The present study included 80 healthy controls and 60 cases each of primary palmar hyperhidrosis (PPH), primary craniomofacial hyperhidrosis (PCH), and primary axillary hyperhidrosis (PAH). All participants were of Han ethnicity and were recruited from the First Affiliated Hospital of Fujian Medical University, with age and gender matched between the two groups. Both groups also met the same inclusion and exclusion criteria, as outlined below:

Inclusion criteria: Participants aged 18 years or older with informed consent.

Exclusion criteria: Individuals with significant damage to vital organs such as heart, liver, or kidneys; those with infectious diseases or autoimmune conditions; individuals on long-term steroid or other immunosuppressive medication; and patients with tumors were excluded from the study.

The study was approved by the Ethics Committee of First Affiliated Hospital of Fujian Medical University ([2021]466), and written informed consent was derived from the participants.

All participants’ fasting blood was collected for the isolation of PBMCs using aseptic techniques. Firstly, 1 ml of Ficoll was added to a 15 ml centrifuge tube, followed by the addition of 1 ml of freshly collected anticoagulated peripheral blood mixed with an equal amount of PBS. Carefully added to the top of the Ficoll layer, the liquid interface was maintained as clear as possible. The mixture was centrifuged at 400×g for 15 minutes at room temperature. After centrifugation, four layers were formed from top to bottom: the plasma layer, mononuclear cell layer, transparent separation fluid layer, and blood cell layer. The second cloudy layer containing the target cells (mononuclear cells) was carefully aspirated and transferred to a centrifuge tube containing PBS. The cells were thoroughly mixed and centrifuged at 2000 rpm for 10 minutes. After two additional washes with PBS, PBMCs were obtained. The PBMCs were suspended in RPMI1640 culture medium, and the cell concentration was adjusted to 1-2×10⁶/ml. For Treg labeling, 500 μl of PBMCs was taken, and for Th17 cell labeling, 600 μl was placed in a 24-well cell culture plate. Stimulation-blocking agents, PMA, and BFA, were added, and the cells were cultured at 37°C, 5% CO₂ in a cell culture incubator for 5 hours.

Cells were resuspended, and 100 μl of the cell suspension was aliquoted into separate 1.5 ml EP tubes, including a blank tube, a compensation tube, and a sample tube. Surface antibodies CD3-FITC (85-11-0037-42, eBioscience) and CD8a-APC (85-17-0086-42, eBioscience) (5 μl each) were added to each tube. Incubation was performed at room temperature in the dark for 30 minutes, followed by centrifugation at 400×g for 5 minutes. The supernatant was discarded, and cells were resuspended. Fixative permeabilization reagent A (100 μl) was added and incubated at room temperature for 15 minutes. Flow cytometry buffer was added, mixed, and centrifuged at 400×g for 5 minutes. The supernatant was discarded, and cells were resuspended. Fixative permeabilization reagent B and IL-17-PE antibody (85-12-7179-42, eBioscience) (100 μl each) were added and incubated at 4°C in the dark for 30 minutes. Flow cytometry buffer was added, mixed, and centrifuged at 400×g for 5 minutes. The supernatant was discarded, and 4% paraformaldehyde (100 μl) was added. Incubation was performed at 4°C in the dark for 15 minutes. After washing, cells were resuspended in 300 μl of PBS and stored at 4°C in the dark for subsequent flow cytometric analysis. The gating strategy used for Th17 cells has been provided in Supplementary Figure S1 .

Cells were resuspended, and 100 μl of the cell suspension was placed in each of the tubes (blank, compensation, and sample). Surface antibodies CD4-FITC (85-11-0047-42, eBioscience) and CD25-PE (85-12-0259-42, eBioscience) (5 μl each) were added to each tube and incubated for 25 minutes at room temperature in the dark. After centrifugation, flow cytometry buffer was added, and the supernatant was discarded. Fixed and permeabilization reagent D (1 ml) was added and incubated in the dark at room temperature for 40 minutes. Diluted washing buffer E (2 ml) was added after centrifugation, and cells were resuspended after discarding the supernatant.

