To understand the mechanisms that promote recruitment and survival of T cells within the pediatric inflamed joint, we have studied the expression of CCR4 and CCR5 on synovial fluid T cells and matched peripheral blood samples from juvenile rheumatoid arthritis (JRA) patients using three-color flow cytometric analysis. Thymus- and activation-regulated chemokine and macrophage-derived chemokine, ligands for CCR4, were measured by ELISA in JRA synovial fluid, JRA plasma, adult rheumatoid arthritis synovial fluid, and normal plasma. IL-4 and IFN-γ mRNA production was assessed in CD4+/CCR4+ and CD4+/CCR4 cell subsets. We found accumulations of both CCR4+ and CCR5+ T cells in JRA synovial fluids and a correlation for increased numbers of CCR4+ T cells in samples collected early in the disease process. Thymus- and activation-regulated chemokine was detected in JRA synovial fluid and plasma samples, but not in adult rheumatoid arthritis synovial fluid or control plasma. Macrophage-derived chemokine was present in all samples. CD4+/CCR4+ synovial lymphocytes produced more IL-4 and less IFN-γ than CD4+/CCR4 cells. These findings suggest that CCR4+ T cells in the JRA joint may function early in disease in an anti-inflammatory capacity through the production of type 2 cytokines and may play a role in determining disease phenotype.

Juvenile rheumatoid arthritis (JRA)3 is a multisystem autoimmune inflammatory disease with three characteristic onset types (pauciarticular, polyarticular, or systemic), which differ both in severity and outcome. Although etiologic agents of JRA are not defined, the resulting chronically inflamed synovium is typified by persistent infiltration of mononuclear effector cells, among which T cells play a prominent role. The failure to identify an arthritogenic Ag or a common disease-relevant T cell has challenged the simple model that disease-associated HLA molecules select and present a single Ag. In JRA, the T cell infiltrate is diverse in terms of TCR gene usage but consistently includes clonally expanded populations that exhibit selective TCRVβ gene segment usage related to disease subtype (1).

Insight into the mechanisms of immunopathogenesis has also been gained through the characterization of the cytokine profiles of infiltrating T cells. Type 1 T cells produce IL-2, IFN-γ, and TNF-β, whereas a type 2 response is characterized by the production of IL-4, IL-5, IL-6, IL-10, and IL-13. The balance between these two subsets regulates the choice between inflammatory cellular and Ab-mediated immune responses while strongly polarized responses can also promote immunopathological reactions (2). As expected in chronic inflammation, adult rheumatoid arthritis (RA) and its animal model collagen-induced arthritis have both been identified as type 1 diseases with only a minor type 2 component (3). Although JRA also has a strong type 1 component (4),4 contrasting cytokine profiles have been identified relative to disease subtype (5, 6, 7). In particular, type 2 cytokines are more readily detectable in synovium of patients with limited joint destruction (7).

Chemokines and their seven-transmembrane domain G protein-coupled receptors determine the selective migration of functional subsets of T cells. Chemokines are often up-regulated in inflammation (8, 9) and act mainly on leukocytes inducing migration and release responses. Together this system is capable of supporting host defense and repair functions (10). In disease, and possibly best defined for asthma (11), this system may also act as an amplifier of inappropriate inflammation.

Chemokines may also be part of the effector and amplification mechanisms of polarized type 1- and 2-mediated immune responses. It has been demonstrated that distinct profiles of chemokine receptors are acquired by T cells after polarization and are modulated by cytokines (11, 12). CCR3, CCR4, and CCR8 expression has been associated with type 2 T cell differentiation in vitro (13, 14, 15, 16). In contrast, CXC chemokine receptor (CXCR) 3 and CCR5 are preferentially expressed on human type 1 T cells, whereas CXCR4 and CCR2 are expressed equally on both type 1 and type 2 cells (14, 17). These observations suggest that this differential expression will be useful in determining T cells important to disease pathogenesis.

Recent studies in RA have investigated the chemokine and corresponding receptors that function in recruitment of T cells to the joint (18, 19, 20). All studies showed a selective accumulation of CCR5+ T cells in RA synovial fluid. In contrast, the single study that examined CCR4+ in RA did not report increases in the inflamed joints (20).

Increased expression of CCR5 and CXCR3 has been found in JRA along with high IFN-γ/IL-4 ratios, suggesting that the type 1 phenotype found within the chronically inflamed joints reflects specific recruitment events (4).

To better understand the mechanisms that promote recruitment and survival of T cells within the pediatric inflamed joint the expression of chemokine receptor CCR4 has been studied. CCR4, rather than CCR3, was chosen as a representative for type 2 T cell populations because <1% of peripheral T cells display CCR3. We found both CCR5+ and CCR4+ T cells accumulate in the inflamed joints of JRA patients with functional consequences for expression of IFN-γ and IL-4. Furthermore, a correlation was found between disease duration and levels of chemokine receptor expression. These findings are particularly interesting in light of the differences in T cell populations and cytokine profiles reported between adult and juvenile arthritis.

