Skip to main content
Download PDF

Low dose methotrexate impaired T cell transmigration through down-regulating CXCR4 expression in rheumatoid arthritis (RA)

Arthritis Research & Therapy volume 26, Article number: 173 (2024) Cite this article

Abstract

Background

CXC chemokine CXCL12 is involved in the pathological development of rheumatoid arthritis (RA) through abnormal migration of peripheral immune cells in the joint. Although low dose methotrexate (MTX) is clinically used to treat RA patients, CXCL12 signaling responses to MTX-mediated treatments is still not well understood.

Methods

In this study, we examined the expression of CXCR4 (cognatic receptor for CXCL12) in peripheral T cells from RA patients and arthritis mice models received from low dose MTX therapies. The effects of low dose MTX on CXCR4 were further determined via both in vitro CD3+ T cells and Cxcr4 conditional knockout (CKO) arthritis mice models.

Results

Our clinical data shows that low dose MTX treatment was clinically associated with down-regulated expression of chemokine receptor CXCR4 on patient peripheral T cells. In vitro, low dose MTX significantly decreased cell transmigration through down-regulated CXCR4’s expression in CD3+ T cells. Consistently, CD3+ T cells treated with low dose MTX demonstrated an increased genomic hypermethylation across the promoter region of Cxcr4 gene. Furthermore, our preclinical studies showed that low dose MTX-mediated downregulation of CXCR4 significantly improved the pathological development in mouse arthritis models. Conditional disruption of the Cxcr4 gene in peripheral immune cells potentially alleviated inflammation of joints and lung tissue in the arthritis mice, though genetic modification itself overall did not change their clinical scores of arthritis, except for a significant improvement on day 45 in CXCR4 CKO arthritis mice models during the recovery phase.

Conclusion

Our findings suggest that the effect of low dose MTX treatment could serve to eliminate inflammation in RA patients through impairment of immune cell transmigration mediated by CXCR4.

Introduction

Rheumatoid arthritis (RA) is a complex autoimmune disorder characterized by chronic inflammation of joint synovia and progressive destruction of cartilage and bone [1]. The pathological development of RA is directly related to the abnormal infiltration of T and B lymphocytes, neutrophils, monocytes, and macrophages into synovial tissues, joints, and other organs [2, 3]. It has been widely accepted that infiltrated cells mainly rely on chemokines and their cognate receptors to functionally mediate their migration from circulation to the organs [4,5,6]. However, targeting chemokine signaling has failed to show clinical improvements though RA-related animal models. On the other hand, agents that target chemokines (receptors) have yielded some promising results [7]. Even so, persistent accumulation of lymphocytic cells within the synovial compartment strongly supports that either chemokines or chemokine receptors play significant roles during the pathological development of RA.

CXC motif chemokine ligand 12 (SDF-1/CXCL12) and its receptor CXC motif chemokine receptor 4 (CXCR4) comprise an important chemotaxis involving both immune response and brain development [8]. In RA patients, it was found that CXCL12 enhanced the accumulation of CXCR4-expressing CD4+, CD8+, and Th1 cells within their synovium [9,10,11,12]. Others suggested that synovial TGF-beta 1 (TGFB1) also played an important role to induce CXCR4’s protein expression on CD4+ T cells [11]. Besides lymphocytes, Lenoir et al. (2004) reported that CXCL12 regulated desensitization of human neutrophil respiratory burst in patient synovial fluid [13]. More importantly, some preclinical studies already showed that targeting CXCR4 with the specific inhibitor AMD3100 resulted in efficient decrease of RA-associated pathogenesis in collagen-induced arthritis (CIA) mice [14, 15]. In the same manner, Chung et al. (2010) reported that the incidence of CIA was significantly reduced in Cxcr4flox/flox/Lck-Cre mice compared with Cxcr4+/+/Lck-Cre mice [16]. These findings indicate that CXCL12-CXCR4 is an important chemotaxis responsible for the abnormal migration of peripheral lymphocytes to the arthritis synovium during the pathological development of RA patients.

As an analog of folic acid, methotrexate (4-amino-10-methylfolic acid, MTX) was originally designed as anticancer medication, given as high as 1 gram in a single dose [17]. High-dose methotrexate antagonized the folate pathway by inhibiting dihydrofolate reductase (DHFR) and purine synthesis, thus leading cancer cells to their apoptotic fate [18]. Low dose MTX (7.5–25 mg weekly) currently serves as one of the most popular first line medications to treat RA patients [19]. However, low-dose MTX may involve more complex molecular mechanisms than folate antagonism in the treatment of RA patients [18, 20,21,22]. Indeed, it has been reported that MTX decreased the expression of inflammatory factors such as tumor necrosis factor alpha (TNF-α), interleukin 1 beta (IL-1β), interleukin 6 (IL-6), interleukin 8 (IL-8), C-C motif chemokine ligand 5 (RANTES/CCL5), chemokine (CXC motif) ligand 1 (GRO/CXCL1), and adhesion molecules in RA patients [23,24,25,26,27,28]. Although the presence of upregulated plasma CXCL12 has been suggested in RA patients, the effect of MTX on this chemokine currently remains controversial [29, 30]. In psoriatic patients, immunohistochemical studies in the skin biopsies suggested that decreased CXCL12 was associated with high dose MTX treatment [31]. In RA patients’ synovial tissue, decreased C-C motif chemokine ligand 2 (MCP-1/CCL2) protein has been suggested to be associated with the clinical MTX treatment [24]. Barsig et al. (2005) also reported that MTX was able to significantly suppress cytokine and chemokine release in inflamed joints in a dose-dependent fashion in an adjuvant arthritis-associated rat model [32]. Other studies suggested that MTX potentially enhanced the receptor interleukin 17 receptor A (IL-17R) expression [33] but decreased interleukin 1 receptor type 1 (IL-1r) expression [27]. Some studies showed that inflammatory arthritis is usually associated with global DNA hypomethylation in peripheral blood mononuclear cells (PBMCs), which could be reverted with MTX treatment in inflammatory arthritis [34,35,36].

In the current study, although enhanced CXCR4 expression on peripheral immune cells was observed in RA patients, we report that CXCR4’s expression was decreased after different clinical treatments. The effects of low dose MTX on CXCR4’s expression and functions were further determined. Our study discloses the mechanism by which MTX depends on to target peripheral lymphocytes during treatment of RA patients. These findings thus shed light on potential roles of CXCR4 on peripheral immune cells that were involved in the pathological development of RA.

Materials and methods

Clinical samples

Forty-eight patients with clinically active RA were diagnosed through fulfillment of the 1987 American College of Rheumatology criteria [37] and were recruited from the outpatient clinic of the Rheumatology department of First Affiliated Hospital of Kunming Medical University of Yunnan Province. Written informed consent for the present study’s use of human peripheral blood was obtained from all forty-eight patients and thirty-seven healthy donors, and ethical approval was obtained from both the Ethics Committee of the First Affiliated Hospital of Kunming Medical University and the Committee on Human Subject Research and Ethics of Yunnan University (Approved number: 20190801). All experiments were carried out according to the relevant guidelines.

Cell sorting and cell culture

For the isolation of PBMCs, the heparinized whole blood samples (4 mL) from each RA patient and healthy donor were diluted with an equal volume of PBS (Biosharp Life Sciences, China), layered onto Ficoll-Hypaque (Solarbio Life Sciences, China) and centrifuged at 2,000 rpm/min, at 20 °C for 20 min by density gradient centrifugation. Mononuclear cells were recovered from the interface and washed twice with PBS. The CD3+ T cells were purified from about 2–10 × 108 PBMCs, which were passaged over a 30 μm nylon wool column, incubated using immunomagnetic microbeads (Miltenyi Biotec, Germany) for 30 min at 4 °C, and finally selected with QuadroMACS™ Separator (Miltenyi Biotec, Germany) via magnetic LS cell separation columns (Miltenyi Biotec, Germany). All assays were performed according to the protocols recommended by the manufacturer. Aliquots of 4–10 × 106 PBMCs and purified CD3+ T cells were optionally stimulated with anti-CD3 (clone UCHT1, 2.5 µg/ml) and anti-CD28 (clone CD28.2, 1.25 µg/ml) antibodies (BD Bioscience, USA) for 48 h in the presence of MTX does (0.001–1 µg/mL or 0.01 µg/mL) or AMD3100 (1 µg/mL) in vitro. Jurkat cells were offered by Professor Shenghong Li from the Insititute of Botany, the Chinese Academy of Sciences, Kunming, China. The PBMCs, purified CD3+ T cells and Jurkat cells were cultured in flat-bottom plates (Corning, USA) in RPMI-1640 medium (Gibco, Invitrogen Corporation, USA) supplemented with 10% fetal calf serum (Viva Cell Biosciences, China), 100 U/mL penicillin and 100 µg/mL streptomycin (BasalMedia Technologies, China) at 37 °C under 5% CO2.

Flow cytometry (FCM)

Expression of chemokine receptor CXCR4 on human T cells was analyzed using flow cytometry. 100 µL of human PBMCs (about 5 × 105 cells) were incubated with mouse anti-human antibodies, including anti-CD184-PE and anti-CD3-FITC (20 µL, BD Biosciences, USA), anti- CD4- and CD8-FITC (5 µL, BioLegend, USA), and their corresponding mouse isotypes as the negative controls. Then, the cells were incubated for 30 min at 4 °C in the dark. Subsequently, the cells were washed twice with 1 mL FACS buffer (1% BSA in PBS) by centrifugation (400 g/min, 5 minutes, 4 °C) to remove unbound antibodies. Cells were then resuspended in 500 µL PBS and sieved through a 40-µm pore size mesh to generate single-cell suspensions. Finally, labeled cells were measured on an Accuri C6 Flow Cytometer (Accuri Cytometers, BD Biosciences, C6, USA) and analyzed by FlowJo software (Treestar FlowJo v10.8.1, Ashland, USA). Negative fluorescence was gated with FITC-conjugated IgG or PE-conjugated IgG. At least 10,000 events collected within the lymphocyte gate. The single fluorochrome-stained cells were used for instrument compensation and the photomultiplier tube (PMT) setup. Analysis of CD3+, CD4+, CD8+ T cell subsets were performed on the total CD3+ T cells within the lymphocyte gate.