Intracellular antibody Foxp3-APC (85-17-4776-42, eBioscience) (5 μl) was added and incubated at room temperature in the dark for 40 minutes. After centrifugation, cells were washed with washing buffer E (2 ml), and the supernatant was discarded. 4% paraformaldehyde (100 μl) was added and incubated at 4°C in the dark for 15 minutes. Following two washes, cells were resuspended in 300 μl of PBS and stored at 4°C in the dark for subsequent flow cytometric analysis. The gating strategy was similar to that used for Th17 cells. Similarly, PBMCs were gated based on FSC/SSC, and doublets were excluded. T lymphocytes were identified by gating CD4+ cells, followed by gating Treg cells based on Foxp3 and CD25 expression. The gating strategy for Treg cells followed the same principles as Th17 cells ( Supplementary Figure S1 ).

At our hospital, sweat gland tissue from patients with PFH was collected through a single-port axillary surgery. A 2 mm wide and 5 mm long full-thickness skin incision was made, and the sweat gland tissue was extracted. Surrounding tissues of a similar volume were collected as the control group. All cases included in the study had no history of palmar hyperhidrosis, axillary hyperhidrosis, or a family history of such conditions. Additionally, there were no reported histories of bromhidrosis (foul-smelling sweat) in any of the patients. The experimental protocol was carried out with the patients’ written consent.

The RT-PCR assay was performed to analyze the gene expression levels of FOXP3, RORγt, and β-actin in the samples. Total RNA was extracted from the desired samples, and cDNA was synthesized using a reverse transcription kit. Specific forward and reverse primers for each target gene (FOXP3, RORγt, and β-actin) were used. Additionally, the Primer-BLAST tool available on the NCBI website was used to evaluate the specificity of the primers. PCR reactions were set up with the cDNA templates and the respective primer pairs, followed by thermal cycling conditions for amplification. The PCR products were visualized through agarose gel electrophoresis, and the bands were analyzed to quantify gene expression levels relative to the reference gene β-actin. The primers were as below:

FOXP3: F: TTTCTGTCAGTCCACTTCACCA

R: CCAGCAGGTCTGAGGCTTTG;

RORγt: F: GTGGGGACAAGTCGTCTGG

R: AGTGCTGGCATCGGTTTCG;

β-actin: F: TGGCACCCAGCACAATGAA

R: CTAAGTCATAGTCCGCCTAGAAGCA

The concentrations of IL-17 (EH3267, sensitivity: 18.75 pg/ml, CV (coefficient of variation) of inter-assay: 4.94%, CV of intra-assay: 5.27%), IL-10 (EH0173, sensitivity: 4.688 pg/ml, CV of inter-assay: 5.05%, CV of intra-assay: 5.19%), IL-6 (EH0201, sensitivity: 2.813 pg/ml, CV of inter-assay: 4.66%, CV of intra-assay: 4.87%) and TGF-β1 (EH0287, sensitivity: 18.75 pg/ml, CV of inter-assay: 5.53%, CV of intra-assay: 5.92%) in the serum of patients were detected by ELISA method (Fine Test, Wuhan, China). Representative standard curves used to calculate cytokine concentrations are provided in Supplementary Figure S2 .

Protein expression of RORγt, FOXP3, IL-17A, and GAPDH in sweat gland tissues was analyzed by Western blotting. Briefly, tissues were lysed in RIPA buffer supplemented with protease inhibitors, and protein concentration was determined by BCA assay. Equal amounts of protein (30 μg) were separated by SDS-PAGE and transferred to PVDF membranes. Membranes were blocked with 5% non-fat milk and incubated overnight at 4°C with the following primary antibodies: RORγt (ab113434, Abcam), FOXP3 (ab215206, Abcam), IL-17A (ab79056, Abcam), and GAPDH (ab8245, Abcam). After washing, membranes were incubated with HRP-conjugated secondary antibodies for 1 hour at room temperature. Protein bands were visualized using enhanced chemiluminescence (ECL) reagents and quantified with ImageJ software.

SR2211, a potent and selective synthetic modulator of RORγ that functions as an inverse agonist, was used for in vivo intervention (19). According to the manufacturer’s data, SR2211 functions as an inverse agonist of RORγ (Ki = 105 nM, IC50 ~320 nM) with minimal off-target activity (>10 μM against RORα, RORβ, TRK, and BRAF). SR2211 was purchased from MedChemExpress (MCE, Cat# HY-16998).

