Radiation‐Resistant Bacteria Deinococcus radiodurans‐Derived Extracellular Vesicles as Potential Radioprotectors

Abstract

The increasing use of radiation presents a risk of radiation exposure, making the development of radioprotectors necessary. In the previous study, it is investigated that Deinococcus radiodurans (R1‐EVs) exert the antioxidative properties. However, the radioprotective activity of R1‐EVs remains unclear. In the present study, the protective effects of R1‐EVs against total body irradiation (TBI)‐induced acute radiation syndrome (ARS) are investigated. To assess R1‐EVs' radioprotective efficacy, ARS is induced in mice with 8 Gy of TBI, and protection against hematopoietic (H)‐ and gastrointestinal (GI)‐ARS is evaluated. The survival rate of irradiated mice group decreases substantially after irradiation. In contrast, pretreatment with R1‐EVs increases the survival rates of the mice. The administration of R1‐EVs provides effective protection against radiation‐induced death of bone marrow cells and splenocytes by scavenging reactive oxygen species (ROS). Additionally, R1‐EVs protect both intestinal stem and epithelial cells from radiation‐induced apoptosis. R1‐EVs stimulate the production of short‐chain fatty acids in the gastrointestinal tract, suppress proinflammatory cytokines, and increase regulatory T cells in pretreated mice versus the irradiation‐only group. Proteomic analysis shows that the R1‐EV proteome is significantly enriched with proteins involved in oxidative stress response. These findings highlight R1‐EVs as potent radioprotectors with applications against radiation damage and ROS‐mediated diseases.

Deinococcus radiodurans‐derived extracellular vesicles (R1‐EVs) provide radioprotection against total‐body irradiation‐induced acute radiation syndrome in mice. R1‐EVs mitigate oxidative damage by scavenging free radicals, promoting intestinal repair, enhancing hematopoietic function, and modulating immune responses. This study highlights the potential therapeutic application of R1‐EVs in managing radiation‐induced gastrointestinal and hematopoietic damage.

1. Introduction

The increasing use of radiation technology has raised concerns about the potential for nuclear accidents and the risk of nuclear terrorism. Exposure to high doses of total‐body irradiation (TBI) can lead to acute radiation syndrome (ARS), which is categorized into hematopoietic (H‐ARS, 1.5–4 Gy), gastrointestinal (GI‐ARS, 6–15 Gy), and neurovascular (NV‐ARS, 15 Gy) syndromes, depending on the dose.^{[ 1 ]} This exposure precipitates the rapid generation of reactive oxygen species (ROS) from radiation‐induced water radiolysis, causing acute or chronic oxidative stress in cells and tissues.^{[ 2 ]} Despite six decades of research, the only FDA‐approved radioprotectant is amifostine, whose clinical use is limited by rapid excretion and toxic side effects such as vomiting and hypotension, thus confining its application to salivary gland protection.^{[ 1} , ^{3 ]} This underscores the urgent need for new radioprotective agents with high ROS scavenging capabilities and minimal toxicity to effectively prevent and treat the severe consequences of high‐dose radiation exposure.

Deinococcus radiodurans is a gram‐positive, red‐pigmented, and non‐pathogenic bacterium that is known for its exceptional resistance to extreme stress conditions, such as radiation, oxidation, and desiccation.^{[ 4 ]} This organism can withstand acute gamma radiation doses of up to 5000 Gy without cell death or mutation, which is significantly higher than the lethal doses for humans (5 Gy) and Escherichia coli (200–800 Gy).^{[ 5 ]} D. radiodurans employs a suite of sophisticated survival mechanisms including a hyperstable surface layer, specialized stress‐response proteins, highly condensed nucleoids, robust DNA repair systems, and advanced protein protection strategies.^{[ 6 ]} Its antioxidant defense is particularly notable, featuring both enzymatic scavengers, such as catalase and superoxide dismutase, and non‐enzymatic antioxidants, such as carotenoids and manganese complexes.^{[ 7 ]} Small‐molecule manganese antioxidants play a crucial role in the oxidative stress response.^{[ 8 ]} Given these unique properties, D. radiodurans has considerable potential for application in bioindustry and extremophile biology, exploiting its durability and resilience in harsh environments.^{[ 9 ]}

Extracellular vesicles (EVs) are small lipid membrane‐enclosed vesicles released by a wide array of cells, containing a diverse group of proteins, lipids, metabolites, and nucleic acids reflective of their cell of origin.^{[ 10 ]} EVs play crucial roles in intercellular and interspecific communication by facilitating the transfer of bioactive molecules that maintain physiological homeostasis and contribute to cellular functions.^{[ 11 ]} Owing to their capacity to transport these substances, EVs are increasingly recognized as versatile therapeutic tools, with applications spanning immune modulation, drug delivery, and diagnostics.^{[ 12 ]} Recent research has expanded to include bacterial EVs, which are released by both gram‐negative and gram‐positive bacteria and range from 20 to 400 nm in size.^{[ 13 ]} These bacterial EVs carry a variety of bioactive components, including lipopolysaccharides, peptidoglycans, and nucleic acids, which show potential for treating diseases such as inflammatory bowel disease and allergies and serve as innovative agents in immunotherapy, nanovaccine development, and drug delivery systems.^{[ 14 ]} Notably, EVs derived from Lactobacillus plantarum have been shown to induce anti‐inflammatory M2 macrophage polarization in vitro, highlighting their therapeutic potential in modulating immune responses and managing inflammation‐driven conditions.^{[ 15 ]}

Our previous study demonstrated the protective roles of EVs derived from D. radiodurans (R1‐EVs) against H₂O₂‐induced oxidative stress in HaCaT cells,^{[ 16 ]} suggesting that the notable antioxidant properties of D. radiodurans are effectively mirrored in its EVs. Based on this, we hypothesized that R1‐EVs could provide significant radioprotection against high‐dose irradiation. This hypothesis is further supported by recent findings that vesiculation is an adaptive mechanism that may enhance radiation resistance in D. radiodurans exposed to low‐Earth orbit conditions.^{[ 17 ]} Therefore, this study aimed to evaluate the efficacy of R1‐EVs as radioprotective agents in a TBI‐induced ARS mouse model, potentially establishing a new avenue for therapeutic interventions in radiation‐related injuries.

2. Experimental Section

2.1. Culture of D. radiodurans

Deinococcus radiodurans R1 (ATCC 13939) was purchased from American Type Culture Collection (ATCC). The bacteria were cultured at 30 °C in tryptone glucose yeast extract (TGY) broth, which consisted of 0.5% tryptone (Difco Laboratories, Detroit, MI, USA), 0.3% yeast extract (Difco Laboratories), and 0.1% glucose (Sigma–Aldrich, St. Louis, MO, USA). For the solid media, 1.5% bacto‐agar (Difco Laboratories) was added to the TGY mixture.

