Primary human dermal fibroblasts were cultured from skin tissues of patients with SSc or healthy donors as previously described (18), under a protocol approved by the Institutional Review Board (IRB) of the University of Pittsburgh. Informed consent was obtained from all participants. Clinical features of the patients included in this study are shown in Supplementary Table 1 . For healthy donors, skin samples were obtained without identifiers and deemed as non-human subject research by the IRB of the Medical University of South Carolina. All research included in this manuscript conforms with the Declaration of Helsinki. Fibroblasts were maintained in Dulbecco’s Modified Eagle’s Medium (DMEM) (Mediatech, Herndon, VA, USA) supplemented with 10% fetal bovine serum (Sigma-Aldrich, St. Louis, MO, USA), penicillin, streptomycin, and antimycotic agent (Invitrogen, Carlsbad, CA, USA) and used in passages 3 to 8.

Primary normal human dermal fibroblasts (NHDF) from healthy donors were treated with the following reagents; PDGF-BB (40 ng/mL) (R&D Systems, Minneapolis, MN, USA), MCH (100nM) (TOCRIS, Minneapolis, MN, USA), PI3K inhibitor (LY294002, 10μM), MEK inhibitor (U0126, 10μM), STAT3 inhibitor (StatticV, 5μM), TGFβ receptor inhibitor (ALK4/5/7 inhibitor, SB431542, 10μM), PDGF receptor inhibitor (CP-673451, 100nM), MCHR1 inhibitor (ATC0065, 50nM), dimethyl sulfoxide (DMSO) or Ethanol as a vehicle control, and used for immunoblotting, PCR and ELISA analyses as appropriate. Inhibitors were used at the indicated concentrations based on previous reports (19–25). Cell viability was determined with Cell Counting Kit-8 (Dojindo, Rockville, MD, USA) assay as previously described (26). Detailed information about the reagents is shown in Supplementary Table 2 .

Total RNA was extracted from each dermal fibroblast cell strain in passage 3 for gene expression analysis using the CsCl-gradient purification method (18). RNA integrity (RINs ≥ 8) was verified using Agilent 2200 TapeStation (Agilent Technologies, Palo Alto, CA). RNAseq libraries were prepared for all dermal fibroblast samples using the TruSeq RNA Sample Prep Kit following the manufacturer’s protocol (Illumina, San Diego, CA) at the Hollings Cancer Center Genomics Core at MUSC ( Supplementary Figure 1A ). Libraries were clustered at a concentration to ensure at least 100 million reads per sample on the cBot as described by the manufacturer (Illumina, San Diego, CA). Clustered RNAseq libraries were paired-end sequenced using version 4 with 2×125 cycles on an Illumina HiSeq2500. Demultiplexing was performed utilizing bcl2fastq-1.8.4 to generate Fastq files.

A second paired-end RNAseq analysis for MCHR1 silenced NHDF and controls treated with/without PDGF-BB was performed at Novogene (Sacramento, CA, USA) with the NEBNext Ultra TM RNA library prep kit (New England Biolabs, MA, USA) on the Illumina NovaSeq 6000 instrument (Illumina) ( Supplementary Figure 1B ).

Total RNA was extracted using TRIzol (Life Tchnologies), and qPCR was performed in duplicate using TaqMan^® gene expression assays using StepOne Plus Real-time PCR machine (Applied Biosystems, Carlsbad, CA), using the following protocol; A. Holding stage: 1) 15 mins at 48°C 2) 10 mins at 95°C. B. Cycling Stage: 1) 1 min at 95°C 2) 1 min at 60°C for a total of 40 cycles. Gene expression levels were normalized to Beta 2 Microglobulin (B2M) and compared using the 2−ΔΔCt method. TaqMan^® probes for human Actin Alpha 2 (ACTA2), Collagen Type I Alpha 1 (Col1α1), CTGF, Fibronectin 1 (FN1), TGFβ1, MCHR1, and B2M were obtained from Applied Biosystems. The assay catalog numbers are shown in Supplementary Table 3 .