Synovial fluid samples were collected from the knee joints of 23 JRA or juvenile spondyloarthropathy patients during routine clinical visits that included joint aspiration for diagnostic purposes or the instillation of corticosteroids. For 15 of the pediatric patients, a peripheral blood sample was obtained at the time of joint aspiration. For three patients, synovial fluid samples were obtained from two knees at a single visit. All pediatric patients met the American College of Rheumatology criteria for a diagnosis of JRA (21) or the European Spondylarthropathy Study Group criteria for juvenile spondyloarthropathy (22) and were attending the pediatric rheumatology clinic at Children’s Hospital Medical Center (Cincinnati, OH). Clinical details were obtained from retrospective chart reviews and are summarized in Table I. Destructive disease was defined as narrowing of joint space or presence of erosion by plain film radiology. Patients were not systematically radiographed, so reports of radiographs performed for clinical purposes only were available. PBLs from nine healthy adult donors (median age 26 years) were included for comparison. Peripheral blood and synovial fluid mononuclear cells were isolated by Ficoll-Hypaque gradient centrifugation and stored in liquid nitrogen until needed. This treatment did not affect staining for any of the cell surface molecules in this study (results not shown). All RA patients met the American College of Rheumatology criteria.

Table I.

Clinical characteristics of JRA patients studied

Patient No.GenderSamples AnalyzedaOnset Age (yr)Age at Sample (yr)Duration at Sample (yr)Onset TypebCoursecDestructive DiseaseHLA DRB1 Allele1HLA DRB1 Allele2CCR5 Δ32 Genotyped
PB, 2 SF No 14 wt /wt 
PB, SF Yes 13 wt /wt 
PB, SF 10 15 Yes 12 wt /wt 
PB, 4 SF 10 No NAe NA wt /wt 
PB, SF 16 12 Yes 98 wt /wt 
PB, SF 18 Yes 11 13 wt /wt 
PB, SF 14 26 12 Yes Δ32 /wt 
PB, 2 SF No NA NA Δ32 /wt 
10 PB, SF 10 No 13 15 wt /wt 
11 PB, SF Yes 11 NA 
12 PB, SF 13 12 Yes wt /wt 
13 PB, SF 13 30 17 NA Yes NA NA wt /wt 
14 PB, SF 10 21 11 No 12 13 wt /wt 
15 SF 14 Yes NA NA Δ32 /Δ32 
16 SF NA Yes NA NA NA 
17 SF 28 19 NA No wt /wt 
18 SF 19 13 NA Yes NA NA NA 
19 SF Yes NA NA wt /wt 
20 SF 14 19 Yes NA NA NA 
21 SF 10 17 Yes 11 wt /wt 
22 SF Yes NA NA NA 
23 PB, 2 SF 10 No 15 wt /wt 
24 PB, SF No wt /wt 
Patient No.GenderSamples AnalyzedaOnset Age (yr)Age at Sample (yr)Duration at Sample (yr)Onset TypebCoursecDestructive DiseaseHLA DRB1 Allele1HLA DRB1 Allele2CCR5 Δ32 Genotyped
PB, 2 SF No 14 wt /wt 
PB, SF Yes 13 wt /wt 
PB, SF 10 15 Yes 12 wt /wt 
PB, 4 SF 10 No NAe NA wt /wt 
PB, SF 16 12 Yes 98 wt /wt 
PB, SF 18 Yes 11 13 wt /wt 
PB, SF 14 26 12 Yes Δ32 /wt 
PB, 2 SF No NA NA Δ32 /wt 
10 PB, SF 10 No 13 15 wt /wt 
11 PB, SF Yes 11 NA 
12 PB, SF 13 12 Yes wt /wt 
13 PB, SF 13 30 17 NA Yes NA NA wt /wt 
14 PB, SF 10 21 11 No 12 13 wt /wt 
15 SF 14 Yes NA NA Δ32 /Δ32 
16 SF NA Yes NA NA NA 
17 SF 28 19 NA No wt /wt 
18 SF 19 13 NA Yes NA NA NA 
19 SF Yes NA NA wt /wt 
20 SF 14 19 Yes NA NA NA 
21 SF 10 17 Yes 11 wt /wt 
22 SF Yes NA NA NA 
23 PB, 2 SF 10 No 15 wt /wt 
24 PB, SF No wt /wt 
a

PB, Peripheral blood; SF, synovial fluid; 2 SF, synovial fluid from two joints on same date.

b

Pauciarticular onset; 2, polyarticular onset; 3, systemic onset; 4, spondylarthropathy; NA, not available.

c

1, Pauciarticular course; 2, polyarticular course.

d

wt, Presence of 174-bp CCR5-specific PCR product; Δ32, presence of 142 bp CCR5-specific PCR product.

e

NA, Not available

Allele-specific PCR for the CCR5 polymorphism was performed (23) for patients with genomic DNA available, and the results are shown in Table I. This assay distinguishes wild-type CCR5 alleles (174-bp PCR product) from alleles containing a 32-bp deletion (142-bp product), which abolishes CCR5 expression.

The primary mAbs with specificity for CCR5 (2D7, IgG2a) and CCR4 (1G1, IgG1) (Millenium Pharmaceuticals, Cambridge, MA) were used for indirect staining and required a FITC- or PE-conjugated (Fab′)2 goat-anti-mouse IgG (H+L) for detection (Jackson ImmunoResearch Laboratories, West Grove, PA). CCR5 (2D7) was also available directly conjugated to FITC (PharMingen, San Diego, CA). Abs to CCR5 from the two sources (identical clone) were indistinguishable and were used interchangeably. PE-conjugated Abs (Immunotech, Marseille, France) with specificity for CD4 (13B8.2, IgG1), CD8 (B9.11, IgG1), CD45RO (UCHL1, IgG2a), TCRγδ (Imm 510, IgG1), and TCRαβ (BMA 031, IgG2b) and tricolor-conjugated Abs (Caltag, Burlingame, CA) to CD3 (S4.1, IgG2a) and CD45RO (UCHL1, IgG2a) were also used in these studies. Isotype-matched control mAb were all derived in mice and included purified and PE-conjugated IgG1 (MOPC-21; PharMingen), FITC-conjugated IgG1 and PE-conjugated IgG2a (679.1Mc7 and U7.27; Immunotech), and tricolor- or FITC-conjugated IgG2a (MG2a06; Caltag or 20102.1; R&D Systems, Minneapolis, MN).