2.2. Isolation and Characterization of R1‐EVs

The methods used in this study closely followed those used in our previous work,^{[ 16 ]} in which the isolation and characterization of R1‐EVs were extensively described. Briefly, D. radiodurans strains were incubated at 30 °C for 72 h under static conditions to facilitate the isolation of R1‐EVs. Following culturing in TGY broth, the bacteria‐free culture supernatants were centrifuged at 10000 × g for 30 min at 4 °C and filtered through a 0.45 µm bottle‐top vacuum filter system (Corning, Merck KGaA, Darmstadt, Germany). Subsequently, EVs in the filtered supernatants were isolated and concentrated from 1 L to 50 mL using tangential flow filtration with a 500 kDa MWCO ultrafiltration membrane filter capsule (Pall Life Sciences, Port Washington, NY, USA). The R1‐EV pellets were collected by ultracentrifugation at 100 000 × g for 2 h at 4 °C, then washed with sterile phosphate‐buffered saline (PBS; pH 7.4), and further purified using an Optiprep density gradient medium (Sigma, #D1556, Steinheim, Germany). The purification involved ultracentrifugation at 170 000 × g for 18 h at 4 °C without brake in a discontinuous 60% Optiprep density gradient (ranging from 10% to 60% w/v). The final EV pellet was re‐suspended in PBS and stored at −80 °C. The protein content of R1‐EVs was measured using a bicinchoninic acid protein assay kit (Thermo Scientific Pierce, Rockford, IL, USA) following the manufacturer's instructions.

The size distribution, zeta potential, and concentration of R1‐EVs were characterized using dynamic light scattering (DLS), scanning electron microscopy (SEM), and nanoparticle tracking analysis (NTA). The hydrodynamic size and zeta potential of the R1‐EVs was measured using a Zetasizer Nano ZS Zen3600 (Malvern, UK). For SEM analysis, the samples were fixed in 3.7% glutaraldehyde (Sigma‐Aldrich GmbH, Taufkirchen, Germany) in PBS for 15 min, washed twice with PBS, and dehydrated using a graded ethanol series (40%, 60%, 80%, and 96–98%). After the ethanol evaporation, the samples were air‐dried at room temperature (RT) for 24 h on a glass substrate. The dried samples were coated with gold‐palladium and examined using a Quanta 400 scanning electron microscope (FEI, Hillsboro, OR, USA). The particle concentration was confirmed using NTA with a QUATT ZetaViewⓇ (PARTICLE METRIX, Meerbusch, Germany).

2.3. Antioxidant Activity

The antioxidant activity of R1‐EVs was evaluated using superoxide dismutase (SOD) activity, catalase (CAT) activity, and hydroxyl radical antioxidant capacity (HORAC) assays.

2.3.1. SOD Activity Assay

The SOD activity was measured using an EZ‐SOD assay kit (DG‐SOD400; DoGenBio, Seoul, Republic of Korea). R1‐EVs were prepared at concentrations of 0.1, 0.2, and 1 mg mL⁻¹. Ascorbic acid (5 mm) was used as the positive control. The reaction mixture consisted of 20 µL of R1‐EV, 200 µL of WST‐1 working, and 20 µL of enzyme working solutions. The mixture was incubated at 37 °C for 20 min, and the absorbance was measured at 450 nm using a microplate reader (Infinite 200Pro, Tecan, Männedorf, Switzerland).

2.3.2. CAT Activity Assay

The CAT activity was determined using an EZ‐catalase assay kit (DG‐CAT400, DoGenBio). R1‐EVs were prepared at concentrations of 0.5, 1, and 5 µg mL⁻¹. Ascorbic acid (0.5 mm) was used as the positive control. Then, 25 µL of R1‐EVs solution was mixed with 25 µL of hydrogen peroxide (H₂O₂) substrate solution. The reaction mixture was then incubated at RT for 30 min. Following incubation, 50 µL of Oxi‐Probe/HRP Working Solution was added to the mixture and then incubated at 37 °C for 30 min. The absorbance was measured at 560 nm using a microplate reader (Infinite 200Pro, Tecan).

2.3.3. HORAC Assay

The HORAC assay was performed using the OxiSelect HORAC activity assay kit (STA‐346, Cell Biolabs, San Diego, CA, USA). R1‐EVs were prepared at concentrations of 0.1, 0.2, and 1 mg mL⁻¹. Gallic acid provided with the kit was used as a positive control. The assay reaction was initiated by adding 140 µL of fluorescein working solution to 20 µL of R1‐EVs, followed by incubation for 30 min at RT. After incubation, 20 µL of hydroxyl radical initiator solution was added, followed immediately by the addition of 20 µL of Fenton reagent. The fluorescence intensity was measured at an excitation wavelength of 480 nm and an emission wavelength of 530 nm using a fluorescence microplate reader (i‐control Infinite 200, Tecan).

2.4. Proteome Analysis of R1‐EVs

2.4.1. Sample Preparation for Mass Spectrometry

A total of 30 µg of purified EV protein was lysed in a lysis buffer (comprising 4% SDS, 100 mm dithiothreitol, and 100 mm Tris‐HCl at pH 7.6) to prepare samples for mass spectrometry. The mixture was then incubated at 37 °C for 45 min. Subsequently, the samples were boiled for the 10 min boiling step to denature the proteins, and were then loaded onto Microcon 30 kDa centrifugal filters (Merck) and centrifuged at 14000 × g for 40 min. The filter was washed thrice with denaturing UA buffer (8 M urea in 100 mm Tris‐HCl, pH 8.5). The samples were treated with an IAA solution (containing 50 mm iodoacetamide in UA buffer) and incubated for 30 min in the dark at RT. The filter was washed thrice with UA buffer and twice with 50 mm ammonium bicarbonate at pH 8.0. Finally, the samples were digested with trypsin at an enzyme‐to‐protein ratio of 1:50 and incubated at 37 °C for 12 h. The resulting tryptic peptides were collected with 50‐mM ammonium bicarbonate at pH 8.0 and purified on a C18 spin column (Thermo Fisher Scientific).

2.4.2. LC‐MS/MS

Desalted peptides were reconstituted in buffer A containing 0.1% formic acid. Subsequently, 1 µg of these peptides was analyzed using an UltiMate 3000 Nano LC system (Thermo Fisher Scientific) coupled to an Orbitrap Fusion Lumos Tribrid mass spectrometer (Thermo Fisher Scientific). The sample separation was performed using a microcapillary column (75 µm internal diameter × 500 mm length) packed with 2 µm average diameter C18 resin. The sample was eluted with a linear gradient of buffers A (0.1% formic acid in distilled water) and B (0.1% formic acid in 80% acetonitrile) for 135 min at a flow rate of 0.3 µL min⁻¹ as follows: 2% buffer B for 5 min, 45% buffer B for 120 min, 100% buffer B for 125 min, 100% buffer B for 130 min, 2% buffer B for 131 min, and 2% buffer B for 135 min. The full MS scans were performed over an m/z range of 300–1800 at a resolution of 120000. The maximum injection time was set to 35 ms, and the automatic gain control was set to 1.0 × 10⁴. For the MS/MS scans, the 20 most intense peptide ions with charge states > 1 were fragmented via higher‐energy collision‐induced dissociation (automatic gain control, 5000; maximum injection time, 35 ms). The dynamic exclusion was set at 30 s.

