Peripheral blood from RA patients was obtained from the Department of Rheumatology and Immunology of the Second Affiliated Hospital of Dalian Medical University in China. All RA patients in this study fulfilled the American College of Rheumatology (ACR) 1987 revised criteria. Age and sex-matched health controls (HC) were obtained from the Medical Examination Center of the Second Hospital of Dalian Medical University. The study has been approved by the ethics committee of the Dalian Medical University (2018–061), and informed consent was obtained from all subjects. The median age of RA patients was 52.40 ± 11.18 years and HC was 40.80 ± 12.75 years.

Iguratimod (IGU) was provided by Simcere Pharmaceutical (Nanjing, China).

Peripheral blood mononuclear cells (PBMCs) were purified from peripheral blood using Ficoll-Paque plus. Human CD4⁺ T and CD19⁺ B cells were purified from PBMCs by Miltenyi Cell Isolation Kit according to the manufacturer’s protocols. The purified CD4⁺ T cells and PBMCs were activated with anti-CD3 and anti-CD28 antibodies (eBioscience) in RPMI-1640 containing 10% FBS and treated with vehicle (DMSO) or IGU (0-200 μg/ml) for 24-72 hours. In several experiments, 2-DG (5 mM) (Solarbio), Rapamycin (100 nM) (Sigma-Aldrich), Echinomycin (5 nM) (Abcam), or Adaptaquin (5 μM) (R&D Systems) were administrated to the cell culture.

CD4⁺ T cells were purified from PBMCs of RA patients using Human CD4⁺ T Cell Isolation Kit (Miltenyi) and then cultured with plate-coated anti-CD3 antibody (5 μg/ml) and anti-CD28 antibody (2 μg/ml) plus IGU or DMSO in RPMI-1640 containing 10% FBS for 3 days. CD19⁺ B cells were purified from PBMCs of healthy donors using Human CD19⁺ B cell Isolation Kit (Miltenyi). In a 96-well round-bottom plate, CD19⁺ B cells were co-cultured with RA-CD4⁺ T cells which were treated with or without IGU at a ratio of 5:1. A total of 2 μg/ml anti-CD3/CD28, 0.5 μg/ml anti-CD40 antibody (R&D Systems), 0.1 μM CpG (Miltenyi) were added to RPMI 1640 complete medium, which was then added to the cells at 200 μl per well. After 7 days, the supernatant was separated from the cells via centrifugation. Both cells and cell culture supernatant were collected for further experiments.

RA-CD4⁺ T cells stimulated by plate-coated anti-CD3 antibody (5 μg/ml) and anti-CD28 antibody (2 μg/ml) with or without IGU for 24 hours were collected and washed with ice-cold PBS twice. Total RNA was extracted from RA-CD4⁺ T cells using AG RNAex Pro Reagent (AG21102, Accurate Biology, Hunan, Co., Ltd), and cDNA was transcribed using 5×All-In-One RT MasterMix (Abm), both according to manufacturers’ instructions. Real-time PCR was carried out with the SYBR Green Premix Pro Taq HS qPCR Kit (AG11701, Accurate Biology, Hunan, Co., Ltd) on the CFX96 Real-Time PCR Detection System (Bio-Rad). The mRNA levels in each sample were normalized to the relative quantity of β-actin gene expression and relative gene expression was determined using the standard 2^-ΔΔCT method. The specific primers used are listed in Supplemental Table 1 .

Glucose uptake was measured following 20 min incubation with a fluorescent D-glucose analog 2-[N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl) amino]-2-deoxy-D-glucose (2-NBDG) (Sigma-Aldrich). PBMCs from RA patients and healthy donors were resuspended in glucose-free RPMI 1640 medium (Gibco) for 30 min, and then 50 μM of 2-NBDG was added. In addition, purified RA-CD4⁺ T cells were treated with IGU or DMSO for 24 hours, then cultured in glucose-free RPMI 1640 medium for 30 min, and followed by adding of 2-NBDG (50 μM). After 20 min, cells were placed on ice and washed twice with ice-cold PBS. Cells were measured immediately by flow cytometry after CD4 staining.