Fibroblast lysates were harvested directly in 2× sodium dodecyl sulfate gel-loading buffer (100 mmol/L Tris-Cl, pH 6.8, 200 mmol/L mercaptoethanol, 4% sodium dodecyl sulfate, 0.2% bromophenol blue, 20% glycerol). Samples were separated by 10% sodium dodecyl sulfate polyacrylamide gel electrophoresis and transferred onto nitrocellulose blotting membranes (GE Healthcare Life science). Membranes were then blocked with 5% milk and incubated with one of the following antibodies; MCHR1, Alpha Smooth Muscle Actin (αSMA), Fibronectin (FN), CTGF, Collagen Type I Alpha 1 (Col1A1), TGFβ1, Caspase 3, and Glyceraldehyde-3-Phosphate Dehydrogenase (GAPDH). Product details are shown in Supplementary Table 2 . Signals were detected using horseradish peroxidase-conjugated secondary antibody and chemiluminescence (Perkin-Elmer, Waltham, MA, USA) on an iBright750 (Thermo Fisher Scientific). Signals were quantified using ImageJ software (designed at the National Institutes of Health) for densitometry (31, 32).

Western ligand blotting was performed as previously described (33) with some modifications. Briefly, 1 × 10⁶ NHDF were cultured and collected using the Subcellular Protein Fractionation Kit (Thermo Fisher Scientific), then membrane fraction samples were electrophoresed under non-reducing conditions and transferred to a nitrocellulose membrane. Similarly, recombinant human MCHR1 (Abnova, Taipei City, Taiwan) was electrophoresed under non-reducing conditions and transferred to a nitrocellulose membrane. The membrane was blocked with 5% nonfat milk in Tris-buffered saline/5% Tween-20 and incubated for 1 hour with biotinylated PDGF-BB (R&D). The membrane was washed and incubated for 1 hour with horseradish peroxidase-conjugated streptavidin (Invitrogen), and the signal was detected using chemiluminescence (Perkin-Elmer) on an iBright750 (Thermo Fisher Scientific).

NHDF were seeded in a 6-well plate at a density of 1 × 10⁵ cells/well in DMEM supplemented with 10% fetal bovine serum. MCHR1-specific small-interfering RNA (ON-TARGET plus) and control RNAi were purchased from Dharmacon (Lafayette, CO, USA). For transfection, Lipofectamine^®2000 (Invitrogen) was used in accordance with the manufacturer’s instructions. A mixture of 10 or 100 nM of each RNAi and Lipofectamine^®2000 was added to cells, and cells were cultured for 72 h. Fibroblasts were serum-starved at least 2 h before further stimulation. MCHR1-silenced or control siRNA fibroblasts were treated with 40 ng/mL PDGF-BB (R&D) and harvested 6hrs or 24hrs after stimulation. The culture supernatants were collected by centrifugation and aliquoted. All samples were stored at - 80°C until further analysis.

All experiments were done at the Biacore Molecular Interaction Shared Resource at Georgetown University using a Biacore T200 instrument (Cytiva, Marlborough, MA, USA) with a sensor chip CM5 (Cytiva) at 25°C. Recombinant human MCHR1 Protein (Abnova) was used as a ligand to capture onto the CM5 chip, using standard amine coupling chemistry. Recombinant human PDGF-BB (Sigma-Aldrich) was used as an analyte to flow over the ligand captured surface. Flow Cell (FC) 1 was used as the reference for FC2. Recombinant human MCHR1 was diluted (1:25 dilution, ~1.2 µg/ml diluted concentration) in 10 mM sodium acetate buffer at pH 4.0 and immobilized onto FC2 to a level of ~5500 RU. PBS-P (20 mM Phosphate buffer pH 7.4, 137 mM NaCl, 2.7 mM KCl, 0.05% v/v surfactant P20) was used as the immobilization running buffer. Based on the Immobilized response value, theoretical R_max values were calculated. The R_max values assume 1:1 interaction mechanism. Overnight kinetics for PDGF-BB binding to MCHR1 were performed in the presence of PBS-P. The contact and dissociation times were 60 s and 300 s, respectively. The flow rates of all analytes solutions were maintained at 50 µL/min. Two 20 s pulses of 1:1000 H₃PO₄ (H₃PO₄:ddH₂O, v/v) were injected for surface regeneration. Injected analyte concentrations were from 100 nM to 3.125 nM (two-fold dilutions). Analytes were injected in duplicate. For analysis, sensorgrams from the overnight kinetics were evaluated using 1:1 kinetics model fitting.

