies in normal tissue radiation damage. Radiation Oncology, 9(1). https://doi.org/10.1186/s13014-014-0266-7  Kozbenko, T., Adam, N., Lai, V., Sandhu, S., Kuan, J., Flores, D., Appleby, M., Parker, H., Hocking, R., Tsaioun, K., Yauk, C., Wilkins, R., & Chauhan, V. (2022). Deploying elements of scoping review methods for adverse outcome pathway development: a space travel case example. International Journal of Radiation Biology, 1–12. https://doi.org/10.1080/09553002.2022.2110306  Krüger-Genge, A., Blocki, A., Franke, R.-P., & Jung, F. (2019). Vascular Endothelial Cell Biology: An Update. International Journal of Molecular Sciences, 20(18), 4411. https://doi.org/10.3390/ijms20184411  Lee, M. S., Finch, W., & Mahmud, E. (2013). Cardiovascular complications of radiotherapy. American Journal of Cardiology, 112(10), 1688–1696. https://doi.org/10.1016/j.amjcard.2013.07.031  Li, J., De Leon, H., Ebato, B., Cui, J., Todd, J., Chronos, N. A. F., & Robinson, K. A. (2002). Endovascular irradiation impairs vascular functional responses in noninjured pig coronary arteries. Cardiovascular Radiation Medicine, 152–162. https://doi.org/10.1016/S1522-1865(03)00096-9  Little, M. P., Azizova, T. V., & Hamada, N. (2021). Low- and moderate-dose non-cancer effects of ionizing radiation in directly exposed individuals, especially circulatory and ocular diseases: a review of the epidemiology. International Journal of Radiation Biology, 97(6), 782–803. https://doi.org/10.1080/09553002.2021.1876955  Luiking, Y. C., Engelen, M. P. K. J., & Deutz, N. E. P. (2010). Regulation of nitric oxide production in health and disease. Current Opinion in Clinical Nutrition and Metabolic Care, 13(1), 97–104. https://doi.org/10.1097/MCO.0B013E328332F99D  Lumniczky, K., Impens, N., Armengol, G., Candéias, S., Georgakilas, A. G., Hornhardt, S., Martin, O. A., Rödel, F., & Schaue, D. (2021). Low dose ionizing radiation effects on the immune system. Environment International, 149, 106212. https://doi.org/10.1016/j.envint.2020.106212  Maier, J. A. M., Cialdai, F., Monici, M., & Morbidelli, L. (2015). The impact of microgravity and hypergravity on endothelial cells. BioMed Research International, 2015. https://doi.org/10.1155/2015/434803  Matsubara, K., Higaki, T., Matsubara, Y., & Nawa, A. (2015). Nitric oxide and reactive oxygen species in the pathogenesis of preeclampsia. International Journal of Molecular Sciences, 16(3), 4600–4614. https://doi.org/10.3390/ijms16034600  Menezes, K. M., Wang, H., Hada, M., & Saganti, P. B. (2018). Radiation Matters of the Heart: A Mini Review. Frontiers in Cardiovascular Medicine, 5, 83. https://doi.org/10.3389/FCVM.2018.00083  Millatt, L. J., Abdel-Rahman, E. M., & Siragy, H. M. (1999). Angiotensin II and nitric oxide: A question of balance. Regulatory Peptides, 81(1–3), 1–10. https://doi.org/10.1016/S0167-0115(99)00027-0  Mitchell, A., Pimenta, D., Gill, J., Ahmad, H., & Bogle, R. (2019). Cardiovascular effects of space radiation: implications for future human deep space exploration. European Journal of Preventive Cardiology, 26(16), 1707–1714. https://doi.org/10.1177/2047487319831497  Mittal, M., Siddiqui, M. R., Tran, K., Reddy, S. P., & Malik, A. B. (2014). Reactive oxygen species in inflammation and tissue injury. Antioxidants and Redox Signaling, 20(7), 1126–1167. https://doi.org/10.1089/ars.2012.5149  Mosca, L., Barrett-Connor, E., & Kass Wenger, N. (2011). Sex/gender differences in cardiovascular disease prevention: What a difference a decade