studies using mouse primary microglial cultures irradiated with X-rays at 10 Gy have shown a 2.5-fold increase in tissue resident cell activation. In vivo mice irradiated with 30 Gy also show a 9-fold increase in microglial activation (Xu et al., 2015). In general, these studies highlight that with an increase in deposited energy larger amounts of primary and secondary tissue damage occurs, which leads to increased proliferation and activation of resident immune cells.

## Time Concordance

Many long-term and a few short-term studies find tissue-resident cell activation after deposition of energy. Mice irradiated with 35 Gy of gamma rays show activated astrocytes and microglia 4 h after irradiation (Kyrkanides et al., 1999). Rats irradiated with X-rays at 15 Gy show activation of astrocytes at 6 h (Hwang et al., 2006). Many studies show 1.2- to 3-fold increased activation one or a few days after irradiation in rats and mice (Casciati et al., 2016; Hwang et al., 2006; Kyrkanides et al., 1999; Moravan et al., 2011; Poulose et al., 2011). Furthermore, it has also been shown that activation continues until 1 year, measured after weeks and months, with a maximum 14-fold increase in marker levels shown after 2 months post-irradiation (Acharya et al., 2016; Allen et al., 2020; Casciati et al., 2016; Chiang, McBride & Withers, 1993; Chen et al., 2016; Cummings et al., 2007; Dey et al., 2020; Hua et al., 2012; Kyrkanides et al., 1999; Mizumatsu et al., 2003; Monje et al., 2002; Moravan et al., 2011; Parihar et al., 2020; Parihar et al., 2018; Parihar et al., 2016; Poulose et al., 2011; Raber et al., 2019; Rola et al., 2008; Rola et al., 2004; Suman et al., 2013).

## Incidence Concordance

No available data.

## Essentiality

Since deposition of energy is a physical stressor, it can be shielded, but chemicals cannot effectively block or decrease it. Thus further research is required to determine the effect of shielding radiation on the activation of tissue resident cells. Since deposited energy initiates events immediately, the removal of deposited energy, a physical stressor, also supports the essentiality of the key event. Studies that do not deposit energy are observed to have no downstream effects.

## Uncertainties and Inconsistencies

* There is no consensus about sex-related responses to radiation exposure and the resulting activation of tissue resident cells. Krukowski et al. (2018a) and Parihar et al. (2020) found that female mice were immune to the effects of 0.3 and 0.5 Gy radiation on cell activation, while Raber et al. (2019) found that only female mice showed increased activated cells after 2 Gy.

* A large amount of uncertainty surrounds the impact of low-dose ionizing radiation on tissue resident cell activation. More evidence is required to determine the relationship between ionizing radiation at doses < 1 Gy and tissue-resident cell activation.

## Known modulating factors

<table><thead><tr><td>Modulating factor</td><td>Details</td><td>Effects on the KER</td><td>References</td></tr></thead><tbody><tr><td>Sex</td><td>Male and female mice had different responses in tissue resident cell activation following irradiation.</td><td>Male mice typically showed an increase in microglia activation, while female mice showed no significant changes. However, not all studies found this trend.</td><td>Krukowski et al., 2018a; Parihar et al., 2020; Raber et al., 2019</td></tr><tr><td>Drug</td><td>Colony stimulating factor 1 receptor inhibitor PLX5622 (eliminates microglia).</td><td>PLX5622 reduced the number of activated microglia.</td><td>Acharya et al., 2016; Allen et al., 2020; Krukowski et al., 2018b</td></tr><tr><td>Drug</td><td>P2X7 receptor (associated with microglial activation) inhibitor Brilliant Blue G.</td><td>Treatment attenuated the increase in microglial activation both in vivo and in vitro after irradiation.</td><td>Xu et al., 2015</td></tr><tr><td>Age</td><td>10-day-old and 10-week-old mice.</td><td>At 10 days old, irradiated mice showed increased glial activation, while at 10 weeks old they did not show significant changes in activation.</td><td>Casciati et al., 2016</td></tr></tbody></table>
<table><tr><td></td><td>7-, 17- and 27-month-old mice.</td><td>Activation of microglia after irradiation decreased as age was increased.</td><td>Hua et al., 2012</td></tr><tr><td>Genetics</td><td>Extracellular SOD knockout mice.</td><td>Microglial activation was increased more in SOD knockout mice than wild-type mice after 5 Gy gamma rays.</td><td>Rola et al., 2007</td></tr></table>

## Quantitative Understanding of the Linkage

### Dose Concordance

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.

<table><thead><tr><td>Reference</td><td>Experiment description</td><td>Result</td></tr></thead><tbody><tr><td>Suman et al., 2013</td><td>In vivo. 6- to 8-week-old female C57BL/6J mice were irradiated with either 1.6 Gy 56Fe ions or 2 Gy gamma rays (137Cs source) both at 1 Gy/min. GFAP levels in the cerebral cortex measured by immunoblotting were used to indicate astrocyte activation.</td><td>GFAP levels increased 2.2-fold in the gamma ray irradiated group and 4.3-fold in the 56Fe ion irradiated group.</td></tr><tr><td>Poulose et al., 2011</td><td>In vivo. 2-month-old male Sprague-Dawley rats were irradiated with 5, 50 and 100 cGy 16O particles. GFAP was measured by western blot in the hippocampus.</td><td>At maximum, GFAP increased 1.2-fold after 5 and 50 cGy and 1.3-fold after 100 cGy.</td></tr><tr><td>Parihar et al., 2016</td><td>In vivo. Male Thy1-EGFP transgenic mice were irradiated with 600 MeV/amu charged particles (16O and 48Ti) (0.05 to 0.25 Gy/min) at 0.05 and 0.3 Gy. Activated glial cells were indicated by ED1 markers through immunostaining.</td><td>Number of activated microglia increased with dose, with a greater fold difference following 48Ti radiation than 16O. A maximum 1.9-fold change was observed at 30 cGy 48Ti at 27 weeks.</td></tr><tr><td>Parihar et al., 2018</td><td>In vivo. Male C57BL/6J mice were irradiated with He ions at 5 and 30 cGy (5 cGy/min). ED1 cells were used as markers for activated microglia by immunostaining and DAPI counterstaining.</td><td>ED1+ cells increased 3.5-fold after 5 cGy and 3.8-fold after 30 cGy, indicating microglial activation.</td></tr><tr><td>Parihar et al., 2020</td><td>In vivo. Mice were irradiated with low doses (≤≤ 30 cGy) of helium ions. Brain tissue sections were stained to identify microglia activation by CD68+ cells.</td><td>A 2.5-fold increase in CD68+ cells was observed in the male irradiated group following 30 cGy irradiation, indicating microglia activation. No increase in CD68+ was seen in the female mice.</td></tr><tr><td>Krukowski et al., 2018a</td><td>In vivo. C57B16/J mice were exposed to GCR at 0, 15, 50 cGy (252 MeV/amu protons, 249.3 MeV/amu helium ions, and 594.4 MeV/amu oxygen ions). Microglia were stained and levels were measured by Iba-1.</td><td>50 cGy male cohorts showed a significant 3- to 5-fold increase in Iba-1+ cells compared to the controls, indicating activated microglia in the dorsal hippocampus. No change was observed in the females.</td></tr><tr><td>Allen et al., 2020</td><td>In vivo. Mice were exposed to 4He irradiation (400 MeV/amu) at 30 cGy. CD68+ cells and Iba-1 immunochemistry were used to measure activated microglia. GFAP was measured to determine astrogliosis.</td><td>1.25-fold change in CD68+ cells was observed in the 30 cGy group compared to the controls. Iba-1+ microglia cells did not change significantly. No change in GFAP expression, a marker indicative of astrogliosis.</td></tr><tr><td>Cummings et al., 2007</td><td>In vivo. Sprague-Dawley rats were irradiated with 56Fe ions (600 MeV/amu) with 4 Gy dose. GFAP immunochemistry was used to identify astrogliosis.</td><td>There was a significant increase in GFAP+ cells 6 and 12 months post irradiation with 4 Gy.</td></tr><tr><td>Rola et al., 2008</td><td>In vivo. Male mice were irradiated with 0.5-4 Gy 56Fe ions. CD68+ cells determined microglia activation and BrdU identified neurogenesis.</td><td>Dose dependent increase was observed in BrdU+/CD68+ cells indicating newly born activated microglia. Significant increases of 3.4, 4.4 and 14-fold were observed at 1 Gy, 2 Gy and 4 Gy, respectively.</td></tr><tr><td>Raber et al., 2019</td><td>In vivo. Mice were irradiated with 1 GeV protons, 16O or 28Si (0, 25, 50, or 200 cGy). ELISA was used to detect levels of CD68 for activated microglia.</td><td>Cortical CD68 levels were increased by 1.7-fold in females irradiated with 200 cGy. This effect was not seen in males.</td></tr><tr><td>Hwang et al., 2006</td><td>In vivo and in vitro. Sprague-Dawley rats were irradiated with X-rays at 2-10 Gy and 15 Gy. Immunostaining, RT-PCR and western blot were used to analyze GFAP protein levels.</td><td>A dose dependent fold increase of cells with processed morphology was found. A 12- to 29.5-fold increase in activated morphology was observed in an astrocyte mixed-culture irradiated with 0-10 Gy.</td></tr><tr><td>Mizumatsu et al., 2003</td><td>In vivo. 2-month-old male C57BL/J6 mice were irradiated with X-rays at 175 cGy/min. New activated microglia in the hippocampus were determined with BrdU and CD68 staining.</td><td>A dose-dependent increase in BrdU/CD68+ cells was observed. No cells were activated without irradiation, 14% were after 2 Gy, 38% were after 5 Gy and 54% were after 10 Gy.</td></tr><tr><td>Hua et al., 2012</td><td>In vivo. Male FxBN rats were irradiated with 10 Gy of 137Cs gamma rays. CD68 was labelled with ED-1 in the hippocampus to show activated microglia.</td><td>The density of ED-1+ microglia increased 2- to 3-fold after 10 Gy.</td></tr></tbody></table>
<table><tr><td>Rola et al., 2007</td><td>In vivo. 8-week-old C57BL/6J mice were irradiated with 5 Gy X-rays. Microglial generation and activation in the hippocampus was determined by BrdU and CD68 staining.</td><td>The number of BrdU/CD68+ cells increased 1.3-fold.</td></tr><tr><td>Chen et al., 2016</td><td>In vitro. Human CHME5 microglia were irradiated with 137Cs gamma rays (LET 0.9 keV/µm) delivered over 1-3 min. Activated cells were determined by morphology as well as western blot for CR3/43 and Glut-5 activation markers.</td><td>CR3/43 and Glut-5 were both observed after 8 Gy, but just slightly or not at all after 0.5 Gy. At 8 Gy, microglia demonstrated an activated morphology.</td></tr><tr><td>Monje et al., 2002</td><td>In vivo. Adult female Fischer 344 rats were irradiated with X-rays at 175 cGy/min with two fractions of 5 Gy. Activated astrocytes and microglia were determined in the hippocampus through GFAP and ED-1 staining, respectively.</td><td>The percent of GFAP+ cells increased from 5.4% to 7.4% after 10 Gy. The percent of ED-1+ cells increased from 0% to 22% after 10 Gy.</td></tr><tr><td>Casciati et al., 2016</td><td>In vivo. Female and male C57BL/6 mice were irradiated with X-rays. Activated astrocytes and microglia were determined in the hippocampus through GFAP and Iba-1 staining, respectively.</td><td>Iba-1+ cells increased 1.5-fold after 2 Gy, while GFAP+ cells increased 1.3-fold after 2 Gy. No changes in GFAP were seen after 0.1 Gy.</td></tr><tr><td>Acharya et al., 2016</td><td>In vivo. 6-month-old male C57BL/6J mice were irradiated with 9 Gy X-rays. Microglia activation was determined by CD68 staining.</td><td>In the hippocampus, CD68+ cells increased 1.75-fold, while in the medial prefrontal cortex, they increased 1.25-fold.</td></tr><tr><td>Rola et al., 2004</td><td>In vivo. 21-day-old male C57BL/J6 mice were irradiated with 5 Gy X-rays at 1.75 Gy/min. Microglial and astrocyte activation was determined by CD68 and GFAP staining, respectively, double-stained with BrdU for gliogenesis in the subgranular zone.</td><td>CD68+ cells increased 2.5-fold, while GFAP+ cells increased 2-fold.</td></tr><tr><td>Dey et al., 2020</td><td>In vivo. 6-month-old male C57BL/6J mice received X-ray irradiation in fractions of 8.67 Gy/day (1.10 Gy/min) on alternating days for a total dose of 26 Gy. Activated microglia were determined through CD68 staining in the hippocampus.</td><td>In the CA1 region, CD68+ cells increased 3-fold, while in the CA3 region they increased 5-fold.</td></tr><tr><td>Chiang, McBride & Withers, 1993</td><td>In vivo. 12-week-old male C3Hf/Sed/Kam mice were irradiated with X-rays (238 cGy/min). Activated astrocytes and microglia were determined through GFAP and Mac-1 immunohistochemistry, respectively.</td><td>GFAP+ cells in the hippocampus and corpus callosum increased 1.5-fold after 30 and 36 Gy, and 2-fold after 45 Gy. Mac-1+ cells in the corpus callosum increased after 2 (1.2-fold), 20 (1.9-fold), 30 (2-fold), 36 (1.8-fold) and 45 Gy (2.8-fold).</td></tr><tr><td>Kyrkanides et al., 1999</td><td>In vivo. Male C3H/HeN mice were irradiated with 60Co gamma rays (35 Gy, 0.9 Gy/min). Activated astrocytes and microglia were determined through GFAP and Mac-1 immunohistochemistry, respectively.</td><td>Mac-1 and GFAP levels both showed greatly increased expression after 35 Gy.</td></tr><tr><td>Moravan et al., 2011</td><td>In vivo. 8- to 10-week-old male C57BL/6J mice were irradiated with 137Cs gamma rays at 1.25 Gy/min. GFAP expression was measured by RT-qPCR.</td><td>GFAP increased 2- to 3-fold after 25 and 35 Gy, but not significantly after 5 and 15 Gy.</td></tr><tr><td>Xu et al., 2015</td><td>In vivo and in vitro. Primary microglia from male BALB/c mice were irradiated with 30 Gy using a 6 MV β-ionizing-ray linear accelerator. Male BALB/c mice were irradiated with 10 Gy X-rays. Microglial cell activation was determined through morphology and Iba-1 immunohistochemistry.</td><td>In vitro, 10 Gy resulted in a 2.5-fold increase in activated microglia. In vivo, 30 Gy resulted in a 9-fold increase in activated microglia.</td></tr></table>