NHDF were plated in 6 well plate at a density of 1 × 10⁵ cells/well and cultured overnight. Cells were washed twice with HBSS and pre-treated with 50nM ATC0065 for 1 hour, followed by addition of PDGF-BB, MCH, or vehicle control for 30min. Fibroblasts were harvested with 0.1M HCL and centrifuged at 1000g for 10 min, and supernatants were used to measure cyclic adenosine monophosphate (cAMP) levels. The levels of cAMP in fibroblasts were measured in duplicate samples using Cyclic AMP ELISA kits (Cayman chemical, Ann Arbor, Michigan, USA) according to the manufacturer’s instructions. The absorbance at 410 nm was measured with a SYNERGY H1 microplate reader (Biotec, Winooski, VT, USA).

Cell proliferation was measured by using the Cell Counting Kit-8 (Dojindo, Rockville, MD, USA). NHDF were seeded in a 96-well plate at a density of 5 × 10³ cells/well in 100 µl culture medium and were allowed to adhere overnight. Cells were treated with 40ng/ml PDGF-BB or 10^-7M MCH or vehicle control with or without 50nM ATC0065, then incubated for 24, 48, or 72 hrs. 10μl of Cell Counting Kit-8 reagent was added to each well 2 hrs prior to measurement of absorbance. The absorbance at 450 nm was measured with a SYNERGY H1 microplate reader (Biotec).

In addition to the described differential expression analysis, statistical comparisons were performed using Mann-Whitney U test, unpaired Student’s t-test, multiple t-test, one-way analysis of variance (ANOVA) (posthoc Tukey or Dunnett), or two-way analysis of variance (post-hoc Sidak) as indicated. All tests were carried out using GraphPad Prism version 8.0 software (GraphPad Software, San Diego, CA). A P value <0.05 was considered significant.

To identify potential genes involved in the pathogenesis of skin fibrosis in SSc, we performed RNA sequencing (RNAseq) using dermal fibroblasts from twins discordant for SSc and healthy subjects. We identified 742 DE genes in dermal SSc fibroblasts (q < 0.1, log2FC < |0.6|). We determined that MCHR1 was a hub gene significantly upregulated in our network analysis ( Supplementary Table 4 and Supplementary Figure 2 ), suggesting MCHR1 might play a pivotal role in the pathogenesis of SSc based on the “centrality principle” stating that highly connected vertices are often functionally important in biological systems (34). Thus, we focused our study on MCHR1.

To confirm the upregulation of MCHR1 in SSc patients, we examined MCHR1 mRNA levels in dermal fibroblasts from SSc patients with early disease compared to fibroblasts from healthy subjects using quantitative PCR (qPCR). MCHR1 expression was significantly higher in the dermal fibroblasts of SSc patients ( Figure 1A ). To determine which growth factors may increase MCHR1, we examined the effects of several fibrosis-promoting factors on MCHR1 expression levels in NHDF, including TGFβ1, interleukin 6 (IL-6), bleomycin (BLM), and PDGF-BB. PDGF-BB and BLM significantly increased MCHR1 levels, while TGFβ1 decreased its expression, albeit not significantly (p = 0.0586) ( Figure 1B ). A time-course experiment showed that PDGF-BB induced an increase in MCHR1 expression levels as early as 2 hrs post-treatment, and the difference reached significance after 24 hrs ( Figure 1C ). After 96 hrs of stimulation, MCHR1 levels returned to basal levels. In parallel, MCHR1 protein abundance was increased in NHDF stimulated with PDGF-BB for 48 and 72 hrs ( Figure 1D ).

To determine which PDGF signaling cascades mediate the induction of MCHR1, NHDF were cultured with PDGF-BB in combination with specific inhibitors of PI3K (LY294002), MEK (U0126), and STAT3 (Stat3 inhibitor V, static) signaling. Inhibition of the PI3K and MEK signaling pathways significantly reduced the PDGF-BB-induced increase in MCHR1 mRNA and protein levels ( Figures 1E, F ), suggesting that these two pathways mediate PDGF-BB induction of MCHR1. We also confirmed that the inhibitors had no off-target effect on MCHR1 levels ( Supplementary Figure 3 ).