Quantitative real-time polymerase chain reaction (qRT-PCR)

Total messenger RNA (mRNA) was extracted using RNAiso Plus Reagent (Clontech Laboratories, Takara Bio Group, Japan) according to the manufacturer’s instructions. Potential genomic DNA contamination was removed from the samples and complement DNA (cDNA) was synthesized from 1 µg total RNA per sample using random primers with PrimeScript™ RT reagent Kit with gDNA Eraser (Clontech Laboratories, Takara Bio Group, Japan) according to the recommended protocol. The qRT-PCR with up to 100 ng cDNA was amplified using the SYBR® Premix Ex Taq™ (Tli RNaseH Plus, Clontech Laboratories, Takara Bio Group, Japan). The ABI Prism 7500 Sequence Detection System (Applied Biosystems, ABI 7500, USA) was used to measure the amplified PCR product. The qRT-PCR thermal cycle included 1 cycle of 95 °C for 10 s, followed by a two-step PCR program: 40 cycles of 95 °C for 5 s, and 60 °C for 34 s, followed by a melt curve analysis. Each sample was performed in triplicate (technical replicates) and repeated at least three times independently. The oligonucleotide primers used for qRT-PCR analyses were listed in Table S2. The relative expression of qRT-PCR data was determined using the 2−∆∆Ct method to compare mRNA expression levels of the target gene with the housekeeping gene (β-actin or 18 S).

CXCR4 methylation-specific PCR analysis based on nucleotide sequencing

The Jurkat cells, or the 0.01 µg/mL MTX treated CD3+ T cells, genomic DNA was extracted by DNA mini isolation kits (Qiagen, Germany) according to the manufacturer’s protocol. Genomic DNA was bisulfite converted with the Zymo EZ DNA methylation kits (Zymo Research, Irvine, CA, USA), and the CpG sites of the bisulfite modified DNA were amplified with bisulfite specific PCR consisted of an initial denaturation step at 95 °C for 5 min, followed by 38 cycles of denaturation at 95 °C for 30 s, annealing at 54 °C for 30 s, extension at 72 °C for 30 s, and final extension step at 72 °C for 7 min by the EpiArt HS Taq Master Mix (Vazyme Biotech, Nanjing, China). The PCR products were verified on 2% agarose gel, followed with sequencing by Sangon Biotech Co., Ltd. (Shanghai, China). For each CpG site, the methylation ratio was calculated as the number of cytosine reads divided by the sum of cytosine and thymine reads [38]. Bisulfite PCR primer sequences were designed by Methyl Primer Express Software v1.0 (Applied Biosystems, USA) and listed in Table S2.

In vitro transmigration

The interaction between CXCR4 and extracellular matrix (ECM, Sigma-Aldrich, USA) assays were conducted on sorting cells with 0.01 µg/mL MTX (Sigma-Aldrich, USA) and CXCR4-specific antagonist 1 µg/mL AMD3100 (Sigma-Aldrich, USA) treatment, using NeuroProbe 48-Well Micro Chemotaxis Chambers (Neuro Probe, Inc. Ap48, USA). Briefly, the Chemotaxis Chambers were separated using 5 μm pore polycarbonate transwell filter membranes (Neuro Probe, Inc. USA) and the lower side was coated with 0.01 mg/mL ECM diluted with RPMI-1640 culture media. In the lower wells, the 100 ng/mL human CXCL12 (SDF-1α) protein (Miltenyi Biotec, Germany) was supplemented as the chemoattractant. And then, 1 × 104 mock or pre-treated cells suspended in 50 µL RPMI-1640 culture media were added into each upper chamber. After 4 hours of incubation at 37 °C and 5% CO2, the membrane was carefully removed. Cells on the upper membrane surface were gently scraped with a cotton swab. The membranes were fixed in 4% (w/v) paraformaldehyde (PFA, Servicebio Technology, China) for 30 min and washed in PBS before staining with 0.1% Hoechst 33342 (Sigma-Aldrich, USA) for 5 min at room temperature, followed by rinsing three times with PBS. The membranes were mounted on glass slides with Fluoro-Gel (Electron Microscopy Sciences, USA) and cells per unit area in triplicate were determined by counting four representative microscopic fields under an epifluorescence microscope at a magnification of ×10 (Leica, DMi8, Germany).

Cell viability assay

To examine the toxicity of low-dose MTX on the activated T cells, PBMCs were seeded into 96-well flat bottom plates and stimulated with anti-CD3 (clone UCHT1, 2.5 µg/ml) and anti-CD28 (clone CD28.2, 1.25 µg/ml) antibodies (BD Bioscience, USA) for 0, 24 and 48 h in PBS or MTX (0.001–1 µg/mL) at the density of 5 × 104 cells per well (100 µL) in quadruplicate. The same volume of PBS was kept consistent with the 0.001–1 µg/mL MTX group added in the vehicle control group for 0, 24 and 48 h, respectively. The effects of MTX on the survival of cells was evaluated with CellTiter 96® Aqueous One Solution Reagent Solution Cell Proliferation Assay kit (Promega, USA), following the manufacturer instructions. Briefly, at 2 h before each of the desired time points, each well was mixed with 20 µL of MTS ((3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2 H-tetrazolium, solution inner salt, and cells were incubated at 37 °C with 5% CO2 for 2 h. The absorbances of the samples were detected at a wavelength of 490 nm using a micro-plate reader (Molecular Devices, SpectraMax 190, USA).

Enzyme-linked immunosorbent assay (ELISA)

The clinical sample blood serums were collected, and enzyme-linked immunosorbent assay (ELISA) kits for detecting human cytokines IL-1, IL-2, IL-6, IL-8, TNF-α (Elsbio Biotec, China) and CXCL12 were obtained (Solarbio Life Sciences, China). ELISA experiments were conducted following the manufacturers’ instructions. Plates were read by a micro-plate reader (Molecular Devices, SpectraMax 190, USA) at a wavelength of 450 nm.

In vivo MTX and AMD3100 treatment

For in vivo assessment of the effect of MTX on the treatment of RA, we used adjuvant-induced arthritis (AA) mouse models that were widely used for drug validation, in particular for T cell-related pathways leading to arthritis [39]. Briefly, thirty-two 10-week-old female C57BL/6J mice were purchased from Vital River Laboratories (Beijing, China) to be used as the experimental model of arthritis. They were kept in healthy conditions with clean air and given plenty of food and water. To avoid superimposed side effects of MTX/AMD3100 treatment and adjuvant immunization on the growth and body weight of arthritis mice, a preventative model with MTX and AMD3100 treatments began on day 1 and lasted 15 days. Briefly, the mice were divided into four groups (eight mice per group). The non-arthritic healthy control mice and untreated arthritis mice received 150 µl normal saline (NS) every three days for 15 days (5 times total) and were then divided into two groups: namely non-arthritic healthy control mice were fed normally, untreated arthritis mice were induced by intradermal tail injection with 150 µl of 4 mg/mL heat-killed Mycobacterium tuberculosis suspended in complete Freund’s adjuvant (CFA) (day16, Sigma Aldrich, USA). The MTX or AMD3100 treatment of them established arthritis group was fed with 2.5 mg/kg MTX or 5 mg/kg AMD3100 from day 1 to day 15 every three days, and arthritis was subsequently induced with the same regimen. Mice were weighed and scored for clinical signs of arthritis once every two days from day 17 to day 48 post-immunization. On day 20, 27, and 38, blood was collected and analyzed for CXCR4+PBMCs. All animal studies and relevant procedures were reviewed and approved by the ethics committee of the institutional animal care and use committee, Yunnan University (IACUC, YNU) (Approved number: 20190801).

Collagen induced arthritis (CIA) mouse model

The type II collagen induced arthritis (CIA) mouse models, which were extensively used to identify potential pathogenic mechanisms of autoimmunity, including the role of individual cell types in disease onset and progression [40], were induced and the effect of CXCR4 on the occurrence and development of RA was assessed. Briefly, 10-week-old female C57BL/6J mice were injected intradermally at the base of the tail with 15 mg/kg chicken type II collagen (CII, Chondrex, USA) dissolved in 10 mM acetic acid and emulsified in an equal volume of CFA (Sigma Aldrich, USA) containing 4 mg/mL Mycobacterium tuberculosis to induce the arthritis mice model (day 0). Twenty-one days (day 21) after the first primary immunization, the mice were given a single booster injection in the same manner. The arthritis index assay, blood collection, and histological assessment were carried out. The expression of TNFα, IL-1β and IL-6 in lung tissue was detected by qRT-PCR and the respective primers used were listed in Table S2.

CXCR4 conditional knockout

Mx1-Cre−/CXCR4flox/flox mice (C57BL/6J) were purchased from the Jackson laboratory (The Jackson laboratory, USA). In these mice, two LoxP sites were genetically inserted in the side of the second exon of the Cxcr4 gene. LoxP site was genotyped using genotyping PCR primers MG-Loxp-F and MG-Loxp-R. A 577-bp and 481-bp DNA band was expected from either flox homozygous mice or wild type/flox heterozygous mice respectively. Meanwhile, Mx1-cre+/CXCR4+/+ mice (C57BL/6J) were purchased from Nanjing Biomedical Research Institute of Nanjing University (Nanjing, China). These mice overexpressed the Cre protein under control of the inducible Mx1 promoter. Mx1-Cre+ was genotyped using genotyping primers MG-Cre-F and MG-Cre-R, and a 408-bp PCR DNA band was expected as well. The Mx1-Cre−/CXCR4flox/flox mice and Mx1-Cre+/CXCR4+/+ mice were founders to be crossbred to obtain Mx1-Cre+/CXCRflox/flox mice. To obtain CXCR4 conditional knockout mice, Mx1-Cre+/CXCR4flox/flox mice were administrated 12.5 mg/kg polyinosinic-polycytidylic acid (PI: PC, Novus Biologicals, USA) by intraperitoneal injection. PI: PC injections were performed every two days, for a total of 3 times. Genetic deletion of Cxcr4 allele genes was verified using genotyping primers MG-CXCR4-F and MG-CXCR4-R. CXCR4+PBMCs and CXCR4+CD3+ T cells on mouse PBMCs were confirmed by flow cytometry. All the primers used were listed in Table S2. The CIA mice model of Cxcr4 conditional knockout was induced as above (Collagen induced arthritis mouse model) mentioned. In brief, 10-week-old female Mx1-Cre+/Cxcr4flox/flox mice were immunized intradermally at the base of the tail with 15 mg/kg chicken type II collagen (CII, Chondrex, USA) dissolved in 10 mM acetic acid and emulsified in an equal volume of CFA (Sigma Aldrich, USA) containing 4 mg/mL Mycobacterium tuberculosis (day 0) after the PI: PC was injected at day -2, day -4, and day -6 respectively. On day 21, the mice were given a single booster injection with the same manner. The arthritis index assay, blood collection, histological assessment, and the expression of TNFα, IL-1β and IL-6 in lung tissue were carried out.

