Transplantation of Donor–Recipient Chimeric Cells Restores Peripheral Blood Cell Populations and Increases Survival after Total Body Irradiation-Induced Injury in a Rat Experimental Model

This study received approval from the Animal Care Committee (ACC Number: 19-167) of the University of Illinois at Chicago, accredited by the American Association for the Accreditation of Laboratory Animal Care. All animals received humane care in compliance with the “Principles of Laboratory Animal Care” formulated by the National Society for Medical Research and the “Guide for the Care and Use of Laboratory Animal Resources” published by the US National Institutes of Health. In this experimental study, a total of 44 male Lewis rats (RT11) (strain: LEW, Research Resource Identifiers Project (RRID): Rat Genome Database (RGD)_737932) and 48 ACI rats (RT1a) (strain: ACI, RRID: RGD_737892), 8–10 weeks of age, purchased from the Charles River Laboratories (Chicago, IL, USA), were used. The animals were housed in an accredited animal facility at the University of Illinois at Chicago, where they were handled humanely according to US Public Health Service Policy. Animals were individually caged at room temperature on a 12-h day/night cycle and provided with standard rodent food and water ad libitum.

In total, 20 male Lewis recipient rats, aged 8–10 weeks, were irradiated with a 7 Gy nonlethal dose of TBI administered at a rate of 0.4–1.0 Gy/min. The ionizing irradiation was evenly delivered using Cs-137 radionuclide via a J.L. Shepherd Irradiator, Model 143-68, ensuring a uniform dose distribution. The irradiation procedure was performed by an experienced technologist at the University of Illinois at Chicago, USA, according to the standard procedure, which required calibration of the instrument prior to irradiation and according to the established doses approved in the protocol. Throughout the irradiation procedure, rats were housed in a specially adapted metal cylinder designed for this purpose (Figure 1). Following irradiation exposure, rats were individually placed in cages with free access to antibiotic-enriched water (trimethoprim–sulfamethoxazole, 40 mg/200 mg/5 mL, Hi-Tech Pharmacal, Amityville, NY, USA) and provided with softened rodent food. Before the application of cellular therapies, rats were anesthetized with 1.5%–2.5% isoflurane inhalation, and blood samples were collected to confirm the 7 Gy TBI-induced hematopoietic injury and development of ARS.

Fig 1.

In total, 24 male fully major histocompatibility complex (MHC) mismatched ACI rats, aged 8–10 weeks, served as allogeneic bone marrow transplant (alloBMT) donors. Additionally, 24 male ACI rats and 24 male Lewis rats, each aged 8–10 weeks, served as BM donors to generate DRCC. The cellular therapies were transplanted via intraosseous injection into the right femur 6 h after the irradiation procedure. Twenty Lewis rat recipients were randomly divided into four experimental groups (n = 5/group) and received: Group 1 control – received 0.1 mL of saline and antibiotic therapy (trimethoprim–sulfamethoxazole, 40 mg/200 mg/5 mL, Hi-Tech Pharmacal); Group 2 – was supported with alloBMT (80 × 10⁶ cells) from the ACI donors; Group 3 – received DRCC (4–6 × 10⁶ cells) therapy created by the fusion of BM cells from ACI and Lewis donors; Group 4 – received alloBMT (80 × 10⁶ cells) from the ACI donors combined with DRCC (4–6 × 10⁶ cells) derived from the fusion of BM cells from ACI and Lewis donors. Summary of experimental groups is presented in Table 1.

Twenty-four male fully MHC mismatched ACI rats, aged 8–10 weeks, were anesthetized with 1.5%–2.5% isoflurane inhalation. After induction of anesthesia, rats were euthanized with a SomnaSol solution (sodium pentobarbital 390 mg + sodium phenytoin 5 mg/mL, Henry Schein Animal Health, Melville, NY, USA) by intraperitoneal injection, with bilateral thoracotomy as an assuring method. The donors’ legs were shaved and washed with iodine solution. Both femurs and tibias were harvested under sterile conditions, then suspended in a sterile phosphate buffered saline (PBS) solution, and transferred to a sterile laminar chamber. The bones were dissected, and the intramedullary cavities were rinsed using a 3 mL syringe and an 18 G needle (305195, BD^® Needle, Becton Dickinson, Franklin Lakes, NJ, USA). The bones were washed with a Dulbecco’s Modified Eagle’s Medium (DMEM, Thermo Fisher Scientific, Waltham, MA, USA) solution in the amount of 20 mL. In the next step, the cell suspension was filtered through a sterile nylon filter (pore size 40 μm) and subjected to a further purification process. Cell viability and abundance were assessed using a microscope with a Vi-CELL^™ XR Cell Viability Analyzer (Beckman Coulter, Brea, CA, USA) after the cells were stained with 0.4% Trypan Blue solution (Thermo Fisher Scientific) and plotted on a microscope slide. The isolated donor BM cells were then labeled with the fluorescent red dye Paul Karl Horan (PKH) 26GL (Sigma-Aldrich, St. Louis, MO, USA), according to the manufacturer’s instructions. The isolated and labeled ACI rat BM cells were then resuspended in 100 μL of PBS staining buffer containing 1% bovine serum albumin (BSA).