makes. Circulation, 124(19), 2145–2154. https://doi.org/10.1161/CIRCULATIONAHA.110.968792  Mozaffarian, D., Wilson, P. W. F., & Kannel, W. B. (2008). Beyond Established and Novel Risk Factors. Circulation, 117(23), 3031–3038. https://doi.org/10.1161/CIRCULATIONAHA.107.738732  Nagane, M., Yasui, H., Kuppusamy, P., Yamashita, T., & Inanami, O. (2021). DNA damage response in vascular endothelial senescence: Implication for radiation-induced cardiovascular diseases. Journal of Radiation Research, 62(4), 564–573. https://doi.org/10.1093/jrr/rrab032  Nathan, C., & Xie, Q. wen. (1994). Nitric oxide synthases: Roles, tolls, and controls. Cell, 78(6), 915–918. https://doi.org/10.1016/0092-8674(94)90266-6  Norbury, J. W., Schimmerling, W., Slaba, T. C., Azzam, E. I., Badavi, F. F., Baiocco, G., Benton, E., Bindi, V., Blakely, E. A., Blattnig, S. R., Boothman, D. A., Borak, T. B., Britten, R. A., Curtis, S., Dingfelder, M., Durante, M., Dynan, W. S., Eisch, A. J., Robin Elgart, S., … Zeitlin, C. J. (2016). Galactic cosmic ray simulation at the NASA Space Radiation Laboratory. Life Sciences in Space Research, 8, 38–51. https://doi.org/10.1016/j.lssr.2016.02.001  North, B. J., & Sinclair, D. A. (2012). The intersection between aging and cardiovascular disease. Circulation Research, 110(8), 1097–1108. https://doi.org/10.1161/CIRCRESAHA.111.246876  Ozasa, K., Shimizu, Y., Suyama, A., Kasagi, F., Soda, M., Grant, E. J., Sakata, R., Sugiyama, H., & Kodama, K. (2012). Studies of the mortality of atomic bomb survivors, report 14, 1950-2003: An overview of cancer and noncancer diseases. In Radiation Research (Vol. 177, Issue 3). https://doi.org/10.1667/RR2629.1  Padgaonkar, V. A., Leverenz, V. R., Bhat, A. V., Pelliccia, S. E., & Giblin, F. J. (2015). Thioredoxin reductase activity may be more important than GSH level in protecting human lens epithelial cells against UVA light. Photochemistry and Photobiology, 91(2), 387–396. https://doi.org/10.1111/php.12404  Patel, Z. S., Brunstetter, T. J., Tarver, W. J., Whitmire, A. M., Zwart, S. R., Smith, S. M., & Huff, J. L. (2020). Red risks for a journey to the red planet: The highest priority human health risks for a mission to Mars. Npj Microgravity, 6(1), 1–13. https://doi.org/10.1038/s41526-020-00124-6  Ping, Z., Peng, Y., Lang, H., Xinyong, C., Zhiyi, Z., Xiaocheng, W., Hong, Z., & Liang, S. (2020). Oxidative Stress in Radiation-Induced Cardiotoxicity. Oxidative Medicine and Cellular Longevity, 2020. https://doi.org/10.1155/2020/3579143  Polak, J. F., Pencina, M. J., Pencina, K. M., O’Donnell, C. J., Wolf, P. A., & Ralph B. D’Agostino, S. (2011). Carotid-Wall Intima–Media Thickness and Cardiovascular Events. The New England Journal of Medicine, 365(3), 213. https://doi.org/10.1056/NEJMOA1012592  Preston, D. L., Shimizu, Y., Pierce, D. A., Suyama, A., & Mabuchi, K. (2003). Studies of Mortality of Atomic Bomb Survivors. Report 13: Solid Cancer and Noncancer Disease Mortality: 1950–1997. Https://Doi.Org/10.1667/RR3049, 160(4), 381–407. https://doi.org/10.1667/RR3049  Pries, A. R., Reglin, B., & Secomb, T. W. (2001). Structural adaptation of microvascular networks: Functional roles of adaptive responses. American Journal of Physiology - Heart and Circulatory Physiology, 281(3). https://doi.org/10.1152/ajpheart.2001.281.3.h1015  Ramadan, R., Baatout, S., Aerts, A., & Leybaert, L. (2021). The role of connexin proteins and their channels in radiation-induced atherosclerosis. Cellular and Molecular Life Sciences, 