To further delineate the role of PDGF-BB in fibrosis, we investigated whether PDGF-BB induces fibrotic gene expression in NHDF. PDGF-BB significantly increased the expression levels of Col1α1 at 6 hrs and 24 hrs, CTGF at 4 and 6 hrs, and TGFβ1 at 6 and 24 hrs ( Figure 2A ). Immunoblotting showed that 48 hrs of PDGF-BB stimulation significantly increased the protein abundance of Col1A1, FN1, CTGF, and TGFβ1 in NHDF ( Figure 2B ). Additionally, PDGF-BB increased the expression levels of the myofibroblast marker ACTA2 ( Figure 2A ) and its corresponding protein αSMA ( Figure 2B ), although the increase was not statistically significant. Together, our data showed that PDGF-BB can induce fibrotic mediators and ECM deposition in NHDF.

To determine if the fibrotic responses elicited by PDGF-BB were induced through PDGF receptor signaling, NHDF were pre-incubated with PDGF receptor inhibitor CP-673451, and treated with PDGF-BB. PDGF-BB induced MCHR1 and TGFβ1 protein abundance, and this effect was significantly attenuated by PDGF receptor inhibition ( Figure 2C ). PDGF-BB-induced FN1 and CTGF, but not Col1A1, were also reduced by PDGF receptor inhibition, albeit not significantly ( Figure 2C ).

TGFβ1 is the prototype fibrotic factor that increases the expression of several profibrotic genes in fibroblasts (35). Since PDGF-BB increased TGFβ1 abundance ( Figure 2B ), we sought to determine if the induction of fibrotic genes by PDGF-BB is mediated by TGFβ1. NHDF were pre-incubated with the ALK5 inhibitor, an inhibitor of TGFβ receptor signaling. PDGF-BB-induced Col1A1 and FN1 protein levels were significantly decreased by ALK5 inhibitor ( Supplementary Figure 4 ), suggesting that PDGF-BB-induced upregulation of Col1A1 and FN1 is dependent on PDGF-BB activation of TGFβ receptor. PDGF-BB-induced CTGF and TGFβ1 levels were modestly reduced by ALK5 inhibition, albeit not significantly. ALK5 inhibition did not affect PDGF-BB regulation of αSMA. Together these results show that PDGF-BB induces the profibrotic factors CTGF and TGFβ1 independently of TGFβ1 receptor signaling, while the induction of Col1A1 and FN1 by PDGF-BB is due to activation of TGFβ1 signaling.

Since PDGF-BB induction of CTGF and TGFβ1 was independent of TGFβ receptor signaling, we sought to determine if this response is mediated by MCHR1. MCHR1 expression in NHDF was silenced using small-interfering RNA prior to stimulation with PDGF-BB (see transfection efficacy in Supplementary Figure 5 ). MCHR1 silencing alone did not affect the expression levels of profibrotic genes, but PDGF-BB-induced CTGF and TGFβ1 gene expression levels were significantly decreased by MCHR1 silencing ( Figure 3A ). In conditioned media of cells treated with PDGF-BB, the protein abundance of TGFβ1 and CTGF were increased compared to vehicle-treated cells, and MCHR1 silencing significantly prevented this increase ( Figure 3B ). We also examined the role of MCHR1 in SSc dermal fibroblasts. Our data show that MCHR1 silencing only reduced PDGF-BB induction of TGFβ1 ( Figures 3C, D ). Interestingly, CTGF was not induced by PDGF-BB in SSc dermal fibroblasts at the time point examined.

The findings in MCHR1 silenced fibroblasts suggest that PDGF-BB is working through MCHR1 to induce the expression of the profibrotic factors CTGF and TGFβ1. We, therefore, examined whether MCH, the ligand of MCHR1, also regulates fibrotic genes in NHDF. Our data show that MCH does not increase profibrotic factor expression in NHDF ( Supplementary Figure 6 ), suggesting that this effect is specific to MCHR1 activation by PDGF-BB. To examine whether PDGF-BB binds to MCHR1, we performed western ligand blotting using NHDF membrane fractions. Our data show that biotinylated PDGF-BB binds to a protein of the same molecular weight as MCHR1 ( Figure 4A ). Higher molecular weight bands correspond to the sizes of PDGFR. We further confirmed the interaction by western ligand blot using recombinant MCHR1. Again, PDGF-BB bound a band corresponding to recombinant human MCHR1 in a dose-dependent manner ( Figure 4B ). To confirm the binding of PDGF-BB to MCHR1, we performed SPR assay. SPR assay revealed that PDGF-BB binds to MCHR1 with an average K_D of 46.6 nM (SEM ±8.1, n=3) ( Figure 4C ). Taken together, our data demonstrate that PDGF-BB directly binds to MCHR1, an association that regulates the expression of CTGF and TGFβ1.