To assess the reactivity of mAbs against peripheral blood or synovial fluid lymphocyte flow cytometry was used. All incubations were performed for 30 min at 4°C. Cells were washed twice in PBS containing 1% BSA (PBS-BSA) and adjusted to 1 × 106 cells/ml in staining buffer (PBS-BSA containing 0.1% sodium azide) and blocked with the appropriate normal serum. For three-color staining, indirect staining was accomplished first by incubation with the purified Abs directed against CCR4, CCR5, or an isotype-matched control (5 μg/ml, 2 × 105 cells/tube), washed two times in PBS-BSA, then incubated with FITC- or PE-conjugated goat anti-mouse IgG (H+L) and washed. In a second step, the appropriate directly conjugated cell surface marker (CD3, CD4, CD8, CD45RO, TCRαβ, TCRγδ) or isotype-matched Abs were then used as directed by the manufacturer. The resulting cells were analyzed by FACScan (BD Biosciences, Mountain View, CA) using electronic gating and compensation. CellQuest software (BD Biosciences) was used for analysis.

The ligands for CCR4, TARC, and MDC were measured in synovial fluid (JRA and RA) and plasma (JRA and control) by sandwich ELISA. Samples were residual supernatants from Ficoll isolation of lymphocytes. To assay for TARC, plates were coated with 1 μg/ml of monoclonal anti-human TARC Ab (clone 54026.11; R&D Systems), and biotinylated anti-human TARC-specific goat IgG was used for detection. Recombinant human TARC (R&D Systems) was used as the standard. For each sample, four serial 2-fold dilutions were analyzed in duplicate and the mean presented for values within the limits of the standard curve (15–1000 pg/ml). MDC was measured using the Quantikine MDC/CCL32 kit (R&D Systems) according to the manufacturer’s instructions. For each sample (same group used in the TARC assay), two serial 2-fold dilutions were analyzed in duplicated and the mean presented for values within the limits of the standard curve (62.5–1000 pg/ml).

After staining with anti-CD4 and anti-CCR4, a FACSVantage (BD Biosciences) was used to separate CD4+/CCR4+ and CD4+/CCR4 populations. For RT-PCR, RNA was isolated from sorted populations using RNAeasy (Qiagen, Chatsworth, CA). The number of cells available for RNA extraction varied dramatically between patients and between subpopulations (range 100,000 to 3 × 106). There were fewer cells in the CCR4+ population than in the CCR4 population for all patient samples. First strand cDNA was synthesized using oligo(dT) as primer and SuperScript II RNase H Reverse Transcriptase (Life Technologies, Gaithersburg, MD) as previously described (24). IL-4 and IFN-γ gene expression was determined relative to β-actin expression levels. Briefly, 2-fold serial dilutions of β-actin competitive fragment (CF; Clontech, Palo Alto, CA) were added to PCR containing constant amounts of cDNA. β-actin PCR products included a 838 bp product and a corresponding 619 bp CF product (primers 5′-ATCTGGCACCACACCTTCTACAATGAGCTGCG-3′ and 5′-CGTCATACTCCTGCTTGCTGATCCACATCTGC-3′). β-actin expression was determined as equal to the number of attomoles of CF used when equivalent band intensities were achieved for sample and CF. For cytokine PCR, template addition based on β-actin equivalents varied between patients but was always identical for CCR4+ and CCR4 populations from a single sample. Sufficient RNA was not available to quantitate IL-4 and IFN-γ by CF PCR. IL-4 mRNA (345 bp product, forward primer 5′-CAACTTTGTCCACGGACAC-3′, reverse primer 5′-TCCAACGTACTCTGGTTGG-3′), and IFN-γ mRNA (427 bp product, forward 5′-GCATCGTTTTGGGTTCTCTTGGCTGTTACTGC-3′, reverse 5′-CTCCTTTTTCGCTTCCCTGTTTTAGCTGCTGG-3′) were detectable by PCR. PCR was performed in a 25 μl volume containing 1.5 mM MgCl2, 10 mM Tris-HCl (pH 8.3), 50 mM KCl, 0.4 μM each primer, 0.2 mM each dNTP, and 1.25 U of Taq Gold DNA polymerase (PE Biosystems, Foster City, CA) using 35 cycles of amplification (94°C 1 min, 57°C 1 min, 72°C 1 min) after an initial hold at 95°C for 10 min.

Student’s t test was used for normally distributed variables. When necessary, natural log transformation was performed. The Pearson correlation was used to assess the correlation between clinical features and the levels of CCR4 or CCR5 expression. Fisher’s exact test was used to assess the frequency of TARC detection between patient and sample subsets. In all instances, p < 0.05 was considered significant.

Paired synovial fluid and peripheral blood samples were available from 15 JRA patients, whereas nine additional patients contributed only a synovial sample (Table I). The mean age at onset was 6.7 years (range 1–13 years), and the mean disease duration was 7 years (range 0–17 years). These patients included pauciarticular, polyarticular, systemic onset types as well as spondyloarthropathies.