Time Concordance

<table><thead><tr><td>Reference</td><td>Experiment description</td><td>Result</td></tr></thead><tbody><tr><td>Suman et al., 2013</td><td>In vivo. 6- to 8-week-old female C57BL/6J mice were irradiated with either 1.6 Gy 56Fe or 2 Gy 137Cs gamma rays both at 1 Gy/min. GFAP levels in the cerebral cortex measured by immunoblotting were used to indicate astrocyte activation.</td><td>GFAP levels were increased 2.2-fold in the gamma irradiated group and 4.3-fold in the 56Fe irradiated group 12 months after exposure.</td></tr><tr><td>Poulose et al., 2011</td><td>In vivo. 2-month-old male Sprague-Dawley rats were irradiated with 5, 50 and 100 cGy 16O particles. GFAP was measured by western blot in the hippocampus.</td><td>After 36 h, GFAP was increased 1.2-fold after 100 cGy. After 75 days, GFAP increased 1.2-fold after 5 and 50 cGy and 1.3-fold after 100 cGy.</td></tr><tr><td>Parihar et al., 2016</td><td>In vivo. Male Thy1-EGFP transgenic mice were irradiated with 600 MeV/amu charged particles (16O and 48Ti) (0.05 to 0.25 Gy/min) at 0.05 and 0.3 Gy. Activated glial cells were indicated by ED1 markers through immunostaining.</td><td>ED1+ cells increased ~1.2- to 1.6-fold 15 weeks post 16O irradiation and ~2-fold 27 weeks post irradiation in all 48Ti irradiated groups.</td></tr><tr><td>Parihar et al., 2018</td><td>In vivo. Male C57BL/6J mice were irradiated with He ions at 5 and 30 cGy (5 cGy/min). ED1 cells were used as markers for activated microglia by immunostaining and DAPI counterstaining.</td><td>52 weeks after radiation, ED1+ cells increased 3.5-fold and 3.8-fold, indicating microglial activation.</td></tr><tr><td>Parihar et al., 2020</td><td>In vivo. Mice were irradiated with low doses (≤≤ 30 cGy) of helium ions. Brain tissue sections were stained to identify microglia activation by CD68+ cells.</td><td>The number of activated microglia increased by 2.5-fold in the male irradiated group 15 weeks after 30 cGy 4He irradiation.</td></tr><tr><td>Allen et al., 2020</td><td>In vivo. Mice were exposed to 4He irradiation (400 MeV/amu) at 30 cGy. CD68+ cells and Iba-1 immunohistochemistry were used to measure activated microglia. GFAP was measured to determine astrogliosis.</td><td>An increase in CD68+ cells was observed 7-8 weeks post-irradiation.</td></tr></tbody></table>
<table>
  <tr>
    <td>Rola et al., 2008</td>
    <td>In vivo. Male mice were irradiated with 0.5-4 Gy 56Fe ions. CD68+ cells determined microglia activation and BrdU identified neurogenesis.</td>
    <td>2 months post-irradiation, BrdU/CD68+ cells increased up to 14-fold.</td>
  </tr>
  <tr>
    <td>Cummings et al., 2007</td>
    <td>In vivo. Sprague-Dawley rats were irradiated with 56Fe ions (600 MeV/amu) with 4 Gy dose. GFAP immunochemistry was used to identify astrogliosis.</td>
    <td>There was a significant increase in GFAP+ cells 6 and 12 months post irradiation. A maximum 1.7-fold change was observed 12 months post irradiation compared to the control.</td>
  </tr>
  <tr>
    <td>Raber et al., 2019</td>
    <td>In vivo. Mice were irradiated with 1 GeV protons, 16O or 28Si (0, 25, 50, or 200 cGy). ELISA was used to detect levels of CD68 for activated microglia.</td>
    <td>1.7-fold increase in CD68 levels in female mice was seen 3 months following 200 cGy exposure.</td>
  </tr>
  <tr>
    <td>Hwang et al., 2006</td>
    <td>In vivo. Sprague-Dawley rats were irradiated with X-rays at 2-10 Gy and 15 Gy. Immunostaining, RT-PCR and western blot were used to analyze GFAP protein levels.</td>
    <td>GFAP levels were slightly higher at 6 h post irradiation compared to the initial levels and significantly increased 24 h post-irradiation.</td>
  </tr>
  <tr>
    <td>Mizumatsu et al., 2003</td>
    <td>In vivo. 2-month-old male C57BL/J6 mice were irradiated with 10 Gy X-rays at 175 cGy/min. New activated microglia in the hippocampus were determined with BrdU and CD68 staining.</td>
    <td>No cells were activated without irradiation or 48 h after irradiation, while up to 50% of cells were activated 2 months after irradiation.</td>
  </tr>
  <tr>
    <td>Hua et al., 2012</td>
    <td>In vivo. Male FxBN rats were irradiated with 10 Gy of 137Cs gamma rays. CD68 was labelled with ED-1 in the hippocampus to show activated microglia.</td>
    <td>The density of ED-1+ microglia increased 2- to 3-fold after 1 week, and was just slightly increased after 10 weeks.</td>
  </tr>
  <tr>
    <td>Chen et al., 2016</td>
    <td>In vitro. Human CHME5 microglia were irradiated with 8 Gy gamma rays (LET 0.9 keV/µm) delivered over 1-3 min. Activated cells were determined by morphology as well as western blot for CR3/43 and Glut-5 activation markers.</td>
    <td>Cells demonstrated a normal morphology until 6 days then an activated morphology starting 7 days post-irradiation. Glut-5 was observed after 7,10 and 14 days, while CR3/43 was observed after 2 weeks.</td>
  </tr>
  <tr>
    <td>Monje et al., 2002</td>
    <td>In vivo. Adult female Fischer 344 rats were irradiated with X-rays at 175 cGy/min with two fractions of 5 Gy. Activated astrocytes and microglia were determined in the hippocampus through GFAP and ED-1 (labels CD68) staining, respectively.</td>
    <td>The percent of GFAP+ cells increased from 5.4% to 7.4% after 2 months. The percent of ED-1+ cells increased from 0% to 22% after 2 months.</td>
  </tr>
  <tr>
    <td>Casciati et al., 2016</td>
    <td>In vivo. Female and male C57BL/6J mice were irradiated with 2 Gy X-rays. Activated astrocytes and microglia were determined in the hippocampus through GFAP and Iba-1 staining, respectively.</td>
    <td>After 1 day, Iba-1+ cells increased 1.5-fold, while after 6 months, GFAP+ cells increased 1.3-fold.</td>
  </tr>
  <tr>
    <td>Acharya et al., 2016</td>
    <td>In vivo. 6-month-old male C57BL/6J mice were irradiated with 9 Gy X-rays. Microglia activation was determined by CD68 staining.</td>
    <td>In the hippocampus, CD68+ cells increased 1.75-fold after 2 and 6 weeks, while in the medial prefrontal cortex they increased 1.25-fold after 6 weeks.</td>
  </tr>
  <tr>
    <td>Rola et al., 2004</td>
    <td>In vivo. 21-day-old male C57BL/6J mice were irradiated with 5 Gy X-rays at 1.75 Gy/min. Microglial and astrocyte activation was determined by CD68 and GFAP staining, respectively, double-stained with BrdU for gliogenesis in the subgranular zone.</td>
    <td>Both 1 and 3 months post-irradiation, CD68+ cells increased 2.5-fold, while GFAP+ cells increased 2-fold 3 months post-irradiation.</td>
  </tr>
  <tr>
    <td>Dey et al., 2020</td>
    <td>In vivo. 6-month-old male C57BL/6J mice received X-ray irradiation in fractions of 8.67 Gy/day at a dose rate of 1.10 Gy/min on alternating days for a total dose of 26 Gy. Activated microglia were determined through CD68 staining in the hippocampus.</td>
    <td>Measured 5 weeks post-irradiation, in the CA1 region, CD68+ cells increased 3-fold, while in the CA3 region they increased 5-fold. No significant changes in these areas were observed 15 weeks post-irradiation.</td>
  </tr>
  <tr>
    <td>Chiang, McBride & Withers, 1993</td>
    <td>In vivo. 12-week-old male C3Hf/Sed/Kam mice were irradiated with X-rays (238 cGy/min). Activated astrocytes and microglia were determined through GFAP and Mac-1 immunohistochemistry, respectively. ELISA was also used to measure GFAP.</td>
    <td>From 30-45 Gy, GFAP was significantly increased 1.2-fold 15 days after irradiation, as well as 120-180 days after irradiation. Similarly, 150 days after irradiation, Mac-1 was increased 1.2- to 3-fold from 2 to 45 Gy.</td>
  </tr>
  <tr>
    <td>Kyrkanides et al., 1999</td>
    <td>In vivo. Male C3H/HeN mice were irradiated with 60Co gamma rays (35 Gy, 0.9 Gy/min). Activated astrocytes and microglia were determined through GFAP and Mac-1 immunohistochemistry, respectively.</td>
    <td>Mac-1 was first seen increased after 4 h, but peaked at 24 h. GFAP was also first increased after 4 h, and increased throughout the 7 days measured.</td>
  </tr>
  <tr>
    <td>Moravan et al., 2011</td>
    <td>In vivo. 8- to 10-week-old male C57BL/6J mice were irradiated with 137Cs gamma rays at 1.25 Gy/min. GFAP expression was measured by RT-qPCR.</td>
    <td>GFAP increased 2- to 3-fold after 25 and 35 Gy, but not significantly after 5 and 15 Gy. The significant increases were observed after 1, 30 and 180 days.</td>
  </tr>
</table>
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## AOPs Referencing Relationship

<table><thead><tr><th>AOP Name</th><th>Adjacency</th><th>Weight of Evidence</th><th>Quantitative Understanding</th></tr></thead><tbody><tr><td>AOP: 470 - Deposition of energy leads to abnormal vascular remodeling</td><td>adjacent</td><td>High</td><td>Low</td></tr><tr><td>AOP: 483 - Deposition of Energy Leading to Learning and Memory Impairment</td><td>adjacent</td><td>High</td><td>Low</td></tr></tbody></table>

## Evidence Supporting Applicability of this Relationship

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.

## Taxonomic Applicability

<table><thead><tr><th>Term</th><th>Scientific Term</th><th>Evidence</th><th>Link</th></tr></thead><tbody><tr><td>human</td><td>Homo sapiens</td><td>Low</td><td>NCBI</td></tr><tr><td>mouse</td><td>Mus musculus</td><td>High</td><td>NCBI</td></tr><tr><td>rat</td><td>Rattus norvegicus</td><td>High</td><td>NCBI</td></tr><tr><td>Pig</td><td>Pig</td><td>Moderate</td><td>NCBI</td></tr></tbody></table>

## Life Stage Applicability

<table><thead><tr><th>Term</th><th>Evidence</th></tr></thead><tbody><tr><td>Juvenile</td><td>High</td></tr><tr><td>Adult</td><td>Moderate</td></tr></tbody></table>

## Sex Applicability

<table><thead><tr><th>Sex</th><th>Evidence</th></tr></thead><tbody><tr><td>Male</td><td>High</td></tr><tr><td>Female</td><td>Low</td></tr><tr><td>Unspecific</td><td>Low</td></tr></tbody></table>

## Key Event Relationship Description

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).

## Evidence Supporting this KER

Overall weight of evidence: High

### Biological Plausibility

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 impaired learning and memory. 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).
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).

## Empirical Evidence

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; Hasan, Radwan & Galal, 2019; El-Missiry et al., 2018; Kenchegowda et al., 2018; Zhao et al., 2013; 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). 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).

## 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).

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 stress response signalling 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. 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).

## Uncertainties and Inconsistencies

* 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) or varying changes (Azimzadeh et al., 2015) in the MAPK pathways.

The assays employed in studies to assess the KEs may lead to variations in the quantitative understanding of observations.

## Known modulating factors

<table><thead><tr><td>Modulating factor</td><td>Details</td><td>Effects on the KER</td><td>References</td></tr></thead><tbody><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></tbody></table>
<table><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>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, or 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></table>

## Quantitative understanding of the Linkage

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><thead><tr><td>Reference</td><td>Experiment Description</td><td>Result</td></tr></thead><tbody><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>Fan et al., 2017</td><td>In vivo. 10-week-old male C57BL/6J mice were irradiated with 60Co gamma rays at 3Gy/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></tbody></table>
<table>
  <tr>
    <td>Azimzadeh et al., 2015</td>
    <td>In vivo. Male 10-week-old C57BL/6J 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 was 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 determine 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>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</td>
    <td>ROS increased 1.8-fold. ERK1/2, p38 and JNK mRNA increased 2- to 3-fold. The ratios of</td>
  </tr>
</table>
<table><tr><td></td><td>cerebral cortex. ROS levels were measured with DCFH-DA fluorescent dye. Signaling molecules were measured with western blotting or RT-qPCR.</td><td>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 hydrogen peroxide 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 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 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>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></table>

Time Concordance

<table><thead><tr><td>Reference</td><td>Experiment Description</td><td>Result</td></tr></thead><tbody><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 determine 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>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</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></tbody></table>
<table><tr><td></td><td>4-HNE levels were determined with immunohistochemical staining. p21 and p53 levels were determined with immunoblotting.</td><td></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 ~10% (1 week) and ~30% at 2 and 3 weeks compared to room temperature control. Phosphorylated ERK increased by ~60% at 1 week, ~150% at 2 weeks and ~140% at 3 weeks. Phosphorylated p38 increased by ~50% at 1 week, ~100% at 2 weeks and ~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></table>

Known Feedforward/Feedback loops influencing this KER

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).
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## AOPs Referencing Relationship

<table><thead><tr><th>AOP Name</th><th>Adjacency</th><th>Weight of Evidence</th><th>Quantitative Understanding</th></tr></thead><tbody><tr><td>AOP: 483 – Deposition of Energy Leading to Learning and Memory Impairment</td><td>adjacent</td><td>Moderate</td><td>Low</td></tr></tbody></table>

## Evidence Supporting Applicability of this Relationship

Evidence for this relationship comes from in vitro human- and mouse-derived models, as well as in vivo rat models. Most of the evidence are in male adult and male models, although sex and age are not always specified.

