Peripheral blood was obtained from volunteers from The Department of Urology of the Third Affiliated Hospital of Soochow University. Human naïve CD4⁺ T cells (CD4⁺CD25^–CD45RA⁺) were isolated and purified from peripheral blood mononuclear cells (PBMCs) using Ficoll-Hypaque separation (Amersham Biosciences) and the two-step magnetic bead separation method of an autoMACS Pro Separator (Miltenyi Biotechnology Company, Germany).

Naïve CD4⁺ T cells were induced to CD4⁺ iTregs with TGF-β (5 ng/ml) (Bio-techne, Abingdon, OX, United Kingdom), recombinant IL-2 (100 IU/ml) (Chiron, Emeryville, CA, United States), and antihuman CD3/CD28 beads (ratio of cells to beads = 1:5) (Life Technology, Carlsbad, CA, United States) in X-Vivo-15 medium (BioWhittaker, Walkersville, MD, United States) containing 10% fetal bovine serum (Cereal Biomedicine) for 6 days. The BA (100 μmol/L) used in cell culture was prepared by dissolving BA sodium powder (Sigma-Aldrich, 303410) in dimethyl sulfoxide (DMSO).

The PC61 α-CD25 antibody (Biolegend Cat#102007) was generated from hybridoma cells (ATCC) in serum-free media (Gibco).

We constructed the renal IRI model according to the conventional method (Hosszu et al., 2017). Male C57BL/6 mice (20–25 g) 7–8 weeks of age were purchased from Kawensi Biotech (Changzhou, China). We established the mouse model of renal IRI by clamping the bilateral renal pedicles for 45 min. A warming pad was used to keep body temperature around 37°C during the surgery. After 45 min of ischemia, the needle was withdrawn to allow reperfusion for 1 day. Similar surgical procedures except clamping of the renal pedicles were applied to the sham-operated group. The animal experiments were approved by the Animal Management and Use Committee of Soochow University.

Mice were given 100 mM BA in their drinking water. The renal IRI model was established after 6 days. Mice were sacrificed on post-operative day 1. Blood samples were collected from the tail vein to examine creatinine (Cr) and blood urea nitrogen (BUN) levels. Spleens and kidneys from the rats were obtained and fixed in 10% neutral formalin for 48–72 h. The specimens were dehydrated by a graded ethanol series and embedded in paraffin. Kidney tissue sections were subjected to hematoxylin-eosin (HE) staining, and the kidney tubule injury was assessed using the Paller score. Ten non-overlapping visual fields (200× microscope) were randomly selected from each rat and observed under a light microscope. For each field of view, 10 kidney tubules were randomly chosen for quantification. A total of 100 renal tubules were scored, and the higher the score was, the more serious the tubular injury was Paller et al. (1984).

We tested the expression of IL-17A, IFN-γ, and IL-10 mRNA in tubules of mice. Total RNA extracted from tissue and cell lysates was further purified using RNeasy Mini Kit (Qiagen, Valencia, CA, United States). We obtained cDNA using Omniscript RT kit (Qiagen, Netherlands). Relative quantification of mRNA expression levels was performed using Absolute QPCR SYBR Green ROX Mix (Thermo Fisher Scientific Inc., Waltham, MA, United States). The relative mRNA levels were assessed (in triplicate) based upon normalization using a reference gene encoding β-actin (Actb).

The total protein was extracted by radioimmunoassay lysate with 1% protease inhibitor on ice. The protein concentration was measured by BCA protein analysis kit (Thermo Fisher Scientific, United States). The proteins were separated by sodium dodecyl sulfate polyacrylamide gel electrophoresis, and transferred to PVDF membrane. After blocking with 5% fresh milk for 1 h, the membrane was cultured overnight with specific primary antibody at 4°C and then cultured with corresponding secondary antibody for 30 min. Finally, the strips on the film were detected by ECL detection system (Thermo Fisher Scientific, United States) and quantified by Quantity One software (v4.3 in the United States).

Enzyme linked immunosorbent assay (ELISA) was performed to detect quantitatively inflammatory cytokine. Blood from the inferior vena cava was left for 20 min and then collected into tubes and centrifuged (2,000 rpm, 15 min, 4°C). Then 3 ml supernatants of kidney homogenates were harvested and stored at −20°C. IL-17A, IFN-γ, and IL-10 were measured by ELISA (R&D Systems) according to the product instruction.