The DRCC were created from BM cells derived from ACI and Lewis rat donors, as depicted in Figure 2a. The isolation of BM cells, creation of DRCC through the ex vivo cell fusion process, and evaluation of DRCC properties were thoroughly described in previous studies (Cwykiel et al. 2021a, b). Briefly, BM cells were isolated from the tibia harvested from randomly selected ACI (RT1^a) and Lewis (RT1¹) rat donors in a sterile manner and subsequently fluorescently stained using PKH26-red (ACI) or PKH67-green (Lewis) membrane dyes (Sigma-Aldrich), respectively, according to manufacturer’s instructions. Next, donor cells were mixed in a 1:1 ratio and suspended in serum-free Roswell Park Memorial Institute (RPMI) 1640 medium (Thermo Fisher Scientific). The cell fusion procedure was performed using a 1.46 g/mL polyethylene glycol (PEG) 4000 solution (EMD, Burlington, MA, USA) containing 16% DMSO (Sigma-Aldrich), as previously reported (Cwykiel and Siemionow 2014; Siemionow et al. 2015, 2018a). Fluorescence-activated cell sorting (FACS, BD FACSAria^™ II cell sorter, Becton Dickinson) was applied to select the PKH26/PKH67-labeled cells, representing the DRCC population.

Fig 2.

The creation of DRCC via the ex vivo PEG-mediated cell fusion procedure of rat BM cells was assessed by FC and CM. Samples of unstained BM cells, the PKH26- and PKH67-stained BM cells, as well as fused PKH26/PKH67-stained DRCC were assessed using LSRFortesa^™ cell cytometer (BD Biosciences, San Jose, CA, USA). PKH26- and PKH67-stained BM cells, and fused PKH26/PKH67-stained DRCC were fixed in 4% paraformaldehyde (EMS, Hatfield, PA, USA) and mounted with VECTASHIELD^® Antifade Mounting Medium with DAPI (Vector Laboratories, Burlingame, CA, USA). Images were taken by upright confocal microscope (Leica TCS-SP2, RRID: SciCrunch Registry (SCR)_020231, Leica Microsystems, Wetzlar, Germany) with a digital camera (QImaging^® Retiga-2000R Charge Coupled Device, QImaging, British Columbia, Canada) and ImagePro Plus software (RRID: SCR_016879, Media Cybernetics, Rockville, MD, USA).

BM cells and DRCC were injected into the right femoral bone of Lewis rats, as previously reported (Siemionow et al. 2005a, b). The procedure was conducted under sterile conditions. While under general anesthesia with 1.5%–2.5% isoflurane inhalation, a 1 cm-long incision was made on the dorsal surface of the right mid-thigh of Lewis rats. Subsequently, the skin, subcutaneous tissue, and surrounding muscles were dissected, exposing the femur. Using a 0.5 mL syringe equipped with a 27 G needle (305109, BD^® Needle, Becton Dickinson), a hole was created in the femoral bone, and 0.1 mL of the recipient’s marrow was aspirated to prevent excessive pressure during cell injection. Then, using a 0.5 mL syringe with a 27 G needle, saline solution as control, alloBMT, or DRCC suspended in sterile saline solution, or a combination therapy comprising alloBMT and DRCC suspension was injected into the femur through a pre-made hole, depending on the belonging to the experimental group. Following the intraosseous injection, the bone opening was sealed with bone wax (Medeline Industries, Mundelein, IL, USA) to prevent cell leakage from the marrow cavity. The muscles and skin were sutured using 4–0 and 5–0 absorbable sutures (Ethicon Inc., Raritan, NJ, USA), respectively. Post-operatively, rats recovered in a heated environment and received necessary care before returning to the colony.