## Taxonomic Applicability

<table><thead><tr><th>Term</th><th>Scientific Term</th><th>Evidence</th><th>Link</th></tr></thead><tbody><tr><td>human</td><td>Homo sapiens</td><td>Low</td><td>NCBI</td></tr><tr><td>mouse</td><td>Mus musculus</td><td>Low</td><td>NCBI</td></tr><tr><td>rat</td><td>Rattus norvegicus</td><td>Moderate</td><td>NCBI</td></tr></tbody></table>

## Life Stage Applicability

<table><thead><tr><th>Term</th><th>Evidence</th></tr></thead><tbody><tr><td>Adult</td><td>Moderate</td></tr><tr><td>Not Otherwise Specified</td><td>Low</td></tr></tbody></table>

## Sex Applicability

<table><thead><tr><th>Sex</th><th>Evidence</th></tr></thead><tbody><tr><td>Male</td><td>Moderate</td></tr><tr><td>Female</td><td>Not Specified</td></tr><tr><td>Unspecific</td><td>Low</td></tr></tbody></table>

## Key Event Relationship Description

Oxidative stress encompasses an increase in the production of free radicals (e.g., superoxide, hydrogen peroxide and hydroxyl radicals) and a loss of antioxidant mechanisms (e.g., superoxide dismutase (SOD), glutathione peroxidase (GSH-Px) and catalase (CAT)). This imbalance can lead to damaging by-products that can activate tissue resident cells. Reactive oxygen and nitrogen species (RONS) are examples of free radicals that may promote oxidative injury (Simpson & Oliver, 2020). In addition, excess free radicals can promote a reduced capacity of the cells to maintain redox balance and prevent ongoing oxidative damage (Huang, Zou & Corniola, 2012; Rojo et al., 2014). Depending on the organ/tissue, different resident cell types may become activated by oxidative stress. For example, in the brain, oxidative stress will specifically activate microglial cells and astrocytes (Lee, Cha & Lee, 2021). Microglia cells are macrophages in the brain that respond to tissue injury, provide surveillance to neurons, and maintain synaptic homeostasis (Zhu et al., 2022). Astrocytes are critical regulators of neurogenesis and synaptogenesis, blood brain barrier permeability, and responsible for maintenance of cellular homeostasis (Zhu et al., 2022). Both microglial cells and astrocytes can change from resting to reactive states, termed gliosis, in response to excess RONS (Lee, Cha & Lee, 2021). In response to RONS, Toll like receptors (TLRs) located on microglia become activated to mediate the immune response (Gill et al., 2010; Mehdipour et al., 2021). These receptors then initiate a cascade of signaling pathways that contribute to the production of pro-inflammatory cytokines and free radicals, resulting in neuroinflammation (Heidari et al., 2022).

Reactive microglia cells increase in size and number, display a reduction in the length and density of their processes, and upregulate their macrophagic processes, marked by expression of proteins related to phagocytic activity such as cluster of differentiation 68 (CD68) (Hol & Pekny, 2015). Astrocytes undergoing astrogliosis exhibit cellular hypertrophy and an upregulation of glial fibrillary acidic
protein (GFAP), an intermediate filament expressed exclusively in astrocytes that plays a critical role
in astroglia cell activation (Hol & Pekny, 2015). Activation of both microglial cells and astrocytes can
accelerate neuroinflammatory pathways that can ultimately promote further formation of ROS creating
a feedforward loop (Lee, Cha & Lee, 2021; Simpson & Oliver, 2020; Zhu et al., 2022).

***Evidence Supporting this KER***

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

**Biological Plausibility**

Biological Plausibility is Moderate. RONS can activate some inflammatory and anti-inflammatory pathways (TLR, TGF-β, NF-κB), and are an essential part of multiple inflammatory and anti-inflammatory pathways (TLR4, TNF-α, TGF-β, NF-κB).

RONS activates or is essential to many inflammatory pathways including TGF-β (Barcellos-Hoff and Dix 1996; Jobling, Mott et al. 2006), TNF (Blaser, Dostert et al. 2016), Toll-like receptor (TLR) (Park, Jung et al. 2004; Nakahira, Kim et al. 2006; Powers, Szaszi et al. 2006; Miller, Goodson et al. 2017; Cavaillon 2018), and NF-κB signaling (Gloire, Legrand-Poels et al. 2006; Morgan and Liu 2011). These interactions principally involve ROS, but RNS can indirectly activate TLRs and possibly NF-κB. Since inflammatory signaling and activated immune cells can also increase the production of RONS, positive feedback and feedforward loops can occur (Zhao and Robbins 2009; Ratikan, Micewicz et al. 2015; Blaser, Dostert et al. 2016).

Damage inflicted by RONS on cells activate TLRs and other receptors to promote release of cytokines (Ratikan, Micewicz et al. 2015). For example, oxidized lipids or oxidative stress-induced heat shock proteins can activate TLR4 (Miller, Goodson et al. 2017; Cavaillon 2018).

ROS is essential to TLR4 activation of downstream signals including NF-κB. Activation of TLR4
promotes the surface expression and movement of TLR4 into signal-promoting lipid rafts (Nakahira,
Kim et al. 2006; Powers, Szaszi et al. 2006). This signal promotion requires NADPH-oxidase and
ROS (Park, Jung et al. 2004; Nakahira, Kim et al. 2006; Powers, Szaszi et al. 2006). ROS is also
required for the TLR4/TRAF6/ASK-1/p38 dependent activation of inflammatory cytokines
(Matsuzawa, Saegusa et al. 2005). ROS therefore amplifies the inflammatory process.

RONS can also fail to activate or actively inhibit inflammatory pathways, and the circumstances
determining response to RONS are not well known (Gloire, Legrand-Poels et al. 2006).

Responses to oxidative stress can vary depending on the organ system. In the central nervous system (CNS), biological plausibility supporting the connection between increased oxidative stress to tissue resident cell activation is moderately supported by evidence compiled from studies using animal and in vitro models. Multiple studies have shown that microglial cells and astrocytes are activated in response to RONS, meaning they change from resting to reactive states by secreting pro-inflammatory mediators and initiating antioxidant defenses mediated through TLRs (Simpson & Oliver, 2020; Heidari et al., 2022). Literature reviews describing the role of oxidative stress imbalances and glial cell activation in the context of general oxidative injury (Lee, Cha & Lee, 2021), stroke (Zhu et al., 2022) and neurodegenerative diseases (Reynolds et al., 2007; Simpson & Oliver, 2020) also suggest a relationship between increased oxidative stress and increased tissue resident cell activation in the CNS.

**Empirical Evidence**

Empirical Evidence is Moderate. Both RONS and inflammation increase in response to agents that increase RONS or inflammation, and antioxidants reduce inflammation. Multiple studies show dose-dependent changes in both RONS and inflammation in response to stressors including ionizing radiation and antioxidants. RONS have been measured at the same or earlier time points as
inflammatory markers, but additional studies are needed to characterize the inflammatory response at the earliest time points to support causation. Uncertainties come from the positive feedback from inflammation to RONS potentially interfering with attempts to establish causality, and from the large number of inflammation-related endpoints with differing responses to stressors and experimental variation.

Oxidative activity is required for or promotes the response to multiple inflammatory stressors, including ionizing radiation, UV radiation (particularly UVB), the endotoxin LPS and other pathogens associated immune activators, and hemorrhagic shock (Park, Jung et al. 2004; Nakahira, Kim et al. 2006; Powers, Szaszi et al. 2006; Zhao and Robbins 2009; Ha, Chung et al. 2010; Hiramoto, Kobayashi et al. 2012; Straub, New et al. 2015).

Both intracellular concentrations of RONS and a wide range of inflammatory markers increase in response to RONS stressors. This paired increase was observed in vivo in rodents in tissue from multiple internal organs following exposure to whole body or abdominal ionizing radiation (Berruyer, Martin et al. 2004; Ha, Chung et al. 2010; Sinha, Das et al. 2011; Sinha, Das et al. 2012; Das, Manna et al. 2014; Ozyurt, Cevik et al. 2014; Khan, Manna et al. 2015; Zetner, Andersen et al. 2016; Haddadi, Rezaeyan et al. 2017; Ezz, Ibrahim et al. 2018) or following UV skin irradiation (Sharma, Meeran et al. 2007; Hiramoto, Kobayashi et al. 2012; Martinez, Pinho-Ribeiro et al. 2016). In vitro, the relationship has been reported in response to IR and UV in keratinocytes (Park, Ju et al. 2006; Kang, Kim et al. 2007; Martin, Sur et al. 2008; Lee, Jeon et al. 2010; Ren, Shi et al. 2016; Hung, Tang et al. 2017; Zhang, Zhu et al. 2017), immune cells (Matsuzawa, Saegusa et al. 2005; Nakahira, Kim et al. 2006; Manna, Das et al. 2015; Soltani, Ghaemi et al. 2016), as well as corneal and conjunctival epithelia, HEK cells, and vocal cord and foreskin fibroblasts (Narayanan, LaRue et al. 1999; Park, Jung et al. 2004; Saltman, Kraus et al. 2010; Black, Gordon et al. 2011; Han, Min et al. 2015). Direct application of micromolar concentrations of H2O2 in vitro also increases inflammatory markers in immune cells (Matsuzawa, Saegusa et al. 2005; Nakao, Kurokawa et al. 2008) and keratinocytes (Zhang, Zhu et al. 2017).

Interventions to reduce oxidative activity also reduce inflammation, further implicating RONS in the inflammatory process. Reduction of inflammation by these interventions has been documented in animals in response to IR (Berruyer, Martin et al. 2004; Sinha, Das et al. 2011; Sinha, Das et al. 2012; Das, Manna et al. 2014; Ozyurt, Cevik et al. 2014; Khan, Manna et al. 2015; Zetner, Andersen et al. 2016; Haddadi, Rezaeyan et al. 2017; Ezz, Ibrahim et al. 2018), UV (Sharma, Meeran et al. 2007; Lee, Jeon et al. 2010; Hiramoto, Kobayashi et al. 2012; Han, Min et al. 2015; Martinez, Pinho-Ribeiro et al. 2016; Ren, Shi et al. 2016; Hung, Tang et al. 2017) and hemorrhagic shock (Powers, Szaszi et al. 2006). In vitro, multiple studies in immune cells (Matsuzawa, Saegusa et al. 2005; Nakahira, Kim et al. 2006; Manna, Das et al. 2015; Soltani, Ghaemi et al. 2016) and keratinocytes (Park, Ju et al. 2006; Kang, Kim et al. 2007; Martin, Sur et al. 2008; Lee, Jeon et al. 2010; Ren, Shi et al. 2016; Hung, Tang et al. 2017; Zhang, Zhu et al. 2017) as well as HEK293, fibroblasts, and epithelial cells (Lee, Dimtchev et al. 1998; Narayanan, LaRue et al. 1999; Park, Jung et al. 2004; Han, Min et al. 2015) provide further evidence for reduction in various inflammatory markers with interventions to reduce RONS. Interventions include antioxidants such as propyl gallate, n-acetylcysteine, or naringin, as well as reduction in the function of NADPH oxidases (NOX/DUOX) via DPI or knockdown of gene expression. In studies using multiple doses of antioxidant, inflammation was reduced dose-dependently with the antioxidant dose (Nakahira, Kim et al. 2006; Manna, Das et al. 2015; Ren, Shi et al. 2016). Interventions reducing nitric oxide were not common, but in one study inhibiting iNOS did not reduce activation of NF-κB by IR (Lee, Dimtchev et al. 1998). The treatment to reduce RONS is administered before, or occasionally immediately after the inflammatory stressor, but studies often continue treatment or do not explicitly report changing media in vitro, so the exact time point at which RONS are required is difficult to pinpoint.

IR and RONS decrease endogenous antioxidant activity (glutathione, superoxide dismutase, and catalase), and antioxidants rescue this suppression in antioxidant activity (Sharma, Meeran et al. 2007; Das, Manna et al. 2014). Mice with more endogenous glutathione have a lower inflammatory response
to IR (Berruyer, Martin et al. 2004), suggesting that IR increases inflammation in part by decreasing
antioxidants.

In response to inflammatory stressors, RONS has been measured at the same (Nakao, Kurokawa et al. 2008; Ha, Chung et al. 2010; Saltman, Kraus et al. 2010; Azimzadeh, Scherthan et al. 2011; Ameziane-El-Hassani, Talbot et al. 2015; Azimzadeh, Sievert et al. 2015; Zhang, Zhu et al. 2017) or earlier time points as inflammatory markers (Nakahira, Kim et al. 2006; Black, Gordon et al. 2011). This suggests that RONS precedes the generation of inflammatory markers, consistent with a role for RONS in promoting inflammation. However, inflammatory markers are not typically measured at the earliest time points, and a more comprehensive survey of the appearance of these events at early time points would help to clarify the timeline and confirm the temporal evidence for causation.

A relatively small number of studies in a variety of cell types have examined both RONS and inflammatory markers across multiple doses. Three of these report dose-dependent increases in both RONS and inflammatory markers; one in which the key events are evaluated immediately after H2O2 application (Nakao, Kurokawa et al. 2008), and two others evaluated them 24 hours or 8-16 weeks after IR (Ha, Chung et al. 2010; Azimzadeh, Sievert et al. 2015). A fourth study reports a dose-dependent reduction in inflammation in response to treatment with antioxidants (Nakahira, Kim et al. 2006). In three other studies, some or all markers of inflammation increase at lower doses but decrease at higher doses (Saltman, Kraus et al. 2010; Black, Gordon et al. 2011; Zhang, Zhu et al. 2017). In two of these studies, RONS is also not consistently increasing with dose (Saltman, Kraus et al. 2010; Zhang, Zhu et al. 2017); however, this finding is consistent with findings from other studies about lack of dose-dependence of ROS measured at intermediate time points after IR. Similarly, 30 minutes after low dose, IR IL8 increases with dose while ROS does not (Narayanan, LaRue et al. 1999). The mixed inflammatory response at higher doses suggests that additional factors such as negative and positive feedback and crosstalk between pathways are also involved in the relationship between RONS and IR.

Dose Concordance

Evidence to support this relationship in the brain is derived from studies using gamma rays (Schnegg et al., 2012), 6-OHDA (Wang et al., 2017), or H2O2 (Daverey & Agrawal, 2016) as the stressor. Oxidative stress and tissue resident cell activation were then assessed within the brain or glial cell cultures (Davery & Agrawal, 2016; Schnegg et al., 2012; Wang, 2017; Daverey & Agrawal, 2016; Schnegg et al., 2012; Wang et al., 2017). Few studies show that oxidative stress occurs at lower or the same dose of a stressor than tissue-resident cell activation. Treatment with 4 µg/µL of the neurotoxin 6-hydroxydopamine caused decreased antioxidant levels as well as increased glial fibrillary acidic protein (GFAP) levels in rats (Wang et al., 2017). In another study, astrocyte activation showed a slight linear increase in response to 50 µM, 100 µM and 200 µM of H2O2 (Daverey & Agrawal, 2016).