2.4.3. Mass Spectrometry Data Analysis

Peptide identification and label‐free quantification were performed using Proteome, utilizing the Sequest HT search engine and the Minora feature algorithm within Discoverer 3.0. The MS/MS spectra were matched to peptide spectra based on the D. radiodurans reference proteome from the National Center for Biotechnology Information. A tolerance of 10 ppm was set for precursor ions and 0.6 Da for fragment ions. The dynamic modifications included oxidation of methionine and acetylation of protein N‐terminal residues, whereas carbamidomethylation of cysteine was set as a fixed modification. For protein identification, the peptide spectrum match and protein false discovery rates were set to 0.05, and each protein had to contain at least one unique peptide. The list of identified proteins was then exported, and protein identifiers were used to perform an over‐representation analysis on the EV proteome using the PANTHER database (20240226 release) and the Gene Ontology classification of functional terms (GO Ontology database https://doi.org/10.5281/zenodo.10536401 Released 2024‐01‐17). For over‐representation analysis, the significance of over‐represented functional terms was assessed using Fisher's exact test with a P‐value of 0.05, with the identified proteins analyzed against a background of known proteins from D. radiodurans genome. Only significantly over‐represented terms are reported.

2.5. Animals

All animal experiments were approved by the Institutional Animal Care and Use Committee of the Korea Atomic Energy Research Institute (approval no. KAERI‐IACUC‐2021‐003) and conducted according to the ethical standards of the KAERI Animal Care Center. Female C57BL/6 mice, aged 7 weeks, with an initial body weight of 18 ± 2 of grams, were obtained from Orient Bio Inc. (Seoul, Republic of Korea). Prior to the experiments, the mice were acclimatized to laboratory conditions with a temperature maintained at 25 ± 2 °C, relative humidity at 50 ± 5%, and a 12‐h light/dark cycle. Throughout the study, mice had ad libitum access to a sterile commercial diet and water.

2.6. Irradiation Study

TBI was conducted using a Gammacell 40 Exactor (Nordion International Inc., Ottawa, Canada), which delivers a dose rate of 0.9 Gy min⁻¹ from a ¹³⁷Cs γ‐ray source. The mice (n = 10) were divided into three groups: control (Con), irradiated (IR), and R1‐EV pretreated irradiated (R1‐EVs + IR). The mice were intraperitoneally injected with 10 mg kg⁻¹ R1‐EVs or PBS 1 h before irradiation. Subsequently, the mice were placed in a chamber and exposed to a single radiation dose (8 Gy). After irradiation, the mice were monitored daily for 30 d to assess their body weight and physical attributes using a cumulative clinical scoring system. The system quantitatively evaluated factors such as weight loss, temperature fluctuations, external appearance, body position, mobility, food consumption, and hydration.^{[ 18 ]} Each parameter was scored based on its severity or deviation from normal, producing an aggregate score that reflected the overall health status of each animal.

2.7. Intracellular ROS Assay of Bone Marrow Cells

The mice were euthanized 1 h after irradiation, and their femurs were collected. All soft tissues attached to the femurs were removed, and the metaphyses were exposed by excising the condyles and epiphyses. Bone marrow cells (BMCs) were pumped from the femurs and centrifuged at 5000 × g for 1 min to harvest cell pellets. The red blood cells were lysed in a lysis buffer (Sigma‐Aldrich, St. Louis). For the intracellular ROS assay, the harvested BMCs were incubated in PBS containing dichlorodihydrofluorescein diacetate (50 µm) for 30 min, followed by washing thrice with PBS. The fluorescence intensity of the oxidized dichlorofluorescein was measured using FACSVerse flow cytometer. Data analysis was performed using FlowJo software (V10, BD Biosciences, V10, BD Biosciences, Ashland, OR, USA).

2.8. Viability Assay of Bone Marrow Cells and Splenocytes

One day after radiation exposure, the mice were euthanized, and their BMCs and splenocytes were collected. The harvested cells were subjected to red blood cell lysis and the number of viable cells was measured using Cellometer Mini (Nexcelom, Lawrence, MA, USA).

2.9. Malondialdehyde Assay for Plasma

The blood plasma was collected from the mice 1 d postirradiation. The samples were centrifuged at 2000 × g for 20 min at 4 °C to separate the supernatant. The protein concentration in the supernatant was measured using the bicinchoninic acid assay. Plasma malondialdehyde (MDA) levels, a marker for oxidative stress, were quantified using OxiSelect TBARS (Cell Biolabs) following the manufacturer's instructions.

2.10. Hematoxylin and Eosin Staining and Immunohistochemistry

The femurs and spleen were harvested 3 d post‐irradiation, while the intestines were harvested 7 d post‐irradiation. The tissues were fixed in 10% neutral‐buffered formalin and embedded in paraffin. Prior to embedding, the femurs underwent an additional decalcification step, and the intestines were thoroughly flushed with PBS to clear the debris from the lumen. Tissue sections of 4 µm thickness were prepared for histological examination.

For routine histopathological assessment, the tissue sections were stained using the standard hematoxylin and eosin (H&E) method. The slides were evaluated by a pathologist who was blinded to the experimental conditions, and semi‐quantitative scores were assigned based on the observed tissue pathology.

Immunohistochemistry was performed using paraffin‐embedded colon sections. The sections were deparaffinized in xylene and rehydrated using a graded series of ethanol solutions. They were then incubated with primary antibodies against Ki67 (1:300 dilution; Novus, Littleton, CO, USA) and leucine‐rich repeat‐containing G‐protein coupled receptor 5 (LGR5, 1:50 dilution; Abcam, Cambridge, MA, USA) at 4 °C overnight. Following primary antibody incubation, the sections were exposed to HRP‐conjugated goat anti‐rabbit IgG (Agilent, Santa Clara, CA, USA) for 30 min at 37 °C. The detection was performed using a DAB kit (Sigma‐Aldrich, St. Louis), and positive staining was visualized and quantified using a microscope equipped with Motic Images Plus 2.0 software (Motic, Hong Kong, China).

2.11. Intestinal Permeability Assay

Five days post‐irradiation, the mice were fasted for 12 h and then orally administered 0.5 mg kg⁻¹ of fluorescein isothiocyanate (FITC)–dextran (4 kDa; Sigma‐Aldrich, St. Louis, MO, USA). Approximately 3 h after drug administration, the blood samples were collected via cardiac puncture. Serum was obtained by centrifuging the blood at 300 × g for 20 min. The fluorescence intensity of the serum samples was quantified using a microplate reader (Zenyth 3100; Anthos Labtec Instruments GmbH, Salzburg, Austria).

2.12. Analysis of Short‐Chain Fatty Acids in Mouse Feces

2.12.1. Fecal Short‐Chain Fatty Acid Extraction and Derivatization

Following the method described by Zhang et al.^{[ 19 ]} with minor modifications, the short‐chain fatty acid (SCFA) analysis was conducted on day 5 postirradiation. Fresh fecal samples were collected and immediately frozen at −80 °C. For analysis, 30 mg of each sample was homogenized in 300 µL of HPLC‐grade water using an Omni Bead Ruptor 24 at 6500 rpm for 20 s. The homogenate was agitated at 4 °C for 30 min and then centrifuged at 13000 × g for another 30 min. Subsequently, 100 µL of the supernatant was acidified with 10 µL of 5 M HCl and mixed with 100 µL of anhydrous diethyl ether. This mixture was vortexed for 5 s, chilled on ice for 5 min, and centrifuged at 10000 × g for 5 min to separate the phases. The ether layer containing SCFAs was transferred to a new tube and dried over Na₂SO₄. Following two further centrifugations, 100 µL of the clear supernatant was transferred to a GC vial for derivatization. The SCFAs were derivatized by adding 5 µL of N,O‐bis(trimethylsilyl)trifluoroacetamide, vortexing briefly, incubating at 70 °C for 40 min, and then at 37 °C for another 120 min. The derivatized SCFAs were then ready for GC–MS analysis.