Animal studies were approved by the First Affiliated Hospital of Fujian Medical University. Hyperhidrosis was induced by intraperitoneal injection of pilocarpine hydrochloride at a dose of 5 mg/kg body weight. Five minutes after pilocarpine administration, the appearance of black sweat dots on the plantar surface was photographed to quantify sweat secretion. For intervention, SR2211 was administered intraperitoneally at 20 mg/kg once daily for seven consecutive days prior to hyperhidrosis induction. The dosage and route of administration were based on previously published methods (20). Two hours post-induction, mice were euthanized, and blood and sweat gland tissues were collected. Serum acetylcholine (Ach) levels were measured using a commercially available ELISA kit (Acetylcholine ELISA Kit, OKEH02568; Aviva Systems Biology, San Diego, CA), according to the manufacturer’s instructions. Flow cytometry was performed on whole blood to assess the proportion of Th17 T cells. Serum IL-17 and IL-6 levels were analyzed by ELISA, and protein expression of IL-17A in sweat gland tissues was evaluated via Western blotting.

The data obtained in the study were subjected to statistical analysis using SPSS 15.0 software. The sample size was calculated based on the following equation: N = φ²(ΣSi²/g)/[Σ( Xi - X)²/(g-1)], N represents the sample size of each group, g is the number of groups, X and Si are the mean and standard deviation of each group, respectively. Estimates of effect size and standard deviation were based on the existing literature and our previous experiences. To calculate the power of analysis we assumed α = 0.05 and β = 0.2. For continuous data, t-tests were performed for comparisons between two groups, while for categorical data, χ² tests were conducted. For multiple comparisons, Brown-Forsythe ANOVA test followed by Dunnett’s T3 multiple comparisons test was employed. During the analysis, if the P-value was less than 0.05 (p< 0.05), it was considered as indicating a significant difference.

A comprehensive cohort of 240 participants was enrolled in the current study, encompassing 80 individuals designated as healthy controls (HC) and 60 cases each of PPH, PCH, and PAH. Notably, no statistically significant disparities emerged in terms of age (p=0.297) and gender distribution (p=0.336) between the healthy control group and the PFH group ( Table 1 ).

The equilibrium of the Th17/Treg cell ratio within the peripheral blood of individuals afflicted with PFH was meticulously explored and juxtaposed against that of 80 healthy controls, alongside a pool of 180 PFH patients. The findings underscored that PFH patients manifested an elevated proportion of Th17 cells ( Figure 1A ), accompanied by a decreased proportion of Treg cells ( Figure 1B ) within the peripheral blood milieu. This dysregulation ultimately culminated in a substantial augmentation of the Th17/Treg ratio ( Figure 1C ). Furthermore, receiver operating characteristic (ROC) analysis unmasked the diagnostic potential of Th17 and Treg lymphocyte subpopulations, as well as the Th17/Treg ratio within the peripheral blood, in the context of PFH. Th17 exhibited a sensitivity of 61.67%, a specificity of 71.25%, an area under the curve (AUC) of 0.72, and p-value < 0.001 ( Figure 1D ). Treg demonstrated a sensitivity of 82.78%, a specificity of 64.75%, an AUC of 0.76, and p-value < 0.001 ( Figure 1E ). Most notably, the Th17/Treg ratio emerged as particularly indicative, with a sensitivity of 71.11%, a specificity of 93.75%, an AUC of 0.86, and p-value < 0.001 ( Figure 1F ). This rigorous evaluation accentuates the diagnostic utility of Th17, Treg lymphocyte subpopulations, and the Th17/Treg ratio within peripheral blood as discerning markers for PFH.

The comparisons of Th17 ( Figure 1G ), Treg ( Figure 1H ) lymphocyte subpopulations, and Th17/Treg ratio ( Figure 1I ) in peripheral blood among 80 healthy controls, 60 patients with PPH, 60 patients with PCH, and 60 patients with PAH. The results suggest that there were no significant differences in Th17/Treg cell ratio among different subtypes of PFH patients. However, all PFH subtypes showed significant differences compared to the healthy controls.