T cell subsets exhibit distinct capacities to migrate to sites of inflammation, and this may in part reflect differential expression of chemokine receptors. The synovial fluid T cell populations contain significantly lower frequencies of total CD4+ cells than paired peripheral blood (Fig. 1,A). In contrast, no statistical difference was found for total CD8+ populations in the paired samples (Fig. 1,B). In addition, both the CD4+ and CD8+ populations show a wider range of values in the synovial compartment (CD4+, 47.0 ± 17.0%, CD8+, 39.8 ± 16.9%; mean ± SD) than peripheral blood (CD4+, 60.4 ± 5.8%; CD8+, 30.8 ± 6.9%). The T cells found in the synovium tend to be CD45RO+, in agreement with previously published studies in JRA. In addition, the synovium has higher percentages of CD3+ cells expressing the γδ TCR and lower percentages of αβ TCR (Table II). Considering CD45RO+ cells alone, there are significantly more CD8+ cells in the synovial fluid than in the blood (33.4 ± 18.0 vs 13.6 ± 4.6%).

FIGURE 1.

Phenotypic characterization of T cells in paired synovial fluid and peripheral blood for CD4 (A) or CD8 (B). The median values for patient peripheral blood samples of CD4+ and CD8+ cells were 60.4 and 30.8% of the CD3+ cells. In contrast, the synovial fluid samples displayed an overall decrease in the CD4+ population (mean = 47.0%) and an accompanying increase in CD8+ (39.8%) relative to the peripheral blood sample. Values of p are shown in the upper right corner. Nonsignificant p values (exceeding 0.05) are designated as “NS”.

FIGURE 1.

Phenotypic characterization of T cells in paired synovial fluid and peripheral blood for CD4 (A) or CD8 (B). The median values for patient peripheral blood samples of CD4+ and CD8+ cells were 60.4 and 30.8% of the CD3+ cells. In contrast, the synovial fluid samples displayed an overall decrease in the CD4+ population (mean = 47.0%) and an accompanying increase in CD8+ (39.8%) relative to the peripheral blood sample. Values of p are shown in the upper right corner. Nonsignificant p values (exceeding 0.05) are designated as “NS”.

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Table II.

Comparison of cell surface markers in JRA-matched peripheral blood and synovial fluid samples

%CD3%CD45RO
TCR αβ+TCR γδ+CD4+CD8+TCR γδ+
Peripheral blood 95.9 ± 2.0a 4.2 ± 3.3 56.2 ± 12.0 13.6 ± 4.6 6.3 ± 5.0 
Synovial fluid 90.4 ± 7.3 7.5 ± 5.6 50.2 ± 23.7 33.4 ± 18.0 4.5 ± 4.1 
p 0.023b 0.006c 0.56 0.004 0.035c 
%CD3%CD45RO
TCR αβ+TCR γδ+CD4+CD8+TCR γδ+
Peripheral blood 95.9 ± 2.0a 4.2 ± 3.3 56.2 ± 12.0 13.6 ± 4.6 6.3 ± 5.0 
Synovial fluid 90.4 ± 7.3 7.5 ± 5.6 50.2 ± 23.7 33.4 ± 18.0 4.5 ± 4.1 
p 0.023b 0.006c 0.56 0.004 0.035c 
a

Values represent mean ± SD percentage of CD3+ or CD45RO+ cells for indicated cell surface markers.

b

Paired comparison of peripheral blood and synovial fluid samples was performed using the Student t test.

c

Non-normally distributed variables were natural log transformed prior to t test.

Because chemokines and their receptors play roles in T cell trafficking to the inflamed joint, we examined expression of CCR4 and CCR5 on lymphocytes present in joint fluid vs the periphery for 15 JRA patients by flow cytometric analysis. For these studies, three-color staining was used to detect CD3, CD4, and CCR4 or CCR5 on the cell surface. In both patients shown in Fig. 2, substantial increases were found in the frequency of CD3+ cells that express CCR5 in the synovial fluid, and this increase was seen for both CD4+ and CD4 populations. These findings are representative of the patients in general, where large but varying degrees of increases between peripheral blood and synovial fluid were found (Fig. 3,A). CCR5 expression in the peripheral blood ranged from ∼1 to 8% of the CD3+ cells (mean = 3.5%), whereas in the synovial fluid a wider distribution ranging from 19 to 94% of the CD3+ cells (mean = 56.2%) was found. It is noteworthy that of the 15 patients illustrated in Fig. 2, the two patients with the lowest levels of CCR5 in their synovial fluid samples were heterozygous for the well-characterized CCR5 Δ32 mutation (23) and therefore had only one functional CCR5 gene.

FIGURE 2.

Ab staining of CD3, CD4, and either CCR4 or CCR5 for paired peripheral blood and synovial fluid samples from two representative JRA patients. The dot plots represent expression of CD4 and either CCR4 (top) or CCR5 (bottom) for peripheral blood and synovial fluid samples from patients 6 and 12 (CD3 gate). At least 20,000 events were collected per tube, and the live lymphocyte population was selected by the light scatter profile for analysis. Values for each quadrant are the percentages of CD3+ gated cells.

FIGURE 2.