The rats underwent daily observations to assess their survival rates over the entire 90-day observation period. The baseline body mass of each animal was recorded at the beginning of the study and was then measured daily during the initial 8 days and subsequently, every other day throughout the entire 90-day follow-up period. To address the weight differences between animals assigned to the four experimental groups at the study beginning before irradiation, adjustments were done using Microsoft Excel (ver.16.0, Reman, WA, USA) software. Therefore, the initial weight on day 0 was set as a baseline, denoted as 100%, and the subsequent weights were then calculated as the percentage relative to this baseline value. These percentage values were calculated in 10-day intervals until the end of the 90-day observation period.

Peripheral blood samples (0.3 mL) were obtained from the jugular vein on day 0 as a baseline reference sample, prior to TBI and the administration of cell therapies. Subsequent samples were collected on days 5, 10, 20, 30, 40, 60, and 90 following TBI for further analysis. During the procedure, rats were anesthetized with 1.5%–2.5% isoflurane inhalation. Blood samples were collected in ethylenediaminetetraacetate collection tubes (BD Microtainer^®, Becton Dickinson) and immediately processed by the accredited Biologic Resources Laboratory at the University of Illinois at Chicago, for complete blood counts, including white blood cells (WBC), WBC differentials (neutrophils, lymphocytes, monocytes, eosinophils, and basophils), red blood cells (RBC), and platelets (PLT). At the 90-day study end-point, rats were euthanized using SomnaSol solution (sodium pentobarbital 390 mg + sodium phenytoin 5 mg/mL, Henry Schein Animal Health) by intraperitoneal injection, followed by the bilateral thoracotomy as an assuring method.

After the intraosseous transplantation of BM cells and DRCC therapies, animals were observed for signs of morbidity and symptoms of GvHD, daily for the first 8 days, and subsequently, every other day over the entire follow-up period of 90 days. The animals had free access to antibiotic-enriched water (trimethoprim–sulfamethoxazole, 40 mg/200 mg/5 mL, Hi-Tech Pharmacal) and softened rodent food. A summary of the assessed clinical parameters and total scores is presented in Table 2. The analysis included implementation of scores for activity in the cage, body posture, fur coverage, presence of blood in the stool, and diarrhea incidence for each individual animal. These scores were then summarized and divided by the total number of observations. Activity in the cage, body posture, and fur coverage were assessed on a scale ranging from 0 to 3 points, where 0 = a normal response and 3 = the most pronounced abnormality. Incidences of blood in stool and diarrhea were evaluated on a scale from 0 to 1, where 0 represented the absence of blood in stool or normal stool consistency, and 1 indicated the presence of blood in stool or diarrhea. All rats underwent evaluation using two total score systems. The first score system, ranging from 0 to 9 points, assessed the overall physical condition, which included parameters such as activity in the cage, body posture, and fur coverage. The second score system, ranging from 0 to 2 points, focused on the digestive distress conditions, considering the presence of blood in the stool and the incidence of diarrhea.

The histological assessment of GvHD included examination of the kidney, skin, and small intestine biopsies. Tissue biopsies were collected on the 90th day and placed in a formalin solution for 48 h. Subsequently, tissue samples were fixed in a 70% ethanol solution and embedded in paraffin blocks. These paraffin blocks were then sliced into 0.5 mm thick sections using a microtome (2088 Ultrotome^® V Microtome, LKB Bromma, Amersham, UK) and affixed to microscope slides. The preparations included Hematoxylin and Eosin staining according to a standard protocol. The histopathological evaluation of GvHD changes was performed by the use of a BX51/IX70 inverted microscope (Olympus, Tokyo, Japan). The presence of histopathological changes in the course of GvHD was assessed in accordance with the Small Criteria presented in Table 3. The criteria assessing the degree of GvHD occurrence, based on the examined organ, are outlined in Table 4. Each tissue biopsy was assigned a numerical score on a scale from 0 to 3 points, depending on the changes suggesting the GvHD occurrence, where the value 0 = no GvHD, the value 1 = GvHD possible, the value 2 = consistent with GvHD, and the value 3 = definite GvHD (Shulman et al. 2015). The closer the score was to zero, the less likely the assessed sample exhibited signs of GvHD.