The MCH-MCHR1 pathway is known to reduce cAMP levels in some cell types (8). To determine whether PDGF-BB activates cAMP signaling downstream of MCHR1, we measured cAMP levels in NHDF by ELISA. PDGF-BB reduced cAMP levels in NHDF, as did MCH ( Figure 5A ). Pre-incubation of NHDF with a selective MCHR1 inhibitor (ATC0065) prevented the reduction of cAMP in response to both PDGF-BB and MCH, indicating that PDGF-BB can also reduce cAMP levels through MCHR1.

Fibroblast cell proliferation in SSc is associated with PDGF-BB signaling (7), and the MCH/MCHR1 pathway is reported to affect cell proliferation in some cell types (14, 36). Thus, we investigated the effects of PDGF-BB and MCHR1 on cell proliferation in NHDF. As shown in Figure 5B , PDGF-BB induced cell proliferation, and the increase was significant 72 hrs after stimulation ( Figure 5B ). MCHR1 silencing in NHDF and inhibition of MCHR1 with ATC0065 did not affect PDGF-BB-induced cell proliferation ( Figures 5C, D ). Furthermore, treatment of NHDF with MCH had no effect on cell proliferation ( Supplementary Figure 7 ). We further validated our data by examining caspase-3 protein abundance in NHDF. Consistent with our cell proliferation assay results, caspase-3 protein abundance significantly increased in PDGF-BB treated NHDF ( Supplementary Figure 8 ). These findings suggest that, although MCHR1 mediates PDGF-BB induction of CTGF and TGFβ1 expression, PDGF-BB regulation of cell proliferation is independent of MCHR1.

This study is the first to demonstrate a role for MCHR1 in mediating the profibrotic effects of PDGF-BB and its reduction of cAMP. The PDGF-BB mediated increase in CTGF and TGFβ1 levels was MCHR1-dependent and TGFβ receptor-independent. We observed that PDGF-BB and BLM stimulation significantly increased MCHR1 expression in NHDF while TGFβ1 decreased its expression, suggesting that TGFβ1 might provide negative feedback for MCHR1 expression since TGFβ1 is induced by PDGF-BB in NHDF ( Figure 6A , arrow #1). BLM is the reagent commonly used to induce SSc-like fibrotic responses in murine and human skin (42). BLM is reported to induce pro-inflammatory cytokines such as TGFβ1 and CTGF in human dermal fibroblasts in vitro (43), suggesting that the combination of several pro-inflammatory mediators, even in the presence of TGFβ1, can increase the expression of MCHR1, consistent with our data in SSc dermal fibroblasts showing high levels of MCHR1. Similar to our findings, Ziogas et al. showed that dextran sodium sulfate (DSS) induces MCHR1 expression and inflammatory colitis via several inflammatory mediators, including TGFβ1, in myofibroblasts (14, 44).

TGFβ is thought to be a master regulator of the fibrotic response and is a prototype fibrotic factor (5). However, PDGF is also important in the pathogenesis of fibrosis. PDGF was initially thought to only promote fibroblast proliferation, but activation of PDGF signaling alone can induce skin and internal organ fibrosis in mice (45). We observed that PDGF-BB induced the fibrotic genes TGFβ1 and CTGF in NHDF ( Figure 6A , arrow #2). The increase in CTGF occurred earlier than TGFβ1, both of which were induced earlier than other fibrotic genes such as Col1α1 and FN1. This led us to speculate that TGFβ1 may mediate, at least in part, the response to PDGF-BB in NHDF. In fact, our data showed that TGFβ1 mediated ECM production in response to PDGF-BB ( Figure 6A , arrows #3,4), but the increase in CTGF and TGFβ1 was independent of TGFβ signaling. Interestingly, we did not see CTGF induction by PDGF-BB in SSc dermal fibroblasts. We speculate this is because SSc fibroblasts produce more CTGF at baseline (46), and thus the amplitude of the response to PDGF stimulation may not be as robust as in NHDF, or because a negative feedback loop is in place in SSc fibroblasts to block further induction of CTGF.