Purified RA-CD4⁺ T cells were treated with IGU and DMSO for 24 hours. Glucose concentrations in the cell culture supernatants were measured by Glucose Assay Kits (Sigma-Aldrich). L-lactate concentrations in cell culture supernatants from RA-CD4⁺ T cells were determined using the CheKine Lactate Assay Kit (Abbkine). The glucose and L-lactate concentrations were determined according to the manufacturer’s protocols. Glucose uptake = glucose concentration in complete medium - glucose concentration in cell culture supernatants.

Real-time measurements of extracellular acidification rate (ECAR) and oxygen consumption rate (OCR) were performed with a Seahorse XF24 Extracellular Flux Analyser (Agilent). To detect the regulation of IGU on CD4⁺ T cell metabolism, RA-CD4⁺ T cells were pre-activated by anti-CD3 antibody and anti-CD28 antibody for 24 hours and then treated with DMSO or IGU 1 hour before the experiment. Collecting and seeding the cells in XF Assay medium in a 24-well microplate pre-coated with 0.01% Poly-L-lysine (Solarbio). Cellular oxidative phosphorylation was determined by Seahorse XF Cell Mito Stress test kit with three injections: 1 μM oligomycin, 1 μM FCCP, and 1 μM rotenone. Glycolysis associated parameters were determined by Seahorse XF Glycolysis Stress test kit with three injections: 10 mM glucose, 1 μM oligomycin, and 50 mM 2-DG.

Cell proliferation assays were performed using the Carboxyfluorescein succinimidyl amino ester (CFSE) Cell Labeling Assay (eBioscience). PBMCs isolated from healthy donors and RA patients were washed two times with PBS and resuspended at pre-warmed PBS. CFSE was added at a final concentration of 1.5 μM to label cells. The CFSE labeled cells were stimulated by anti-CD3/CD28 antibodies (2 μg/ml) in the presence of DMSO or IGU for 3 or 5 days. Finally, the cells were washed with PBS twice and measured by flow cytometry after CD4 staining.

Cell apoptosis was analyzed by using the Propidium Iodide (PI) and Annexin V-FITC Apoptosis Test Kit (eBioscience). PBMCs isolated from healthy donors or RA patients were stimulated by anti-CD3/CD28 antibodies (2 μg/ml) with or without IGU for 24 hours. Then, the cells were harvested and washed with cold PBS twice. Next, the cells were resuspended in 100 μl binding buffer to stain CD4 and Annexin V-FITC in the dark. After 20 min, the cells were rewashed, and PI was added. The early apoptotic cells were detected by flow cytometry.

Cell surface staining was performed using BD Biosciences or eBioscience reagents. The dead cells were excluded from the analysis by fixable viability dye (FVD) (eBioscience) before labeling the surface antibody except for cell apoptosis analysis. For intracellular cytokine and phosphorylation mTOR (p-mTOR) staining, cells were permeabilized using the Cytofix/Cytoperm Intracellular Staining Kit (BD Biosciences) and stained with cytokine-specific mAbs. For Bcl-6, Hif1α, and Foxp3 staining, cells were fixed and permeabilized using the Transcription Factor Fixation/Permeabilization Buffer Set (eBioscience). For HK2 staining, the cells were fixed in 4% paraformaldehyde and permeabilized in 0.5% Triton X-100 before antibody staining. For LDH staining, the cells were fixed in 4% paraformaldehyde and permeabilized in 90% methanol before antibody staining. The flow antibodies are listed in Supplemental Table 2 . All stained cells were analyzed on the Flow Cytometer (NovoCyte 2040R) and data were analyzed with NovoExpress software.

After the T-B cell was co-cultured for 7 days, the supernatant was separated from the cells via centrifugation. The concentrations of IgM and IgG in the supernatant were measured using the Human IgM ELISA Kit and the Human IgG ELISA Kit (SenBeiJia Biological Technology, China) according to their instructions. The plates were read at 450 nm by a microplate reader (Thermo).