Histological assessments of arthritis

At the end of the mice experimental procedure, the knee joints and paws were analyzed for histological assessment after the mice were sacrificed. Animals were perfused with 4% phosphate-buffered paraformaldehyde (PFA). Then, animal joints, paws and lungs were removed, dissected, and sliced into 5-µm cross-sections. Sections were stained by hematoxylin and eosin (H&E) (Beyotime Life Sciences, China) according to standard procedures. The severity of arthritic changes, such as articular cartilage destruction and bone erosion were histologically scored as described previously [41]. Briefly, we scored cartilage destruction from 0 to 3 and bone erosion from 0 to 5. The positive inflammatory areas of mouse lung sections were measured on each slide and quantified using ImageJ software [42].

Statistical analysis

All reported values were expressed as mean ± SEM (Standard Error of Mean), unless otherwise stated. The arthritis mice and MTX and AMD3100 treated experimental groups included at least five mice. Statistical analyses were done with SPSS 22.0 computer software (SPSS Inc, Chicago, USA). The statistical significance between healthy controls and patient, sham and MTX treated mice, sham and arthritis mice were analyzed using unpaired Student t-test. The statistical significance between two groups of qRT-PCR, in vitro transmigration and cell viability assay were analyzed with a paired Student’s t-tests. In all statistical analyses, the probability (P) values less than 0.05 were statistically significant.

Results

CXCR4-expressing peripheral cells was increased in RA patients

From 48 patients (Table 1) with longstanding RA, the CXCR4-expressing (CXCR4+PBMCs) was analyzed by flow cytometry. As shown in the Fig. S1D and Table S1, about 12.35 ± 1.57% of CXCR4+PBMCs were detected in RA patients. However, only 3.48 ± 0.25% PBMCs in the heathy control were found to express CXCR4. Similar observations were obtained when examining CXCR4’s protein expression on patients’ CD3+ T cells. About 7.11 ± 1.15% of CXCR4+CD3+ T cells were detected in RA patients, while only 1.81 ± 0.13% of CXCR4+CD3+ T cells were observed in the heathy control (Fig. S1C and S1E). Considering the averaged CD3+ T cell (Fig. S1B and S1G) and PBMCs cell numbers (Fig. S1A and S1F) from both groups were almost the same, it was concluded that CXCR4’s expression was significantly increased in the RA patients compared with the healthy controls. The results showed a significant increase in the expression of CXCR4 on PBMCs and CD3+ T cells, with an averaged 3.55-fold change (P < 0.001) and 3.93-fold change (P < 0.001) respectively between RA patients and the healthy control. We also found that the frequency of CXCR4+CD3+ T cells in RA exhibited a significantly positive correlation with the patients’ clinical scores of DAS28/ESR, SDAI, TJC in 28 joints, PtGA and PhGA (Fig. S2A, C, D, F, G, P < 0.05 or 0.001), and a moderate positive correlation with DAS28/CRP (Fig. S2B, P = 0.091) and almost no correlations with SJC in 28 joints (Fig. S2E, P = 0.423). These results reveal that CXCR4 on CD3+ T cells is an activating factor associated with RA disease activity.

Table 1 Summary of RA patients and healthy controls
Full size table

The serum protein level of inflammation cytokines from RA patients was further analyzed by ELISA. Interestingly, we found that protein concentrations of cytokines IL-1, IL-2, IL-8, and TNFα were upregulated in RA compared with the healthy control (Fig. S3A-D). The protein concentration of IL-6 was increased in RA, but there was no statistical significance (Fig. S3E). Consistent with previous results [30], we observed serum chemokine CXCL12 was significantly increased by 68.40 ± 1.54% (P = 0.004) in the RA patients as compared to the healthy control (Fig. S3F). Our results showed an increased serum level of inflammatory and pro-inflammatory factors in RA patients. These observations suggested that either CXCL12’s serum level or CXCR4’s expression on CD3+ T cells was potentially associated with the enhanced inflammatory environment found during the pathological development of RA.

CXCR4-expressing peripheral CD3+ T cells was decreased in RA patients after clinical treatments

To analyze the effects of clinical treatments on RA patients, the same 48 RA patients were further regrouped into three different subgroups (Table 2): (1) RA-Untreated (RA patients without clinical treatments, n = 9); (2) RA + MTX (RA patients clinically treated with MTX, n = 15); (3) RA-MTX (RA patients clinically treated without MTX, n = 24). As shown in Fig. 1C and E, about 15.98 ± 3.42% of CXCR4+CD3+ T cells were detected in RA-Untreated patients. Meanwhile, 1.81 ± 0.13% of CXCR4+CD3+ T cells were detected from the healthy control, consistently demonstrating an 8.8-fold (P < 0.001) increase of CXCR4’s expression on CD3+ T cells in RA-Untreated as compared to the healthy control. Furthermore, we noticed that only 5.67 ± 1.43% of CXCR4+CD3+ T cells were detected from RA-MTX patients and 4.08 ± 0.21% CXCR4+CD3+ T cells in RA + MTX patients. Compared to RA-Untreated patients, expression of CXCR4 on CD3+ T cells decreased significantly by a 2.8-fold change (P < 0.001) and 3.9-fold change (P < 0.001) in RA-MTX and RA + MTX patients respectively. CXCR4+PBMCs was also observed to be decreased in RA patients after different treatments (Fig. 1D). However, both CXCR4+PBMCs and CXCR4+CD3+ T cells was not different between the RA + MTX and RA-MTX treatment groups (Fig. 1D-E). The total number of PBMCs between RA-Untreated and RA + MTX patient groups was not changed, while the total number of CD3+ T cells was increased between RA-Untreated and RA + MTX patient groups (Fig. 1A-B, F-G). We further regrouped the RA-MTX group into GC monotherapy, DMARDs and biology agent, DMARDs and prednisone, DMARDs and biology agent and prednisone and others (leflunomide monotherapy, hydroxychloroquine monotherapy) groups according to their clinical treatment. We found that the proportions of CXCR4+PBMCs (Fig. S4A), CXCR4+CD3+ cells (Fig. S4B) were decreased in these groups, with an exception for the other group due to it including only three samples. PBMCs (Fig. S4C) and CD3+ T cells (Fig. S4D) overall did not significantly change between RA-Untreated and the various RA-MTX clinic treatment groups. Patient’s therapeutic outcomes were further evaluated according to the clinical diagnostic index (Fig. S5A-G). We found that patients’ clinic scores of DAS28/ESR, DAS28/CRP, SDAI, TJC in 28 joints, PtGA, and PhGA showed significantly decrease after treatment with MTX and non-MTX drugs. Thus, these findings indicated that downregulation of CXCR4’s expression in RA patients was associated with clinical treatments, including MTX.

Table 2 Clinical characteristics of different RA groups (n = 48). Data are mean ± SEM
Full size table
Fig. 1
figure 1

Low dose MTX-treatment was clinically downregulated the frequency of CXCR4+ immune cells in RA patients. (A-C) Representative flow charts demonstrating CXCR4+ and CD3+ T cells from both RA patients and healthy control. (A) Gating of human PBMCs. (B) CD3+ T cells (stained with anti-human CD3 antibody, red) and Isotype controls (black). (C) Double staining with both anti-human CXCR4 and anti-human CD3 antibodies. Quantification of CXCR4+PBMCs (D) and CXCR4+CD3+ T cells (E) from A-C. **p < 0.01, ***p < 0.001 and error bars indicate SEM. Quantification of PBMCs (F) and CD3+ T cells (G) from A-C. Healthy, (Normal control group, n = 37); RA-Untreated (RA patients without clinical treatments, n = 9); RA + MTX (RA patients treated with low dose MTX, n = 15); RA-MTX (RA patients treated without low dose MTX, n = 24)

Full size image

Low dose MTX downregulates CXCR4’s expression on lymphocytes

The use of low dose MTX has served as one of the most efficient drug treatments for RA patients in the clinic. Decreased CXCR4’s expression in patients following MTX treatment indicated that low dose MTX could downregulate the expression of CXCR4. We tested the effects of 0.001–1 µg/mL MTX on the cell survival of human PBMCs. As shown in Fig. S6A-D, we found that 0.001 and 0.01 µg/mL MTX concentrations did not inhibit PBMCs survival in vitro, while the MTX doses between 0.1 and 1 µg/mL showed toxic effects on human PBMCs within 48 h. Hence, we chose 0.01 µg/mL MTX treat cells in the following experiments.

PBMCs and CD3+ T cells from healthy subjects were optionally stimulated with anti-CD3 and anti-CD28 antibodies for 48 h in the absence or presence of 0.01 µg/mL MTX in vitro. In the non-stimulated cells, CXCR4+PBMCs and CXCR4+CD3+ T cells were significantly decreased by 35.82 ± 2.65% (P < 0.05) and 40.48 ± 5.65% (P < 0.05) respectively after treatment with 0.01 g/mL MTX (Fig. 2A). Treatment with 0.01 µg/mL MTX in vitro did not result in significant changes of PBMCs and CD3+ T cell numbers between vehicle and treatment groups (Fig. 2B). The qRT-PCR consistently showed that 0.01 µg/mL MTX-treatments decreased CXCR4 mRNA levels in both PBMCs and CD3+ T cells by 70.75 ± 1.30% (P < 0.001) and 71.68 ± 3.33% (P < 0.01) (Fig. 2C). We also noticed that CXCR4 mRNA levels decreased by 54.40 ± 5.66% (P < 0.01) in 0.01 µg/mL MTX treated Jurkat cells (Fig. 2D), and increased DNA methylation in the -186 to -19 bp region of CXCR4 promoters in Jurkat cell after 0.01 µg/mL MTX treatment (Fig. 2E) for 48 h. Similarly, in the stimulated cells, CXCR4+PBMCs and CXCR4+CD3+ T cells were significantly decreased by 23.32 ± 1.80% (P < 0.05) and 28.44 ± 3.47% (P < 0.05), (Fig. 2F), and the PBMCs and CD3+ T cells numbers showed no significant changes respectively after treatment with 0.01 µg/mL MTX (Fig. 2G) for 48 h. We also detected CXCR4+CD4+ T cells and CXCR4+CD8+ T cells treated with 0.01 µg/mL MTX under activation states. The results showed that CXCR4+CD4+ T cells (Fig. S7B) and CXCR4+CD8+ T cells (Fig. S7D) were significantly decreased (P < 0.05), and the percentage of CD4+ T cells (Fig. S7A) and CD8+ T cells (Fig. S7C) showed no significant changes, suggesting that the decrease of CXCR4+CD3+ T cells may be partly due to the decrease of CXCR4+CD4+ T cells and CXCR4+CD8+ T cell treatment with 0.01 µg/mL MTX. Furthermore, CXCR4 mRNA levels decreased by 74.00 ± 4.36% (P < 0.001) and 62.46 ± 5.45% (P < 0.01), respectively (Fig. 2H) in PBMCs and CD3+ T cells, and DNA methylation increased in the -186 to -19 bp region of CXCR4 promoters in the activated CD3+ T cells treated with 0.01 µg/mL MTX. Taken together, these results suggest that the decrease in CXCR4 mRNA levels are partly due to increased DNA methylation of its promoter after 0.01 µg/mL MTX treatment.