Data are expressed as the mean ± standard error of the mean (SEM). GraphPad Prism (ver.9.2.1, RRID: SCR_002798, Dotmatics, Boston, MA, USA) software and Microsoft Excel (ver.16.0, Microsoft Corporation) were used to perform statistical analysis. Two-way analysis of variance (ANOVA) for group comparison was used to define statistical significance. Results were considered statistically significant for p < 0.05. The graphs represent mean values with SEM, statistical significance is marked with asterisks: *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

The study design of the ex vivo PEG-mediated cell fusion procedure of the rat BM cells derived from ACI and Lewis donors is presented in Figure 2a. The DRCC population demonstrated >95% purity and the efficacy of the donor and recipient cell fusion was assessed at 32.2%, as confirmed by FC analysis (Figure 2b). The efficacy of the fusion procedure and the quality of the created chimeric cells were further confirmed by CM, revealing the presence of overlapping PKH26/PKH67 fluorescent staining confirming the chimeric state of the created DRCC (Figure 2c). The presented data confirmed the successful creation of DRCC from ACI and Lewis BM cells by FC and CM analyses.

Following 7 Gy TBI, there was a 60% mortality rate in the group receiving alloBMT therapy, whereas a 100% survival rate was observed in the other therapy groups and the control group (Figure 3). Animals that did not survive until the end of the observation period displayed a gradual decline in activity, pallor, changes in fur condition, a slumped body posture, and the presence of blood in the stool. Rat mortality occurred naturally on days 10, 11, and 12 following TBI and alloBMT administration; therefore, no euthanasia procedures were performed in these cases.

Fig 3.

The changes in rats’ body weight over a 90-day observation period following 7 Gy TBI and administration of different cellular therapies are presented in Figure 4. There was a decrease in body weight from across all experimental groups from the beginning of the observation period until the 5th day, followed by a subsequent increase in body weight during the remaining observation period up to 90 days following irradiation exposure. Additionally, at the 30-day observation, a decrease in body weight was observed in the DRCC therapy group, which was subsequently regained in the following weeks. In summary, aside from the initial post-irradiation drop in body weights, animals across all experimental groups regained body weight, surpassing the initial body weight values. This recovery was evidenced by body weight changes exceeding 100%. The regained weight was then maintained throughout the 90-day observation period.

Fig 4.

The summary of the peripheral blood cells populations changes observed in the WBC, lymphocytes, monocytes, neutrophils, basophils, and eosinophils as well as in RBC and PLT values over the 90-day observation period following 7 Gy TBI and the administration of different cellular therapies is presented in Figure 5. The baseline values represent measurements taken before TBI and the administration of cellular therapies and served as a reference for comparative analysis.

Fig 5.

On day 0 before TBI and application of cell therapies, assessment of WBC counts revealed higher baseline WBC values in the DRCC therapy group when compared to the control group (8.116 ± 0.179 vs. 6.190 ± 0.888, respectively, p < 0.0001). Similarly, higher WBC counts were observed in the alloBMT group compared to the control group (7.864 ± 0.582 vs. 6.190 ± 0.888, respectively; p < 0.001). Moreover, higher WBC counts were also seen in the alloBMT + DRCC therapy group when compared to the control group (7.383 ± 0.229 vs. 6.190 ± 0.888, respectively; p < 0.001) .

On day 5 after TBI and application of cell therapies, the DRCC therapy group exhibited almost a three-fold increase in WBC counts compared to the alloBMT group (3.428 ± 0.402 vs. 1.238 ± 0.110, respectively; p < 0.0001). In addition, increased WBC counts were observed in the DRCC group compared to the control group (3.428 ± 0.402 vs. 2.363 ± 0.146; respectively,p < 0.05). Furthermore, a three-fold increase in WBC counts was observed in alloBMT + DRCC compared to alloBMT therapy (3.073 ± 0.421 vs. 1.238 ± 0.110, respectively;p < 0.0001). Finally, the WBC counts recorded in the control group were almost twice as high as those in the alloBMT therapy group (2.363 ± 0.146 vs. 1.238 ± 0.0110, respectively; p < 0.05) (Figure 5a).

On day 0 before TBI and application of cell therapies, lymphocyte baseline values in the alloBMT group were significantly higher compared to the control group (6.609 ± 0.467 vs. 5.460 ± 0.746, respectively; p < 0.01) (Figure 5b).