We carried out the concentration detection of the relevant cytokines of two kinds of cells. Notably, IL-17 and IL-6, recognized as Th17-associated cytokines (21), along with IL-10 and TGF-β1, known to be Treg-associated cytokines (22), underwent careful scrutiny. The discerned serum levels of IL-17 and IL-6 were notably and significantly escalated across all three PFH subtypes in comparison to the healthy control cohort ( Figures 2A, B ). Conversely, a strikingly converse pattern emerged for IL-10 and TGF-β1, with both these cytokines registering marked reductions in the serum of patients encompassing all three PFH subtypes as opposed to healthy controls ( Figures 2C, D ). These findings mirror previous outcomes, further underscoring the prevailing Th17/Treg imbalance within PFH patients.

Th17 and Treg cells are characterized by the selective expression of distinct transcriptional factors. RORγt is characteristic for Th17 cells (23), whereas FOXP3 is characteristic for Treg cells (24). The current investigation ventured into scrutinizing the expression variations of RORγt and FOXP3 within sweat gland tissues extracted from PFH patients. The results unveiled substantial elevation at both the mRNA and protein levels of RORγt ( Figures 3A, C, D ) across all three PFH subtypes in comparison to the healthy control group. Conversely, for FOXP3, both mRNA and protein levels exhibited noteworthy diminutions within sweat gland tissues across all three PFH subtypes relative to healthy controls ( Figures 3B, C, E ). Collectively, these findings reinforce the assertion that PFH is intrinsically associated with a discernible Th17/Treg immune imbalance. This comprehensive evaluation contributes crucial insights into the intricate immunological underpinnings of PFH, underscoring the pivotal role played by Th17 and Treg cells alongside their associated cytokine profiles in shaping the pathophysiological landscape of this condition.

To evaluate the therapeutic effect of SR2211, an inverse agonist of RORγ (25), on sweat secretion in a mouse model of hyperhidrosis, mice were intraperitoneally administered with SR2211 (20 mg/kg) or vehicle daily for one week prior to pilocarpine-induced hyperhidrosis ( Figure 4A ). Pilocarpine hydrochloride (5 mg/kg) was injected intraperitoneally to induce sweat secretion, and the sweat output was quantified by counting black dots formed on the paw surface (26). As shown in Figure 4B , the number of black dots was significantly increased in hyperhidrosis model mice compared to the vehicle-treated control group, confirming successful model establishment. Notably, pretreatment with SR2211 markedly reduced the number of black dots (p < 0.001), indicating that SR2211 effectively attenuated excessive sweat secretion.

Further histological analysis revealed that the number of sweat secretory granules in the sweat glands was significantly decreased following SR2211 treatment compared with hyperhidrosis model mice ( Figure 4C , p < 0.01), supporting the inhibitory effect of SR2211 on sweat gland activity. Additionally, ELISA of serum acetylcholine concentrations demonstrated a significant reduction in acetylcholine levels in the SR2211-treated group compared with the untreated hyperhidrosis group ( Figure 4D , p < 0.01), suggesting that SR2211 may alleviate hyperhidrosis by modulating cholinergic signaling. Collectively, these results demonstrate that SR2211 ameliorates hyperhidrosis-like symptoms in mice, as evidenced by decreased sweat secretion, reduced secretory granules, and lower serum acetylcholine levels.

To investigate the immunomodulatory effects of SR2211 on Th17-related inflammation in hyperhidrosis, we analyzed peripheral blood lymphocyte subpopulations and pro-inflammatory cytokines. Flow cytometry revealed a significant increase in Th17 lymphocyte proportions in hyperhidrosis model mice compared with vehicle-treated controls. Importantly, SR2211 treatment markedly reduced the proportion of circulating Th17 cells ( Figure 4E , p < 0.01), indicating a suppressive effect on Th17 cell differentiation or expansion. Consistent with these findings, ELISA analysis showed elevated serum levels of IL-17 and IL-6 in hyperhidrosis mice, both of which were significantly reduced following SR2211 administration ( Figures 4F, G , p < 0.01 and p < 0.001, respectively). These results suggest that SR2211 dampens systemic Th17-driven inflammatory responses.

To further validate the local inflammatory status within sweat glands, IL-17A protein expression was examined by Western blotting. IL-17A expression in sweat gland tissues was upregulated in hyperhidrosis mice but significantly decreased after SR2211 treatment ( Figures 4H, I , p < 0.01), while GAPDH served as a loading control. Collectively, these data demonstrate that SR2211 effectively suppresses Th17 cell-associated inflammatory responses both systemically and locally in hyperhidrosis mice, aligning with clinical observations of Th17 involvement in hyperhidrosis pathophysiology.