78(7), 3087–3103. https://doi.org/10.1007/s00018-020-03716-3  Ramadan, R., Vromans, E., Anang, D. C., Goetschalckx, I., Hoorelbeke, D., Decrock, E., Baatout, S., Leybaert, L., & Aerts, A. (2020). Connexin43 Hemichannel Targeting With TAT-Gap19 Alleviates Radiation-Induced Endothelial Cell Damage. Frontiers in Pharmacology, 11. https://doi.org/10.3389/fphar.2020.00212  Rehman, M. U., Jawaid, P., Uchiyama, H., & Kondo, T. (2016). Comparison of free radicals formation induced by cold atmospheric plasma, ultrasound, and ionizing radiation. Archives of Biochemistry and Biophysics, 605, 19–25. https://doi.org/10.1016/j.abb.2016.04.005  Rios, A., Joshi, R., & Shin, H. (2020). Quantifying 60 Years of Gender Bias in Biomedical Research with Word Embeddings. 1–13. https://doi.org/10.18653/v1/2020.bionlp-1.1  Rodgers, J. L., Jones, J., Bolleddu, S. I., Vanthenapalli, S., Rodgers, L. E., Shah, K., Karia, K., & Panguluri, S. K. (2019). Cardiovascular risks associated with gender and aging. Journal of Cardiovascular Development and Disease, 6(2). https://doi.org/10.3390/jcdd6020019  Sadhukhan, R., Leung, J. W. C., Garg, S., Krager, K. J., Savenka, A. V., Basnakian, A. G., & Pathak, R. (2020). Fractionated radiation suppresses Kruppel-like factor 2 pathway to a greater extent than by single exposure to the same total dose. Scientific Reports, 10(1), 1–13. https://doi.org/10.1038/s41598-020-64672-3  Sakata, K., Kondo, T., Mizuno, N., Shoji, M., Yasui, H., Yamamori, T., Inanami, O., Yokoo, H., Yoshimura, N., & Hattori, Y. (2015). Roles of ROS and PKC-βII in ionizing radiation-induced eNOS activation in human vascular endothelial cells. Vascular Pharmacology, 70, 55–65. https://doi.org/10.1016/j.vph.2015.03.016  Santamaría, R., González-Álvarez, M., Delgado, R., Esteban, S., & Arroyo, A. G. (2020). Remodeling of the Microvasculature: May the Blood Flow Be With You. Frontiers in Physiology, 11, 1256. https://doi.org/10.3389/FPHYS.2020.586852/XML/NLM  Sárközy, M., Gáspár, R., Zvara, A., Kiscsatári, L., Varga, Z., Kővári, B., Kovács, M. G., Szűcs, G., Fabian, G., Diószegi, P., Cserni, G., Puskás, L. G., Thum, T., Kahán, Z., Csont, T., & Batkai, S. (2019). Selective heart irradiation induces cardiac overexpression of the pro-hypertrophic miR-212. Frontiers in Oncology, 9(JUN), 598. https://doi.org/10.3389/fonc.2019.00598  Schaue, D., Micewicz, E. D., Ratikan, J. A., Xie, M. W., Cheng, G., & McBride, W. H. (2015). Radiation and Inflammation. In Seminars in Radiation Oncology (Vol. 25, Issue 1, pp. 4–10). W.B. Saunders. https://doi.org/10.1016/j.semradonc.2014.07.007  Schiffrin, E. L. (2008). Oxidative stress, nitric oxide synthase, and superoxide dismutase: A matter of imbalance underlies endothelial dysfunction in the human coronary circulation. Hypertension, 51(1), 31–32. https://doi.org/10.1161/HYPERTENSIONAHA.107.103226  Schmidt-Ullrich, R. K., Dent, P., Grant, S., Mikkelsen, R. B., & Valerie, K. (2000). Signal transduction and cellular radiation responses. Radiation Research, 153(3), 245–257. https://doi.org/10.1667/0033-7587(2000)153[0245:STACRR]2.0.CO;2  Schultz, W. M., Kelli, H. M., Lisko, J. C., Varghese, T., Shen, J., Sandesara, P., Quyyumi, A. A., Taylor, H. A., Gulati, M., Harold, J. G., Mieres, J. H., Ferdinand, K. C., Mensah, G. A., & Sperling, L. S. (2018). Socioeconomic Status and Cardiovascular Outcomes: Challenges and Interventions. Circulation, 137(20). https://doi.org/10.1161/circulationaha.117.029652  Senoner, T., & Dichtl, W. (2019). Oxidative