On day 5 after TBI and application of cell therapies, lymphocyte counts showed a six-fold increase following administration of DRCC therapy when compared to the alloBMT (1.148 ± 0.031 vs. 0.185 ± 0.12; respectively, p < 0.05). Similarly, the lymphocyte counts in the alloBMT + DRCC therapy group exhibited a six-fold increase compared to the alloBMT group (1.108 ± 0.047 vs. 0.185 ± 0.121, respectively; p < 0.05).

On day 0 before TBI and application of cell therapies, the only notable increase in monocyte counts was observed in the alloBMT + DRCC group when compared to the DRCC therapy group, relative to the baseline values (0.169 ± 0.021 vs. 0.110 ± 0.019, respectively; p < 0.0001) (Figure 5c). No significant differences were observed in neutrophil or eosinophil counts among the administered cellular therapies (Figures 5d and 5e).

On day 0 before TBI and application of cell therapies, the assessment of baseline basophil counts revealed significantly higher values in the alloBMT + DRCC group when compared to the control group (0.064 ± 0.012 vs. 0.006 ± 0.004, respectively; p < 0.0001) (Figure 5f). Following TBI, the monocyte counts revealed initial decline between days 5 and 10, followed by a gradual increase from day 20 onward throughout the remaining observation period, reaching the highest values in the DRCC therapy group on days 30, 40, 60, and 90.

On day 0 before TBI and application of cell therapies, the mean baseline RBC values in the alloBMT therapy group were higher compared to the control group (8.449 ± 0.120 vs. 6.709 ± 1.200, respectively; p < 0.05) (Figure 5g).

On day 5 after TBI and application of cell therapies, the RBC values were higher in the alloBMT compared to the alloBMT + DRCC therapy group (8.770 ± 0.146 vs. 6.973 ± 0.291, respectively; p < 0.05).

On day 10 after TBI and application of cell therapies, there was a 1.5-fold increase in RBC values recorded in the control group compared to the alloBMT + DRCC group (8.038 ± 0.808 vs. 5.423 ± 0.175, respectively; p < 0.001).

On day 20 after TBI and application of cell therapies, a 1.3-fold increase in RBC values was observed in the DRCC group compared to the alloBMT + DRCC group (7.086 ± 0.302 vs. 5.400 ± 0.920, respectively; p < 0.05) (Figure 5g).

On day 5 after TBI and application of cell therapies, PLT counts analysis revealed two-fold increase in the DRCC therapy group compared to the alloBMT + DRCC group (586.0 ± 59.093 vs. 287.75 ± 18.759, respectively; p < 0.0001). Also, PLT counts were significantly higher in the DRCC group compared to the control group (586.0 ± 59.093 vs. 395.50 ± 62.8, respectively; p < 0.05) and in the alloBMT group, compared to the alloBMT + DRCC therapy group (494.0 ± 149.346 vs. 287.7 ± 18.75, respectively; p < 0.01) (Figure 5h).

On day 40 after TBI and application of cell therapies, a 1.4- fold increase in the PLT values was recorded in the DRCC therapy group when compared to the alloBMT + DRCC group (650.5 ± 21.683 vs. 432.5 ± 12.906, respectively; p < 0.01) (Figure 5h).