Ab staining of CD3, CD4, and either CCR4 or CCR5 for paired peripheral blood and synovial fluid samples from two representative JRA patients. The dot plots represent expression of CD4 and either CCR4 (top) or CCR5 (bottom) for peripheral blood and synovial fluid samples from patients 6 and 12 (CD3 gate). At least 20,000 events were collected per tube, and the live lymphocyte population was selected by the light scatter profile for analysis. Values for each quadrant are the percentages of CD3+ gated cells.

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FIGURE 3.

Comparison of frequencies of CD45RO, CCR5, and CCR4 between normal peripheral blood, patient peripheral blood, and synovial fluid samples. Graphs represent frequency of the CD45RO or chemokine receptor in paired peripheral blood and synovial fluid samples (connected by lines) or normal peripheral blood (nPB) as determined by three-color flow cytometric analysis. Solid bars represent the means. A, Percentage of CD3+ cells that express CCR5. Samples from patients with wild-type/wild-type CCR5 genotypes (♦) or heterozygous (⋄) for the Δ32 mutation are designated. B, Percentage of CD3+ cells that express CCR4. C, Percentage of CD4+ cells that express CCR4. D, Percentage of CD3+ cells that express the memory cell phenotype (CD45RO+). E, Percentage of CD45RO+ cells that express CCR5. F, Percentage of CD45RO+/CD4+ cells that express CCR4. Values of p using Student’s t test between normal peripheral blood and patient peripheral blood, and between the patient peripheral blood and synovial fluid, are indicated above the respective sample type. Variables were natural log transformed when necessary to achieve normal distributions.

FIGURE 3.

Comparison of frequencies of CD45RO, CCR5, and CCR4 between normal peripheral blood, patient peripheral blood, and synovial fluid samples. Graphs represent frequency of the CD45RO or chemokine receptor in paired peripheral blood and synovial fluid samples (connected by lines) or normal peripheral blood (nPB) as determined by three-color flow cytometric analysis. Solid bars represent the means. A, Percentage of CD3+ cells that express CCR5. Samples from patients with wild-type/wild-type CCR5 genotypes (♦) or heterozygous (⋄) for the Δ32 mutation are designated. B, Percentage of CD3+ cells that express CCR4. C, Percentage of CD4+ cells that express CCR4. D, Percentage of CD3+ cells that express the memory cell phenotype (CD45RO+). E, Percentage of CD45RO+ cells that express CCR5. F, Percentage of CD45RO+/CD4+ cells that express CCR4. Values of p using Student’s t test between normal peripheral blood and patient peripheral blood, and between the patient peripheral blood and synovial fluid, are indicated above the respective sample type. Variables were natural log transformed when necessary to achieve normal distributions.

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CCR4 is found almost exclusively on CD4+ T cells (Fig. 2) in both peripheral blood and synovial fluid samples. The percentage of CD3+ cells expressing CCR4 (Fig. 3,B) is uniformly low in patient blood samples (4.9 ± 1.3%), whereas synovial fluid values (9.2 ± 6.2%) were significantly higher (p = 0.012, paired t test, two-tailed). Representative patients with high (patient 6) or low (patient 12) levels of CCR4+ cells are illustrated in Fig. 2. Of the 15 patients examined, 12 show higher numbers of CD3+ cells expressing CCR4 in their fluid compared with blood. This increase is even more striking when CCR4 expression is considered on CD4+ cells (Fig. 3,C, peripheral blood, 7.9 ± 2.1; synovial fluid, 22.9 ± 14.7, p = 0.001, paired t test, two-tailed). Comparison of findings between patient and normal blood (n = 9) revealed significantly higher percentages of CCR4+ and CCR5+ in the normal samples (Fig. 3, A–C).

As expected, the percentage of T cells (CD3+) expressing the CD45RO isoform was higher in patient synovial fluid than in the paired blood samples. In addition, CD45RO+ cells are also found at higher frequency in normal than in patient peripheral blood (Fig. 3 D). This comparison included only the normal controls (n = 5) that were <25 years of age. The patients ranged in age from 2 to 26 years at the time of sample with only five of the patients under 10 years old. No relationship was noted for patient age and CD45RO+ cell frequency. In patient synovial fluid most T cells expressed CD45RO+ (83.4 ± 11.7%), and this population included essentially all CD4+ cells. Because most T cell chemoattractants selectively attract memory/activated T cells (25, 26, 27), we next examined the CCR4+ and CCR5+ populations within the memory cell compartment.

CCR5 is expressed largely within the CD45RO+ subpopulation of T cells in normal adults (12, 19) and within JRA peripheral blood and synovial fluid samples as previously demonstrated (4). In normal adults CCR4 expression is further limited to the CD4+/CD45RO+ subpopulation of T cells (28). We have found the CCR4 cell surface expression is limited to this subpopulation in both our patient fluid and blood samples. Although increases in CCR5 expression were found for synovial fluid relative to peripheral blood CD45RO+ cells (Fig. 3,E), increases in CCR4 expression for CD45RO+ T cells were not uniformly seen (Fig. 3,F). This may not be surprising, due to the difference in the magnitude of increase between peripheral blood and synovial fluid of CCR4+ cells relative to CCR5+ cells (i.e., comparing Fig. 3, A and B).