For western blot analyses, proteins were extracted by lysing anti-CD3/CD28 activated RA-CD4⁺ T cells in RIPA buffer. Equal amounts of protein in each sample were analyzed by SDS-PAGE and electrophoretically transferred to nitrocellulose filter (NC) membranes. The membranes were incubated with antibodies including HK2 (anti-rabbit, Cell Signaling Technology), LDHA (anti-rabbit, Cell Signaling Technology), β-actin (anti-rabbit, Cell Signaling Technology) at 4 °C overnight, then incubated with a fluorescent secondary antibody (Abclonal) for 1.5 hours at room temperature. The results were detected by Odyssey CLx Infrared Scanner (Odyssey CLx). The expression levels of HK2 and LDH were normalized to β-actin and adjusted to the levels in the control group (served as 1).

Statistical analysis was performed with GraphPad Prism 9 software. Unless indicated otherwise, data are expressed as mean ± standard error of mean (SEM). Statistical differences were analyzed by unpaired Student t-test, paired Student’s t-test, Mann-Whitney test, and one-way ANOVA. The P-values < 0.05 were considered significant. Asterisks mark the significant differences between different groups (*, P < 0.05; **, P < 0.01; ***, P < 0.001 and ****, P < 0.0001).

It is generally believed that IGU plays an important immunomodulatory role by inhibiting the production of immunoglobulins on B cells (16, 17). However, B cells require interaction with helper CD4⁺ T cells to become activated (18). The interactions of CD4⁺ T cells and B cells are fundamental for the generation of antibody responses, as well as for the development of harmful autoimmune diseases (19, 20). We wondered whether IGU might regulate T-B interactions. We first screened the optimal concentration of IGU in CD4⁺ T cells, detecting the proliferation and apoptosis of HC CD4⁺ T cells that were treated with different concentrations of IGU (0, 10, 50, 100, 150, 200 μg/ml). The gating strategy of CD4⁺ T cells was shown in Figure S1A . The results showed that 100 μg/ml of IGU could significantly inhibit the proliferation of CD4⁺ T cells ( Figure S1B ) but had no obvious effect on their apoptosis ( Figure S1C ). Therefore, we selected 100 μg/ml IGU for the following studies.

To examine whether IGU impinges on T-B interactions, we conducted in vitro conjugation assay. We first treated RA-CD4⁺ T cells with IGU or the equal volume of DMSO for 3 days, then co-cultured with CD19⁺ B cells for 7 days. The purity of CD4⁺ T cells was higher than 95% and the purity of CD19⁺ B cells was higher than 90% ( Figure 1A ). As compared with the B cell-control RA-CD4⁺ T cell co-culture group (B + T_CON), the percentage of CD138⁺ plasma cells decreased in the B cell + IGU treated RA-CD4⁺ T cell co-culture group (B + T_IGU) ( Figure 1B ). Simultaneously, we also found that although there was no significant difference in IgG level in the supernatant of the co-culture system, IgM level decreased when CD19⁺ B co-cultured with IGU treated RA-CD4⁺ T cells ( Figure 1C ). These results hint that IGU can inhibit T cell-dependent antibody production.

Next, we want to explore what function of T cells is affected by IGU. Firstly, we examined whether IGU regulated RA-CD4⁺ T cell proliferation and apoptosis. We found that IGU inhibited the proliferation of RA-CD4⁺ T cells ( Figure 2A ) but had no significant effect on their apoptosis ( Figure 2B ). We then tested the effects of IGU on the activation of RA-CD4⁺ T cells. We found that activation markers, including CD25 and CD69, decreased significantly on day 3 in the presence of IGU ( Figures 2C , D ). Collectively, these results indicate that IGU inhibits RA-CD4⁺ T cells proliferation and activation.