Time Concordance

Few studies show that oxidative stress occurs before or at the same time as tissue-resident cell activation in a time course. BV-2 microglia irradiated with 10 Gy gamma rays showed an increase in ROS 1h post-irradiation, while an increase in NF-κB and AP-1 DNA binding was also observed 1h post-irradiation (Schnegg et al., 2012). Treatment of human glioblastoma astrocytes (A172 cell) with 50 µM of H2O2 for various times, showed a small linear increase in GFAP levels from 2h to 24h of treatment (Daverey & Agrawal, 2016).

Incidence Concordance

Schnegg et al. (2012) irradiated BV-2 microglia with gamma rays and showed increased ROS as well as increased NF-κB and AP-1 DNA binding, indicating activated glial cells, at the same dose of 10 Gy.

Essentiality

Studies examining the use of antioxidants to inhibit free radicals demonstrate the essentiality of oxidative stress and tissue resident cell activation. This has been observed in microglial cells using
multiple types of inhibitors. In BV-2 microglia, activation of PPARδ (involved in anti-inflammatory responses) with the agonist L-16504, reduced formation of reactive oxygen species and microglial activation (Schnegg et al., 2012). Treatment with multiple doses of an antioxidant Kukoamine A (KuA) elicited a dose-dependent partial attenuation of radiation-induced markers of microglial cell activation in rats (Zhang et al. 2017). After administration of another antioxidant, curcumin, levels of SOD and GSH-Px were restored and GFAP levels were decreased (Daverey & Agrawal, 2016; Wang et al., 2017). Furthermore, a knockout model of mitochondrial SOD (SOD2) resulted in an increase in reactivity of microglial cells post-irradiation (Fishman et al. 2009).

## Uncertainties and Inconsistencies

Although ROS can activate NF-KB (Gloire, Legrand-Poels et al. 2006), not all studies consistently show NF-kB activation after RONs stressor IR. It is possible that the link between ROS and NF-kB depends on the local environmental context, with different studies not adequately controlling all influential variables. One study offers a possible explanation based on temporal response: in macrophages, NF-kB was activated by shorter exposures to H2O2 (30 min), but the response disappeared with longer exposures (Nakao, Kurokawa et al. 2008).

While many models in vivo and in vitro showed a decreased inflammatory response to RONS stressors IR in combination with antioxidants, in endothelial cells in culture the increase in IL6 and IL8 after IR was not reduced by antioxidants. Although a synergistic increase in those cytokines occurring with combined TNF-a and IR treatment was reduced by antioxidants (Meeren, Bertho et al. 1997). Multiple mechanisms can increase inflammation, that inflammatory factors participate in positive feedback loops, and that responses to stimuli vary between cells.

Many studies do not report direct measures of RONS. As RONS are quickly scavenged, the quantitative understanding of this relationship can be inconsistent, due to varied response of antioxidant enzymes across experimental conditions and time measurements.

Limited data is available to support an understanding of this relationship at low doses (<0.1 Gy).

## Known modulating factors

<table><thead><tr><td>Modulating factor</td><td>Details</td><td>Effects on the KER</td><td>References</td></tr></thead><tbody><tr><td>Drug</td><td>KuA (antioxidant)</td><td>After 30 Gy X-ray whole-brain irradiation of rats, activated microglia increased to over 320% of control. KuA at 5 mg/kg decreased this to 240%, at 10 mg/kg decreased it to 180% and at 20 mg/kg decreased it to 170%.</td><td>Zhang et al., 2017</td></tr><tr><td>Drug</td><td>L-16504 (PPARδ agonist, involved in anti-inflammatory responses)</td><td>Treatment prevented the increase in ROS and reduced NF-kB and AP-1 DNA binding.</td><td>Schnegg et al., 2012</td></tr><tr><td>Drug</td><td>Curcumin (antioxidant)</td><td>Treatment increased SOD and GSH-Px levels and decreased the number of GFAP-positive cells.</td><td>Wang et al., 2017; Daverey & Agrawal, 2016</td></tr><tr><td>Age</td><td>Increased age</td><td>Increased age can cause susceptibility to ROS accumulation and tissue-resident cell activation.</td><td>Liguori et al., 2018; Hanslik, Marino & Ulland, 2021</td></tr><tr><td>Diet</td><td>High antioxidant diet</td><td>Increased antioxidants in diet can lead to reduced oxidative stress.</td><td>Ávila-Escalante et al., 2020</td></tr><tr><td>Diet</td><td>Hypocaloric diet</td><td>Caloric restriction has been shown to lead to reduced markers of oxidative stress.</td><td>Ávila-Escalante et al., 2020</td></tr><tr><td>Smoking</td><td>Active smokers</td><td>Active smokers show reduced GSH-Px activity compared to non-smokers (measured in patients with coronary artery disease).</td><td>Kamceva et al., 2016</td></tr><tr><td>Prior Disease</td><td>Neurodegenerative diseases like Alzheimer's and Parkinson's</td><td>These diseases can generate an environment of increased oxidative stress and promotes the activation of glial cells.</td><td>Hanslik, Marino & Ulland, 2021</td></tr></tbody></table>
<table><tr><td>Genotype</td><td>SOD knockout mice</td><td>SOD2 knockout mice experienced increased microglia activation following irradiation, indicating an impact of genotype on tissue resident cell activation.</td><td>Fishman et al., 2009</td></tr></table>

## Quantitative Understanding of the Linkage

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><thead><tr><td>Reference</td><td>Experiment description</td><td>Result</td></tr></thead><tbody><tr><td>Wang, 2017</td><td>In vivo. Adult male Sprague-Dawley rats were treated with 4 µg/µL of the neurotoxin 6-hydroxydopamine. Oxidative stress was measured by SOD and GSH-Px levels through a bicinchoninic acid protein assay kit. GFAP was used as a marker of astrocytes and was detected using immunohistochemistry.</td><td>SOD decreased 0.64-fold and GSH-Px decreased 0.34-fold. GFAP increased 1.7-fold.</td></tr><tr><td>Daverey & Agrawal, 2016</td><td>In vitro. Human A172 (glioblastoma astrocytes) and HA-sp (spinal cord astrocytes) cell lines were treated with H2O2. GFAP expression was detected through immunofluorescence.</td><td>After 50 µM of H2O2, both cell types showed increased GFAP expression about 1.5-fold. GFAP was also increased 1.5- to 2-fold after 100 and 200 µM of H2O2.</td></tr></tbody></table>

### Time Concordance

<table><thead><tr><td>Reference</td><td>Experiment description</td><td>Result</td></tr></thead><tbody><tr><td>Schnegg et al., 2012</td><td>In vitro. BV-2 immortalized microglia were irradiated with 10 Gy of <sup>137</sup>Cs gamma rays at 3.56 Gy/min. Measured 1h after irradiation, intracellular ROS generation was measured by the fluorescent DCFH-DA probe, and activation of NF-κB and AP-1 was determined by immunoblotting as a measure of cell activation.</td><td>Both measured 1h after irradiation, ROS increased about 7-fold while NF-κB and AP-1 DNA binding was increased 2.5- and 2-fold, respectively.</td></tr><tr><td>Daverey & Agrawal, 2016</td><td>In vitro. Human A172 (glioblastoma astrocytes) and HA-sp (spinal cord astrocytes) cell lines were treated with 50 µM of the ROS H2O2. GFAP expression was detected through immunofluorescence after various durations of H2O2 treatment.</td><td>Both cell types showed increased GFAP about 1.5-fold, measured after treatment with H2O2. H2O2 administered for 2, 6 and 12h showed slight increases at each timepoint, while after 24h of H2O2 treatment, GFAP was only increased in A172 cells.</td></tr></tbody></table>

### Incidence Concordance

<table><thead><tr><td>Reference</td><td>Experimental description</td><td>Result</td></tr></thead><tbody><tr><td>Schnegg et al., 2012</td><td>In vitro. BV-2 immortalized microglia were irradiated with 10 Gy of <sup>137</sup>Cs gamma rays at 3.56 Gy/min. Intracellular ROS generation was measured by the fluorescent DCFH-DA probe, and activation of NF-κB and AP-1 was determined by immunoblotting as a measure of cell activation.</td><td>ROS increased about 7-fold while NF-κB and AP-1 DNA binding was increased 2.5- and 2-fold, respectively.</td></tr></tbody></table>

## Know Feedforward/Feedback loops influencing this KER

Since inflammatory signaling and activated immune cells can also increase the production of RONS, positive feedback and feedforward loops can occur (Zhao and Robbins 2009; Ratikan, Micewicz et al. 2015; Blaser, Dostert et al. 2016). Similarly, positive feedforward and feedback loops regarding RONS, cellular activation, and inflammation also occur in the CNS. Both RONS and microglial cell activation can accelerate neuroinflammatory pathways that can ultimately promote further formation of RONS (Lee, Cha & Lee, 2021; Simpson & Oliver, 2020; Zhu et al., 2022).
Ávila-Escalante, M. L. et al. (2020), "The effect of diet on oxidative stress and metabolic diseases—Clinically controlled trials", Journal of Food Biochemistry, Vol. 44/5, https://doi.org/10.1111/jfbc.13191.

Daverey, A. and S. K. Agrawal. (2016), "Curcumin alleviates oxidative stress and mitochondrial dysfunction in astrocytes", Neuroscience, Vol. 333, https://doi.org/10.1016/j.neuroscience.2016.07.012.

Fishman, K. et al. (2009), "Radiation-induced reductions in neurogenesis are ameliorated in mice deficient in CuZnSOD or MnSOD", Free Radical Biology and Medicine, Vol. 47/10, https://doi.org/10.1016/j.freeradbiomed.2009.08.016.

Gill, R., A. Tsung and T. Billiar. (2010), "Linking oxidative stress to inflammation: Toll-like receptors", Free Radical Biology and Medicine, Vol. 48/9, https://doi.org/10.1016/j.freeradbiomed.2010.01.006.

Hanslik, K. L., K. M. Marino and T. K. Ulland. (2021), "Modulation of Glial Function in Health, Aging, and Neurodegenerative Disease", Frontiers in Cellular Neuroscience, Vol. 15, https://doi.org/10.3389/fncel.2021.718324.

Heidari, A., N. Yazdanpanah and N. Rezaei. (2022), "The role of Toll-like receptors and neuroinflammation in Parkinson's disease", Journal of Neuroinflammation, Vol. 19/1, https://doi.org/10.1186/s12974-022-02496-w.

Hol, E. M. and M. Pekny. (2015), "Glial fibrillary acidic protein (GFAP) and the astrocyte intermediate filament system in diseases of the central nervous system", Current Opinion in Cell Biology, Vol. 32, https://doi.org/10.1016/j.ceb.2015.02.004.

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## AOPs Referencing Relationship

<table><thead><tr><th>AOP Name</th><th>Adjacency</th><th>Weight of Evidence</th><th>Quantitative Understanding</th></tr></thead><tbody><tr><td>AOP: 483 – Deposition of Energy Leading to Learning and Memory Impairment</td><td>adjacent</td><td>Moderate</td><td>Low</td></tr></tbody></table>

## Evidence Supporting Applicability of this Relationship

Evidence for this relationship comes from in vitro mouse- and human-derived models, as well as in vivo mouse and rat models. The relationship is not sex or life stage specific.

### Taxonomic Applicability

<table><thead><tr><th>Term</th><th>Scientific Term</th><th>Evidence</th><th>Link</th></tr></thead><tbody><tr><td>human</td><td>Homo sapiens</td><td>Low</td><td>NCBI</td></tr><tr><td>mouse</td><td>Mus musculus</td><td>Moderate</td><td>NCBI</td></tr><tr><td>rat</td><td>Rattus norvegicus</td><td>Moderate</td><td>NCBI</td></tr></tbody></table>

### Life Stage Applicability

<table><thead><tr><th>Term</th><th>Evidence</th></tr></thead><tbody><tr><td>All life stages</td><td>Moderate</td></tr></tbody></table>

### Sex Applicability

<table><thead><tr><th>Sex</th><th>Evidence</th></tr></thead><tbody><tr><td>Unspecific</td><td>Moderate</td></tr></tbody></table>

## Key Event Relationship Description

Tissue-resident cell activation refers to the stimulation of resident cells in organ systems. Tissue-resident immune cells can be found throughout the body, each tissue and organ containing specific resident immune cells (Chen et al., 2018; Gray & Farber, 2022). Monocytes, found in the blood, and macrophages, found in all tissues in the body, are the main components of the immune system (Ivanova & Orekhov, 2016). In the brain, the primary tissue-resident macrophages are microglia, while astrocytes are also important cells found in the brain (Bourgognon & Cavanagh, 2020; Greene-Schloesser et al., 2012; Wang et al., 2020). Activated tissue-resident cells can undergo gliosis, whereby they adopt a hypertrophic morphology and proliferate, exhibiting rounding of the cell body and retraction of cell processes (Greene-Schloesser et al., 2012; Phatnani & Maniatis, 2015). It is well-characterized that activated tissue-resident cells can increase expression of pro-inflammatory mediators (Hladik & Tapio, 2016; Lumniczky, Szatmari & Safrany, 2017; Kaur et al., 2019). Acute inflammation from controlled biosynthesis of pro-inflammatory mediators protects tissue and promotes healing (Kim & Joh, 2006; Vezzani & Viviani, 2015). Prolonged tissue-resident cell activation leads to dysregulation in production or secretion of pro-inflammatory mediators, which results in chronic inflammation and damage to tissue (Kim & Joh, 2006; Vezzani & Viviani, 2015). Additionally, activated tissue-resident cells can show increased levels of transcription factor nuclear factor κB (NF-κB) and activated protein 1 (AP-1) DNA binding due to increased oxidative stress or DNA damage (Betlazar et al., 2016; Lumniczky, Szatmari & Safrany, 2017). Through the activity of NF-κB, AP-1 and other signaling pathways, activated immune cells can together produce/secrete a variety of cytokines and chemokines (Betlazar et al., 2016; Chen et al., 2018; Greene-Schloesser et al., 2012; Kim & Joh, 2006; Phatnani & Maniatis, 2015; Smith et al., 2012; Wang et al., 2020). Chronic secretion of these inflammatory proteins can lead to downstream detriments, such as in the brain, altering blood-brain barrier permeability (Lumniczky, Szatmari & Safrany, 2017).
*Evidence Supporting this KER*

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

**Biological Plausibility**

Tissue-resident cells recognize pathogens or molecules released by injured or activated cells (Vezzani & Viviani, 2015). In response, resident cells become activated and release various pro-inflammatory mediators (Chen et al., 2018; Gray & Farber, 2022). There is an abundance of studies which explore this relationship using the brain microenvironment, where astrocytes and microglia are the primary tissue-resident cells. After activation, these cells increase in number (whether through proliferation or recruitment), undergo morphological changes and release cytokines (Greene-Schloesser et al., 2012; Kim & Joh, 2006).