stress in cardiovascular diseases: Still a therapeutic target? Nutrients, 11(9). https://doi.org/10.3390/nu11092090  Shen, Y., Jiang, X., Meng, L., Xia, C., Zhang, L., & Xin, Y. (2018). Transplantation of bone marrow mesenchymal stem cells prevents radiation-induced artery injury by suppressing oxidative stress and inflammation. Oxidative Medicine and Cellular Longevity, 2018. https://doi.org/10.1155/2018/5942916  Shi, F., Wang, Y. C., Zhao, T. Z., Zhang, S., Du, T. Y., Yang, C. Bin, Li, Y. H., & Sun, X. Q. (2012). Effects of simulated microgravity on human umbilical vein endothelial cell angiogenesis and role of the PI3K-Akt-eNOS signal pathway. PLoS ONE, 7(7). https://doi.org/10.1371/journal.pone.0040365  Shimizu, Y., Kodama, K., Nishi, N., Kasagi, F., Suyama, A., Soda, M., Grant, E. J., Sugiyama, H., Sakata, R., Moriwaki, H., Hayashi, M., Konda, M., & Shore, R. E. (2010). Radiation exposure and circulatory disease risk: Hiroshima and Nagasaki atomic bomb survivor data, 1950-2003. The BMJ, 340(7739), 193. https://doi.org/10.1136/BMJ.B5349  Siamwala, J. H., Reddy, S. H., Majumder, S., Kolluru, G. K., Muley, A., Sinha, S., & Chatterjee, S. (2010). Simulated microgravity perturbs actin polymerization to promote nitric oxide-associated migration in human immortalized Eahy926 cells. Protoplasma, 242(1), 3–12. https://doi.org/10.1007/s00709-010-0114-z  Slezak, J., Kura, B., Babal, P., Barancik, M., Ferko, M., Frimmel, K., Kalocayova, B., Kukreja, R. C., Lazou, A., Mezesova, L., Okruhlicova, L., Ravingerova, T., Singal, P. K., Bacova, B. S., Viczenczova, C., Vrbjar, N., & Tribulova, N. (2017). Potential markers and metabolic processes involved in the mechanism of Radiation-Induced heart injury. Canadian Journal of Physiology and Pharmacology, 95(10), 1190–1203. https://doi.org/10.1139/cjpp-2017-0121  Soloviev, A. I., & Kizub, I. V. (2019). Mechanisms of vascular dysfunction evoked by ionizing radiation and possible targets for its pharmacological correction. Biochemical Pharmacology, 159, 121–139. https://doi.org/10.1016/j.bcp.2018.11.019  Sonveaux, P., Brouet, A., Havaux, X., Grégoire, V., Dessy, C., Balligand, J. L., & Feron, O. (2003). Irradiation-induced angiogenesis through the up-regulation of the nitric oxide pathway: Implications for tumor radiotherapy. Cancer Research, 63(5), 1012–1019. https://doi.org/10.1016/s0167-8140(03)80572-8  Soucy, K. G., Lim, H. K., Attarzadeh, D. O., Santhanam, L., Kim, J. H., Bhunia, A. K., Sevinc, B., Ryoo, S., Vazquez, M. E., Nyhan, D., Shoukas, A. A., & Berkowitz, D. E. (2010). Dietary inhibition of xanthine oxidase attenuates radiation-induced endothelial dysfunction in rat aorta. Journal of Applied Physiology, 108(5), 1250–1258. https://doi.org/10.1152/japplphysiol.00946.2009  Soucy, K. G., Lim, H. K., Benjo, A., Santhanam, L., Ryoo, S., Shoukas, A. A., Vazquez, M. E., & Berkowitz, D. E. (2007). Single exposure gamma-irradiation amplifies xanthine oxidase activity and induces endothelial dysfunction in rat aorta. Radiation and Environmental Biophysics, 46(2), 179–186. https://doi.org/10.1007/s00411-006-0090-z  Soucy, K. G., Lim, H. K., Kim, J. H., Oh, Y., Attarzadeh, D. O., Sevinc, B., Kuo, M. M., Shoukas, A. A., Vazquez, M. E., & Berkowitz, D. E. (2011). HZE 56Fe-ion irradiation induces endothelial dysfunction in rat aorta: Role of xanthine oxidase. Radiation Research, 176(4), 474–485. https://doi.org/10.1667/RR2598.1  