The proportion of cTfh (CD4⁺CXCR5⁺PD-1⁺ T) cells in RA patients was higher than that in HC ( Figure S2A ). Next, we evaluated the percentages of cTfh cells after IGU treatment. As shown in Figure 3A , the percentages of cTfh cells significantly decreased in the presence of IGU. Tfh cells are regulated by a complicated network of transcription factors, including B cell lymphoma 6 protein (Bcl-6) and basic leucine zipper ATF-like transcription factor (BATF) (3). We purified CD4⁺ T cells from the PBMCs of RA patients and treated them with or without IGU. We observed that the expression of Bcl-6 and BATF significantly decreased at the mRNA level after IGU treatment ( Figure S2B ). Flow cytometry results showed that the percentages of CD4⁺Bcl-6⁺ T cells were also down-regulated after IGU treatment ( Figure 3B ). Tfh cells provide help to cognate B cells via their expression of CD40 ligand (CD40L), interleukin (IL)-21, IL-4, and other characteristic markers (21). Thus, we examined the surface molecules and cytokines which are related to Tfh cell function. We found that the expression of ICOS, CD40L, and CD84 ( Figures 3C –E ) on RA-CD4⁺ T cells significantly decreased, and we found that IL-21, IL-4, and IL-10 secreted by CD4⁺ T cells also reduced ( Figures 3F –H ). These results indicate that IGU can inhibit the RA-cTfh cell function.

In addition, peripheral helper T (Tph), Th1, Th17, and Treg cells are also involved in the pathogenesis of RA by secreting different cytokines (22, 23). As shown in Figures S2C–F , we found that IGU could also inhibit the percentage of Tph (CD4⁺PD-1⁺CXCR5^- T) cells, CD4⁺IFN-γ⁺ T cells, CD4⁺IL-17A⁺ T cells, but had no significant effect on Treg (CD4⁺Foxp3⁺ T) cells.

Cellular glucose metabolism controlling the functions of immune cells has greatly evolved in the past few years. Moreover, inhibition of glycolysis can significantly decrease the severity of joint inflammation in a model of RA (10). We further consider whether IGU can regulate the glucose metabolism in RA-CD4⁺ T cells. Firstly, we characterized the glucose metabolism ability of RA and HC CD4⁺ T cells. We found no significant difference in glucose uptake ability in RA and HC groups ( Figure S3A ). However, our results demonstrated that the expression of key enzymes such as glucose transporter type 1 (GLUT1) and HK2 in RA-CD4⁺ T cells were higher than in HC-CD4⁺ T cells ( Figures S3B, C ), although there was no significant difference in lactate dehydrogenase (LDH) expression ( Figure S3D ). These phenomena suggest that RA-CD4⁺ T cells may have more active glucose metabolism than HC.

To explore whether IGU can also regulate the glucose metabolism of RA-CD4⁺ T cells, we determined the potential suppression of the glucose uptake ability in RA-CD4⁺ T cells during IGU-mediated function inhibition. We found that IGU treatment markedly inhibited the glucose uptake ability in RA-CD4⁺ T cells ( Figure 4A ). We also determined the key enzymes of the glycolytic pathway, including HK2, muscle-type phosphofructokinase (PFKM), pyruvate kinase M1/2 (PKM1/2), and lactate dehydrogenase A (LDHA) by using qPCR. The results showed that the mRNA expression of HK2 and LDHA decreased in the IGU treated group, while no effect was observed in PFKM or PKM1/2 ( Figure 4B ). As some studies have reported that RA T cells are characterized by the increased availability of the pentose phosphate pathway (24), we next examined two key enzymes that regulate the pentose phosphate pathway, 6-phosphofructo-2-kinase/fructose-2,6-biphosphatase 3 (PFKFB3) and glucose-6-phosphate dehydrogenase (G6PD). However, IGU did not affect these two enzymes ( Figure S3E ). Furthermore, we analyzed the protein levels of GLUT1, HK2, and LDH ( Figures 4C –E ). Consistent with the mRNA levels, the protein levels of HK2 and LDH were significantly down-regulated after IGU treatment, while the expression of GLUT1 had no significant change. Concurrently, we also detected the supernatant of CD4⁺ T cells treated with IGU or DMSO and found that the glucose uptake ( Figure 4F ) and lactate production ( Figure 4G ) of CD4⁺ T cells were significantly inhibited in the presence of IGU. To further verify the direct effect of IGU on CD4⁺ T cell glucose metabolism, we measured the real-time bioenergetic profiles: ECAR, an indicator of glycolysis, and OCR, an indicator of OXPHOS ( Figures 4H , I ). Notably, compared with the control groups, IGU treated CD4⁺ T cells showed lower glycolysis and glycolytic capacity. However, there was no significant difference in basal OCR, ATP production, and spare respiratory capacity. Collectively, these results indicate that IGU inhibits glucose metabolism in RA-CD4⁺ T cells.