Several pathways and molecules are involved in the inflammatory response to activate tissue-resident cells. These molecules include certain pro-inflammatory cytokines and various inflammatory stimuli, such as lipopolysaccharide (LPS), thrombin (a protease), β-amyloid (Aβ), interferon (IFN)-γ, CD40 and gangliosides (Dheen, Kaur & Ling, 2007; Kim & Joh, 2006). For example, LPS can diffuse into brain parenchyma and activate microglia, which then expresses inflammatory mediators and ROS to initiate inflammation (Dheen, Kaur & Ling, 2007; Kim & Joh, 2006). As well, pattern recognition receptor activation, through molecules such as damage-associated molecular pattern molecules (DAMPs), can activate tissue-resident cells and, in turn, lead to pro-inflammatory mediator secretion (Chen et al., 2018).

Once resident cells become activated, various pathways, including the NF-κB transcription factor pathway and the mitogen-activated protein kinase (MAPK)-AP-1 signaling pathway, can result in pro-inflammatory mediator production (Chen et al., 2018; Vezzani & Viviani, 2015; Wang et al., 2020). When activated, microglia and astrocytes are sources of cytokines in the central nervous system (CNS) (Bourgognon & Cavanagh, 2020; Kim & Joh, 2006; Smith et al., 2012; Vezzani & Viviani, 2015; Boyd et al., 2021). Furthermore, uniquely in the brain, astrocyte activation can lead to blood-brain barrier permeability through decreased astrocyte function, which can allow pro-inflammatory mediators to enter from the blood (Lumniczky, Szatmari & Safrany, 2017).

An abundance of studies supports the connectivity of the two key events using activated glial cells in the brain microenvironment. Activated astrocytes express high levels of glial fibrillary acidic protein (GFAP) (Greene-Schloesser et al., 2012), while microglia activation results in the expression of OX-42, Iba1 (activated and non-activated), Mac-1, CD68, F4-80, Glut-5 and CR3/43. These markers play an important role in the phagocytic activity and morphological changes of activated microglia (Jurga, Paleczna & Kuter, 2020). Upon expression of these markers, cells release pro-inflammatory mediators, these can further activate other glial cells (Bourgognon & Cavanagh, 2020). This results in an inflammatory state, that initiates the further release of cytokines including IL-1 and TNF-α, IL-6, Cox-2 (Betlazar et al., 2016; Kim & Joh, 2006; Smith et al., 2012) and chemokines MCP-1 and ICAM-1 (Greene-Schloesser et al., 2012; Kyrkanides et al., 2002). Chronic activation of these cells can result in neurodegenerative disease due to their continuous release and the overexpression of potentially cytotoxic molecules, which may eventually lead to cognitive decline (Dheen, Kaur & Ling, 2007; Kaur et al., 2019; Smith et al., 2012). To suppress this activity, anti-inflammatory proteins need to be released such as tumor growth factor (TGF)-β and IL-10; these act to reduce neuron activity (Kim & Joh, 2006; Phatnani & Maniatis, 2015). In a state of chronic inflammation, the production of pro-inflammatory proteins increases, while anti-inflammatory proteins decrease, and this imbalance results in a reduced stress response (Jeon & Kim, 2016).

**Empirical Evidence**
The evidence supporting the relationship between tissue-resident cell activation leading to increased pro-inflammatory mediators was gathered from studies using human (Chen et al., 2016; Lodermann et al., 2012) and mouse (Dong et al., 2015; Komatsu et al., 2017; Liu et al., 2010; Ramanan et al., 2008; Scharpfenecker et al., 2012; van Neerven et al., 2010; Welser-Alves & Milner, 2013) in vitro cell cultures as well as rat (Lee et al., 2010; Liu et al., 2010; Zhou et al., 2017) and mouse (Dong et al., 2015; Kyrkanides et al. 2002; Parihar et al., 2018) in vivo models. The stressors used were gamma rays (Kyrkanides et al., 2002; Lee et al., 2010), X-rays (Liu et al., 2010; Lodermann et al., 2012; Scharpfenecker et al., 2012), 4He ions (Parihar et al., 2018), nominal photon energy (Zhou et al., 2017) and LPS treatment (Komatsu et al., 2017; van Neerven et al., 2010; Welser-Alves & Milner, 2013). Tissue-resident cell activation is assessed by examining the morphology of activated cells (Dong et al., 2015; Zhou et al., 2017) and the detection of activation markers (Chen et al., 2016; Dong et al., 2015; Liu et al., 2010; Parihar et al., 2018; Scharpfenecker et al., 2012; van Neerven et al., 2010; Welser-Alves & Milner, 2013; Zhou et al., 2017) and AP-1 and NF-κB DNA binding (Komatsu et al., 2017; Lee et al., 2010; Lodermann et al., 2012; Ramanan et al., 2008). Pro-inflammatory mediators, including cytokines, adhesion markers, and inflammatory enzymes were assessed at both the protein and mRNA level although transcriptional responses may not necessarily translate to changes in active signaling molecules. Such studies provide indirect evidence to support the relationship.

## Dose Concordance

Many studies demonstrate that activation of tissue-resident cells occurs at lower or the same doses as increased pro-inflammatory mediators. Although, at doses of ionizing radiation less than 1 Gy, evidence suggests anti-inflammatory activation of macrophages instead of towards inflammation (Wu et al., 2017). This was shown after irradiation of human monocytes from 0.1 to 0.7 Gy which resulted in decreased NF-κB nuclear translocation and decreased IL-1β levels (Lodermann et al., 2012). Dong et al. (2015) used various models and endpoints to study this relationship. In vitro mouse microglia showed an activated morphology, increased F4-80 (activated macrophage indicator), and increased levels and expression of TNF-α and IL-1β all after 16 Gy X-rays (Dong et al., 2015). With the same stressor and dose but in mice kidneys, F4-80 and various pro-inflammatory cytokines were increased (Scharpfenecker et al., 2012). In vivo mice with 10 Gy cranial X-ray irradiation showed a similar response (Dong et al., 2015). Activation of glial cells in rat brains as well as levels of IL-1β and TNF-α in BV2 murine microglial cells increased linearly after 2, 4, 6, 8 and 10 Gy X-rays (Liu et al., 2010). Multiple studies found that LPS treatment at 1 µg/mL resulted in both macrophage activation and TNF-α production (Lodermann et al., 2012; Welser-Alves & Milner, 2013). Rats with 10 Gy gamma ray irradiation showed increases in both NF-κB and AP-1 DNA binding activity, indicators of activated microglia. At the same dose, pro-inflammatory mediators TNF-α, IL-1β and MCP-1 also increased (Lee et al., 2010). When human microglia were irradiated with gamma rays, they showed a characteristic activated morphology after 8 Gy, while CR3/43 and Glut-5 (microglial activation markers) showed increased expression after 8 Gy, and IL-1α and TNF-α also showed increased expression after 8 Gy (Chen et al., 2016).

## Time Concordance

Many studies observed that tissue-resident cell activation occurs earlier or at the same time as increased pro-inflammatory mediators. Mice irradiated with 4He particles showed increases in both activated microglia and pro-inflammatory chemokine CCL-3 after 1 year (Parihar et al., 2018). In vitro murine microglial cells and in vivo rat brains were irradiated with X-rays and showed increases in activated glial cells, as well as IL-1β and TNF-α pro-inflammatory mediators 24h post-irradiation (Liu et al., 2010). After mouse cranial X-ray irradiation, the number of F4-80 positive cells/mm2 was significantly increased from 3h to 2 weeks with a maximum after 24h, while TNF-α and IL-1β both showed significantly increased levels and expression at many times from 3h to 6 weeks post-irradiation, with maximum levels occurring after 24h (Dong et al., 2015). Gamma ray irradiation of rats showed increased NF-κB and AP-1 DNA binding at 4 and 8h after irradiation, with levels reduced to control after 24h, and mRNA and protein levels of pro-inflammatory mediators TNF-α, IL-1β and MCP-1 in the hippocampus and cerebellum showed a similar trend (Lee et al., 2010). Juvenile rats
irradiated with 4MV photon energy showed that microglial density increased at 6 and 24 h post-irradiation in the external germinal layer of the cerebellum, while IL-1α, IL-1β, IL-6, IL-18, GRO/KC and CCL-2 were all significantly increased after 24h (Zhou et al., 2017). Human microglia irradiated with gamma rays showed a characteristic activated morphology after 7 days (Chen et al., 2016). In addition, activation markers CR3/43 and Glut-5 were expressed after 2 weeks and Glut-5 continued expression after 1 week and 10 days. IL-1α and TNF-α showed increased expression after 7 days, which was slightly lower after 2 weeks, but still significant (Chen et al., 2016).

### Incidence Concordance

Two studies identified an incidence-concordant relationship between tissue resident cell activation and increase in pro-inflammatory mediators. Parihar et al. (2018) found increased microglial activation after mice were irradiated with 5 and 30 cGy 4He particles. Chemokine CCL-3 was increased slightly after both 5 and 30 cGy. Zhou et al. (2017) irradiated juvenile rats with a single dose of 6 Gy nominal photon energy (4MV) and showed microglial proliferation in the external germinal layer of the cerebellum, while cytokines (IL-6, IL-18) and chemokines (CCL-2, GRO/KC) increased significantly.

### Essentiality

In the absence of tissue-resident cell activation, an increase in pro-inflammatory mediators is not expected. The activation of tissue-resident cells can be attenuated by tamoxifen. Tamoxifen, an estrogen receptor modulator that can serve as a radiosensitizer, attenuated microglial activation and significantly decreased inflammatory cytokine production, including IL-1β and TNF-α, compared to the X-ray irradiated samples (4, 6, 8, 10 Gy) in absence of tamoxifen (Liu et al., 2010). A study by van Neerven et al. (2010) pretreated cells of mice cerebral cortices with a transcriptional activator, retinoic acid (RA) containing anti-inflammatory effects, for 12 h, and then exposed the cells for another 12 h to RA and LPS, an endotoxin which induces production of IL-1β, IL-6, TNFα in astrocytes. RA, when used with LPS, led to a significant reduction of the LPS-induced release of IL-1β, IL-6, TNF-α (van Neerven et al., 2010). Similarly, N-acetyl-L-cysteine (NAC) at 40 and 80 µM was able to reduce the LPS-induced levels of IL-6, TNF-α while preventing NF-κB nuclear translocation (Komatsu et al., 2017). NF-κB and AP-1 transcription factors binding to DNA indicate activated microglia, which was observed post-irradiation in the study by Ramanan et al. (2008). Radiation led to an increase in TNFα and IL-1β expression in microglia cultures, as well as an increase in Cox-2 protein levels. Treatment with SP 600125 (SP), a specific c-jun kinase inhibitor, which prevented AP-1 binding to DNA, was found to inhibit the radiation-induced increase in TNF-α, IL-1β, and Cox-2 expression. In contrast, treating BV-2 cells with 6-amino-4-(4-phenoxyphenylethylamino) quinazoline (Q), NF-κB activation inhibitor, prevented the increase in NF-κB binding to DNA, thus blocking the activation of tissue-resident cells, which then inhibited the radiation-induced increase in IL-1β expression but allowed the radiation-induced increases in Cox-2 and TNF-α. (Ramanan et al., 2008). Proinflammatory marker, ICAM-1, increased by 2.2-fold 72 h following 35 Gy irradiation (Kyrkanides et al. 2002). However, inhibition of Cox-2, a microglial activator, by NS-398 led to a 0.6-fold decrease in ICAM-1 levels induced by radiation in the brain (Kyrkanides et al. 2002).

### Uncertainties and Inconsistencies

* More work could be done to observe this relationship in human models.

* Limited data is available to support an understanding of this relationship at low doses (<0.1 Gy).

### Known modulating factors

<table><thead><tr><td>Modulating factor</td><td>Details</td><td>Effects on the KER</td><td>References</td></tr></thead><tbody><tr><td>Drug</td><td>Flavonoids</td><td>Flavonoids can inhibit NF-κB, preventing transcription of pro-inflammatory mediators in active glial cells.</td><td>Wang et al., 2020</td></tr></tbody></table>
<table><tr><td>Drug</td><td>Tamoxifen (estrogen receptor modulator commonly used in breast cancer treatment)</td><td>Treatment with Tamoxifen decreased the radiation-induced activation of glial cells. It also consistently decreased the amount of TNF-α and IL-1β and blood-brain barrier permeability after irradiation at various doses.</td><td>Liu et al., 2010</td></tr><tr><td>Drug</td><td>RA (modulates inflammatory effects in different cell types)</td><td>RA treatment completely inhibited the increase in pro-inflammatory mediators after LPS-induced glial activation.</td><td>van Neerven et al., 2010</td></tr><tr><td>Drug</td><td>SP (JNK, c-jun N-terminal kinase, inhibitor)</td><td>AP-1 DNA binding (glial activation) was reduced by SP treatment after irradiation. TNF-α, Cox-2 and IL-1β were reduced by SP treatment after irradiation or viral infection.</td><td>Ramanan et al., 2008</td></tr><tr><td>Drug</td><td>Q (NF-κB inhibitor)</td><td>NF-κB DNA binding (glial activation) was reduced by Q treatment after irradiation. IL-1β was also reduced by Q treatment after irradiation.</td><td>Ramanan et al., 2008</td></tr><tr><td>Drug</td><td>NS-398 (Cox-2 inhibitor)</td><td>Treatment with NS-398 reduced TNF-α, IL-1β, IL-6, ICAM-1 and MCP-1 expression after irradiation.</td><td>Kyrkanides et al., 2002</td></tr><tr><td>Age</td><td>Increased age</td><td>Aging tissue becomes more sensitive to immune signals and increases inflammation. In the aging brain, microglia will produce more pro-inflammatory mediators.</td><td>Patterson, 2015</td></tr><tr><td>Drug</td><td>NAC</td><td>NAC treatment inhibited pro-inflammatory mediator production in macrophages.</td><td>Komatsu et al., 2017</td></tr></table>