Fig. 2
figure 2

Low dose MTX downregulated CXCR4’s expression in normal human peripheral immune cells and mouse. (A-C) PBMCs from normal humans were non-stimulated with anti-CD3 and anti-CD28 for 48 h in the absence or presence of 0.01 µg/mL low dose MTX in vitro. (A) Quantification of human CXCR4+PBMCs and CXCR4+CD3+ T cells (non-stimulated). (B) Quantification of human PBMCs and CD3+ T cells (non-stimulated). (C-D) The qRT-PCR showing CXCR4’s mRNA levels in human PBMCs and CD3+ T cells (non-stimulated), Jurkat cells. (E) DNA methylation levels of CXCR4 promoter (-186 to -19 bp) in Jurkat cells after treatment with 0.01 µg/mL low dose MTX in vitro. (F-I) PBMCs from normal humans were stimulated with anti-CD3 and anti-CD28 for 48 h in the absence or presence of 0.01 µg/mL low dose MTX in vitro. (F-G) Quantification of human CXCR4+/PBMCs and CXCR4+/CD3+, PBMCs and CD3+ T cells (stimulated). (H) CXCR4’s mRNA levels in human PBMCs and CD3+ T cells (stimulated). (I) DNA methylation levels of CXCR4 promoter (-186 to -19 bp) in CD3+ T cells (stimulated). Values on graphs represent mean ± SEM, *p < 0.05, **p < 0.01, ***p < 0.001. (J-L) Representative flow charts showing CXCR4+CD3+ T cells after treatment with 2.5 mg/Kg MTX in vivo (see the details in Fig. S8). (J) Gating of mouse PBMCs. (K) Mouse CD3+ T cells (stained with anti-mouse CD3 antibody, red, and Isotype controls, black). (L) Double staining with both anti-mouse CXCR4 and anti-mouse CD3 antibodies. (M) Quantification of CXCR4+CD3+ mouse T cells (left) and CXCR4-expressing mouse PBMCs (right) from J-L. (N) Quantification of mouse PBMCs (left) and mouse CD3+ T cells (right) from J-L. (M-N) Data represented as mean ± SEM (n = 3 normal mice, **p < 0.01, ***p < 0.001)

Full size image

We further tested the effect of low dose MTX on CXCR4’s expression on PBMCs in normal mice. To address this question, healthy C57BL/6J mice were treated with the low dose of 2.5 mg/Kg MTX via gastric infusion (GI) three times per week, for a total of two weeks (Fig. S8A). The general animal survival (Fig. S8B) and animal weights (Fig. S8C) did not show any direct impact by low dose MTX. However, we observed low dose MTX significantly decreased the proportion of CXCR4+PBMCs by 56.39 ± 1.27% (P < 0.001) at day 7 and by 50.11 ± 3.82% (P < 0.01) at day 14 (Fig. 2M-right). In the same manner, we found that CXCR4+CD3+ T cells was decreased by 39.07 ± 7.72% (P < 0.01) at day 14 due to the administration of low dose MTX (Fig. 2L and M-left). Similarly, the numbers of PBMCs (Fig. 2J and N-left) and CD3+ T cells (Fig. 2K and N-right) cells did not show any significant changes between the vehicle and low does MTX treated mice. Here, we have provided evidence from both in vitro and in vivo studies to support the direct downregulating the expression of CXCR4 in peripheral immune cells by low dose MTX treatment.

Low dose MTX inhibits the transmigration of lymphocytes by downregulating CXCR4 expression

Due to CXCR4 widely functionally regulates the cell transmigration, hence, downregulation of CXCR4 by MTX suggests that low dose MTX may functionally regulate the transmigration of peripheral lymphocytes. To address this question, 100 ng/mL CXCL12 was applied to induce in vitro transmigration of activated healthy human PBMCs, followed by treating with 0.001–1 µg/mL MTX (Fig. 3A). Compared to the PBS control, we found that 0.01–1 µg/mL MTX decreased PBMCs’ transmigration by approximately 35.29 ± 5.20% (P < 0.05) to 22.27 ± 3.30% (P < 0.05), indicating that 0.01–1 µg/mL MTX could efficiently inhibit the in vitro transmigration of human PBMCs. To verify the specificity of CXCL12’s induction, we further applied a CXCR4-specific inhibitor AMD3100 to activated PBMCs’ transmigration and observed dose-dependent inhibition of CXCL12-induced transmigration by AMD3100 (0.001–1 µg/mL) (Fig. 3B). These findings suggested that low dose MTX may regulate the transmigration of immune cells through downregulating the CXCR4’s expression. Given that immune cells from RA patients expressed higher CXCR4, we further tested whether patients’ immune cells contained more transmigration potency. Compared to the healthy control, we found that CXCL12-induced transmigration from both PBMCs and CD3+ T cells were significantly increased by 57.40 ± 9.88% (P < 0.05) and 37.40 ± 0.94% (P < 0.01) respectively (Fig. 3C). More importantly, we observed a synergistic inhibitory effect with 0.01 µg/mL MTX and 1 µg/mL AMD3100 combined treatment on activated PBMCs and CD3+ T cells isolated from healthy people (Fig. 3D-E) and RA patients (Fig. 3F-G). In summary, our results supported the inhibitory effect of low dose MTX-treatment on the transmigration of peripheral lymphocytes through downregulation of their CXCR4 expression.

Fig. 3
figure 3

Low dose MTX impaired CXCL12-induced CD3+ T cell transmigration in vitro. (A-B) In vitro CXCL12-induced activated PBMC transmigration was blocked by MTX (A) and AMD3100 (B). (A-B) Added CXCL12 and Vehicle (100 ng/mL recombinant CXCL12-induced activated PBMCs treated with control vehicle), and added CXCL12 and MTX/AMD3100 (100 ng/mL recombinant CXCL12-induced activated PBMCs treated with 0.001 to 1 µg/mL MTX/AMD3100), and n = 3 individual experiments. (C) PBMCs isolated from healthy and RA patients demonstrating different transmigration capacity (n = 3 patients or healthy controls). (D-G) Combination of 0.01 µg/mL MTX and 1 µg/mL AMD3100 showing a synergistic inhibition for healthy controls and RA patients PBMCs and CD3+ T cells, n = 3 patients or healthy controls. (A-G) *P < 0.05, **P < 0.01, ***p < 0.001. Data represented as mean ± SEM

Full size image

MTX-mediated downregulation of CXCR4 to the treatment of AA mouse

Our studies suggested that downregulation of CXCR4 caused by low dose MTX may contribute to the effective clinical treatment of RA patients. To address this question, an AA mouse model was used to test the treatment effects from low dose MTX and AMD3100 (Fig. 4A). Consistent with previous studies [43], the AA mice developed typical severity scores of arthritic limbs (Fig. 4C). C57BL/6J wild type mice were preventatively administrated the doses of either 2.5 mg/Kg MTX or 5 mg/Kg AMD3100 per time for two weeks total treatment (5 times) (Fig. 4A). At day 15, mice were immunized with 150 µl CFA containing 4 mg/mL heat-killed Mycobacterium tuberculosis. Intradermal injection was conducted at three sites near the base of the tail according to the protocol from Brand et al. [40]. AA model mice were treated with low dose MTX or AMD3100 and 100% survival was observed at day 48 and the body weight were not affected between the four group mice (Fig. 4B, D).

Fig. 4
figure 4

Downregulation of the frequency of CXCR4+ peripheral immune cells were associated with low dose MTX-mediated treatments in AA mice model. (A) Experimental design where wild type (WT) mice were pre-treated with 2.5 mg/Kg MTX or 5 mg/Kg AMD3100 before immunization. (B-D) Clinical characteristics of AA-related mice from A. (B) Survival, (C) Arthritic clinical scores and severity, (D) Weight. (E-F) Flow cytometry analysis of CXCR4-expressing mouse PBMCs (n = 3 mice). CXCR4+PBMCs (E) and PBMCs (F). (C-F) Sham (Mice without immunization), AA Untreated (Mice with immunization), AA + MTX (Mice with immunization after pretreating with 2.5 mg/Kg MTX), AA + AMD3100 (Mice with immunization after pretreating with 5 mg/Kg AMD3100). *P < 0.05, **P < 0.01, ***P < 0.001, and data represented as mean ± SEM

Full size image

At the pathological “peak” stage (day 25 to day 37), we observed that severity scores of arthritic limbs were significantly decreased after treatment with low dose MTX (Fig. 4C). For example, compared to untreated mice, clinical scores on day 31 had decreased by a maximum 8.7-fold change (P < 0.001) after treatment with low dose MTX. However, no obvious clinical differences were observed from the pathological stages of “onset” (day 17–25). The effects of low dose MTX on CXCR4+PBMCs were further examined at day 20 (“onset” stage), day 27 (“peak” stage), and day 38 (“recovery”, day 38–48, stage). CXCR4+PBMCs at day 20 was decreased by 33.50 ± 5.88% (P < 0.05) after treatment with low dose MTX compared with untreated AA mice (Fig. 4E). Because their total cells numbers were not changed (Fig. 4F), these observations suggested early preventatively effects from low dose MTX during the treatments. Conversely, we found that enhanced CXCR4’s expression in untreated AA mice mainly appeared at later timepoints of day 27 compared with the Sham’s (Sham versus AA Untreated). From the above observations, our results suggested that low dose MTX-mediated treatments may possibly rely on the suppression of CXCR4’s function.