We explored whether the profibrotic effects of PDGF-BB were mediated by MCHR1 for two reasons: 1) PDGF-BB increased MCHR1 levels in NHDF and 2) MCH stimulation alone did not induce the expression of fibrotic genes nor MCHR1. However, previous studies suggested that MCH had profibrotic properties. MCH infusion increased profibrotic genes including TGFβ1 in mouse liver (16), and co-stimulation of MCH with IGF1 or TGFβ1 enhanced cell proliferation rate or collagen production, respectively, in CCD-18Co human myofibroblasts (14). Human microvascular endothelial cells produced MCH following stimulation with Th2 cytokines such as IL4 and IL13 (47). IL4 and IL13 contribute to fibrosis by promoting cell differentiation and collagen production and are found in both serum and lesional skin tissues of SSc patients (48). These studies suggest that IL4 and IL13 can induce MCH, contributing to the pathogenesis of fibrosis in SSc patients. However further investigation will be needed since MCH levels of peripheral tissue or plasma in SSc patients have not been reported, and the contribution of MCH in SSc is unexplored.

The interaction between MCHR1 and PDGF-BB or PDGF receptors has not been reported, however, it is quite possible that PDGF signaling activates MCHR1 since transactivation of GPCR by growth factor receptor-tyrosine kinase has been shown (49, 50). We determined that MCHR1 silencing in NHDF significantly reduced PDGF-BB-induced TGFβ1 and CTGF levels, indicating that PDGF-BB signaling could be modulated by MCHR1. GPCRs are involved in the transmission of PDGF signaling (51), leading us to speculate that MCHR1 may be required to activate downstream signaling of PDGF. We assume that MCHR1 upregulation by PDGF-BB is the result of a positive feedback of PDGF signaling, consistent with other reports showing that a ligand can induce the expression of its receptor (52, 53). Moreover, our western ligand blot and SPR assay results suggest that PDGF-BB directly binds to MCHR1 to activate downstream pathways. MCHR1 then interacts with the Gi/o/q protein and regulates intracellular signaling (8).

In MCHR1 transfected cells, MCH binding to MCHR1 can inhibit the accumulation of cAMP, activate MAP kinase signaling, induce IP3 production, and increase intracellular Ca²⁺ (8). We observed a reduction in cAMP levels following PDGF-BB and MCH treatment in NHDF, and MCHR1 chemical inhibition neutralized cAMP reduction due to both PDGF-BB and MCH, suggesting that PDGF-BB can directly activate the MCHR1 signaling pathway. This data also suggests that MCHR1 inhibition has anti-fibrotic effects by increasing cAMP levels. Elevated intracellular cAMP levels have been shown to exert anti-fibrotic effects, decrease fibroblast proliferation, promote fibroblast cell death, and inhibit ECM production (54). Increased cAMP levels in MDCK cells prevented TGFβ−mediated increases in αSMA levels, suggesting that increased cAMP can inhibit the fibrotic response induced by TGFβ, likely as a result of inhibition of Smad3 activation (54). Increased cAMP levels in lung fibroblasts can inhibit PDGF-BB-induced CTGF and TGFβ1 (55). Based on our findings and those of others, we speculate that increased MCHR1 expression levels might modulate cAMP levels in NHDF and enhance the fibrotic response to PDGF-BB ( Figures 6A, B arrow #5, 6).

Interestingly, Janus kinase-1 (JAK1) and PDGF-receptor-α (PDGFRA) were among the list of 34 genes predicted to be upstream regulators of DE genes regulated by PDGF-BB via MCHR1 [“PDGF-BB vs. VC” in siCtr NHDF] ( Supplementary Table 8 ). JAK1 and PDGFRA are involved in PDGF signaling (56, 57), indicating that MCHR1 might modulate PDGF-BB signaling. In support of this observation, Zigoas et al. showed that the inhibition of MCH/MCHR1 signaling attenuated Smad3 expression levels in mouse fibroblasts of a colitis murine model (14). In contrast, the cAMP signaling pathway was activated by PDGF in several cell types as a result of a negative feedback loop (58, 59). Our observation of decreased cAMP levels could be due to the use of different cell types, PDGF-BB concentrations, and experimental time points.