A recent study has shown that high glucose utilization is a unique requirement of autoreactive Tfh cells and has implied that inhibiting glycolysis can uniquely target pathogenic autoreactive Tfh cells while preserving protective immunity against pathogens (11). Both pentose phosphate pathway and glycolysis will work through HK2, which is the first-rate limiting enzyme of glucose metabolism. We determined whether the inhibitory effect of IGU on RA-CD4⁺ T cells was dependent on the inhibition of HK2 by using a specific pharmacological inhibitor, 2-DG, which had no effect on cell apoptosis ( Figure S4 ). Consistent with the previous reports (25), the frequencies of cTfh cells, the activated molecules ICOS on CD4⁺ T cells, and IL-21 producing CD4⁺ T cells were significantly reduced by the 2-DG treatment ( Figures 5A –C ). Importantly, when HK2 was inhibited, the combined use of IGU did not further reduce the inhibitory effect ( Figures 5A –C ), indicating that IGU inhibited HK2 resulting in cTfh cell function inhibition.

Recent experiments have shown that mTOR and Hif1α perform important functions in glucose metabolism (26). In our study, we evaluated the expression of mTOR and Hif1α in RA and HC CD4⁺ T cells. We found that the level of p-mTOR in RA and HC CD4⁺ T cells was similar ( Figure S5A ). However, the expression of Hif1α in RA-CD4⁺ T cells was higher than in HC ( Figure S5B ).

We further dissected whether mTOR-Hif1α signaling was involved in IGU-mediated cTfh cells inhibition. We found that IGU treatment down-regulated both p-mTOR and Hif1α in RA-CD4⁺ T cells ( Figures 6A , B ). We then used the specific pharmacological inhibitors Rapamycin and Echinomycin against mTOR and Hif1α respectively to confirm the functional importance of mTOR-Hif1α signaling in RA-CD4⁺ T cells inhibition induced by IGU. We observed that Echinomycin strongly reduced the frequencies of cTfh cells, the expression of ICOS, and the IL-21 produced by CD4⁺ T cells ( Figures 6C –E ). Importantly, When Hif1α was inhibited, combined treatments with Echinomycin and IGU did not further reduce the inhibitory effect ( Figures 6C –E ). The frequencies of cTfh cells, ICOS, and IL-21 were also decreased by mTOR antagonist Rapamycin ( Figure S6 ), but the frequency of cTfh cells, the activated molecules on CD4⁺ T cells, and the IL-21 production by CD4⁺ T cells were further decreased by adding IGU. We ensured that Echinomycin and Rapamycin did not affect cell apoptosis ( Figure S4 ). These results indicate that Hif1α is critically involved in IGU mediated RA-cTfh cell suppression and IGU could decrease mTOR which might partially contribute to the reduction of cTfh cells. Moreover, we found that treatment with Hif1α inhibitor significantly decreased HK2 but not LDH expression ( Figures 6F , G ). In addition, we activated the Hif1α function with Adaptaquin. We found that activation of Hif1α signaling dramatically upregulated the expression of HK2 in RA-CD4⁺ T cells and IGU treatment reconstituted HK2 expression ( Figure 6H ). These results collectively suggest that IGU restrains RA cTfh cell function by inhibiting the Hif1α-HK2 axis ( Figure 6I ).