## Quantitative Understanding of the Linkage

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><thead><tr><td>Reference</td><td>Experiment Description</td><td>Result</td></tr></thead><tbody><tr><td>Liu et al., 2010</td><td>In vivo. BV2 murine microglia were irradiated with 0, 2, 4, 6, 8 or 10 Gy X-rays to measure microglial activation, and OX-42 and GFAP staining was performed on 15 Gy irradiated rat brains to measure microglial and astrocyte activation, respectively. In vitro. BV-2 murine microglia were irradiated with 0, 2, 4, 6, 8 or 10 Gy X-rays to determine cytokines IL-1β and TNF-α production. Glial activation was identified by light microscopy and immunohistochemistry. ELISA was used to assess cytokine levels of IL-1β and TNF-α.</td><td>Irradiation of in vitro microglia cultures caused a dose-dependent increase in microglial activation from 0 to 10 Gy. At 15 Gy, in vivo astrocyte activation increased 5-fold, while microglial activation increased 3-fold. Levels of IL-1β and TNF-α production were also dose-dependently increased following 0 to 10 Gy irradiated microglia, resulting in an 8.6-fold increase for IL-1β and a 6.8-fold increase for TNF-α after 10 Gy.</td></tr><tr><td>Welser-Alves & Milner, 2013</td><td>In vitro. Cultures of microglia and astrocytes from postnatal mouse central nervous system were stimulated with 1 µg/mL LPS. ELISA and immunocytochemistry were used to measure glial cytokine (TNF-α) production with Mac-1 as a microglial marker.</td><td>Microglia activation with 1 µg/mL LPS led to increased TNF-α production from 40.6 pg/ml to 1875.0 pg/ml, and TNF-α showed co-localization with Mac-1 positive microglia. TNF-α was not present in astrocytes.</td></tr><tr><td>Lee et al., 2010</td><td>In vivo. Rats received whole-brain gamma ray irradiation at 10 Gy. Levels of AP-1 and NF-κB (microglial activation) as well as pro-inflammatory mediators TNF-α, IL-1β, IL-6, and MCP-1 were determined in the hippocampus and cortex. AP-1 and NF-κB DNA binding was determined through electrophoretic mobility shift assay (EMSA), and pro-inflammatory mediator levels were determined using enzyme-linked immunosorbent assay (ELISA).</td><td>After 10 Gy, DNA binding of NF-κB and AP-1 increased a maximum of 3.6-fold and 2.8-fold, respectively. Hippocampus: After 10 Gy at maximum, TNF-α was increased 23-fold, IL-1β increased 10-fold, IL-6 did not significantly change, and MCP-1 increased 1.6-fold. Cortex: After 10 Gy at maximum, TNF-α increased 30-fold, IL-1β increased 7-fold. IL-6 did not significantly change, and MCP-1 increased 2.2-fold.</td></tr><tr><td>Chen et al., 2016</td><td>In vitro. Human CHME5 microglia were irradiated with various doses of 137Cs gamma radiation delivered acutely over 1-3 min. Microglial activation markers CR3/43 and Glut-5 were determined by Western blot, morphology of microglia was determined through</td><td>After 8 Gy, microglia showed a characteristic activated morphology, but not after 0.5 Gy. CR3/43 and Glut-5 were both expressed after 8 Gy, but not 0.5 Gy. mRNA levels measured at 8 Gy were found to increase a maximum of 7.8-fold for IL-1α and 5.8-fold for TNF-α.</td></tr></tbody></table>
<table><tr><td></td><td>fluorescence microscopy, and expression of cytokines IL-1α and TNF-α were determined through RT-PCR.</td><td></td></tr><tr><td>Dong et al., 2015</td><td>In vivo and in vitro. BV2 mouse microglial cells and C57BL/6J mice brains were irradiated with various doses of X-rays. Iba1 staining was performed to determine cell morphology, while anti-F4-80 antibodies were used to determine microglial activation. TNF-α and IL-1β levels were determined through RT-PCR, ELISA (in vitro), and Western blot (in vivo).</td><td>In vitro:<br/>At 16 Gy, microglia adopted a characteristic activated morphology while F4-80 was greatly upregulated. Also at 16 Gy, IL-1β expression increased a maximum of 23-fold, TNF-α expression increased a maximum of 13-fold, IL-1β increased from 0 to a maximum of 530 pg/mL, and TNF-α increased from almost 0 to a maximum of 115 pg/mL.<br/>In vivo:<br/>The number of F4-80 positive cells/mm2 increased from 9 to a maximum of 40 after 10 Gy. Also at 10 Gy, IL-1β expression increased a maximum of 7-fold, TNF-α expression increased a maximum of 5-fold, IL-1β levels increased a maximum of 10-fold, and TNF-α levels increased a maximum of 5-fold compared to controls.</td></tr><tr><td>Komatsu et al., 2017</td><td>In vitro. Murine macrophage RAW264 cell line activation was induced by 1 µg/mL LPS treatment. Activation was determined through NF-κB nuclear translocation measured by western blot. Pro-inflammatory mediators TNF-α and IL-6 were measured by ELISA.</td><td>After LPS treatment, cytosolic NF-κB decreased 0.64-fold, nuclear NF-κB increased 7.2-fold and IκB (NF-κB inhibitor) decreased 0.21-fold. Both TNF-α and IL-6 were increased from around 0 ng/mL to about 70 and 30 ng/mL, respectively.</td></tr><tr><td>Lodermann et al., 2012</td><td>In vitro. Human monocytic leukemia cell lines were irradiated with X-rays at 0.1, 0.3, 0.5, 0.7 and 1 Gy. Activation was determined through NF-κB nuclear translocation by western blot. IL-1β was measured by ELISA.</td><td>NF-κB and IL-1β both showed linear decreases from 0.1 to 0.7 Gy, resulting in maximum decreases of 0.6- to 0.7-fold. Although, the decrease for NF-κB was nonsignificant. No changes were observed at 1 Gy.</td></tr><tr><td>Scharpfenecker et al., 2012</td><td>In vitro. Mice kidneys were irradiated with 16 Gy of X-rays. Immunofluorescence staining was performed for IL-6, IL-1β and macrophage marker F4-80.</td><td>F4-80 positive area increased by 6.7 and 3.8-fold in Eng+/- and Eng+/- irradiated mice respectively. Macrophages in irradiated mouse kidney led to IL-6 and IL-1β production.</td></tr></table>

Time Concordance

<table><thead><tr><td>Reference</td><td>Experiment Description</td><td>Result</td></tr></thead><tbody><tr><td>Parihar et al., 2018</td><td>In vivo. C57BL/6 J mice were irradiated with 4He particles 1 year after irradiation. The level of CCL-3 was determined in the brain, and microglial activation was determined by immunohistochemistry in the perihedral cortex.</td><td>At maximum, there was a 3.8-fold increase in activated microglia, CCL-3 increased 2.2-fold.</td></tr><tr><td>Liu et al., 2010</td><td>In vitro. BV2 murine microglia were irradiated with X-rays to measure microglial activation after 24h, and OX-42 and GFAP staining was performed on irradiated rat brains to measure microglial and astrocyte activation, respectively, after 3 days. BV-2 murine microglia were irradiated with X-rays to determine cytokines IL-1β and TNF-α production after 24h. Glial activation was identified by light microscopy and immunohistochemistry. ELISA was used to assess cytokine levels of IL-1β and TNF-α.</td><td>10 Gy irradiation of microglia cultures caused an increase in microglial activation after 24h. At 15 Gy after 3 days, astrocyte activation increased 5-fold, while microglial activation increased 3-fold. Levels of IL-1β and TNF-α were also increased in 10 Gy irradiated microglia, resulting in an 8.6-fold increase for IL-1β and a 6.8-fold increase for TNF-α after 24h.</td></tr><tr><td>Lee et al., 2010</td><td>In vivo. Rats received whole-brain gamma ray irradiation at 10 Gy. Levels of AP-1 and NF-κB (microglial activation) as well as pro-inflammatory mediators TNF-α, IL-1β, IL-6, and MCP-1 were determined 4, 8, and 24h after irradiation in the hippocampus and cortex. AP-1 and NF-κB DNA binding was determined through electrophoretic mobility shift assay (EMSA), and pro-inflammatory mediator levels were determined using enzyme-linked immunosorbent assay (ELISA).</td><td>DNA binding of NF-κB and AP-1 increased a maximum of 3.6-fold and 2.8-fold, respectively, after 8h. Binding activity returned to control levels after 24h.<br/>Hippocampus:<br/>At control, TNF-α was 3.6 pg/mg protein, which increased 23-fold after 4 hours, 8.3-fold after 8h, and 3.6-fold after 24h. IL-1β increased linearly from 4 to 24h, reaching a 10-fold maximum increase. IL-6 did not significantly change other than a non-significant decrease over 24h, even though an increase in IL-6 mRNA was observed at 4h with RT-qPCR. MCP-1 showed a maximum increase of 1.6-fold after 8h.<br/>Cortex:<br/>At control, TNF-α was 4.4 pg/mg protein, which increased 30-fold after 4 hours, 13-fold after 8h, and 4.1-fold after 24h. IL-1β showed a maximum increase of 7-fold after 4h. IL-6 did not significantly change other than a non-significant decrease over 24h, even though an increase in IL-6 mRNA was observed at 4h</td></tr></tbody></table>
<table><tr><td>Chen et al., 2016</td><td>In vitro. Human CHME5 microglia were irradiated with 8 Gy gamma radiation (137Cs source) delivered acutely over 1-3 min. Microglial activation markers CR3/43 and Glut-5 were determined by Western blot, morphology of microglia was determined through fluorescence microscopy, and expression of cytokines IL-1α and TNF-α were determined through RT-PCR.</td><td>with RT-qPCR. MCP-1 showed a maximum increase of 2.2-fold after 8h.<br>Beginning after 7 days, microglia showed a characteristic activated morphology. CR3/43 was expressed after 2 weeks, while Glut-5 was expressed after 1 week, 10 days, and 2 weeks. After 7 days, mRNA levels were found to increase a maximum of 7.8-fold for IL-1α and 5.8-fold for TNF-α. mRNA levels dropped slightly but were still above controls after 2 weeks.</td></tr><tr><td>Zhou et al., 2017</td><td>In vivo. Juvenile rats were irradiated with 4MV nominal photon energy and a single dose of 6 Gy (2.3 Gy/min). At 6 or 24h, the molecular and cellular changes in the EGL of the cerebellum was studied. Immunohistochemistry staining was used to measure Iba1 (microglia marker) with morphometry analysis performed on microglia. Luminex assay measured cytokines, chemokines and growth factors for the inflammatory response.</td><td>Microglia density increased by 2.3-fold after 6 h and 6.77-fold 24 h post-irradiation. Most of the iba1-positive cells had a bushy or amoeboid morphology, signifying an activated state. At 6 h of irradiation, IL-1α and CCL-2 increased by 2-2.8-fold. IL-1β decreased at 6 h then slightly increased at 24 h post-irradiation. IL-6, IL-18, GRO/KC, VEGF, and GM-CSF all increased significantly at 24 h compared to the control group.</td></tr><tr><td>Dong et al., 2015</td><td>In vivo and in vitro. BV2 mouse microglial cells and C57BL/6J mice brains were irradiated with X-rays. Iba1 staining was performed to determine cell morphology, while anti-F4-80 antibodies were used to determine microglial activation. TNF-α and IL-1β levels were determined through RT-PCR, ELISA (in vitro), and Western blot (in vivo) from 3h to 6 weeks after irradiation.</td><td>In vivo, the number of F4-80 positive cells/mm2 increased from 9 to a maximum of 40 after 24h, decreased over 2 weeks, and returned to control levels at 4 and 6 weeks. IL-1β expression increased to a maximum after 72h and was significantly increased from 6h to 6 weeks. TNF-α expression was a maximum after 3 and 6h, was at controls from 24h to 1 week, and was increased again from 2 to 6 weeks. IL-1β levels were high after 3h, lower at 6h, increased to a maximum at 2 weeks, then decreased at 4 and 6 weeks. TNF-α levels increased to a maximum after 72h, then decreased until 4 weeks, where they increased again after 6 weeks.<br>In vitro. IL-1β levels reached a maximum at 3h, but then decreased at 6h, before rising again at 12h. TNF-α levels remained elevated up to 24h, although its peak was at 6h.</td></tr></table>

Incidence Concordance

<table><thead><tr><td>Reference</td><td>Experimental Description</td><td>Result</td></tr></thead><tbody><tr><td>Parihar et al., 2018</td><td>In vivo. C57BL/6 J mice were irradiated with 4He particles at either 5 or 30 cGy (5 cGy/min). The level of CCL-3 was determined in the brain, and microglial activation was determined by ED-1 immunohistochemistry in the perirhinal cortex.</td><td>Microglial activation increased 3.5-fold after 5 cGy and 3.8-fold after 30 cGy. CCL-3 increased 1.4-fold (non-significant, ns) after 5 cGy and 2.2-fold after 30 cGy.</td></tr><tr><td>Zhou et al., 2017</td><td>In vivo. Juvenile rats were irradiated with 4 MV nominal photon energy and a single dose of 6 Gy (2.3 Gy/min). At 6 or 24h, the molecular and cellular changes in the EGL of the cerebellum was studied. Immunohistochemistry staining was used to measure Iba1 (microglia marker) with morphometry analysis performed on microglia. Luminex assay measured cytokines, chemokines and growth factors for the inflammatory response.</td><td>Microglia density increased by 2.3-fold after 6 h and 6.77-fold 24 h post-irradiation. Most of the iba1-positive cells had a bushy or amoeboid morphology, signifying an activated state. At 6 h of irradiation, IL-1α and CCL-2 increased by 2-2.8-fold. IL-1β decreased at 6 h then slightly increased at 24 h post-irradiation. IL-6, IL-18, GRO/KC, VEGF, and GM-CSF all increased significantly at 24 h compared to the control group.</td></tr></tbody></table>

Known Feedforward/Feedback loops influencing this KER

It is well-characterized that activated tissue-resident cells can increase expression of pro-inflammatory mediators (Hladik & Tapio, 2016; Lumniczky, Szatmari & Safrany, 2017; Kaur et al., 2019). However, there exists a feedforward loop for this key event relationship as pro-inflammatory mediators can also activate tissue-resident cells within the brain and perpetuate the inflammatory response (Kim & Joh, 2006; Vezzani & Viviani, 2015). Thus, after stimulation by cytokines, chemokines or inflammogens such as from damaged neurons, microglia and astrocytes activate inflammatory signaling pathways, which result in increased expression and/or release of inflammatory mediators such as cytokines, eicosanoids, and metalloproteinases (Dong & Benveniste, 2001; Bourgognon & Cavanagh, 2020). Various studies have shown that overexpression of IL-1β in mouse models resulted in the appearance of inflammatory markers including activated glial cells and increased pro-inflammatory cytokine and chemokine mRNAs (Hein et al., 2010; Moore et al., 2009). Additionally, IL-6 plays a role in activating
glial cells as mouse models with IL-6 knocked out showed reduced astrocytic population, as well as a reduced ability in activating microglia (Klein et al., 1997). Cytokines and chemokines can also increase the permeability of the blood-brain barrier, further increasing pro-inflammatory mediator levels (Lumniczky, Szatmari & Safrany, 2017).
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## AOPs Referencing Relationship

<table><thead><tr><td>AOP Name</td><td>Adjacency</td><td>Weight of Evidence</td><td>Quantitative Understanding</td></tr></thead><tbody><tr><td>AOP: 483 – Deposition of Energy Leading to Learning and Memory Impairment</td><td>adjacent</td><td>Moderate</td><td>Low</td></tr></tbody></table>

## Evidence Supporting Applicability of this Relationship

Evidence for this relationship comes from rat and mouse models. There is in vivo evidence in both male and female animals, with more evidence in males. Animal age is occasionally not indicated in studies, but most evidence is in adult rodent models.