In experimental colitis, MCHR1 antagonist reduced colonic inflammation, probably by blocking IL10 upregulation, suggesting that inhibition of MCH/MCHR1 signaling could be a novel anti-inflammatory therapeutic approach (60, 61). Anti-fibrotic effects of MCH/MCHR1 inhibition have been reported. Anti-MCH antibody suppressed the production of fibrotic genes in experimental colitis (14), and oral administration of MCHR1 antagonist decreased Col1α1 and TGFβ1 expression levels in a dose-dependent manner in the liver of C57BL/6 J mice with severe hepatic steatosis (16). Previous reports suggest that the anti-fibrotic effects of alpha-melanin stimulating hormone (α-MSH) are actually due to inhibition of MCH/MCHR1 signaling (14, 61), since α-MSH can neutralize the functional effects of MCH (62, 63). The anti-fibrotic effects of α-MSH were reported in NHDF and in a murine model of skin fibrosis induced by TGFβ1 or BLM (64, 65), suggesting that inhibition of MCH/MCHR1 signaling has anti-fibrotic effects in skin.

The functional enrichment analysis of our RNAseq data revealed the involvement of MCHR1 in the fibrotic and inflammatory responses induced by PDGF-BB. Genes and biological processes involved in fibrosis were exclusively enriched by PDGF-BB in NHDF. For example, PDGF-BB decreased ITGB4 expression, a signature that is associated with enhanced lipopolysaccharide-induced inflammation (38). GRPR is the G protein-coupled receptor that binds to gastrin-releasing peptide (GRP). GRP was shown to induce the fibrotic response in a murine model of lung fibrosis and in human cell lines (37, 66), and GRPR antagonism can reverse the effect of GRP on cell proliferation (67), indicating increased GRPR may be involved in the fibrotic response.

GO term analysis revealed some biological processes are deregulated by PDGF-BB via MCHR1. Consistent with previous reports, our data indicated MCHR1 could modulate the immune system (12) and inflammatory responses (15). The biological processes related to biosynthetic, metabolic, and development are also perturbed. This is not surprising as MCHR1 modulates energy metabolism (11), likely in association with leptin (68). The biological process related to ubiquitination may affect SSc pathogenesis by modulating TGFβ signaling and TLR-dependent signaling (69, 70). We also observed that the biological processes related to ECM remodeling and PDGF-BB signaling were perturbed when MCHR1 expression is silenced in NHDF, indicating that silencing of MCHR1 mediated the effects of PDGF-BB in the regulation of these genes, namely PDGFRA, PDGFRB, PDGFD, and MAPK9. Interestingly, we found several biological processes related to vascular development. Vasculopathy is one of the most common features of SSc (1), and MCHR1 has not been previously reported to contribute to angiogenesis or vasculopathy.

In the present study, we observed that PDGF-BB induced cell proliferation of NHDF, in agreement with previous studies (45), whereas MCH did not. In addition, MCHR1 silencing or chemical inhibition did not affect cell proliferation. It is reported that PDGF-BB-induced cell proliferation is suppressed by inhibition of PDGF receptor (71). Taken together, these findings lead us to conclude that PDGF-BB induces cell proliferation in an MCHR1-independent manner. MCH/MCHR1 signaling in cell proliferation is still controversial. In some cell types, MCH/MCHR1 signaling inhibited cell proliferation, e.g., MCH had inhibitory effects on CD3+ lymphocyte proliferation (12) and MCHR1 antagonism induced proliferation of progenitor cells in mouse brain (72). In contrast, MCH induced cell proliferation in DSS-treated human myofibroblasts (14). There are some possible explanations for these conflicting results: MCH/MCHR1 signaling can promote different responses depending on cell type, or cell proliferation rate might depend on the basal expression levels of MCHR1 since DSS treated human myofibroblasts showed higher levels of MCHR1. However, our study has some limitations. Although we found that transcription levels of MCHR1 were upregulated in SSc dermal fibroblasts, we did not examine the role of MCHR1 in vivo. We showed that PDGF-BB induces MCHR1 and confirmed that blocking PDGF-BB signaling by inhibiting PDGF receptor prevented the upregulation of MCHR1, but there is a possibility that MCHR1 is further activated by PDGF-BB-induced molecules or other factors. We also showed that PDGF-BB directly binds to MCHR1 by western ligand blot and SPR, however, we must also consider the possibility that PDGFR and MCHR1 are co-localized and PDGF-BB signals via both receptors simultaneously or sequentially.