A summary of the assessed clinical parameters is presented in Table 2. The analysis included the assessment of scores for the animals’ activity in the cage, the presence of blood in the stool, the body posture, incidence of diarrhea, and the fur coverage for each individual animal, and the scores were summarized and divided by the total number of observations (Table 5). Activity in the cage, the body posture, and fur coverage were assessed on a scale ranging from 0 to 3 points, where 0 = a normal response and 3 = the most pronounced abnormality. Presence of blood in the stool and the incidence of diarrhea were evaluated on a scale from 0 to 1, where 0 = the absence of blood in stool or normal stool consistency, and 1 = the presence of blood in stool or diarrhea. The closer the score was to zero, the more closely the assessed feature aligned with the normal clinical conditions. Subsequently, all rats were evaluated using two total score systems. The first score system, ranging from 0 to 9 points, assessed the overall physical condition, which included parameters such as activity in the cage, body posture, and fur coverage. The second score system, ranging from 0 to 2 points, focused on digestive distress conditions, considering the presence of blood in the stool and incidence of diarrhea. The poorest activity was recorded in the control saline group (0.96 ± 0.453) and the alloBMT + DRCC therapy group (0.29 ± 0.302) (Table 5). The activity closest to normal was observed in the alloBMT group (0.13 ± 0.031). The difference between the alloBMT and the control group was statistically significant (0.13 ± 0.031 vs. 0.96 ± 0.453, respectively; p < 0.05). The body posture exhibiting minimal changes was observed in the alloBMT + DRCC (0.00 ± 0.000) and DRCC therapy (0.02 ± 0.027) groups, whereas the most significant changes in body posture were seen in animals receiving alloBMT (0.34 ± 0.227). The differences between the alloBMT and the alloBMT + DRCC therapy group were statistically significant (0.34 ± 0.227 vs. 0.00 ± 0.000, respectively; p < 0.05). Assessment of fur coverage in the DRCC therapy group revealed scores close to normal fur coverage (0.51 ± 0.118). Similar trend in fur coverage was observed in animals receiving alloBMT + DRCC therapy (0.60 ± 0.094). In contrast, animals supported with the alloBMT showed the most advanced changes in fur coverage (1.15 ± 0.273). There was a significant difference in the fur coverage score in the DRCC compared to the alloBMT group (0.51 ± 0.118 νs. 1.15 ± 0.27, respectively; p < 0.05). The presence of blood in the stool was notably more frequent in rats receiving alloBMT compared to the alloBMT + DRCC therapy (0.19 ± 0.194 vs. 0.07 ± 0.023, respectively; p < 0.05). The most frequent occurrence of diarrhea was observed in the control (0.16 ± 0.118) and the alloBMT + DRCC therapy (0.08 ± 0.027) groups, and least frequently among animals receiving DRCC therapy (0 ± 0.009). There were significant differences in the frequency of diarrhea occurrence between the control and DRCC therapy group (0.16 ± 0.118 vs. 0 ± 0.009, respectively; p < 0.05). Overall, the clinical changes closest to normal were observed in the DRCC therapy group, with an overall physical condition total score of 0.78 and a digestive distress condition total score of 0.06. In the remaining experimental groups, mild changes were observed, with the most pronounced changes noted in the control group, which exhibited an overall physical condition total score of 1.68 and a digestive distress condition total score of 0.34. Following closely, the alloBMT group displayed total scores of 1.62 and 0.28 for overall physical condition and digestive condition, respectively. In comparison, the alloBMT + DRCC group demonstrated lower total scores, with values of 0.89 for overall physical condition and 0.15 for digestive distress condition.

Comparisons of histopathological changes assessed in the tissue biopsies collected from kidney, skin, and small intestine suggesting presence of GvHD are presented in Figure 6a. For histological assessment, each tissue sample was assigned a numerical score on a scale from 0 to 3 points, depending on the changes suggesting the occurrence of GvHD, where score 0 = no GvHD, score 1 = GvHD possible, score 2 = consistent with GvHD, and score 3 = definite GvHD (Shulman et al. 2015). The closer the score was to zero, the less likely the assessed sample exhibited signs of GvHD. The criteria assessing the degree of the GvHD occurrence, based on the examined organ, are outlined in Table 4. The lowest score suggesting the occurrence of GvHD on kidney tissue histological slides was noted in animals receiving DRCC therapy (0.2 ± 0.447), while the highest score was found in the control group (0.6 ± 0.548) (Figure 6b). Based on the histological assessment of skin slides, the highest score for GvHD occurrence was found in animals receiving alloBMT (0.6 ± 0.894), and the lowest score was found in the DRCC therapy group (0.0 ± 0.000). Additionally, the highest score indicating the possible presence of GvHD on the histological slides of small intestine tissue was observed in animals subjected to alloBMT (1.4 ± 0.548) and alloBMT + DRCC therapy (1.4 ± 0.548), whereas the lowest score was found in the control group (0.6 ± 0.548). A similar score indicating GvHD occurrence in small intestine slides was observed in the DRCC therapy group (0.7 ± 0.645). Overall, no significant histological changes indicative of GvHD were observed in the tissue samples of the kidney, skin, and small intestine in the DRCC group. In contrast, changes indicative of GvHD were present in all three tissue biopsies in the alloBMT group, including the skin. Although the scores in kidney and small intestine samples were higher in the alloBMT group when compared to the DRCC group, the differences were not significant, but the DRCC therapy reduced signs indicative of GvHD.

Fig 6.