Furthermore, we observed that AMD3100 can efficiently treat AA mice (Fig. 4C). Compared with untreated AA mice, clinical scores at day 31 were decreased by a maximum 3.84-fold change (P < 0.001) after treatment with AMD3100. The effect of AMD3100 was most significantly detected between day 25 to day 37 as well (AA Untreated versus AA + AMD3100). However, AMD3100 did not directly affect CXCR4’s expression, but resulted in an abnormal accumulation of peripheral CXCR4-expressing cells. For instance, compared with untreated arthritis model mice, we noticed that the number of CXCR4-expressing cells increased by a 2.31-fold change (P < 0.01) after AMD3100 treatments at day 20 (Fig. 4E, AA Untreated versus AA + AMD3100). Our findings suggest that both low dose MTX and CXCR4-specific inhibitor AMD3100 may share a similar mechanism to suppress immune cell migration during the treatment of arthritis mice.

Conditional knockout (CKO) of CXCR4 in peripheral immune cells in mouse

To explore the potential roles of CXCR4 in the arthritis mice model, we generated CXCR4 CKO (CXCR4−/−) mice in which floxed Cxcr4 gene was disrupted through the inducible Cre in peripheral blood cells. To accomplish this goal, Mx1-Cre transgenic mice (Mx1-Cre+/CXCR4+/+) and Mx1-Cre−/CXCR4flox/flox mice both have a C57BL/6J background were founders to be crossbred to obtain Mx1-Cre+/CXCR4flox/flox female mice (Fig. 5A). Between exon1 and exon2 of the Cxcr4 gene, genotype primers were designed to amplify a 481-bp DNA for wild type (WT) Cxcr4 allele and a 577-bp DNA floxed Cxcr4 allele. In addition, primers to amplify a 408-bp DNA of Cre were designed to detect exogenous Cre gene. We detected a 577-bp of floxed Cxcr4 band and a 408-bp of Cre band in the same mouse, supporting the successful generation of Mx1-Cre+/CXCR4flox/flox mice (Fig. 5B). To disrupt the Cxcr4 gene, Mx1-Cre+/CXCR4flox/flox female mice were administrated with 12.5 mg/Kg PI: PC to induce Cre’s expression through its inducible Mx1 promoter and generated a 569-bp Cxcr4 genome DNA fragment (Fig. 5B-D). In addition, mice with different genotypes of either WT (Mx1-cre−/CXCR4+/+), Mx1-Cre+ (Mx1-cre+/CXCR4+/+), or CXCR4flox/flox (Mx1-Cre−/CXCR4flox/flox) were injected by 12.5 mg/Kg PI: PC, and PI: PC did not affect their survival (Fig. 5E).

Fig. 5
figure 5

Conditional deletion of Cxcr4 in mouse’s PBMCs. (A) Breeding and PCR-based genotyping designs for CXCR4 CKO mice. Mx1-Cre+ transgenic mice (Mx1-Cre+/CXCR4+/+) and Mx1-Cre−/CXCR4flox/flox mice both have a C57BL/6J background were founders to be crossbred to obtain Mx1-Cre+/CXCR4flox/flox mice. (B) Genomic PCR of mice tail biopsies. (C-E) Mx1-Cre+/CXCR4flox/flox mice were administrated 12.5 mg/Kg PI: PC to induce Cre activity to specifically delete Cxcr4 gene in PBMCs (See the details in Materials and methods). (C) Experimental design. (D) 12.5 mg/Kg PI: PC injection, and (E) Animal survival (all females, n = 6 mice). (F) The qRT-PCR analysis of Cxcr4’s mRNA level in PBMCs from CXCR4 CKO mice. (G-H) Flow cytometry analysis of CXCR4+PBMC from CXCR4 CKO mice. (G) Representative flow charts and (H) Quantitative analysis from G. (I) Cxcr4-deleted PBMCs loss of transmigration induced by 100 ng/mL recombinant CXCL12 in vitro. (A-I) Mouse genotypes indicted by Wild type (WT, Mx1-Cre−/CXCR4+/+), Mx1-Cre+ (Mx1-Cre+/CXCR4+/+), CXCR4flox/flox (Mx1-Cre−/CXCR4flox/flox), and Mx1-Cre+/CXCR4flox/flox (Cxcr4-CKO). (F-I) Data represented as mean ± SEM, *p < 0.05, **p < 0.01 and ***p < 0.001

Full size image

To verify the effects of PI: PC-induced CXCR4 CKO, peripheral PBMCs were isolated from the above mice and qRT-PCR was used to detect the mRNA levels of CXCR4 from their cells. Compared to WT, Mx1-Cre+, or CXCR4flox/flox mice, expression of CXCR4’s mRNA in Mx1-Cre+/CXCR4flox/flox mice was decreased by 66.94 ± 7.46% (P < 0.05), 33.26 ± 3.03% (P < 0.05), and 72.32 ± 1.59% (P < 0.05) respectively (Fig. 5F). To confirm these observations, we further detected the CXCR4+PBMCs by flow cytometry (Fig. 5G-H). Consistently, our results showed that CXCR4+PBMCs from Mx1-Cre+/CXCR4flox/flox mice was decreased by 61.38 ± 0.96% (P < 0.01), 71.19 ± 3.41% (P < 0.001), and 68.46 ± 3.95% (P < 0.01) when compared respectively with counterparts from WT, Mx1-Cre+, or CXCR4flox/flox mice (Fig. 5H). More importantly, we found that transmigration of PBMCs from Mx1-Cre+/CXCR4flox/flox mice was decreased by 48.58 ± 3.08% (P < 0.001), 33.94 ± 4.60% (P < 0.01), and 36.99 ± 3.97% (P < 0.01) when compared respectively with counterparts from WT, Mx1-Cre+, or CXCR4flox/flox mice (Fig. 5I). Taken together with genotyping results, these findings supported a successful CXCR4 CKO mouse line by which the Cxcr4 gene could be disrupted in peripheral PBMCs.

Loss of CXCR4 function in PBMCs resulted in decreased joint and lung inflammation in CIA mice

Above results indicate that the CXCL12-CXCR4 chemotaxis may play important roles during the pathological development of arthritis model mice. To address this hypothesis, we further explored whether disrupting CXCR4 in PBMCs could affect the pathogenesis of CIA mice (Fig. 6A). We noticed that although the WT CIA mice developed a three core arthritic stages (onset, peak and recovery stages) (Fig. S9A), loss of Cxcr4 gene in peripheral blood cells did not directly impact this trend. However, the incidence of arthritis was decreased in Mx1-Cre+/CXCR4flox/flox mice compared to Mx1-Cre+ mice (Fig. 6B). The clinical scores of Mx1-Cre+/CXCR4flox/flox mice were overall decreased compared with the Mx1-Cre+ mice during the recover stage, and there was significantly decreased (P < 0.05) at day 45 in Mx1-Cre+/CXCR4flox/flox mice compared with Mx1-Cre+ mice in CIA model mice (Fig. 6C). We further observed that cartilage destruction and bone erosion were significantly decreased (Fig. S10B-C), and local inflammation within the lungs were decreased after disrupting Cxcr4 (Fig. S10D-E). To confirm this observation, inflammatory cytokines, such as TNFα, IL-6, and IL-1β from mice lung tissues, were further examined using qRT-PCR. In CIA mice, both TNFα’s and IL-6’s levels were increased by 21.81-fold change (P < 0.001) and 2.72-fold change (P < 0.05) when compared with Sham mice (Fig. 6D). We also found that IL-6’s and IL-1β’s mRNA levels from Mx1-Cre+/CXCR4flox/flox CIA mice decreased significantly by a 6.36-fold change (P < 0.05) and a 4.27-fold change (P < 0.01) when compared with Mx1-Cre+ CIA mice counterparts (Fig. 6E). Again, we noticed a decrease in CXCR4+PBMCs and CXCR4+CD3+ T cells after disrupting the Cxcr4 gene (Fig. 6F-H). These findings supported the potential effect of CXCR4 genetic disruption in eliminating both peripheral and local tissue inflammation in arthritis mice, suggesting a key role of the CXCL12-CXCR4 chemotaxis during the pathological development of RA diseases.

Fig. 6
figure 6

Conditionally deleting Cxcr4 gene in PBMCs decreased peripheral inflammation. (A) Experimental design of CIA mouse model using Mx1-Cre+ (I, CXCR4-WT, Mx1-Cre+/CXCR4+/+) or CXCR4 CKO (II, Mx1-Cre+/CXCR4flox/flox) mice. (B) The incidence of arthritis. (C) The clinic scores of arthritis. (D) Enhanced lung inflammation observed in wild type CIA mice (Sham versus CIA mice). (E) The qRT-PCR analysis demonstrating decreased mRNA levels of inflammatory factors in the lung of CXCR4 CKO CIA mice compared with CXCR4-WT CIA mice at day 35. (F-H) Flow-based quantitative analysis demonstrating decreased the frequency of CXCR4+ peripheral immune cells in CXCR4 CKO CIA mice. (F) CXCR4+PBMCs. (G) CXCR4+CD3+ T cells. (H) Total CD3+ T cells. (D) *P < 0.05 and ***P < 0.001, and n = 4 mice. (E-H) *p < 0.05, **p < 0.01 and ***p < 0.001, and n = 5 mice. Data represented as mean ± SEM

Full size image

Discussion

Abnormal accumulation of immune cells within the synovial membrane [44], DNA hypomethylation [45, 46] are the major reason for the pathological development RA-related clinic scores. Under the inflammatory microenvironment, activated lymphocytes, dendritic cells and monocytes migrating from the peripheral blood cause the swollen synovial membrane, increased synovial fluid, and degraded cartilage in the joint. It has been suggested that migration of immune cells directly relies on multiple important chemotactic signals. For example, it has been reported that the expression of CXC chemokine CXCL12 is enhanced [47,48,49], both globally and in gene-specific hypomethylation of CXCL12 [50] in synovial cells in the joints of RA patients and is correlated with patient’s pathological development [51, 52]. Recent studies have also suggested that both inflammatory cytokines and hypoxia could be the major factors that upregulated the expression of CXCL12 in patient’s joints [53,54,55]. Other than the local tissue environment, it has been reported that the plasma level of the CXCL12 protein was increased in RA patients as well [30]. These findings potentially support the hypothesis that elevated CXCL12 within the synovium may provide the important chemoattractant for immune cell migration during the pathological development of RA diseases.