2.12.2. GC‐MS Analysis

The SCFAs were quantified using GC‐MS system (QP 2010 Ultra, Shimadzu Corporation, Kyoto, Japan) equipped with DB‐5MS UI capillary column (30 m × 0.25 mm × 0.25 µm film thickness; J&W Scientific, CA, USA). The temperatures of the injector, ion source, and interface were set at 260 °C, 230 °C, and 280 °C, respectively. The sample (1 µL) was injected in a 10:1 split mode at a constant flow rate of 1 mL min⁻¹ using helium (99.999%) as the carrier gas.

The column oven temperature was initially set to 40 °C and held for 2 min, then increased to 150 °C at a rate of 15 °C min⁻¹ and maintained for 1 min, and finally ramped up to 300 °C at a rate of 30 °C min⁻¹, where it was held for 5 min. The electron impact ionization was performed at a voltage of 70 eV. The total run time was 20 min, and the mass spectral data were collected in full scan mode from a mass range of 40–400 m/z at an acquisition speed of 12.8 scans per second.

To identify SCFAs, the selected ion monitoring mode was used to confirm the target and qualifier ions. The target ions for acetic, propionic, isobutyric, butyric, isovaleric, and valeric acids were identified at m/z values of 117, 131, 145, 145, 159, and 159, respectively. The qualifier ions for all acids were at m/z 75, with an additional m/z of 117 for acids containing 4–5 carbons. The SCFA concentrations were calculated by confirming the retention times and comparing the mass spectra with those of external standards.

2.13. Assay of Cytometric Bead Array

The serums were collected 7 d post‐irradiation, and the Mouse Th1/Th2/Th17 Kit (BD, Cat# 560485, San Jose, CA, USA) was used to detect the concentrations of IL‐2, IL‐4, IL‐6, IFN‐γ, TNF, IL‐17A, and IL‐10 in serum. All related reagents, flow cytometry protocols, and results were analyzed according to the kit instructions. The cytokine standards were prepared to generate standard curves. The capture beads of the above seven cytokines were mixed rigorously to prepare the mixture of capture beads. Subsequently, 50 µL of the mixture was added to each sample, closely followed by adding 50 µL of PE detection reagent. All array tubes were incubated for 2 h at RT in the dark. The array tubes were rinsed once with wash buffer, centrifuged, and re‐suspended before flow cytometry. FACSVerse flow cytometer was used for the content of cytokines. Data analysis was performed using FlowJo software (V10, BD Biosciences).

2.14. Analysis of Splenic T Cells in a Mouse Model of TBI‐Induced ARS

The spleens were harvested 7 d postirradiation. After euthanasia, splenocytes from each mouse group were isolated and stimulated for 4 h at 37 °C using 1X Cell Stimulation Cocktail (Thermo Fisher Scientific) supplemented with PMA, ionomycin, and protein transport inhibitors. For Th1, Th2, and Th17 cell analyses, the cells were initially stained with anti‐CD3 antibody (Alexa488; eBioscience, San Diego, CA, USA), anti‐CD4 antibody (V450; eBioscience), and L/D reagent (Invitrogen, Carlsbad, CA, USA) for 15 min at RT. Post fixation and permeabilization using BD Cytofix/Cytoperm kit reagents (Thermo Fisher Scientific), the cells were washed and subsequently stained with anti‐IFN‐γ (PE; BD Biosciences), anti‐IL‐5 (APC; BD Biosciences), and anti‐IL‐17A (PE‐Cy7; BD Biosciences) antibodies for 20 min at RT. For regulatory T‐cell analysis, the splenocytes were directly stained with anti‐CD3 (Alexa488), anti‐CD4 (V450), and anti‐CD25 (PE‐Cy7; Thermo Fisher Scientific) antibodies along with the L/D reagent for 15 min at RT. After washing with cold PBS, the cells were fixed and permeabilized using Foxp3/Transcription Factor Staining Buffer Set (Thermo Fisher Scientific). Following several washes with a fixation/permeabilization diluent (Thermo Fisher Scientific), the cells were stained with an anti‐Foxp3 (PE; Thermo Fisher Scientific) antibody for 20 min at RT. Flow cytometry analysis of regulatory T cells (CD3+CD4+CD25+Foxp3+) was performed using a FACSVerse™ flow cytometer, and data were processed using FlowJo software (Version 10, BD Bioscience). The absolute number of regulatory T cell subsets was determined by multiplying the percentage of each cell type by the total cell count obtained from the single‐cell suspension for each group.

2.15. Statistical Analysis

Statistical analyses were conducted using GraphPad Prism 8 (GraphPad Software Inc., San Diego, CA, USA). Data are expressed as mean ± standard deviation (SD). For comparisons involving multiple groups, one‐way analysis of variance (ANOVA) followed by Tukey's multiple comparison test was employed. For pairwise comparisons, an unpaired t‐test was utilized. Statistical significance was determined at a p‐value of less than 0.05. Significance levels are indicated in the figures as follows: #P < 0.05, ##P < 0.01, ###P < 0.001 versus control group; *P < 0.05, **P < 0.01, ***P < 0.001 versus IR group.

2.16. Ethics Approval Statement

All animal experiments were approved by the Institutional Animal Care and Use Committee of the Korea Atomic Energy Research Institute (approval no. KAERI‐IACUC‐2021‐003) and conducted according to the ethical standards of the KAERI Animal Care Center.

3. Results

3.1. Isolation and Characterization of R1‐EVs

The R1‐EVs were isolated and characterized in accordance with the Minimal Information for Studies of EVs guidelines.^{[ 20 ]} The EVs were isolated from cultured D. radiodurans supernatants using a combination of differential centrifugation, tangential flow filtration, and density gradient ultracentrifugation. Following isolation, the R1‐EVs were comprehensively characterized. The dynamic light scattering analysis demonstrated that the isolated R1‐EVs had a mode size of 320 nm with a zeta potential of −8.37 mV (Figure  1A,B). The nanoparticle tracking analysis results indicated a concentration of 1.7 × 10⁸ particles per 0.2 µg mL⁻¹ of R1‐EVs (Figure 1C). The SEM confirmed that the diameter of R1‐EVs was approximately 330 nm, revealing the round‐shaped structural characteristics of EVs (Figure 1D).

Figure 1.

Characterization of EVs isolated from Deinococcus radiodurans. A) Size distribution analysis of R1‐EVs by dynamic light scattering (DLS). B) Zeta potential of R1‐EVs was analyzed through DLS. C) Particle concentration and size of R1‐EVs were analyzed by nanoparticle tracking analysis (NTA). D) Scanning electron microscopy (SEM) image of R1‐EVs. Scale bar = 300 nm. ROS‐scavenging performance of R1‐EVs eliminate E) O2‐, F) H₂O₂, and G) OH•. Data are shown as mean ± SD of three independent experiments.