(1) Flow cytometry was used to measure the percentage of Th17 cells. A 250-μl whole blood sample was mixed with 250 μl RPMI-1640 medium without fetal bovine serum to a volume of 500 μl. Individual control and experimental tubes were set up. Then 250 μl diluent, 1 μl PMA (0.1 mg/ml), 5 μl ionomycin (1 mg/ml), and 1 μl monensin (50 mg/ml) were added to each tube, incubated at 37°C for 12 h in a 5% CO₂ incubator. The mixture was centrifuged again and the supernatant was discarded. Then red cells were lysed by adding a 1-ml lysing buffer (FACS lysing solution; Bio Legend Co.). Cells were incubated for 5 min at room temperature. The mixture was centrifuged again and the supernatant again discarded. The cells were resuspended in PBS and transferred into a 1.5-ml Eppendorf tube. The cells were centrifuged at 2,500 rpm for 5 min, and the supernatant was discarded. The wells were washed twice with PBS, added with 1 μl anti-mouse CD3FITC (0.5 mg/ml) and 1 μl anti-rabbit CD8aPE (0.2 mg/ml), and kept at room temperature away from light for 30 min. The wells were washed once with PBS and the supernatant was discarded, then added with 0.5 ml fixatives and protected from light for 20 min for incubation at room temperature. The cells were washed once with 1 ml PBS, centrifuged at 1,000 rpm for 5 min, and the supernatant was discarded. The cells were added with 1 ml permeabilization wash buffer and protected from light for 20 min for incubation at room temperature, centrifuged at 1,000 rpm for 5 min, and the supernatant was discarded. After washing with PBS, 1 μl anti-rabbit IL-17AAPC (0.2 mg/ml) was added into experimental tubes and 1 μl APC-lgG1 was added into control tubes. The cells were protected from light for 30 min for incubation at 4°C, washed once with PBS, centrifuged at 1,000 rpm for 5 min, and the supernatant was discarded. The cells were resuspended in 0.5 ml PBS and analyzed for CD3⁺CD8^–IL-17⁺ expression by flow cytometry.

(2) Flow cytometry was used to measure the percentage of Treg cells. Individual control and experimental tubes were set up. Then 100 μl whole blood, 1 μl anti-mouse CD4ECD, and 1 μl anti-mouse CD25PE were added to each tube, kept at room temperature away from light for 30 min, centrifuged at 1,000 rpm for 5 min, and the supernatant was discarded. The cells were washed once with 1 ml PBS, centrifuged at 1,000 rpm for 5 min, and the supernatant was discarded. Then red cells were lysed by adding a 1-ml lysing buffer (FACS lysing solution; Bio Legend Co.). Cells were incubated for 5 min at room temperature. The cells were added with 0.5 ml Foxp3 fixatives and protected from light for 20 min for incubation at room temperature. The cells were washed once with 1 ml PBS, centrifuged at 1,000 rpm for 5 min, and the supernatant was discarded. The cells were added with 1 ml permeabilization wash buffer and protected from light for 20 min for incubation at room temperature, centrifuged at 1,000 rpm for 5 min, and the supernatant was discarded. After washing with PBS, 1 μl anti-mouse Foxp3Percpcy5 was added into experimental tubes and 1 μl Percpcy-lgG2 was added into control tubes. The cells were protected from light for 30 min for incubation at room temperature, washed once with PBS, centrifuged at 1,000 rpm for 5 min, and the supernatant was discarded. The cells were resuspended in 0.5 ml PBS and analyzed for CD4⁺CD25⁺Foxp3⁺ expression by flow cytometry.

Stata software (version 11.0) was used to perform the analysis. Data were expressed as mean ± SD; the differences between groups were analyzed by either the paired t-test or ANOVA test (both one-way ANOVA test and two-way ANOVA test). In addition, we use Bonferroni or LSD as a post hoc test. P-values < 0.05 (two-tailed) were considered statistically significant.

Mice were orally administered BA and DMSO 3 days before surgery as indicated in Figure 1A. We tested the BUN and Cr of mice on the first day after surgery and obtained kidney tissue specimens. BA significantly reduced the increase of BUN and Cr in mice caused by IRI, suggesting the protective effect of BA on the renal function of IRI in mice (Figures 1B,C). HE staining of kidneys in the control group showed apparent renal tubular damage, renal tubular epithelial cell atrophy, degeneration, necrosis, and extensive lesion range. In the BA treatment group, the renal tubular epithelial injury was significantly improved compared with the control group, the lesion was lighter, and the lesion range was smaller (Figure 1D). BA treatment significantly reduced the tubular injury Paller score in IRI mice (Figure 1E). All of the aforementioned indicate that BA has a significant effect in alleviating the decline of renal function and tubular damage caused by IRI.

Next, renal cortex homogenate was collected to assay the anti-inflammatory effects of BA. The inflammatory cytokines IL-17A and IL-6 (Proinflammatory factor) were significantly decreased after the treatment of BA in kidney IRI while IL-10 (anti-inflammatory factor) was significantly increased (Figures 2A–C). Treg was involved in the process of liver and kidney IRI (Zheng et al., 2018; Luan et al., 2020). Thus, we designed a series of in vitro experiments to test the effect of BA on the regulation of iTreg generation. Human naïve CD4⁺CD25^–CD45RA⁺ T cells (purity ≥ 95%) were extracted from peripheral blood samples provided by healthy donors and then induced naïve CD4⁺ T cells to iTregs by TGF-β, IL-2, and CD3/CD28 beads.

Next, we explored at the cellular level whether BA regulated Treg differentiation in vitro. For this reason, CD4⁺ cells (CD4⁺CD62L^HighCD44^Low) isolated from C57/B6J mouse spleens were cultured under Treg polarizing conditions with BA or placebo.