## Taxonomic Applicability

<table><thead><tr><td>Term</td><td>Scientific Term</td><td>Evidence</td><td>Link</td></tr></thead><tbody><tr><td>mouse</td><td>Mus musculus</td><td>Moderate</td><td>NCBI</td></tr><tr><td>rat</td><td>Rattus norvegicus</td><td>Moderate</td><td>NCBI</td></tr></tbody></table>

## Life Stage Applicability

<table><thead><tr><td>Term</td><td>Evidence</td></tr></thead><tbody><tr><td>Adult</td><td>Moderate</td></tr><tr><td>Not Otherwise Specified</td><td>Low</td></tr><tr><td>Juvenile</td><td>Low</td></tr></tbody></table>

## Sex Applicability

<table><thead><tr><td>Sex</td><td>Evidence</td></tr></thead><tbody><tr><td>Male</td><td>Moderate</td></tr><tr><td>Female</td><td>Low</td></tr><tr><td>Mixed</td><td>Moderate</td></tr><tr><td>Unspecific</td><td>Low</td></tr></tbody></table>

## Key Event Relationship Description

Inflammatory mediators such as IL-1β, TNF-α, and IL-6 can affect the normal behavior of neuronal cells through alterations in: (a) the neuronal architecture and (b) synaptic activity. Overexpression of these pro-inflammatory mediators can disrupt the integrity of neurons through increased necrosis and demyelination, decreased neurogenesis, neural stem cell proliferation and synaptic complexity (Cekanaviciute et al., 2018; Fan & Pang, 2017). Structurally, the neuron is comprised of the cell body, dendrites, axon, and axon terminals, all of which are critical in the normal functioning of the central nervous system. Another important component of the neuron is its signaling properties, which uses chemical neurotransmitters to transfer messages in the synaptic cleft (Cekanaviciute et al., 2018; Hladik & Tapio, 2016). Disruption to these structures or signaling properties results in abnormal neural remodeling.

Under physiological conditions, cytokine levels are low, but these can increase in response to various insults. Cytokines mediate immune response through ligand binding to cell surface receptors, which activate signaling cascades such as the JAK-STAT or MAPK pathways to produce or recruit more cytokines. Once organs initiate inflammatory reactions, the cytokines are capable of impairing neural function through direct effects on neurons or by indirect mechanisms mediated by microglia, astrocytes or vascular endothelial cells (Mousa & Bakhiet, 2013; Prieto & Cotman, 2018).
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

**Biological Plausibility**

Various studies provide support for the biological plausibility of the link between an increase in pro-inflammatory mediators and abnormal neural remodeling. It is known that cytokines and their receptors are constitutively expressed by neurons in the central nervous system, and even in normal or pathological states, these cytokines can be produced by individual cells and act on neurons. At the tissue level, the collective action and distribution of the cytokines can influence the overall inflammatory response within the brain tissue, impacting its function (Kishimoto et al., 1994). One important factor in the pathogenesis of neurotoxicity is the overexpression of pro-inflammatory mediators, or Th1-type, cytokines. These cytokines bind to their receptors to induce a conformational change, which triggers the activation of intracellular signaling pathways that alter cell structure and function (Mousa and Bakhiet, 2013). Several sources have reported a change in physical and electrophysiological properties of neurons in either whole-brain samples, specific brain regions such as the hippocampus or dentate gyrus, or neuronal cell cultures in response to increased expression of cytokines (Jenrow et al., 2013; Fan and Pang, 2017; Wong et al., 2004). The main cytokines presenting detrimental effects are IL-1β, TNF-α, and IL-6, which can cause alterations in neuronal architecture such as morphological changes in dendrites and synapses (Tang et al., 2017; Cekanaviciute et al., 2018; Shi et al., 2017). Many studies have also reported decreased proliferation and differentiation in progenitor cells, inhibited neural stem cell differentiation and decreased neurogenesis following increases in cytokines (Zonis et al., 2015; Wong et al., 2004; Tang et al., 2017). IL-6 can affect neurogenesis through various distinct mechanisms. One mechanism is through the stimulation of hypothalamic-pituitary-adrenal axis, which then increases circulating glucocorticoids. These steroids can then inhibit cell proliferation and neurogenesis in the dentate gyrus (Turnbull and Rivier, 1999; Gould et al., 1992; Cameron and Gould, 1994). Decreased neurogenesis in the hippocampus is also well documented as a result of pro-inflammatory mediators, and one possible mechanism for this detrimental effect is through the interaction of IL-1β and orphan nuclear receptor, TLX. This receptor is required to maintain the neural precursor cell pool in neurogenic brain regions, and it has been shown that IL-1β can reduce the expression of TLX and consequently cell proliferation (Ryan et al., 2013). TNF-α affects neuronal fate by interacting with its receptor, TNFR1, which is expressed on neural stem cells. It has been reported that TNFR1-mediated signaling pathway inhibits growth therefore, a reduction in neuron production after TNF-α injection (Chen and Palmer, 2013). A clear mechanistic relationship has not yet been established, although it is accepted that pro-inflammatory mediators can alter the structure and function of neurons.

**Empirical Evidence**

The empirical evidence collected for this KER comes from both in vitro and in vivo studies. Abnormal neural remodeling can be defined by changes to the physical, functional and/or electrophysiological properties of neurons. Most of the evidence examines effects of inflammation induced by either dextran sodium sulfate (DSS) treatment, seizure induction, heavy ion or X-ray irradiation at higher doses (>5 Gy) and varying dose-rates from 78.0 cGy/min to 3.2 Gy/min, or lipopolysaccharide injection on both in vivo and in vitro rodent models. Injections or treatments with pro-inflammatory cytokines were also used. These studies provide evidence to support a causal association between an increase in pro-inflammatory mediators and decreased neurogenesis, but also provide evidence of reduced neuronal signaling, neural stem cell and progenitor cell proliferation, as well as a change in overall neuronal shape (Zonis et al., 2015; Vallieres et al., 2002; Green et al., 2012; Wu et al., 2012; Chen and Palmer, 2013; Saraiva et al., 2019; Zhang et al., 2017; Dong et al., 2015; Kalm et al., 2013; Wong et al., 2004; Jenrow et al., 2013; Liu et al., 2010).
**Time Concordance**

Multiple studies suggest time-concordant relationship between increases in pro-inflammatory mediators leading to abnormal neural remodeling. The proliferation and survival of stem and progenitor cells, collectively known as neural precursor cells, was decreased or inhibited in response to increase pro-inflammatory cytokines at 4 days, 7 days and 1-month post-treatment (Zonis et al., 2015; Wong et al., 2004, Green et al., 2012; Ryan et al., 2013; Vallieres et al., 2002). Various studies also report a reduction in neurogenesis in the hippocampus and dentate gyrus 1-month post-treatment that was well-correlated with increased IL-1β, TNF-α, and IL-6 levels (Zonis et al., 2015; Vallieres et al., 2002; Wu et al., 2012; Chen and Palmer, 2013). In addition to a significant decrease in neurogenesis 1-month post-treatment, a study found that sustained IL-1β expression caused a significant reduction in neurogenesis that persisted for 3 months (Wu et al., 2012). Another study observed that X ray-induced increases in IL-1β and TNF-α at three and six hours resulted in a time-dependent decrease in DCX+ cells, a marker for neurogenesis, which persisted for 2 weeks. This was seen after a single dose of 10 Gy (Dong et al., 2015). Thus, these studies provide evidence to support time concordance of the relationship using both in vivo and in vitro models.

**Dose Concordance**

No available data.

**Incidence Concordance**

No available data.

**Essentiality**

Several treatments that directly alter pro-inflammatory mediators preserve neuronal integrity in the dentate gyrus and positively modulate hippocampal neurogenesis. The treatments included MW-151, a selective inhibitor of pro-inflammatory cytokine production and Kukoamine A, an alkaloid that inhibits neuronal oxidative stress and hippocampal apoptosis (Jenrow et al., 2013; Zhang et al., 2017). Another treatment used was histamine, which has been shown to ameliorate the loss of neuronal complexity and synaptic plasticity (Saraiva et al., 2019). Multiple studies use cytokine receptor antagonists or knock-out key receptors to block the effects of IL-1β, TNF-α, and CCL2, which have preservatives neuron survival (Green et al., 2012; Ryan et al., 2013; Wu et al., 2012; Chen and Palmer, 2013). Another knockout model of complement component 3 (C3) have also been used to demonstrate essentiality. C3 is a central molecule in the complement cascade, and its roles include microglia-mediated synapse elimination. C3 knockout models have been shown to cause reduced pro-inflammatory cytokines, increased synaptic number, reduced neuron loss and synaptic morphology impairment (Shi et al., 2017).

**Uncertainties and Inconsistencies**

* Various in vitro studies have reported a stimulation of neural precursor cell proliferation and differentiation or increased neurogenesis by different cytokines such as IL-6 and IFN-γ (Islam et al., 2009; Wong et al., 2004). Another study found increased proliferation within the hippocampus after repeated IL-6 and IL-1β infusion (Seguin et al., 2009). Although a clear mechanism has not yet been elucidated, it is thought that these cytokines have contradictory effects from the differential activation of various signaling cascades (Borsini et al., 2015). For example, hyper-IL-6, a fusion of IL-6 and IL-6 receptor, was found to increase neurogenesis through the activation of MAPK/CREB (mitogen-activated protein kinase/cAMP response element binding protein) cascade (Islam et al., 2009).

* It has also been reported that TNF-α exhibits neuroprotective effects as their transmembrane receptors can influence different signaling pathways (Figiel, 2008; Masli & Turpie, 2009)

* Kalm et al. (2013) found a higher inflammatory response in lipopolysaccharide (LPS) treated females compared with males after irradiation. Specifically, increased levels of pro-inflammatory cytokines IL-1β, IL-12, and IL-17, as well as pro-inflammatory chemokines CCL4, CCL3 and
CCL2 were detected relative to vehicle-treated animals and LPS-treated males. This was associated with a 32% decrease in DCX+ cells, a marker for neurogenesis, in females. However, in LPS-treated males, a 64% reduction in DCX+ cells compared to vehicle-treated males following irradiation was reported (Kalm et al., 2013). Further research is required to elucidate the exact effects of increased pro-inflammatory mediators on neural integrity between males and females.

## Known modulating factors

<table><thead><tr><td>Modulation Factor</td><td>Details</td><td>Effects on the KER</td><td>References</td></tr></thead><tbody><tr><td>Drug Therapy</td><td>MW01-2-151SRM (MW-151) – water soluble, nontoxic, bioavailable compound that mitigates pro-inflammatory cytokine production, glial activation and inflammation in rat hippocampus.</td><td>MW-151 reduced the neuroinflammation caused by 10 Gy of heavy ion exposure, thus preserving the integrity of neurogenic signaling in the dentate gyrus.</td><td>Jenrow et al., 2013</td></tr><tr><td>Genetic Manipulation</td><td>IL-1 receptor antagonist to prevent the interaction between IL-1β with IL-1R1.</td><td>After 7 days in vitro, IL-1β significantly decreased the percentage of DCX-positive neurons, but pre-treatment and subsequent co-treatment with IL-1RA abolished this anti-neurogenic effect of IL-1β.</td><td>Green et al., 2012</td></tr><tr><td>Neuromodulator</td><td>Histamine – an endogenous amine that can regulate both brain inflammation and neurogenesis.</td><td>Histamine treatment significantly increased the total number cells, positively modulates hippocampal neurogenesis, ameliorates the loss of neuronal complexity of hippocampal neuroblasts and reverts synaptic plasticity loss caused by LPS.</td><td>Saraiva et al., 2019</td></tr><tr><td>Drug</td><td>Tamoxifen – synthetic, non-steroidal estrogen receptor modulator with anti-inflammatory and neuroprotective properties.</td><td>Tamoxifen decreased the production of inflammatory cytokines released from irradiated microglia, attenuating glial activation and decreasing neuronal apoptosis.</td><td>Liu et al., 2010</td></tr><tr><td>Drug</td><td>Kukoamine A (KuA) – alkaloid extracted from traditional Chinese herb cortex lycii radicis that has been previously reported to have antioxidant properties.</td><td>KuA inhibited radiation-induced increases in pro-inflammatory cytokines, alleviated the activation of hippocampal microglia and ameliorated the suppression of hippocampal neurogenesis.</td><td>Zhang et al., 2017</td></tr><tr><td>Genetics</td><td>Polymorphism that increases the expression of APOE4 increases the risk of developing Alzheimer's diseases, which generally consists of a decline in memory, thinking and language.</td><td>In homozygous human APOE4 knock-in mice, a dramatic increase in pro-inflammatory cytokines TNF-α, IL-1β and IL-6 was seen after LPS injection compared to the APOE2 and APOE3 alleles, suggesting that APOE4 is implicated in a greater inflammatory response.</td><td>Hunsberg et al., 2019; Zhu et al., 2012</td></tr></tbody></table>

## Quantitative Understanding of the Linkage

The table below provides some representative examples of quantitative linkages between the two key events as presented in the paper. 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.