Given that EVs are known to reflect the characteristics of their source cells, we hypothesized that R1‐EVs would exhibit significant antioxidant properties. We measured the activities of SOD, CAT mimetics, and HORAC, which represent the radical‐scavenging effects of superoxide (O₂ ⁻), hydrogen peroxide (H₂O₂), and hydroxyl radicals (OH•), respectively. The results demonstrated a concentration‐dependent increase in the antioxidant activity (Figure 1E–G). Specifically, the R1‐EVs effectively scavenged ROS in all the assays. The SOD and CAT assays indicated that R1‐EVs could effectively neutralize O₂ ⁻ and H₂O₂ radicals. Additionally, the OH• radical scavenging ability of R1‐EVs, as assessed using the HORAC assay, was comparable to that of the positive control, 1.5 mm gallic acid at a concentration of 1 mg mL⁻¹. These findings suggested that R1‐EVs possess potent ROS‐scavenging effects, underscoring their potential antioxidant and radioprotective roles.

3.2. . Proteome Profiles of R1‐EVs

We performed a proteomic analysis of two independently isolated EV samples by subjecting them to tryptic digestion, followed by LC‐MS/MS analysis. The resulting mass spectrometry data were analyzed using Proteome Discoverer (v3.0) for protein identification and further processed through the PANTHER database for over‐representation analysis of functional terms using Gene Ontology's functional classification system. A total of 169 proteins were identified, with a false discovery rate of 5% (APPENDIX /list of proteins). The functional over‐representation analysis of the identified proteins showed that the vesicular proteins were significantly over‐represented in response to oxidative stress (GO:0042744, GO:0042743, GO:0042542, and GO:0072593), glycolipid transport (GO:0046836), and protein import (GO:0017038). The precise H₂O₂ catabolic process (GO:0042744) was the most enriched biological process term (Figure  2 ). Similarly, oxidoreductase (GO:0016715), glycolipid transfer (GO:0017089), and CAT activity (GO:0004096) were the most significantly over‐represented molecular function terms. Other significantly enriched molecular function terms included peptidoglycan glycosyl transferase (GO: 0008955), penicillin‐binding (GO 0008658), and ligand‐gated channel (GO:0022834) activities. As shown in Figure 2 (cellular component), we found significant enrichment of proteins belonging to the periplasmic space, cell envelope, cell membrane, and transmembrane transport complex, with the outer membrane‐bound periplasmic space (GO:0030288) being the most significantly enriched cellular component term.

Figure 2.

Functional over‐representation analysis of R1‐EVs proteome. Identified proteins were analyzed against a background of all the proteins in Deinococcus radiodurans (R1) using Fisher's exact test for significance threshold; p value = 0.05. Only gene ontology functional terms that were significant are shown. The data is presented as a bubble plot with x‐axis showing the fold enrichment and y‐axis the functional term from all the three gene ontology categories. The size of the dot shows the count of proteins from each pathway identified in the R1‐EV proteome while the significance threshold (‐log10 p value) is indicated by the color of each dot (red indicates the smallest p value while green shows larger p values).

3.3. Administration of R1‐EVs Protects against Radiation‐Induced Mortality in Mice and Enhances Survival and Mitigates Radiation‐Induced Mortality in Mice

One hour after R1‐EV injection, the mice were subjected to TBI (8 Gy), and assays were performed to evaluate H‐ARS and GI‐ARS at the indicated time points (Figure  3A). Notably, the survival rate of mice in the irradiated group drastically decreased after irradiation, resulting in 85% mortality rate after 30 d. However, pretreatment with R1‐EVs (10 mg kg⁻¹) significantly increased the survival rates to 85% following TBI (Figure 3B). Correspondingly, mice that received R1‐EV inoculation (R1‐EVs + IR) also showed significant improvements in clinical scores post‐TBI, reflecting a direct correlation with increased survival rates (Figure 3C). Importantly, this treatment demonstrated excellent biocompatibility, showing no toxicity to major organs and no significant changes in body weight (Figure S1, Supporting Information). Additionally, R1‐EV treatment provided long‐term protection 150 d without toxicity after TBI (Figure S2, Supporting Information). These findings highlighted the safety and therapeutic potential of R1‐EVs.

Figure 3.

Effects of administration of R1‐EVs on survival rates of mice from radiation‐induced death. A) Timeline for in vivo radioprotection assays. B) Kaplan–Meier survival rates over 30 d after 8 Gy TBI (n = 10 for control, IR, and R1‐EVs + IR groups). C) Clinical score for radiation sickness after 8 Gy TBI. Data are shown as mean ± SD of three independent experiments. *p < 0.05 or ***p < 0.001 versus the IR group.

3.4. R1‐EVs Improve Hematopoietic Toxicity after 8 Gy TBI

The IR‐induced ROS generation and cell viability in BMCs were measured to assess the hematopoietic damage caused by H‐ARS. One hour after TBI, BMCs were harvested, and endogenous ROS levels were assessed using flow cytometry. In the IR group, the fluorescence intensity of BMCs indicated a higher production of intracellular ROS compared with that in the control group. Conversely, pretreatment with R1‐EVs (10 mg kg⁻¹) before TBI significantly reduced the generation of intracellular ROS (Figure  4A). Additionally, ROS damage was assessed by quantifying MDA levels, a byproduct of lipid peroxidation.^{[ 21 ]} One day post‐TBI, the R1‐EVs + IR group exhibited a 56.2% reduction in plasma MDA levels compared with the IR group (Figure 4B). Furthermore, BMCs and spleen cells isolated and quantified 1 d after TBI showed significant depletion in the IR group, with approximately 84.9% of femur BMCs and 82.9% of splenocytes being affected. In contrast, the R1‐EVs + IR group showed considerably lower depletion, with only 56% and 47.1% depletion, respectively (Figure 4C). The protective effect of R1‐EVs against radiation‐induced bone marrow and spleen damage was validated using histological analyses. The H&E staining of the bone marrow from the R1‐EVs + IR group showed substantially less destruction than that from the IR group, confirming the enhanced preservation of hematopoietic cellularity (Figure 4D). Additionally, as shown in Figure 4E, R1‐EVs + IR group presented the reduced inflammatory damage to the spleen, as indicated by the preservation of white pulp integrity compared to the IR group. Taken together, pretreatment with R1‐EVs may exert a protective effect against radiation‐induced hematopoietic damage.

Figure 4.

Effects of administration of R1‐EVs on suppressing the TBI‐induced hematopoietic ARS. Mice were pretreated with or without R1‐EVs before 8 Gy of TBI. A) Intracellular ROS levels in BMCs were measured 1 h after TBI using flow cytometry. B) Plasma MDA concentration 1 d after TBI. C) Total number of remaining BMCs in femur and splenocytes was measured 1 d following TBI. D) H&E staining of femur sectioned 3 d after TBI (scale bar = 200 µm). E) H&E staining of spleen sectioned 3 d after TBI (scale bar = 200 µm). White pulp (WP): region enclosed by dashed line, red pulp (RP): surrounding region. Data are shown as mean ± SD (n = 5 mice per group) of three independent experiments. ###p < 0.001 versus control group; ***p < 0.001 versus IR group.