Sridharan, V., Seawright, J. W., Landes, R. D., Cao, M., Singh, P., Davis, C. M., Mao, X. W., Singh, S. P., Zhang, X., Nelson, G. A., & Boerma, M. (2020). Effects of single-dose protons or oxygen ions on function and structure of the cardiovascular system in male Long Evans rats. Life Sciences in Space Research, 26, 62–68. https://doi.org/10.1016/j.lssr.2020.04.002  Srinivasan, L., Harris, M. C., & Kilpatrick, L. E. (2017). Cytokines and Inflammatory Response in the Fetus and Neonate. In Fetal and Neonatal Physiology, 2-Volume Set (pp. 1241–1254). https://doi.org/10.1016/B978-0-323-35214-7.00128-1  Stanojković, T. P., Matić, I. Z., Petrović, N., Stanković, V., Kopčalić, K., Besu, I., Đorđić Crnogorac, M., Mali&scaron;ić, E., Mirjačić-Martinović, K., Vuletić, A., Bukumirić, Z., Žižak, Ž., Veldwijk, M., Herskind, C., & Nikitović, M. (2020). Evaluation of cytokine expression and circulating immune cell subsets as potential parameters of acute radiation toxicity in prostate cancer patients. Scientific Reports, 10(1). https://doi.org/10.1038/s41598-020-75812-0  Su, Y. T., Cheng, Y. P., Zhang, X., Xie, X. P., Chang, Y. M., & Bao, J. X. (2020). Acid sphingomyelinase/ceramide mediates structural remodeling of cerebral artery and small mesenteric artery in simulated weightless rats. Life Sciences, 243. https://doi.org/10.1016/j.lfs.2019.117253  Summers, S. M., Nguyen, S. V., & Purdy, R. E. (2008). Hindlimb unweighting induces changes in the RhoA-Rho-kinase pathway of the rat abdominal aorta. Vascular Pharmacology, 48(4–6), 208–214. https://doi.org/10.1016/j.vph.2008.03.006  Sylvester, C. B., Abe, J. I., Patel, Z. S., & Grande-Allen, K. J. (2018). Radiation-Induced Cardiovascular Disease: Mechanisms and Importance of Linear Energy Transfer. Frontiers in Cardiovascular Medicine, 5, 1. https://doi.org/10.3389/FCVM.2018.00005  Tahimic, C. G. T., & Globus, R. K. (2017). Redox signaling and its impact on skeletal and vascular responses to spaceflight. International Journal of Molecular Sciences, 18(10). https://doi.org/10.3390/ijms18102153  Takahashi, I., Shimizu, Y., Grant, E. J., Cologne, J., Ozasa, K., & Kodama, K. (2017). Heart disease mortality in the life span study, 1950-2008. In Radiation Research (Vol. 187, Issue 3). https://doi.org/10.1667/RR14347.1  Tapio, S. (2016). Pathology and biology of radiation-induced cardiac disease. Journal of Radiation Research, 57(5), 439–448. https://doi.org/10.1093/jrr/rrw064  Tsao, C. W., Aday, A. W., Almarzooq, Z. I., Alonso, A., Beaton, A. Z., Bittencourt, M. S., Boehme, A. K., Buxton, A. E., Carson, A. P., Commodore-Mensah, Y., Elkind, M. S. V., Evenson, K. R., Eze-Nliam, C., Ferguson, J. F., Generoso, G., Ho, J. E., Kalani, R., Khan, S. S., Kissela, B. M., … Martin, S. S. (2022). Heart Disease and Stroke Statistics-2022 Update: A Report From the American Heart Association. Circulation, 145(8), e153–e639. https://doi.org/10.1161/CIR.0000000000001052/FORMAT/EPUB  Ungvari, Z., Podlutsky, A., Sosnowska, D., Tucsek, Z., Toth, P., Deak, F., Gautam, T., Csiszar, A., & Sonntag, W. E. (2013). Ionizing radiation promotes the acquisition of a senescence-associated secretory phenotype and impairs angiogenic capacity in cerebromicrovascular endothelial cells: Role of increased dna damage and decreased dna repair capacity in microvascular radiosens. Journals of Gerontology - Series A Biological Sciences and Medical Sciences, 68(12 A), 1443–1457. https://doi.org/10.1093/gerona/glt057  UNSCEAR. (2008). UNSCEAR 2006 report. Annex B. Epidemiological evaluation of cardiovascular disease and other non_cancer diseases following radiation exposure.  