### Time Concordance

<table><thead><tr><td>Reference</td><td>Experiment Description</td><td>Results</td></tr></thead><tbody><tr><td>Zonis et al., 2015</td><td>In vitro. Adult female C57Bl/6 mice were treated with 3% wt/vol dextran sodium sulfate (DSS) using a multiple-cycle administration for up to 26 days. ELISA and qRT-PCR were used to measure cytokine levels in neural precursor cells and neural progenitor cell cultures. Western blot and immunostaining were used to measure Ki67 (neuronal cell proliferation marker), DCX (marker for neurogenesis), BLBP (marker for stem/early progenitor cells) and nestin (neural stem cell marker and involved in radial growth of axons). p21 (regulator of cell cycle progression at G1 and S phase) was also measured using these methods.</td><td>Neural precursor cells (NPCs) treated with 50 ng/mL of IL-6 led to a decline in the % of neuroblasts, with declines from 41% ± 6.4% in untreated cells to 22.39% ± 5.2% in IL-6-treated cells). Thus, decreasing neurogenesis was observed in in NPCs after DSS treatment. Chronic intestinal inflammation (29 days post-treatment) reduced hippocampal neurogenesis. Evidence has highlighted that IL-1β and TNF-α mRNA levels were increased more than 2-fold, as well as a fourfold increase in p21 mRNA levels, which were accompanied by an approximate 2-fold down-regulation of progenitor cell proliferation and neurogenesis. Protein levels of IL-1β and TNF-α were not assessed.</td></tr><tr><td>Jenrow et al., 2013</td><td>In vivo. Adult male Fischer 344 rats were exposed to 10 Gy of whole-brain irradiation (WBI) using a 137Cs irradiator at a dose rate of 3.2 Gy/min. Rats also underwent mitigating therapy with MW01-2-151SRM (MW-151), a selective inhibitor of β amyloid-induced glial pro-inflammatory cytokine production that was initiated</td><td>MW-151 mitigated radiation-induced neuroinflammation, measured by OX-6+ cell densities, an indicator of pro-inflammatory level of activation, at 2- and 9-months post-irradiation. The 10 Gy group had mean OX-6+ cell densities of 1,966 (±218) and 1,493 (±270) cells/mm3, respectively. In the 10 Gy MW-151 group, the mean OX-6+ cell densities were</td></tr></tbody></table>
<table>
  <tr>
    <td></td>
    <td>24 h post-WBI and was continued for 28 days post-WBI by daily injection. Immunofluorescence assays for cell proliferation and neurogenesis were performed at 2 months and for neuroinflammation at 2 and 9 months post-WBI.</td>
    <td>significantly reduced to 849 (±350) and 437 (±119) cells/mm3, respectively. In addition, 2 months post-irradiation, the mean DCX+ cell density, a marker for neurogenesis, was 1,375 (±535) cells/mm3 in the 10 Gy group, but was significantly increased to 4,702 (±622) cells/mm3 after MW-151 therapy.</td>
  </tr>
  <tr>
    <td>Vallieres et al., 2002</td>
    <td>In vitro. Seizure was induced in transgenic mice expressing IL-6 under control of the glial fibrillary acidic protein (GFAP) promoter. Bromodeoxyuridine (BrdU) assay and immunofluorescence was used to detect proliferating neural progenitor cells and fluoro-jade staining was used to detect degenerating neurons.</td>
    <td>IL-6 compromised proliferation, survival, and differentiation of hippocampal progenitor cells when expressed over the long term (31 days) in young adult transgenic mice expressing the IL-6 transgene, causing a 63% decrease in the production rate of new neurons. The BrdU assay revealed a 27% reduction in progenitor cell proliferation, 53% reduction in progenitor cell survival in the dorsal hippocampus.</td>
  </tr>
  <tr>
    <td>Wong et al., 2004</td>
    <td>In vitro. Neural stem cell lines derived from adult C57BL/6 mice were allowed to proliferate and differentiate for 1 h to 3 days in the presence of IFN-γ and TNF-α. Proliferation and cytotoxicity assays were used to assess neural stem cell survival and their ability to differentiate and proliferate.</td>
    <td>IFN-γ (100 U/mL) and TNF-α (10 ng/mL) inhibited neural stem cell proliferation. The reduction was first significant at 4 days when control cells began to proliferate rapidly, and until day 6, control cells continued to proliferate while IFN-γ and TNF-α inhibited proliferation.</td>
  </tr>
  <tr>
    <td>Green et al., 2012</td>
    <td>In vitro. Rat hippocampal NPC cultures were treated with IL-1β (10 ng/mL) in the presence or absence of IL-1 receptor antagonist (IL-1RA), which prevents the interaction of IL-1β with IL-1R1. Cell proliferation analysis was performed, and RT-PCR and immunoblotting were used to measure DCX and cytokines. MTT assay was used to assess cell viability.</td>
    <td>Treatment with 10 ng/mL of IL-1β significantly decreased neurosphere circumference in a time-dependent manner. Compared to untreated cultures, a difference was seen at day 4, which continued until day 7. Neural integrity was assessed by examining the neural cell viability and proliferation IL-1β treatment for 24 hours after 4 days in vitro significantly decreased cell proliferation. IL-1β treatment for 7 days in vitro caused a significant decrease in cell viability compared to untreated cultures.</td>
  </tr>
  <tr>
    <td>Wu et al., 2012</td>
    <td>In vitro. Wild-type and IL-1βXAT C57BL/6 mice were exposed to Cre recombinase to induce IL-1β overexpression. Immunohistochemistry was used to measure IL-1β and DCX, a marker for migrating neuroblasts and neurogenesis.</td>
    <td>There was a significant effect of sustained IL-1β overexpression on adult neurogenesis one month after injection. A 94% decrease in migrating neuroblasts and neurogenesis was found, mirroring the pattern of neuroinflammation. 3 months post-injection, there was also an 87% reduction in neurogenesis.</td>
  </tr>
  <tr>
    <td>Chen and Palmer, 2013</td>
    <td>In vitro. Microglial cultures isolated from neonatal pups of C57BL/6 mice were treated with 1 µg/mL LPS, then incubated in neural stem cell (NSC) differentiation medium. The conditioned medium was then collected and applied to NSCs. Immunohistochemistry was used to measure cytokine levels and DCX, a marker for neurogenesis.</td>
    <td>After 72 h, neurogenesis significantly reduced in the differentiation culture after TNF-α injection of 20 ng/mL. One month after injection, proliferation and survival of endogenous neural stem cells decreased, as well as a reduction in neurogenesis.</td>
  </tr>
  <tr>
    <td>Saraiva et al., 2019</td>
    <td>In vitro. C57BL/6J male mice were injected with 1 or 2 mg/kg LPS. Histamine was also injected in the hippocampal dentate gyrus. The BrdU assay was used to quantify proliferating cells, while immunohistochemical analysis and western blot was used to measure IL-1β and DCX.</td>
    <td>In mice exposed to LPS, histamine significantly increased the total amount of proliferative cells. Mice exposed to 1 mg/kg of LPS had 98.8 ± 4.7 BrdU+ cells, a marker for proliferating cells, whereas 1 mg/kg of LPS + His had 197.6 ± 28.2 BrdU+ cells. 2 mg/kg of LPS yielded 96.7 ± 9.7 BrdU+ cells and 2 mg/kg of LPS + Histamine had 154.1 ± 23.8 BrdU+ cells. Histamine was also able to revert the LPS-induced loss of neuronal volume from 238.6 ± 19.7 (1 mg/kg LPS) to 331.1 ± 33.4 (1 mg/kg LPS + His) µm3. And 248.7 ± 18.8 (2 mg/kg LPS) to 334.3 ± 24.8 (2 mg/kg LPS + His) µm3. Postsynaptic density protein 95 (PSD-95) levels were also elevated due to histamine treatment (2 mg/kg LPS, 55.8 ± 9.9; 2 mg/kg LPS + His, 110.1 ± 12.9).</td>
  </tr>
  <tr>
    <td>Ryan et al., 2013</td>
    <td>In vitro. Adult Sprague Dawley rat dentate gyrus NPC cultures were prepared and treated with IL-1β (100 ng/mL) for 7 days. Immunohistochemistry was used to detect NPCs, proliferating cells and newly born cells.</td>
    <td>24 h after IL-1β treatment (100 ng/mL), there was no difference in proliferating neural cells relative to untreated cells. However, at 7 days post-treatment, there was a significant reduction in proliferating neural cells. There was also a decrease in neurogenesis.</td>
  </tr>
  <tr>
    <td>Dong et al., 2015</td>
    <td>In vitro. The mouse microglial cell line, BV-2, were irradiated with a single 16 Gy dose of X-rays, then assessed at various time points up to 6 weeks. ELISA was used to measure levels of IL-1β and TNF-α, whereas immunohistochemical staining was used to detect DCX+ cells.</td>
    <td>DCX+ cells, a marker of neurogenesis, were significantly reduced at 6 h until 2 weeks post-irradiation, which was accompanied by increased levels of IL-1β and TNF-α. TNF-α levels peaked at 3 h post-irradiation, decreased at 6 h, then increased in a time-dependent manner until 2 weeks. IL-1β levels increased in a time-dependent manner until peaking at 72 h, then spiked again at 6 weeks.</td>
  </tr>
</table>
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## AOPs Referencing Relationship

<table><thead><tr><th>AOP Name</th><th>Adjacency</th><th>Weight of Evidence</th><th>Quantitative Understanding</th></tr></thead><tbody><tr><td>AOP: 483 - Deposition of Energy Leading to Learning and Memory Impairment</td><td>adjacent</td><td>Moderate</td><td>Low</td></tr></tbody></table>

## Evidence Supporting Applicability of this Relationship

Evidence for this relationship comes from rat and mouse models. There is in vivo evidence in both male and female animals, with more evidence in males. Animal age is occasionally not indicated in studies, but most evidence is in adult rodent models.

## Taxonomic Applicability

<table><thead><tr><th>Term</th><th>Scientific Term</th><th>Evidence</th><th>Link</th></tr></thead><tbody><tr><td>mouse</td><td>Mus musculus</td><td>Moderate</td><td>NCBI</td></tr><tr><td>rat</td><td>Rattus norvegicus</td><td>Moderate</td><td>NCBI</td></tr></tbody></table>

## Life Stage Applicability

<table><thead><tr><th>Term</th><th>Evidence</th></tr></thead><tbody><tr><td>Adult</td><td>Moderate</td></tr><tr><td>Not Otherwise Specified</td><td>Low</td></tr><tr><td>Juvenile</td><td>Low</td></tr></tbody></table>

## Sex Applicability

<table><thead><tr><th>Sex</th><th>Evidence</th></tr></thead><tbody><tr><td>Male</td><td>Moderate</td></tr><tr><td>Female</td><td>Low</td></tr></tbody></table>

## Key Event Relationship Description

Neural stem cells (NSCs) come from different sources, such as the subgranular zone (SGZ) located in the dentate gyrus (DG) of the hippocampal formation, and the subventricular zone (SVZ) region of the brain. NSCs give rise to mature neurons that are then able to receive signals from other neurons (Bálentová & Adamkov, 2020; Hladik & Tapio, 2016; Monje & Palmer, 2003; Romanella et al., 2020; Tomé et al., 2015). Changes in neuronal architecture can lead to altered synaptic activity, necrosis, demyelination, neurogenesis, neurodegeneration, and dendrite morphology, all of which encompass abnormal neural remodeling (Hladik & Tapio, 2016; Monje & Palmer, 2003; Tomé et al., 2015). These alterations can then cause cognitive deficits in the form of impaired learning and memory (Barker & Warburton, 2011; Hladik & Tapio, 2016; Monje & Palmer, 2003; Tomé et al., 2015). Impaired learning consists reduced ability to create new associative or non-associative relationships, whereas impaired memory consists of decreased ability to establish sensory, short-term or long-term memories (Desai et al., 2022; Kiffer et al., 2019). Multiple brain areas are involved these processes, such as the hippocampal region, amygdala, prefrontal cortex, basal ganglia, and other areas of the neocortex (Cucinotta et al., 2014; Desai et al., 2022; NCRP

## Evidence Supporting this KER

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
Several reviews provide support for the biological plausibility of the link between abnormal neural
remodeling and impaired learning and memory. It is generally accepted that abnormal neural
remodeling can disrupt learning and memory through changes to neurogenesis, neurodegeneration,
neuronal excitability and synaptic plasticity, demyelination and dendritic spine density (Hladik &
Tapio, 2016; Bálentová & Adamkov, 2020; Tomé et al., 2015; Monje & Palmer, 2003; Romanella et
al., 2020).

Neurogenesis in the DG creates new neurons in the hippocampus that can make connections to Cornu Ammonis (CA) neurons involved in learning and memory (Monje & Palmer, 2003; Tomé et al., 2015). Accordingly, learning and memory can be impaired through reduced neurogenesis in the neurogenic SGZ of the DG (Bálentová & Adamkov, 2020; Monje & Palmer, 2003). Many studies also associate decreased neurogenesis to impaired hippocampal-dependent cognitive function, indicating that it is a common mechanism for the relationship (Tomé et al., 2015). Similarly, apoptosis and neurodegeneration impair cognitive function (Bálentová & Adamkov, 2020; Hladik & Tapio, 2016). In the hippocampus, the degree of atrophy corresponds to the degree of impaired learning and memory (Tomé et al., 2015).

Synaptic strength and neuronal excitability are important components of learning and memory. Decreased hippocampal excitability and disrupted long-term potentiation (LTP), a form of long-term synaptic plasticity, are associated with reduced learning and memory (Romanella et al., 2020). Changes in the expression of synaptic receptors and other synaptic proteins may also result in impaired learning and memory (Hladik & Tapio, 2016).

Demyelination correlates with decreased long-term memory formation (Tomé et al., 2015) and along
with white matter necrosis, these lead to impaired learning and memory (Bálentová & Adamkov,
2020). Although demyelination is a factor in learning and memory, sub-threshold demyelination may
still cause impaired learning and memory (Monje & Palmer, 2003).

Reduced dendritic complexity and spine density are also associated with impaired learning and
memory (Bálentová & Adamkov, 2020; Hladik & Tapio, 2016; Romanella et al., 2020). The
complexity of signal processing in the hippocampus can be reduced by a loss in dendritic spines, which
results in impaired learning and memory (Romanella et al., 2020). In addition, various studies show
reduced dendritic branching, length and area in hippocampal neurons associated with learning and
memory (Hladik & Tapio, 2016).

Empirical Evidence

The empirical evidence surrounding this KER stems from research using in vivo models. When compared to the control, irradiated subjects demonstrated significant changes in neuronal integrity and cognitive deficits (Achanta, Fuss & Martinez, 2009; Acharya et al., 2019; Akiyama et al., 2001; Hodges et al., 1998; Howe et al., 2019; Krukowski et al., 2018; Madsen et al., 2003; Miry et al., 2021; Parihar et al., 2015, 2016; Raber et al., 2004; Rola et al., 2004; Simmons et al., 2019; Sorokina et al., 2021; Whoolery et al., 2017; Winocur et al., 2006).

Dose Concordance

There is strong evidence suggesting dose-concordance between abnormal neural remodeling and impaired learning and memory. Dose-dependent changes in dendritic complexity are correlated with behavioral deficits at doses from 0.05 to 30 Gy (Bálentová & Adamkov, 2020; Howe et al., 2019; Parihar et al., 2015, 2016; Simmons et al., 2019; Whoolery et al., 2017). Mice exposed to 0.05 and 0.3 Gy of 48Ti showed dose-dependent decreases in total dendritic length and number of dendritic branch points. When subjected to cognitive testing, mice exposed to 0.3 Gy demonstrated greater impairment in the novel object recognition (NOR) and object in place (OiP) tests compared to 0.05 Gy exposed mice (Parihar et al., 2016). These results for OiP were correlated positively to the number of dendritic spines and negatively to the number of postsynaptic density protein 95, also a marker for abnormal neural remodeling. Similarly, another study identified a correlation between the total number of