3.5. R1‐EVs Alleviate TBI‐Induced Intestinal Structural Damage

Seven days post‐TBI, we assessed the colon lengths of mice treated with or without R1‐EVs to determine the protective effects of R1‐EVs on radiation‐induced gastrointestinal damage. The administration of R1‐EVs (10 mg kg⁻¹) effectively restored the colon length, reversing radiation‐induced shortening (Figure  5A). Radiation‐induced gastrointestinal syndrome, characterized by the erosion of GI tract integrity, can lead to severe outcomes, including fluid loss, uncontrolled diarrhea, and electrolyte imbalances.^{[ 22 ]} We assessed the integrity of the GI epithelium using an FITC‐dextran assay, which measures the permeability of the intestinal barrier. Three hours after gavage, FITC‐dextran levels in the blood were analyzed. The R1‐EVs + IR group showed a 40% reduction in FITC‐dextran uptake compared to the IR group (Figure 5B). Furthermore, the survival of Lgr5+‐expressing intestinal stem cells (ISCs), which are crucial for the regeneration of the GI tract following radiation exposure, was evaluated.^{[ 23 ]} Immunohistochemical staining of the large intestinal crypts revealed that although ISCs in the IR group were extensively damaged, those in the R1‐EVs + IR group demonstrated markedly improved survival rates (Figure 5C). Additionally, the number of Ki67+ proliferative zones in the R1‐EVs + IR group exceeded that in the IR group (Figure 5D). The histopathological examination of the colon via H&E staining further underscored the protective effects of R1‐EVs, showing significantly reduced damage in the R1‐EVs + IR group compared to that in the IR group (Figure 5E). Collectively, these findings confirm that R1‐EVs substantially mitigate the adverse effects of ionizing radiation on the colon.

Figure 5.

Effects of administration of R1‐EVs on promoting intestinal structure regeneration in mice after 8 Gy TBI. A) Colon tissues and length of mice in the control, IR, and R1‐EVs + IR groups 7 d after 8 Gy TBI (n = 5 mice per group). B) FITC‐dextran in blood serum to evaluate intestinal permeability of mice in the three groups was evaluated 5 d after radiation. C,D) Colon sections were analyzed using immunohistochemical methods. Representative immunostaining images showing the expressions of Lgr5⁺ (C, scale bar = 200 µm) and Ki67 (D, Scale bar = 100 µm) of the control, IR, and R1‐EVs + IR groups (n = 5 mice per group). Proportional quantization histogram of Lgr5⁺ and Ki67‐positive areas in each field. E) H&E staining of the cross‐sectioned colon 7 d after irradiation (scale bar = 500 µm). Data are shown as mean ± SD (n = 5 mice per group) of three independent experiments. ###P < 0.001 versus control group; ***P < 0.001 versus IR group.

3.6. R1‐EVs Induced the Production of SCFAs in TBI‐Induced ARS Mice

Radiation exposure disrupts the balance of the gut microbiota, leading to reduced diversity of intestinal bacteria and subsequent impairment of intestinal homeostasis. This imbalance significantly reduces the production of SCFAs such as acetic, propionic, and butyric acids, which are critical for maintaining gut health.^{[ 24 ]} Research has demonstrated that SCFAs can mitigate colitis by decreasing pro‐inflammatory cytokines and boosting anti‐inflammatory cytokines, which are essential for regulating radiation‐induced damage.^{[ 25 ]} To explore the potential radioprotective role of the enhanced SCFA production by R1‐EVs, we measured the levels of acetic, propionic, and butyric acids in fecal samples collected over 5 d post‐irradiation. The results showed that SCFA levels were significantly lower in the IR group than those in the control group. Conversely, the administration of R1‐EVs markedly increased the levels of acetic and butyric acids (Figure  6A–C). Furthermore, we employed principal component analysis to evaluate the effects of R1‐EVs on SCFA secretion in the control, IR, and R1‐EVs + IR groups (Figure 6D). The principal component analysis revealed distinct clustering patterns of SCFA production in fecal samples, demonstrating that R1‐EVs likely mitigate radiation‐induced damage by enhancing SCFA production.

Figure 6.

Effects of administration of R1‐EVs on production of short‐chain fatty acid. At 5 d after 8 Gy TBI, feces were harvested from each group to analyze the abundance of short‐chain fatty acids (SCFAs). A–C) The amount of SCFAs in feces (n = 5 mice per group). A) Acetic acids, B) propionic acids, and C) butyric acids. (D) Visualization of principal component analysis in the control, IR, and R1‐EVs + IR groups. Data are shown as mean ± SD of three independent experiments. ##P < 0.01 or ###P < 0.001 versus control group; ***P < 0.001 versus the IR group.

3.7. R1‐EVs Exhibit Immunoregulatory Properties against Radiation‐Induced Inflammation

Exposure to high doses of IR can severely affect various tissues and organs by triggering the cytokine cascades that modulate the immune system. In this context, proinflammatory cytokines are critical in the pathogenesis of radiation‐induced diseases, and the regulation of these cytokines is essential for survival.^{[ 26 ]} To explore the regulatory effects of R1‐EVs on radiation‐induced inflammation, we measured the serum cytokine levels 7 d post‐TBI. The IR group exhibited increased levels of IL‐6 and IL‐17A compared to those in the control group. Notably, IL‐6, known for its role in exacerbating radiation‐induced damage,^{[ 27 ]} was significantly elevated in the IR group but substantially reduced in the R1‐EVs + IR group, thus underscoring the protective role of R1‐EVs against inflammatory responses (Figure 7A). Tregs play a crucial role in maintaining immune homeostasis by modulating immune responses. They play a pivotal role in inducing and maintaining peripheral tolerance by suppressing the activation and function of effector T cells, thereby mitigating excessive immune reactions and inflammation.^{[ 28 ]} To assess the influence of R1‐EVs on T‐cell dynamics 7 d post‐TBI, splenocytes from various mouse groups were incubated for 4 h with or without PMA/ionomycin. Subsequently, we conducted flow cytometry to quantify Th1 (CD4+ IFN‐γ+), Th2 (CD4+ IL‐5+), and Th17 (CD4+ IL‐17A+) cell populations using a specified gating strategy. As shown in Figure 7B, pretreatment of R1‐EVs modestly decreased the levels of Th1, Th2, and Th17 cells compared to those in the IR group. Remarkably, R1‐EVs administration significantly increased the Treg levels (Foxp3⁺CD25⁺CD4⁺) compared to those in the IR group (Figure 7C). These results indicated that the R1‐EVs may enhance radioprotection by moderating inflammatory responses and facilitating Treg induction.

Figure 7.