Meanwhile, CXCL12 and its cognate receptor CXCR4 mediate a key chemotaxis responsible for immune cell migration [8]. As such, it is possible that CXCR4-expressing immune cells from both peripheral blood and synovium may contribute to the pathogenesis of RA. Recent studies have observed that expression of CXCR4 is upregulated in dendritic cells and memory T cells in the synovium from RA patients when compared against healthy controls [9, 56, 57]. In the current study, we examined the frequency of CXCR4+PBMCs and CXCR4+CD3+ T cells on RA patients. Compared to healthy group, RA patients exhibited a higher frequency of both CXCR4+PBMCs and CXCR4+CD3+ T cells. Enhanced CXCR4+ immune cells is potentially correlated with the inflammatory cytokines in patients’ plasma. These observations further supported previous findings reporting elevated expression of CXCR4 in peripheral CD4+ Th17+ T cells in RA patients [56, 58]. Increased CXCR4+ immune cells not only appeared in synovial fluid within the joint but also in the peripheral blood in RA patients. These findings indicate that enhanced CXCR4+ immune cells in peripheral blood may represent some early events that potentially contribute to the pathological development of RA diseases.

In the current study, we also found that clinical treatments potentially downregulate the frequency of CXCR4+ immune cells in RA patients. High-dose and low-dose MTX have been widely used to treat cancer or autoimmune diseases. However, the potential targets for this drug are still not well understood. A study indicating that global DNA hypomethylation can be reversed with MTX treatment in bulk T cells from patients with RA [35]. Recent studies suggested that CXCL12 was increased in blood in rats treated with high dose MTX. However, a high dose of MTX did change the CXCL12 levels in bone marrow from the same groups of rats [59]. Different from high dose MTX, Hansen et al. (2006) reported that low dose MTX did not change the blood levels of CXCL12 in RA patients [30]. These findings suggest the effect of MTX on CXCL12 truly depends on drug’s concentrations and tissue types treated.

As for CXCR4, it has been reported that CXCR4’s expression in rat’s hematopoietic stem cells was decreased after treatment with high dose MTX [59]. High dose MTX upregulated the expression of CXCR4 in colon cancer cells [60]. Conversely, low dose MTX downregulated CXCR4’s expression in treated synovium of rats [61]. In the current study, we observed that the downregulation of CXCR4+PBMCs and CXCR4+CD3+ T cells was associated with the treatments of low dose MTX in RA patients. Our in vitro studies showed that low dose MTX reverted hypomethylation of promoter CpG sites in Jurkat and CD3+ T cell, which lead to the downregulated mRNA expression of CXCR4. Consistently, low dose MTX inhibited the transmigration that was induced by CXCL12, a similar observation obtained from using CXCR4-specific inhibitor AMD3100. Our studies further showed that both low dose MTX and AMD3100 can treat RA-related diseases in the arthritis mouse model. The treatments using low dose MTX may rely on the functions of CXCR4, particularly in peripheral immune cells. However, the potential regulation of other chemokine receptors by low dose MTX cannot be fully excluded. These results suggest that low dose MTX decreased the migration capacity of immune cells through downregulation of CXCR4 expression due to the hypermethylation. However, these effects were not limited to low dose MTX, as other clinical treatments were also found to downregulate the frequency of CXCR4+ peripheral immune cells in RA patients. The CXCL12-CXCR4 chemotaxis may represent the universal molecular pathway involved in both pathological development and treatment of RA patients. Therefore, the detailed mechanism underlying low dose MTX-mediated treatments in RA patients still remains relatively unknown and open for further future investigation.

Chung et al. (2010) reported that the incidence, but not the severity, of CIA was significantly reduced in T cell-specific Cxcr4-deficient mice [16]. Since expression of CXCR4 is not limited to T cells, we further explored the roles of CXCR4 in peripheral immune cells by generating mice with Mx1-Cre/CXCR4flox/flox genotype. A broader genetic disruption of Cxcr4 gene in different peripheral immune cells from Mx1-Cre/CXCR4flox/flox mice was induced by PI: PC. We also noticed a little improvement in CXCR4 CKO mice compared with Cxcr4 WT mice during the recovery phase. In addition, we observed that both peripheral and lungs’ inflammations were significantly eliminated in CXCR4 CKO mice. Our loss of function studies supported the significance of CXCR4’s role during the pathological development of RA. Even so, most human trials of anti-chemokine signaling pathways have failed to show clinical improvements [7], suggesting CXCL12-CXCR4 is not the only chemotaxis related to RA’s pathogenesis. Therefore, to fully grasp the detailed mechanism associated with chemokine signaling underlying low dose MTX and other clinic treatments in RA patients, a more thorough investigation must be conducted in the future.

Conclusions

In summary, this study found that the frequency of CXCR4+PBMCs and CXCR4+CD3+ T cells were increased in RA patients. Following, CXCR4’s expression or the frequency of CXCR4+PBMCs and CXCR4+CD3+ T cells on peripheral immune cells was significantly downregulated after treated with low dose MTX treatment in vitro and in vivo. Consistently, the capacity of immune cell transmigration was inhibited by down-regulated expression of chemokine receptor CXCR4. Briefly, in vitro, low dose MTX significantly decreased cell transmigration through down-regulated CXCR4’s expression caused by an increased genomic hypermethylation across the promoter region of Cxcr4 gene in CD3+ T cells. In vivo, low dose MTX-mediated downregulate the frequency of CXCR4+PBMCs and CXCR4+CD3+ T cells significantly improved the pathological development of mouse arthritis models. Moreover, the conditional disruption of the Cxcr4 gene in peripheral immune cells potentially alleviated inflammation of joints and lung tissues in the arthritis mice (Fig. 7). Our results taken together suggested that the effect of low dose MTX treatment could serve to eliminate inflammation in RA patients through impairment of immune cell transmigration due to the down-regulated expression of chemokine receptor CXCR4.

Fig. 7
figure 7

The Proposed schematic model for low dose MTX impaired T cell transmigration through down-regulating CXCR4 expression in RA patients. The increased CXCR4+PBMCs and CXCR4+CD3+ T cells in RA patients’ peripheral immune cells caused more cell migration capacity to promote the development of RA. CXCR4’s expression on peripheral immune cells was significantly downregulated after low dose MTX treatment due to the increased genomic hypermethylation across the promoter region of Cxcr4 gene, mediating the decrease of cell transmigration in vitro, which significantly improved the pathological development of RA partly through potentially alleviated inflammation of joints and lung tissue in RA patients

Full size image

Data availability

Data and full uncropped Gels are provided within the related information files.

Abbreviations

cDNA:

Complement DNA

CFA:

Complete freund’s adjuvant

CIA:

Collagen-induced arthritis

CII:

Type II collagen

CKO:

Conditional knockout

CXCR4:

CXC motif chemokine receptor 4

DHFR:

Inhibiting dihydrofolate reductase

ECM:

Extracellular matrix

ELISA:

Enzyme-linked immunosorbent assay

FCM:

Flow cytometry

GRO/CXCR1:

Chemokine (CXC motif) ligand 1

H&E:

Hematoxylin and eosin

IL-1β:

Interleukin 1 beta

IL-1r:

Interleukin 1 receptor type 1

IL-6:

Interleukin 6

IL-8:

Interleukin 8

IL-17R:

Interleukin 17 receptor A

MCP-1/CCL2:

C-C motif chemokine ligand 2

MTX:

Methotrexate

NS:

Normal saline

PBMC:

Peripheral blood mononuclear cells

PFA:

Paraformaldehyde

PI:

PC: Polyinosinic-polycytidylic acid

PMT:

Photomultiplier tube

qRT-PCR:

Quantitative real-time polymerase chain reaction

RA:

Rheumatoid arthritis

RANTES /CCL5:

C-C motif chemokine ligand 5

SDF-1/CXCL12:

CXC motif chemokine ligand 12

SEM:

Standard error of mean

TNF-α:

Tumor necrosis factor alpha

TGFB1:

TGF-beta 1

References

  1. McInnes IB, Schett G. The pathogenesis of rheumatoid arthritis. N Engl J Med. 2011;365(23):2205–19.

    Article  CAS  PubMed  Google Scholar 

  2. Buckley CD. Science, medicine, and the future. Treatment of rheumatoid arthritis. BMJ. 1997;315(7102):236–8.

    Article  CAS  PubMed  Google Scholar 

  3. Salmon M, Gaston JS. The role of T-lymphocytes in rheumatoid arthritis. Br Med Bull. 1995;51(2):332–45.

    Article  CAS  PubMed  Google Scholar 

  4. Elemam NM, Hannawi S, Maghazachi AA. Role of chemokines and Chemokine receptors in Rheumatoid Arthritis. Immunotargets Ther. 2020;9:43–56.

    Article  CAS  PubMed  Google Scholar 

  5. Vergunst CE, van de Sande MG, Lebre MC, Tak PP. The role of chemokines in rheumatoid arthritis and osteoarthritis. Scand J Rheumatol. 2005;34(6):415–25.

    Article  CAS  PubMed  Google Scholar 

  6. Szekanecz Z, Kim J, Koch AE. Chemokines and chemokine receptors in rheumatoid arthritis. Semin Immunol. 2003;15(1):15–21.

    Article  CAS  PubMed  Google Scholar 

  7. Szekanecz Z, Koch AE. Successes and failures of chemokine-pathway targeting in rheumatoid arthritis. Nat Rev Rheumatol. 2016;12(1):5–13.

    Article  CAS  PubMed  Google Scholar 

  8. Li M, Ransohoff RM. Multiple roles of chemokine CXCL12 in the central nervous system: a migration from immunology to neurobiology. Prog Neurobiol. 2008;84(2):116–31.

    Article  CAS  PubMed  Google Scholar 

  9. Nanki T, Hayashida K, El-Gabalawy HS, Suson S, Shi K, Girschick HJ, Yavuz S, Lipsky PE. Stromal cell-derived factor-1-CXC chemokine receptor 4 interactions play a central role in CD4 + T cell accumulation in rheumatoid arthritis synovium. J Immunol. 2000;165(11):6590–8.

    Article  CAS  PubMed  Google Scholar 

  10. Bradfield PF, Amft N, Vernon-Wilson E, Exley AE, Parsonage G, Rainger GE, et al. Rheumatoid fibroblast-like synoviocytes overexpress the chemokine stromal cell-derived factor 1 (CXCL12), which supports distinct patterns and rates of CD4 + and CD8 + T cell migration within synovial tissue. Arthritis Rheum. 2003;48(9):2472–82.