As the next step in determining a functional role for CCR4 in the JRA joint, we looked for its ligands, TARC and MDC. TARC is a recently identified lymphocyte-directed CC chemokine that specifically chemoattracts CD4+ T cells in humans through the CCR4 molecule. The samples assayed by ELISA included 17 synovial fluid supernatants and six blood supernatants with origins identical with flow cytometry samples, and two fluid and three blood samples obtained at different dates from study patients. For comparison, TARC levels were also measured in synovial fluid supernatants from five RA patients and in nine healthy adult plasma samples. As shown in Fig. 4 A, TARC was detectable at similar frequencies in JRA patient synovial fluid (16/19 samples) and plasma (9/9 samples, p = 0.30, Fisher’s exact test). The finding of TARC in JRA plasma contrasts with detection in healthy adults (1/9 samples, p = 0.0001, Fisher’s exact test). In contrast to JRA, TARC was only found in one of the five RA synovial fluid samples, consistent with the low levels of CCR4 in RA fluid (p = 0.014, Fisher’s exact test). Although much variation in levels exists between individuals, TARC was consistently detected in the JRA, but not RA or normal samples, suggesting a role for the CCR4/TARC receptor/ligand pair in JRA.

FIGURE 4.

CCR4 ligands TARC and MDC are present in JRA synovial fluid supernatants and JRA plasma. TARC and MDC were measured by ELISA for JRA synovial fluid supernatants (19 samples for TARC and 18 for MDC), nine JRA plasma samples (lines connect paired samples), 10 normal control plasma samples, and five RA synovial fluid samples. A, TARC median values for JRA synovial fluid and peripheral blood, normal plasma, and RA synovial fluid were 105, 139, 0, and 0 pg/ml, respectively. Statistical differences were found between the normal plasma and JRA plasma (p = 0.001) and JRA synovial fluid vs RA synovial fluid (p = 0.019). Statistical evaluation of either the matched synovial fluid and peripheral blood pairs or all JRA samples yielded no evidence of significant differences in TARC levels between the JRA synovial fluid and peripheral blood. MDC was detected in JRA synovial fluid (591.8 ± 260.9, 550.0 pg/ml; mean ± SD, median), JRA peripheral blood (756.1 ± 339.0, 839.0 pg/ml), normal plasma (609.6 ± 195.8, 608.5 pg/ml), and RA synovial fluid (400.8 ± 164.0, 418 pg/ml). No significant differences were found between groups of samples.

FIGURE 4.

CCR4 ligands TARC and MDC are present in JRA synovial fluid supernatants and JRA plasma. TARC and MDC were measured by ELISA for JRA synovial fluid supernatants (19 samples for TARC and 18 for MDC), nine JRA plasma samples (lines connect paired samples), 10 normal control plasma samples, and five RA synovial fluid samples. A, TARC median values for JRA synovial fluid and peripheral blood, normal plasma, and RA synovial fluid were 105, 139, 0, and 0 pg/ml, respectively. Statistical differences were found between the normal plasma and JRA plasma (p = 0.001) and JRA synovial fluid vs RA synovial fluid (p = 0.019). Statistical evaluation of either the matched synovial fluid and peripheral blood pairs or all JRA samples yielded no evidence of significant differences in TARC levels between the JRA synovial fluid and peripheral blood. MDC was detected in JRA synovial fluid (591.8 ± 260.9, 550.0 pg/ml; mean ± SD, median), JRA peripheral blood (756.1 ± 339.0, 839.0 pg/ml), normal plasma (609.6 ± 195.8, 608.5 pg/ml), and RA synovial fluid (400.8 ± 164.0, 418 pg/ml). No significant differences were found between groups of samples.

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MDC, also a potent chemoattractant for chronically activated Th2 lymphocytes, was measured for the same group of samples described above (with the exception of one sample that was no longer available). In contrast to our findings with TARC, MDC was detected in all JRA, RA, and normal synovial fluid and peripheral blood samples tested. Furthermore, no differences were found between these sample subsets (Fig. 4 B), and no correlation was found between TARC and MDC levels.

To determine the relationship between type 1/type 2 cytokine mRNA expression and chemokine receptor expression, synovial fluid lymphocytes from three patients were sorted into CD4+CCR4+ and CD4+CCR4 populations. β-actin expression was rigorously quantitated using CF RT-PCR, allowing adjustment of cDNA stock to provide for equal β-actin PCR products. This is illustrated for patient 7 (Fig. 5, top); 2 μl of a 1/20 stock dilution for both CCR4 and CCR4+ samples yielded PCR product equivalent to 0.05 amol of the β-actin CF. For IL-4 cytokine detection, cDNA equivalent to 1.5 and 3.0 amol β-actin served as template for patients 7 and 19. For both patients, IL-4 product could be detected for CCR4+ but not CCR4 subpopulations. Sufficient cDNA for patient 6 was not available for IL-4 RT-PCR. For IFN-γ, cDNA equivalent to 0.4 amol of β-actin was used as PCR template. This analysis revealed lower levels of IFN-γ expression in the CD4+CCR4+ than in CD4+CCR4 cells in all three patients.

FIGURE 5.

Cytokine expression patterns vary relative to CCR4 cell surface expression. cDNA derived from synovial fluid lymphocytes separated by FACS into CD4+/CCR4+ and CD4+/CCR4 populations was subjected to β-actin-specific CF PCR to provide equal templates for cytokine-specific PCR. As shown in the top panel for patient 7, β-actin PCR product is present in amounts equal to the product obtained using 0.05 amol of the β-actin CF for both CCR4+ and CCR4 fractions. For cytokine mRNA detection (IL-4, middle; IFN-γ, bottom), cDNA equal to 1.5 or 3.0 amol (IL-4 for patients 7 and 19, respectively) or 0.4 amol (IFN-γ, all patients) of β-actin served as template for cytokine PCR for CD4+/CCR4+ (4+) or CD4+/CCR4 (4−) fractions. dH2O as template in PCR served as negative control (neg). Marker (100 bp ladder) was always included; the 600 bp fragment is indicated.