Valerie, K., Yacoub, A., Hagan, M. P., Curiel, D. T., Fisher, P. B., Grant, S., & Dent, P. (2007). Radiation-induced cell signaling: Inside-out and outside-in. Molecular Cancer Therapeutics, 6(3), 789–801. https://doi.org/10.1158/1535-7163.MCT-06-0596  Van Varik, B. J., Rennenberg, R. J. M. W., Reutelingsperger, C. P., Kroon, A. A., De Leeuw, P. W., & Schurgers, L. J. (2012). Mechanisms of arterial remodeling: Lessons from genetic diseases. Frontiers in Genetics, 3(DEC), 290. https://doi.org/10.3389/FGENE.2012.00290/BIBTEX  Varma, S. D., Kovtun, S., & Hegde, K. R. (2011). Role of ultraviolet irradiation and oxidative stress in cataract formation-medical prevention by nutritional antioxidants and metabolic agonists. Eye and Contact Lens, 37(4), 233–245. https://doi.org/10.1097/ICL.0b013e31821ec4f2  Venkatesulu, B. P., Mahadevan, L. S., Aliru, M. L., Yang, X., Bodd, M. H., Singh, P. K., Yusuf, S. W., Abe, J. ichi, & Krishnan, S. (2018). Radiation-Induced Endothelial Vascular Injury: A Review of Possible Mechanisms. JACC: Basic to Translational Science, 3(4), 563–572. https://doi.org/10.1016/j.jacbts.2018.01.014  Verma, S., Buchanan, M. R., & Anderson, T. J. (2003). Endothelial Function Testing as a Biomarker of Vascular Disease. Circulation, 108(17), 2054–2059. https://doi.org/10.1161/01.CIR.0000089191.72957.ED  Vernice, N. A., Meydan, C., Afshinnekoo, E., & Mason, C. E. (2020). Long-term spaceflight and the cardiovascular system. Precision Clinical Medicine, 3(4), 284–291. https://doi.org/10.1093/PCMEDI/PBAA022  Versari, S., Longinotti, G., Barenghi, L., Maier, J. A. M., & Bradamante, S. (2013). The challenging environment on board the International Space Station affects endothelial cell function by triggering oxidative stress through thioredoxin interacting protein overexpression: the ESA-SPHINX experiment. FASEB Journal, 27(11), 4466–4475. https://doi.org/10.1096/fj.13-229195  Wang, H., Wei, J., Zheng, Q., Meng, L., Xin, Y., Yin, X., & Jiang, X. (2019). Radiation-induced heart disease: a review of classification, mechanism and prevention. International Journal of Biological Sciences, 15(10), 2128. https://doi.org/10.7150/IJBS.35460  Wang, Y., Boerma, M., & Zhou, D. (2016). Ionizing Radiation-Induced Endothelial Cell Senescence and Cardiovascular Diseases. Radiation Research, 186(2), 153–161. https://doi.org/10.1667/RR14445.1  Winham, S. J., de Andrade, M., & Miller, V. M. (2014). Genetics of cardiovascular disease: Importance of sex and ethnicity. Atherosclerosis, 241(1), 219–228. https://doi.org/10.1016/j.atherosclerosis.2015.03.021  Yakerson, A. (2019). Women in clinical trials: A review of policy development and health equity in the Canadian context. International Journal for Equity in Health, 18(1), 56. https://doi.org/10.1186/s12939-019-0954-x  Yan, T., Zhang, T., Mu, W., Qi, Y., Guo, S., Hu, N., Zhao, W., Zhang, S., Wang, Q., Shi, L., & Liu, L. (2020). Ionizing radiation induces BH4 deficiency by downregulating GTP-cyclohydrolase 1, a novel target for preventing and treating radiation enteritis. Biochemical Pharmacology, 180, 114102. https://doi.org/10.1016/j.bcp.2020.114102  Yang, Y. et al. (1998), “The Effect of Catalase Amplification on Immortal Lens Epithelial Cell Lines”, Experimental Eye Research, Vol. 67/6, Elsevier, Amsterdam https://doi.org/10.1006/exer.1998.0560  