    Article  CAS  PubMed  Google Scholar 

  11. Buckley CD, Amft N, Bradfield PF, Pilling D, Ross E, Arenzana-Seisdedos F, et al. Persistent induction of the chemokine receptor CXCR4 by TGF-beta 1 on synovial T cells contributes to their accumulation within the rheumatoid synovium. J Immunol. 2000;165(6):3423–9.

    Article  CAS  PubMed  Google Scholar 

  12. Shadidi KR, Aarvak T, Henriksen JE, Natvig JB, Thompson KM. The chemokines CCL5, CCL2 and CXCL12 play significant roles in the migration of Th1 cells into rheumatoid synovial tissue. Scand J Immunol. 2003;57(2):192–8.

    Article  CAS  PubMed  Google Scholar 

  13. Lenoir M, Djerdjouri B, Perianin A. Stroma cell-derived factor 1alpha mediates desensitization of human neutrophil respiratory burst in synovial fluid from rheumatoid arthritic patients. J Immunol. 2004;172(11):7136–43.

    Article  CAS  PubMed  Google Scholar 

  14. Matthys P, Hatse S, Vermeire K, Wuyts A, Bridger G, Henson GW, De Clercq E, Billiau A, Schols D. AMD3100, a potent and specific antagonist of the stromal cell-derived factor-1 chemokine receptor CXCR4, inhibits autoimmune joint inflammation in IFN-gamma receptor-deficient mice. J Immunol. 2001;167(8):4686–92.

    Article  CAS  PubMed  Google Scholar 

  15. De Klerck B, Geboes L, Hatse S, Kelchtermans H, Meyvis Y, Vermeire K, et al. Pro-inflammatory properties of stromal cell-derived factor-1 (CXCL12) in collagen-induced arthritis. Arthritis Res Ther. 2005;7(6):R1208–20.

    Article  PubMed  Google Scholar 

  16. Chung SH, Seki K, Choi BI, Kimura KB, Ito A, Fujikado N, Saijo S, Iwakura Y. CXC chemokine receptor 4 expressed in T cells plays an important role in the development of collagen-induced arthritis. Arthritis Res Ther. 2010;12(5):R188.

    Article  PubMed  Google Scholar 

  17. Bedoui Y, Guillot X, Selambarom J, Guiraud P, Giry C, Jaffar-Bandjee MC, Ralandison S, Gasque P. Methotrexate an Old Drug with New tricks. Int J Mol Sci 2019; 20(20).

  18. Visentin M, Zhao R, Goldman ID. The antifolates. Hematol Oncol Clin North Am. 2012;26(3):629–48. ix.

    Article  PubMed  Google Scholar 

  19. Weinblatt ME, Maier AL, Coblyn JS. Low dose leucovorin does not interfere with the efficacy of methotrexate in rheumatoid arthritis: an 8 week randomized placebo controlled trial. J Rheumatol. 1993;20(6):950–2.

    CAS  PubMed  Google Scholar 

  20. Whittle SL, Hughes RA. Folate supplementation and methotrexate treatment in rheumatoid arthritis: a review. Rheumatology (Oxford). 2004;43(3):267–71.

    Article  CAS  PubMed  Google Scholar 

  21. Friedman B, Cronstein B. Methotrexate mechanism in treatment of rheumatoid arthritis. Joint Bone Spine. 2019;86(3):301–07.

    Article  CAS  PubMed  Google Scholar 

  22. Cronstein BN, Aune TM. Methotrexate and its mechanisms of action in inflammatory arthritis. Nat Rev Rheumatol. 2020;16(3):145–54.

    Article  CAS  PubMed  Google Scholar 

  23. Deleuran B, Kristensen M, Paludan K, Zachariae C, Larsen CG, Zachariae E, Thestrup-Pedersen K. The effect of second-line antirheumatic drugs on interleukin-8 mRNA synthesis and protein secretion in human endothelial cells. Cytokine. 1992;4(5):403–9.

    Article  CAS  PubMed  Google Scholar 

  24. Boiardi L, Macchioni P, Meliconi R, Pulsatelli L, Facchini A, Salvarani C. Relationship between serum RANTES levels and radiological progression in rheumatoid arthritis patients treated with methotrexate. Clin Exp Rheumatol. 1999;17(4):419–25.

    CAS  PubMed  Google Scholar 

  25. Yoshida M, Kanno Y, Ishisaki A, Tokuda H, Hirade K, Nakajima K, Katagiri Y, Shimizu K, Kozawa O. Methotrexate suppresses inflammatory agonist induced interleukin 6 synthesis in osteoblasts. J Rheumatol. 2005;32(5):787–95.

    CAS  PubMed  Google Scholar 

  26. Gao IK, Leins C, Bohlen H, Heilig B, Lemmel EM. Inhibition of interleukin-8 synthesis by intraarticular methotrexate therapy in patients with rheumatoid arthritis. Z Rheumatol. 1998;57(2):95–100.

    Article  CAS  PubMed  Google Scholar 

  27. Seitz M, Loetscher P, Dewald B, Towbin H, Rordorf C, Gallati H, Gerber NJ. Interleukin 1 (IL-1) receptor antagonist, soluble tumor necrosis factor receptors, IL-1 beta, and IL-8–markers of remission in rheumatoid arthritis during treatment with methotrexate. J Rheumatol. 1996;23(9):1512–6.

    CAS  PubMed  Google Scholar 

  28. Seitz M, Dewald B, Ceska M, Gerber N, Baggiolini M. Interleukin-8 in inflammatory rheumatic diseases: synovial fluid levels, relation to rheumatoid factors, production by mononuclear cells, and effects of gold sodium thiomalate and methotrexate. Rheumatol Int. 1992;12(4):159–64.

    Article  CAS  PubMed  Google Scholar 

  29. Brandalise SR, Assis R, Laranjeira ABA, Yunes JA, de Campos-Lima PO. Low-dose methotrexate in sickle-cell disease: a pilot study with rationale borrowed from rheumatoid arthritis. Exp Hematol Oncol. 2017;6:18.

    Article  PubMed  Google Scholar 

  30. Hansen IB, Ellingsen T, Hornung N, Poulsen JH, Lottenburger T, Stengaard-Pedersen K. Plasma level of CXC-chemokine CXCL12 is increased in rheumatoid arthritis and is independent of disease activity and methotrexate treatment. J Rheumatol. 2006;33(9):1754–9.

    CAS  PubMed  Google Scholar 

  31. Abdelaal NH, Elhefnawy NG, Abdulmonem SR, Sayed S, Saleh NA, Saleh MA. Evaluation of the expression of the stromal cell-derived factor-1 alpha (CXCL 12) in psoriatic patients after treatment with Methotrexate. J Cosmet Dermatol. 2020;19(1):253–58.

    Article  PubMed  Google Scholar 

  32. Barsig J, Yam G, Lehner MD, Beume R. Methotrexate treatment suppresses local cytokine and chemokine production in rat adjuvant arthritis. Drugs Exp Clin Res. 2005;31(1):7–11.

    CAS  PubMed  Google Scholar 

  33. Kehlen A, Thiele K, Riemann D, Langner J. Expression, modulation and signalling of IL-17 receptor in fibroblast-like synoviocytes of patients with rheumatoid arthritis. Clin Exp Immunol. 2002;127(3):539–46.

    Article  CAS  PubMed  Google Scholar 

  34. Kim YI, Logan JW, Mason JB, Roubenoff R. DNA hypomethylation in inflammatory arthritis: reversal with methotrexate. J Lab Clin Med. 1996;128(2):165–72.

    Article  CAS  PubMed  Google Scholar 

  35. de Andres MC, Perez-Pampin E, Calaza M, Santaclara FJ, Ortea I, Gomez-Reino JJ, Gonzalez A. Assessment of global DNA methylation in peripheral blood cell subpopulations of early rheumatoid arthritis before and after methotrexate. Arthritis Res Ther. 2015;17(1):233.

    Article  PubMed  Google Scholar 

  36. Guderud K, Sunde LH, Flam ST, Maehlen MT, Mjaavatten MD, Norli ES, et al. Methotrexate Treatment of newly diagnosed RA patients is Associated with DNA methylation differences at Genes Relevant for Disease Pathogenesis and Pharmacological Action. Front Immunol. 2021;12:713611.

    Article  CAS  PubMed  Google Scholar 

  37. Arnett FC, Edworthy SM, Bloch DA, McShane DJ, Fries JF, Cooper NS, et al. The American Rheumatism Association 1987 revised criteria for the classification of rheumatoid arthritis. Arthritis Rheum. 1988;31(3):315–24.

    Article  CAS  PubMed  Google Scholar 

  38. Sun C, McAndrew T, Smith BC, Chen Z, Frimer M, Burk RD. Characterization of HPV DNA methylation of contiguous CpG sites by bisulfite treatment and massively parallel sequencing-the FRAGMENT approach. Front Genet. 2014;5:150.

    Article  PubMed  Google Scholar 

  39. Kollias G, Papadaki P, Apparailly F, Vervoordeldonk MJ, Holmdahl R, Baumans V, et al. Animal models for arthritis: innovative tools for prevention and treatment. Ann Rheum Dis. 2011;70(8):1357–62.

    Article  PubMed  Google Scholar 

  40. Brand DD, Latham KA, Rosloniec EF. Collagen-induced arthritis. Nat Protoc. 2007;2(5):1269–75.

    Article  CAS  PubMed  Google Scholar 

  41. Weinberger A, Halpern M, Zahalka MA, Quintana F, Traub L, Moroz C. Placental immunomodulator ferritin, a novel immunoregulator, suppresses experimental arthritis. Arthritis Rheum. 2003;48(3):846–53.

    Article  CAS  PubMed  Google Scholar 

  42. Gao B, Lin J, Jiang Z, Yang Z, Yu H, Ding L, et al. Upregulation of chemokine CXCL10 enhances chronic pulmonary inflammation in tree shrew collagen-induced arthritis. Sci Rep. 2018;8(1):9993.

    Article  PubMed  Google Scholar 

  43. Inglis JJ, Simelyte E, McCann FE, Criado G, Williams RO. Protocol for the induction of arthritis in C57BL/6 mice. Nat Protoc. 2008;3(4):612–8.

    Article  CAS  PubMed  Google Scholar 

  44. Leblond A, Allanore Y, Avouac J. Targeting synovial neoangiogenesis in rheumatoid arthritis. Autoimmun Rev. 2017;16(6):594–601.