FIGURE 5.

Cytokine expression patterns vary relative to CCR4 cell surface expression. cDNA derived from synovial fluid lymphocytes separated by FACS into CD4+/CCR4+ and CD4+/CCR4 populations was subjected to β-actin-specific CF PCR to provide equal templates for cytokine-specific PCR. As shown in the top panel for patient 7, β-actin PCR product is present in amounts equal to the product obtained using 0.05 amol of the β-actin CF for both CCR4+ and CCR4 fractions. For cytokine mRNA detection (IL-4, middle; IFN-γ, bottom), cDNA equal to 1.5 or 3.0 amol (IL-4 for patients 7 and 19, respectively) or 0.4 amol (IFN-γ, all patients) of β-actin served as template for cytokine PCR for CD4+/CCR4+ (4+) or CD4+/CCR4 (4−) fractions. dH2O as template in PCR served as negative control (neg). Marker (100 bp ladder) was always included; the 600 bp fragment is indicated.

Close modal

CCR5+ and CCR4+ T cell levels were examined relative to gender, onset type, course, age at onset, destructive disease, and disease duration to determine clinical relationships to the observed variation in chemokine receptor levels. Chemokine receptor and T cell surface marker data were available from 23 synovial fluid samples. There were three patients (total of 18 genotyped) who possessed the CCR5 Δ32 mutation; these also had the lowest levels of CCR5. Interestingly, the highest level of CCR4 was found in the single patient who was homozygous for the CCR5 Δ32 mutation. Based on these biases, patients who possessed CCR5 Δ32 were excluded from statistical analysis for both CCR4 and CCR5.

A correlation was found for CCR4 and disease duration (r = 0.4533, p = 0.034) but not between CCR4 levels and gender, onset age, onset type, course, or destructive disease. Specifically, CCR4 levels tended to be higher in samples obtained at early stages of disease (Fig. 6,A). In contrast to CCR4, no correlation was found for synovial fluid CCR5 expression levels and disease duration (Fig. 6,B) or any other clinical feature. It is interesting that concurrent joints for single patients reveal nearly identical findings for all markers examined (Fig. 7).

FIGURE 6.

Correlation of chemokine receptor expression and disease duration. A, Correlation of percentage of CD4+ cells that express CCR4 and the duration of JRA (includes data from all synovial samples analyzed for CCR4 expression regardless of availability of matching blood sample). B, Correlation of the percentage of CD3+ cells that express CCR5 and the duration of JRA at sampling. Symbols indicate the CCR5 genotype (♦, wild-type/wild-type; •, wild-type/Δ32; ▴, Δ32/Δ32). Spearman correlation and p values are shown in the bottom right corner of each graph and included only wild-type/wild-type samples.

FIGURE 6.

Correlation of chemokine receptor expression and disease duration. A, Correlation of percentage of CD4+ cells that express CCR4 and the duration of JRA (includes data from all synovial samples analyzed for CCR4 expression regardless of availability of matching blood sample). B, Correlation of the percentage of CD3+ cells that express CCR5 and the duration of JRA at sampling. Symbols indicate the CCR5 genotype (♦, wild-type/wild-type; •, wild-type/Δ32; ▴, Δ32/Δ32). Spearman correlation and p values are shown in the bottom right corner of each graph and included only wild-type/wild-type samples.

Close modal
FIGURE 7.

Analysis of concurrent synovial fluid samples. During the course of the study, samples from three patients (patients 9, 23, and 1) provided the opportunity to analyze cell surface expression in multiple knee joints (designated L, left knee and R, right knee) for CD4 and CD8 (A) or CCR4 and CCR5 (B). CCR4 information was not available for patient 1. For each patient, expression frequencies for all markers were similar in both joint samples obtained at single clinic visits.

FIGURE 7.

Analysis of concurrent synovial fluid samples. During the course of the study, samples from three patients (patients 9, 23, and 1) provided the opportunity to analyze cell surface expression in multiple knee joints (designated L, left knee and R, right knee) for CD4 and CD8 (A) or CCR4 and CCR5 (B). CCR4 information was not available for patient 1. For each patient, expression frequencies for all markers were similar in both joint samples obtained at single clinic visits.

Close modal

This study explores CCR4 and CCR5 expression in JRA and the functional consequences in terms of type 1/type 2 cytokine expression. We report the accumulation of CD4+ cells expressing CCR4 in JRA joints. These findings are in contrast to RA, where selective accumulation of CCR4+ cells in the synovial fluid was not found (20). CCR5+ T cells were found at greater frequencies in JRA synovial fluid samples compared with peripheral blood, in agreement with previous work in adult (18, 19, 20) and juvenile forms of rheumatoid arthritis (4).

Because chemokines selectively attract memory cells (25, 26, 27), we compared CCR4 and CCR5 expression within this subset of T cells. We found that the expression of CCR5, but not CCR4, was higher in the joint for CD45RO+ cells. Although this raises the possibility that the observed increases in CCR4+ when cells are gated on CD3 or CD4 are secondary to the increase in CD45RO+ cells, we provide abundant evidence for a significant role for CCR4+ T cells in JRA.