Effects of administration of R1‐EVs on suppressing pro‐inflammatory cytokines and inducing immunoregulatory properties. At 7 d after 8 Gy TBI, serum or splenic T cells were isolated from each group. Cells were immunostained for L/D viability dye, anti‐CD3, anti‐CD4, anti‐CD25, anti‐IFN‐γ, anti‐IL‐5, anti‐IL‐17A, and anti‐Foxp3 mAbs using an intracellular staining protocol. A) Cytokine bead array CBA analysis of serum samples illustrating the levels of IL‐2, IL‐4, IL‐6, IL‐10, TNF‐α, IFN‐γ, and IL‐17A. B) Flow cytometry gating strategy and representative data for Th1 (CD3⁺CD4⁺IFN‐γ⁺ cells), Th2 (CD3⁺CD4⁺IL‐5⁺ cells), and Th17 (CD3⁺CD4⁺IL‐17A⁺ cells) populations. Quantitative data show the percentage of Th1, Th2, and Th17 cells. C) Flow cytometry gating strategy and representative data for regulatory T cells (Tregs, CD3⁺CD4⁺CD25⁺Foxp3⁺ cells). The figure shows the percentage and absolute number of Treg cells. Data are presented as means ± SD (n = 5 mice per group) and are based on a representative plot from two independent experiments. #P < 0.05 or ###P < 0.001 versus control group; *P < 0.05 or ***P < 0.001 versus the IR group.

4. Discussion

Studies on D. radiodurans have reported that its tolerance to radiation, desiccation, and oxidative stress results from a combination of physiological determinants and well‐regulated molecular mechanisms.^{[ 17 ]} Given the extreme radioresistance of D. radiodurans, we investigated the radioprotective effects of EVs isolated from D. radiodurans. The administration of R1‐EVs increases the survival rates by alleviating radiation‐induced injury in mice. The R1‐EVs protect BMCs and splenocytes by scavenging ROS. In addition, they alleviate gastrointestinal syndrome by maintaining intestinal integrity, stemness, and barrier function. These protective effects are closely related to the immunoregulatory properties of R1‐EVs, along with the increasing production of SCFAs, such as acetic, propionic, and butyric acids. To our knowledge, this is the first study to report that EVs derived from D. radiodurans exert substantial protective effects against TBI‐induced ARS in a mouse model.

ARS is a broad term that describes a range of specific injuries to organ systems based on their sensitivity to radiation doses of 1–15 Gy, of which the most immediate problems are bone marrow failure and gastrointestinal tract damage.^{[ 29 ]} We reported that 85% of the mice died from multi‐organ injuries induced by 8 Gy of TBI. In contrast, 85% of mice pretreated with R1‐EVs survived the same irradiation dose (Figure 3B). H‐ARS typically manifests with radiation doses exceeding 1–3 Gy. It is characterized by the rapid death of bone marrow‐derived hematopoietic stem and progenitor cells, which are highly radiosensitive due to their rapid division.^{[ 30 ]} Notably, the administration of R1‐EVs mitigated radiation‐induced H‐ARS by scavenging radiation‐induced ROS in the bone marrow and inhibiting cell death in the bone marrow and splenocytes (Figure 4). GI‐ARS typically manifests with radiation doses exceeding 4–12 Gy. Additionally, it is characterized by deterioration of villi and crypts containing ISCs, which is associated with a decline in the regenerative ability of cells and disruption of the intestinal epithelial barrier.^{[ 23 ]} The relentless renewal of intestinal crypt‐based stem cells renders the intestine in the gastrointestinal tract more vulnerable to radiation‐induced damage.^{[ 31 ]} If a specific number of ISCs expressing LGR5 can survive radiation exposure, they may repair damaged intestinal structures.^{[ 32 ]} Herein, pretreatment with R1‐EVs restored the integrity and function of the gastrointestinal tract after radiation‐induced damage. The R1‐EVs + IR group considerably protected LGR5+ ISCs, which are considered the most critical population for recovery after radiation‐induced intestinal damage, compared with the IR group. In addition, H&E and Ki67 staining results confirmed that the R1‐EVs + IR group had significantly alleviated radiation‐induced gastrointestinal toxicity compared to the IR group (Figure 5). These findings indicated that R1‐EVs are effective radioprotectants for preventing TBI‐induced ARS.

SCFAs such as acetic, propionic, and butyric acids are metabolites derived from the gut microbiota that play essential roles in maintaining intestinal homeostasis and promoting immune stability.^{[ 24} , ^{33 ]} These metabolites can enter systemic circulation directly and function as signaling molecules in peripheral tissues, thereby regulating inflammation and the immune system.^{[ 34 ]} These properties are pivotal for protection against radiation‐induced dysbiosis and damage.^{[ 35 ]} This study demonstrated that the TBI‐induced reduction in acetate, propionate, and butyrate levels was significantly increased by pretreatment with R1‐EVs (Figure 6). Acetate, known for its anti‐inflammatory capacity, increases Tregs and regulates immunity via Foxp3 acetylation.^{[ 36 ]} Propionate exerts protective effects against radiation damage, particularly by providing long‐term radioprotection, reducing H‐ and GI‐ARS levels, and decreasing proinflammatory responses in mice.^{[ 18 ]} Additionally, butyrate alleviates radiation‐induced intestinal damage and inflammation, restores microbial balance, and strengthens gut integrity via activation of the GPR41/43 pathway.^{[ 37 ]} Consequently, increased levels of SCFAs may be correlated with survival rates after radiation‐induced injury through their immunomodulatory effects and maintenance of gut homeostasis.^{[ 18} , ^{36 ]} Thus, enhancing SCFA production by modulating the gut microbiome could serve as an effective strategy for radioprotection, and the increased production of SCFAs by R1‐EVs may represent one of the mechanisms underlying the radioprotective roles of R1‐EVs.

Another important finding was that the administration of R1‐EVs exerted immunoregulatory effects on radiation‐induced systemic damage. Following the initial physicochemical and free radical events that occur immediately after radiation exposure, a prolonged inflammatory response is characterized by the excessive generation of ROS, cytokines, chemokines, and growth factors. This sustained inflammatory process leads to the accumulation of inflammatory infiltrates, thereby disrupting homeostasis and balance of the immune system.^{[ 38 ]} Consequently, strategies aimed at identifying agents that can modulate oxidative stress and inflammation hold promise in the fight against TBI‐induced ARS.^{[ 39 ]} Our results demonstrated that R1‐EVs effectively suppressed the pro‐inflammatory cytokine IL‐6, which is pivotal for exacerbating tissue damage and triggering the inflammatory cascade following radiation exposure^{[ 40 ]} (Figure 7A). This suppression is particularly significant given the central role of dysregulated inflammation in the pathogenesis of ARS, which contributes to tissue damage and systemic complications.^{[ 41 ]} By attenuating the inflammatory response, R1‐EVs underscore their potential as a therapeutic strategy to mitigate the adverse effects of radiation exposure. Furthermore, our findings revealed a notable increase in the number of Tregs, which are critical for maintaining immune system homeostasis by modulating excessive inflammatory responses^{[ 42 ]} (Figure 7B,C). The expansion of Tregs following treatment with R1‐EVs indicates a shift toward immune tolerance, potentially linked to reduced production of the pro‐inflammatory cytokine IL‐6. This relationship is crucial for preserving tissue integrity and function following radiation exposure.^{[ 43 ]} Additionally, the dual regulatory actions of R1‐EVs, both suppressing IL‐6 and inducing Tregs, were significantly correlated with enhanced SCFA production, as shown in Figure 6. These findings highlight the synergistic effects of SCFAs, which are known to boost the production of the anti‐inflammatory cytokine IL‐10, reduce pro‐inflammatory cytokine expression, and maintain Treg balance, further supporting the immunoregulatory potential of R1‐EVs.^{[ 44 ]} These findings highlight the critical role of R1‐EVs in modulating immune responses, an effect that is significantly mediated by SCFA production. Despite the intricate connection between SCFA production and immunoregulation observed in our study, further investigation into SCFA‐mediated pathways as potential targets for radioprotective therapies is not only warranted but also essential. This study paves the way for developing innovative treatments that leverage the immunoregulatory capacities of R1‐EVs and SCFAs to mitigate the effects of ARS.