    Article  PubMed  Google Scholar 

  45. Javierre BM, Hernando H, Ballestar E. Environmental triggers and epigenetic deregulation in autoimmune disease. Discov Med. 2011;12(67):535–45.

    PubMed  Google Scholar 

  46. Meda F, Folci M, Baccarelli A, Selmi C. The epigenetics of autoimmunity. Cell Mol Immunol. 2011;8(3):226–36.

    Article  CAS  PubMed  Google Scholar 

  47. Kim HR, Kim KW, Kim BM, Jung HG, Cho ML, Lee SH. Reciprocal activation of CD4 + T cells and synovial fibroblasts by stromal cell-derived factor 1 promotes RANKL expression and osteoclastogenesis in rheumatoid arthritis. Arthritis Rheumatol. 2014;66(3):538–48.

    Article  CAS  PubMed  Google Scholar 

  48. Nanki T, Nagasaka K, Hayashida K, Saita Y, Miyasaka N. Chemokines regulate IL-6 and IL-8 production by fibroblast-like synoviocytes from patients with rheumatoid arthritis. J Immunol. 2001;167(9):5381–5.

    Article  CAS  PubMed  Google Scholar 

  49. Watanabe N, Ando K, Yoshida S, Inuzuka S, Kobayashi M, Matsui N, Okamoto T. Gene expression profile analysis of rheumatoid synovial fibroblast cultures revealing the overexpression of genes responsible for tumor-like growth of rheumatoid synovium. Biochem Biophys Res Commun. 2002;294(5):1121–9.

    Article  CAS  PubMed  Google Scholar 

  50. Karouzakis E, Rengel Y, Jungel A, Kolling C, Gay RE, Michel BA, et al. DNA methylation regulates the expression of CXCL12 in rheumatoid arthritis synovial fibroblasts. Genes Immun. 2011;12(8):643–52.

    Article  CAS  PubMed  Google Scholar 

  51. Kanbe K, Chiba J, Inoue Y, Taguchi M, Yabuki A. SDF-1 and CXCR4 in synovium are associated with disease activity and bone and joint destruction in patients with rheumatoid arthritis treated with golimumab. Mod Rheumatol. 2016;26(1):46–50.

    Article  CAS  PubMed  Google Scholar 

  52. Grassi F, Cristino S, Toneguzzi S, Piacentini A, Facchini A, Lisignoli G. CXCL12 chemokine up-regulates bone resorption and MMP-9 release by human osteoclasts: CXCL12 levels are increased in synovial and bone tissue of rheumatoid arthritis patients. J Cell Physiol. 2004;199(2):244–51.

    Article  CAS  PubMed  Google Scholar 

  53. Santiago B, Calonge E, Del Rey MJ, Gutierrez-Canas I, Izquierdo E, Usategui A, Galindo M, Alcami J, Pablos JL. CXCL12 gene expression is upregulated by hypoxia and growth arrest but not by inflammatory cytokines in rheumatoid synovial fibroblasts. Cytokine. 2011;53(2):184–90.

    Article  CAS  PubMed  Google Scholar 

  54. Iwamoto T, Okamoto H, Toyama Y, Momohara S. Molecular aspects of rheumatoid arthritis: chemokines in the joints of patients. FEBS J. 2008;275(18):4448–55.

    Article  CAS  PubMed  Google Scholar 

  55. Hitchon C, Wong K, Ma G, Reed J, Lyttle D, El-Gabalawy H. Hypoxia-induced production of stromal cell-derived factor 1 (CXCL12) and vascular endothelial growth factor by synovial fibroblasts. Arthritis Rheum. 2002;46(10):2587–97.

    Article  CAS  PubMed  Google Scholar 

  56. Lande R, Giacomini E, Serafini B, Rosicarelli B, Sebastiani GD, Minisola G, et al. Characterization and recruitment of plasmacytoid dendritic cells in synovial fluid and tissue of patients with chronic inflammatory arthritis. J Immunol. 2004;173(4):2815–24.

    Article  CAS  PubMed  Google Scholar 

  57. Perretti M, Ingegnoli F, Wheller SK, Blades MC, Solito E, Pitzalis C. Annexin 1 modulates monocyte-endothelial cell interaction in vitro and cell migration in vivo in the human SCID mouse transplantation model. J Immunol. 2002;169(4):2085–92.

    Article  CAS  PubMed  Google Scholar 

  58. Burska AN, Thu A, Parmar R, Bzoma I, Samans B, Raschke E, et al. Quantifying circulating Th17 cells by qPCR: potential as diagnostic biomarker for rheumatoid arthritis. Rheumatology (Oxford). 2019;58(11):2015–24.

    Article  CAS  PubMed  Google Scholar 

  59. Georgiou KR, Scherer MA, King TJ, Foster BK, Xian CJ. Deregulation of the CXCL12/CXCR4 axis in methotrexate chemotherapy-induced damage and recovery of the bone marrow microenvironment. Int J Exp Pathol. 2012;93(2):104–14.

    Article  PubMed  Google Scholar 

  60. Cutler MJ, Lowthers EL, Richard CL, Hajducek DM, Spagnuolo PA, Blay J. Chemotherapeutic agents attenuate CXCL12-mediated migration of colon cancer cells by selecting for CXCR4-negative cells and increasing peptidase CD26. BMC Cancer. 2015;15:882.

    Article  PubMed  Google Scholar 

  61. Zhang M, Gao M, Chen J, Song L, Wei W. CP-25 exerts anti-angiogenic effects on a rat model of adjuvant-induced arthritis by promoting GRK2-induced downregulation of CXCR4-ERK1/2 signaling in endothelial cells. Mol Med Rep. 2019;20(6):4831–42.

    CAS  PubMed  Google Scholar 

Download references

Acknowledgements

We thank professor Shenghong Li (Institute of Botany, the Chinese Academy of Sciences, Kunming, China) kindly providing Jurkat cells. We thank Dr. Wei Wang and Dr. Guiyuan Chen for their assistance to collect patient sample collection, isolate PBMCs, establish arthritis mice model and conduct flow cytometry experiments. We thank Dr. Wei Wang who worked in the Department of Rheumatology from First Affiliated Hospital of Kunming Medical University and studied in the School of Life Sciences of Yunnan University for kindly providing human blood specimens and clinical RA diagnosis.

Funding

This work was financially supported by the Talent Program of Yunnan Province (No. 2011HA009), the Youth Project of the Science and Technology Department of Yunnan Province (No. 202001BB050041), and the National Natural Science Foundation of China (No. 32360165, 31660273).

Author information

Authors and Affiliations

  1. School of Life Sciences, Yunnan University, Kunming, Yunnan, 650091, China

    Lei Ding & Bo Gao

  2. Pediatric Emergency Medicine, Johns Hopkins University, Baltimore, MD, 21218, USA

    Daniel H. Park

  3. College of Bioengineering, Sichuan University of Science and Engineering, Yibin, Sichuan, 644000, China

    Bo Gao

  4. Emory University School of Medicine, Atlanta, GA, 30322, USA

    Lingyuan Wu

  5. Department of Pathology and Laboratory Medicine, University of Kansas Medical Center, Kansas City, KS, 66103, USA

    Meizhang Li

  6. Division of Hematologic Malignancies and Cellular Therapeutics, University of Kansas Medical Center, 3901 Rainbow Blvd. 4019 Wahl Hall East, MS 3040, Kansas City, KS, 66103, USA

    Haitham Abedelhakim

  7. Institute of Neuroscience, Kunming Medical University, 931 Basic Medical Blvd. 1168 Chunrong West Road, Kunming, Yunnan, 650500, China

    Ming Zhang

Authors
  1. Lei Ding
    View author publications

    You can also search for this author in PubMed Google Scholar

  2. Daniel H. Park
    View author publications

    You can also search for this author in PubMed Google Scholar

  3. Bo Gao
    View author publications

    You can also search for this author in PubMed Google Scholar

  4. Lingyuan Wu
    View author publications

    You can also search for this author in PubMed Google Scholar

  5. Meizhang Li
    View author publications

    You can also search for this author in PubMed Google Scholar

  6. Haitham Abedelhakim
    View author publications

    You can also search for this author in PubMed Google Scholar

  7. Ming Zhang
    View author publications

    You can also search for this author in PubMed Google Scholar

Contributions

Conception and design (Lei Ding, Daniel H. Park, Bo Gao, Meizhang Li, Haitham Abedelhakim, and Ming Zhang). Experiments and data analysis (Lei Ding, Daniel H. Park, Bo Gao, Meizhang Li, Haitham Abedelhakim, and Ming Zhang). Manuscript preparation (Lei Ding, Daniel H. Park, Meizhang Li, Haitham Abedelhakim, and Ming Zhang). Reviewed and edited the manuscript (Lei Ding, Daniel H. Park, Bo Gao, Lingyuan Wu, Meizhang Li, Haitham Abedelhakim, and Ming Zhang). All authors of this paper have read and approved the final version submitted.

Corresponding authors

Correspondence to Haitham Abedelhakim or Ming Zhang.

Ethics declarations

Ethics approval and consent to participate

The study was performed in accordance with the ethical standards of the institutional and national research committee and with the 1964 Helsinki Declaration and its later amendments. This study was approved by both the Ethics Committee of the First Affiliated Hospital of Kunming Medical University and the Committee on Human Subject Research and Ethics of Yunnan University (Approved number: 20190801). All experiments were carried out according to the relevant guidelines. All animal studies and relevant procedures were reviewed and approved by the ethics committee of the institutional animal care and use committee, Yunnan University (IACUC, YNU) (Approved number: 20190801). Written informed consent for the present study’s use of human peripheral blood was obtained from all forty-eight patients and thirty-seven healthy donors (HDs).

Consent for publication

Written informed consent for the publication was obtained from the study participants.

Competing interests

The authors declare no competing interests.

Additional information

Publisher’s note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Electronic supplementary material

Below is the link to the electronic supplementary material.

Supplementary Material 1

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International License, which permits any non-commercial use, sharing, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if you modified the licensed material. You do not have permission under this licence to share adapted material derived from this article or parts of it. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by-nc-nd/4.0/.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Ding, L., Park, D.H., Gao, B. et al. Low dose methotrexate impaired T cell transmigration through down-regulating CXCR4 expression in rheumatoid arthritis (RA). Arthritis Res Ther 26, 173 (2024). https://doi.org/10.1186/s13075-024-03403-9

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: https://doi.org/10.1186/s13075-024-03403-9

Keywords

Arthritis Research & Therapy

ISSN: 1478-6362

Contact us