The expression of FoxP3 was significantly increased in the cells treated by BA after induction (Figure 2D). In addition, compared with the control group, the absolute number of CD4⁺ iTregs in the BA group was slightly increased on day 8 while the difference was not significant on day 8 (Figure 2E). The suppressive ability of BA-CD4⁺ iTregs was estimated by carboxyfluorescein succinimidyl amino ester (CFSE) coculture assays. The washed CD4⁺ iTregs were cocultured with CFSE-labeled fresh PBMCs at different ratios (Tregs: PBMCs = 1:1 or 1:8) in the presence of expansion beads. Figure 2F shows the expansion of CD4⁺ effector T cells. Although there was no difference in suppressive ability at 1:1 (Treg:CD4⁺ effector T cells), BA-CD4⁺ iTregs showed stronger inhibition at 1:8. Studies have confirmed that there is a significant correlation between pro-inflammatory/anti-inflammatory cytokines and Treg function (Pandiyan and Zhu, 2015; Qiu et al., 2018). These results suggest that BA inhibits the differentiation of naïve CD4⁺ T cells into other inflammatory cells and partly increases the function of CD4⁺ iTregs by regulating the expression of cytokines. The demethylation of CpG islands caused by epigenetic regulation of the Foxp3 loci is considered to be an important marker of the stability and functionality of Tregs (Floess et al., 2007). We used bisulfite sequence analysis to detect the methylation status of CD4⁺ iTregs induced by BA, and compared with the control group, there were fewer methylated CpG sites in BA-CD4⁺ iTregs (Figure 2G). Thus, BA regulates Treg cell differentiation and keeps its regulatory phenotype in vitro.

Treg and Th17 cells are critical immune cells and play an essential role in various inflammatory diseases including renal IRI (Dellepiane et al., 2020). Flow cytometry was then used to detect the percentage of Th17 cells in mice blood. The addition of BA significantly reduced the frequency and number of CD4⁺IL-17A⁺ cells, promoting the inflammatory responses in renal IRI (Figure 3A). Our study found that the frequency of Th17 cells in the BA treatment group was significantly reduced, suggesting that BA regulates Treg/Th17 balance and regulates CD4⁺ T cell differentiation. Suppressor of cytokine signaling family (SOCS1/SOCS3) is necessary for the balance of Treg/Th17 population in different organ diseases (Qu et al., 2017; Wang D. et al., 2018; Kmiolek et al., 2020; Saleh et al., 2020). Therefore, we detected the expression of RORγt, SOCS1, and SOCS3 expression. As indicated in Figures 3B,C, the expression of RORγt and SOCS3, but not SOCS1, was decreased after BA treatment. These results suggest that BA regulates Treg/Th17 balance by targeting SOCS3 but not SOCS1 in renal IRI.

Foxp3 and RORγt mRNA and JAK2, STAT3, and SOCS3 protein are essential for the balance of Treg/Th17. We further tested the expression of p-JAK2, JAK2, p-STAT3, and STAT3 by western blot. As shown in Figures 4A,B, BA treatments attenuated the expression of p-STAT3 but not p-JAK2.

Since BRL52537, a kappa-opioid receptor agonist, could regulate p-STAT3 but not STAT3 expression in cerebral IRI and is involved in kidney injury, we used BRL52537 to assess whether p-STAT3 is necessary for the protective effects of BA (Fang et al., 2013; Golosova et al., 2020). After BRL52537 treatment, the expression of p-STAT3 but not p-JAK2 was increased, which conformed to the p-STAT3 agonist role of BRL52537 (Figures 4C,D). The previous studies show that overexpression of STAT-3 would increase the percentages of Th17 and relevantly reduce the rates of Treg cells in vivo and in vitro (Wang Y. et al., 2018; Liu et al., 2020). We observed similar results after BRL52537 treatment in our results (data not shown). Renal cortex homogenate was collected to assay the inflammatory effects after the increased expression of p-STAT3. Figures 4E–G suggests that BRL52537 significantly attenuated the anti-inflammatory effects of BA on renal IRI. Also, HE staining indicated that BRL52537 significantly attenuated the protective effects of BA on renal IRI (Figure 4H). Taken together, BA exerts therapeutic and protective effects on renal IRI through p-STAT3.

Butyric acid is an effective activator of Nrf2, while Nrf2/HO-1 signaling pathway is an essential regulator of oxidative stress. The researches show that HO-1 mediates STAT3 pathway in different cells and diseases (Tang et al., 2018; Kuang et al., 2019). Therefore, we decided to identify whether the signaling factor HO-1 was implicated in Treg/Th17 cell differentiation mediated by BA. As shown in Figures 5A,B, SnPP, the inhibitor of HO-1, significantly reduced the upregulation of BA-related p-STAT3 expression. In CD4⁺ T cells under Treg/TH17 polarizing conditions, BA/SnPP co-treatment reversed the immunological balance of Treg/Th17 cells by BA only (Figure 5C). Also, the pathological results show that SnPP treatment significantly, but not wholly, reversed the regulating effect of BA (Figure 5D). To conclude, HO-1/p-STAT3 signaling pathway was implicated in Treg/Th17 balance mediated by BA in renal IRI.