High doses of ionizing radiation induce rapid generation of ROS in tissues via radiolysis of water, causing both acute and chronic oxidative damage to cells and tissues.^{[ 2 ]} Thus, R1‐EVs can scavenge rapidly generated O₂ ⁻, H₂O₂, and OH^• (Figure 1) and protect cells and organs against oxidative injuries. The radioprotective effect of R1‐EVs can be attributed to their powerful ROS scavenger capacity, facilitated by both antioxidant molecules obtained from the parental bacterial cells and enzymes that detoxify harmful ROS into unharmful substances. For instance, deinoxanthin, a unique carotenoid synthesized by D. radiodurans, has a characteristic red color^{[ 45 ]} and exerts a strong antioxidant effect owing to its distinct chemical structure comprising extended conjugated double bonds and the presence of a hydroxyl group at the C‐1′ position.^{[ 46 ]} Several studies have demonstrated that the S‐layer deinoxanthin‐binding complex is resistant to UV radiation and thermostable, thereby playing a protective role in D. radiodurans.^{[ 47 ]} Interestingly, the R1‐EVs exhibited a red color, indicating the presence of deinoxanthin within their membranes (Figure S3, Supporting Information). Additionally, the structural organization of D. radiodurans cell envelope, including the S‐layer, plays an important role in protecting D. radiodurans against oxidative stress.^{[ 48 ]} Functional over‐representation analysis showed that proteins belonging to the cell envelope were significantly over‐represented in the R1‐EV proteome (Figure 2). This was also supported by mass spectrometry proteomic data, which identified the S‐layer protein (accession: ANC71643.1) as having the greatest number of peptide spectral matches (PSMs) among all identified proteins in the R1‐EVs proteome (APPENDIX). Three other proteins that are directly involved in the cell envelope structure, particularly the S‐layer organization, were also found to be among the top 10 proteins with the highest number of peptide spectral matches. These included the outer membrane protein assembly factor‐accession (QEM72334.1; 110 PSMs), another S‐layer protein‐accession (ANC70400.1; 79 PSMs), and the hexagonally packed intermediate‐layer surface protein‐protein accession (WP_010889133.1; 58 PSMs). Our data also revealed that the R1‐EV proteome was significantly enriched in enzymes that respond to oxidative stress, with the response to hydrogen peroxide being the most significantly over‐represented biological process carried out by the identified proteins (Figure 2). D. radiodurans genome encodes various CAT proteins, including katA and DR_A0259, both of which were identified in the R1‐EV proteome (APPENDIX). CAT detoxifies H₂O₂ by facilitating its decomposition into H₂O and O₂, thereby preventing the Fenton reaction and which would otherwise lead to the formation of highly reactive OH^• radicals.^{[ 49 ]} Thus, R1‐EVs’ protective effect against radiation induced oxidative damage can be attributed to the presence of antioxidant molecules such as deinoxanthin and a proteome enriched with proteins that protect against ROS species.

Although our study provides compelling evidence for the radioprotective effects of EVs derived from R1‐EVs, there were several limitations that warrant further investigation. A primary limitation was the lack of detailed identification of the bioactive molecules within R1‐EVs that mediate their protective effects. Although we demonstrated the enrichment of proteins associated with oxidative stress response, the specific molecular entities responsible for these effects remain unclear. To address this gap, future research should employ a comprehensive multi‐omics approach that includes lipidomics, metabolomics, and transcriptomics. This approach will facilitate a thorough characterization of R1‐EVs, enabling the identification of specific lipids, metabolites, RNAs, and proteins that contribute to their radioprotective properties. Another limitation was that we could not completely elucidate the systemic mechanisms through which R1‐EVs confer radioprotection. To overcome this limitation, future studies should focus on transcriptomic analyses in TBI mouse models. By comparing mice pretreated with R1‐EVs to untreated controls, these analyses can reveal host‐level changes and pathways activated by R1‐EVs, providing insights into their role in mitigating radiation‐induced damage.

5. Conclusion

In conclusion, this study demonstrated that R1‐EVs play a critical role in protecting against radiation‐induced H‐ and GI‐ARS through multiple mechanisms. Proteomic analysis of the protein profiles of R1‐EVs highlighted the significant increase in SCFAs production. Elucidating the immunoregulatory roles of R1‐EVs provided robust evidence for the radioprotective capacities of R1‐EVs (Figure 8 ). Moreover, the ability of R1‐EVs to modulate oxidative stress and enhance cellular repair mechanisms highlighted their potential as effective therapeutic agents. These findings suggest that R1‐EVs could be developed into effective countermeasures not only for radiation‐induced injuries, but also for other ROS‐mediated diseases. Future research should also explore the potential applications of R1‐EVs in combination with other therapeutic agents and as delivery vehicles for radioprotective compounds. These strategies could enhance their efficacy and broaden their applicability, offering promising avenues for advancing radioprotection strategies.

Figure 8.

Schematic diagram of the protective role of R1‐EVs on total‐body irradiation (TBI)‐induced acute radiation syndrome (ARS) in mice. This schematic illustrates the radioprotective mechanisms of Deinococcus radiodurans‐derived extracellular vesicles (R1‐EVs) against TBI‐induced ARS in mice. R1‐EVs mitigate the effects of ionizing radiation by scavenging reactive oxygen species (ROS) generated through water radiolysis, thereby protecting tissues from oxidative damage. The diagram highlights the ROS scavenging efficacy of R1‐EVs, specifically against superoxide, hydrogen peroxide, and hydroxyl radicals. Additionally, R1‐EVs enhance the production of short‐chain fatty acids (SCFAs) and regulatory T cells (Tregs), leading to the regeneration of intestinal structure and hematopoietic function. This figure underscores the potential therapeutic application of R1‐EVs in managing radiation‐induced injuries by showcasing their multifaceted protective effects in both gastrointestinal (GI‐ARS) and hematopoietic (H‐ARS) ARS. The comprehensive antioxidant properties and immunomodulatory capabilities of R1‐EVs demonstrate their potential as effective radioprotective agents. The illustration was created using Biorender (BioRender.com).

Conflict of Interest

The authors declare no conflict of interest.

Supporting information

Supporting Information

Supporting Information

Han J. M., Mwiti G., Yeom S.‐J., Lim J., Kim W. S., Lim S., Lim S.‐T., Byun E.‐B., Radiation‐Resistant Bacteria Deinococcus radiodurans‐Derived Extracellular Vesicles as Potential Radioprotectors. Adv. Healthcare Mater. 2025, 14, 2403192. 10.1002/adhm.202403192

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.