Yang, E. H., Marmagkiolis, K., Balanescu, D. V., Hakeem, A., Donisan, T., Finch, W., Virmani, R., Herrman, J., Cilingiroglu, M., Grines, C. L., Toutouzas, K., & Iliescu, C. (2021). Radiation-Induced Vascular Disease—A State-of-the-Art Review. Frontiers in Cardiovascular Medicine, 8, 223. https://doi.org/10.3389/FCVM.2021.652761/XML/NLM  Yao, L., Romero, M. J., Toque, H. A., Yang, G., Caldwell, R. B., & Caldwell, R. W. (2010). The role of RhoA/Rho kinase pathway in endothelial dysfunction. Journal of Cardiovascular Disease Research, 1(4), 165–170. https://doi.org/10.4103/0975-3583.74258  Yentrapalli, R., Azimzadeh, O., Barjaktarovic, Z., Sarioglu, H., Wojcik, A., Harms-Ringdahl, M., Atkinson, M. J., Haghdoost, S., & Tapio, S. (2013). Quantitative proteomic analysis reveals induction of premature senescence in human umbilical vein endothelial cells exposed to chronic low-dose rate gamma radiation. Proteomics, 13(7), 1096–1107. https://doi.org/10.1002/pmic.201200463  Yentrapalli, R., Azimzadeh, O., Sriharshan, A., Malinowsky, K., Merl, J., Wojcik, A., Harms-Ringdahl, M., Atkinson, M. J., Becker, K. F., Haghdoost, S., & Tapio, S. (2013). The PI3K/Akt/mTOR Pathway Is Implicated in the Premature Senescence of Primary Human Endothelial Cells Exposed to Chronic Radiation. PLoS ONE, 8(8). https://doi.org/10.1371/journal.pone.0070024  Yu, T., Parks, B. W., Yu, S., Srivastava, R., Gupta, K., Wu, X., Khaled, S., Chang, P. Y., Kabarowski, J. H., & Kucik, D. F. (2011). Iron-ion radiation accelerates atherosclerosis in apolipoprotein E-Deficient mice. Radiation Research, 175(6), 766–773. https://doi.org/10.1667/RR2482.1  Zakrzewicz, A., Secomb, T. W., & Pries, A. R. (2002). Angioadaptation: Keeping the vascular system in shape. News in Physiological Sciences, 17(5). https://doi.org/10.1152/nips.01395.2001  Zhang, L. F. (2013). Region-specific vascular remodeling and its prevention by artificial gravity in weightless environment. European Journal of Applied Physiology, 113(12), 2873–2895. https://doi.org/10.1007/S00421-013-2597-8  Zhang, R., Bai, Y. G., Lin, L. J., Bao, J. X., Zhang, Y. Y., Tang, H., Cheng, J. H., Jia, G. L., Ren, X. L., & Jin, M. (2009). Blockade of at 1 receptor partially restores vasoreactivity, NOS expression, and superoxide levels in cerebral and carotid arteries of hindlimb unweighting rats. Journal of Applied Physiology, 106(1), 251–258. https://doi.org/10.1152/japplphysiol.01278.2007  Zielinski, J. M., Ashmore, P. J., Band, P. R., Jiang, H., Shilnikova, N. S., Tait, V. K., & Krewski, D. (2009). Low dose ionizing radiation exposure and cardiovascular disease mortality: Cohort study based on Canadian national dose registry of radiation workers. International Journal of Occupational Medicine and Environmental Health, 22(1), 27–33. https://doi.org/10.2478/v10001-009-0001-z  Zieman, S. J., Melenovsky, V., & Kass, D. A. (2005). Mechanisms, Pathophysiology, and Therapy of Arterial Stiffness. Arteriosclerosis, Thrombosis, and Vascular Biology, 25(5), 932–943. https://doi.org/10.1161/01.ATV.0000160548.78317.29  Zigman, S., McDaniel, T., Schultz, J., & Reddan, J. (2000). Effects of intermittent UVA exposure on cultured lens epithelial cells. Current Eye Research, 20(2), 95–100. https://doi.org/10.1076/0271-3683(200002)2021-DFT095  Zou, B., Schuster, J. P., Niu, K., Huang, Q., Rühle, A., & Huber, P. E. (2019). Radiotherapy-induced heart disease: a review of the literature. Precision Clinical Medicine, 2(4), 270–282. https://doi.org/10.1093/pcmedi/pbz025 AOPWiki 2022-09-28T11:20:48

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