The higher values of CCR4+ cells in normal than in patient blood suggest that trafficking of the CCR4+ cells from the periphery to the joint has occurred. This is also true for CCR5+ and CD45RO+ cells (Fig. 3, A–D). Although increases in CCR4+ T cells were not demonstrated in the synovial fluid of RA patients, a selective decrease of CCR4+ (and CCR5+) T cells in blood of RA patients compared with normal or osteoarthritis was reported (20). Analysis of concurrent joint samples provided nearly identical results for both knees in the three patients examined. Together these findings suggest that CCR4+ cells (and also CCR5+ cells) play a role in a systemic disease component. This is consistent with the finding that arthritis can be generated by systemic recognition of self-MHC-peptide complexes by autoreactive T cells that trigger B cells to produce arthritogenic Abs in a mouse model of spontaneous arthritis (29, 30).

MDC, RANTES, and TARC have all been identified as ligands for CCR4. Assays for two of these, TARC and MDC, were used to demonstrate these ligands in JRA. TARC was found in nearly all synovial fluid and peripheral blood of JRA patients but not in synovial fluid of RA patients or normal plasma. In addition, it is noteworthy that no correlations were found between either TARC or MDC levels and the number of CCR4+ cells. The fact that TARC detection was limited to JRA samples suggests a disease-specific role.

In contrast, MDC was detected in all samples assayed; furthermore, there were no statistically significant differences in the levels of MDC between JRA patients and normal blood samples. This may not be surprising because JRA does have a strong type 1 component, and increases in MDC serum levels relative to healthy adults have been reported for type 2-dominated disorders such as atopic dermatitis and mycosis fungoides/Sézary syndrome but not for type 1-dominated disorders such as multiple sclerosis or Crohn’s disease (31). Although only a relatively small number of samples were studied, two interesting observations are apparent. First, for the six JRA patients with information available for paired blood and fluid samples, levels of MDC were higher in the blood for five patients. Second, although only approaching statistical significance (p = 0.073), MDC levels in JRA synovial fluid (591.8 ± 260.9) tend to be higher than in RA synovial fluid (400.8 ± 164.0). Although these findings do not allow us to define a role for MDC in JRA, we can speculate that JRA and RA use different mechanisms of T cell recruitment into the synovium that can only be elucidated by further study.

The CCR5 ligands, macrophage-inflammatory protein-1α and RANTES, are abundantly produced in inflamed RA but not osteoarthritic joints (20). Furthermore, in support of an important role for CCR5 in disease, macrophage-inflammatory protein-1α and RANTES mRNAs have been detected at higher levels in JRA compared with control nonautoimmune synovial tissue samples by RNase protection assays.4

Cytokine mRNA was measured in CD4+ T cells sorted for CCR4 expression to determine the functional relevance of this population of cells. RT-PCR with rigorous quantitation of template was used to avoid the nonspecific stimulation required for intracellular cytokine staining. The small number of CD4+/CCR4+ cells available in the pediatric synovial fluid samples limited the amount of RNA available for RT-PCR and prevented the use of CF PCR for cytokine quantitation.

Even with the limited cell numbers available, our findings show a clear difference in cytokine mRNA between CD4+/CCR4 and CD4+/CCR4+ T cell populations. IL-4 detection was especially difficult because, in general, IL-4 is produced by synovial CD4+ cells at much lower levels (20- to 400-fold lower, our unpublished observation) than IFN-γ. This is highlighted by the much brighter band intensities found for IFN-γ than IL-4, even though 4- to 7-fold more cDNA was used for IL-4 detection. These cytokine mRNA expression studies suggest that the CD4+CCR4+ cells exhibit a type 2 cytokine profile providing evidence for an anti-inflammatory role in JRA.

CCR4 expression has been associated with a type 2 profile but has not been previously reported in arthritis to our knowledge. In addition, this appears to be unique to the juvenile form of arthritis, because a study of RA joints found no increase in the synovial fluid CCR4+ T cells (20) or TARC. Interestingly, and consistent with CCR4 as a type 2 marker, IL-4 mRNA has been detected in a subset of JRA samples (7) but is less prominent in RA synovium (32, 33) and end stage JRA synovium.4 These observations are compatible with the apparent down-regulation of IFN-γ and possible up-regulation of IL-4 by CCR4-expressing cells.

We report increased levels of CCR5 on synovial T cells in a well-defined group of JRA patients. Increased expression of CCR5 has been associated with a type 1 cytokine profile (12, 14, 17) and is dependent on recent activation events (9, 12, 34). Increased levels of CCR5+ cells have been demonstrated for synovial T cells in RA (19, 20) as well. The CCR5 Δ32 mutation was found in three of the patients included in this study (two heterozygous, one homozygous), consistent with the published gene frequency of 0.14 (35). It is interesting that the patient with the highest frequency of CD4+/CCR4+ T cells was homozygous for the Δ32 mutation, even though individuals that are homozygous or heterozygous for mutant CCR5 alleles do not appear to exhibit defective T cell responses (36, 37, 38).

We speculate that the elevated levels of CCR4 may be a counterreaction to type 1-driven inflammation. In this case, the majority of T cells within the JRA inflamed joint are polarized toward a type 1 phenotype with a variable anti-inflammatory component present, especially in early disease. Furthermore, our data suggest that this component may not be present in RA. This view is consistent with a generally better outcome for JRA than is the case for RA.

There has been recent evidence that, at least in a mouse model of spontaneous RA, a joint-specific process is not the initiating event (29). The very similar frequencies of CCR4+ and CCR5+ T cells found in different joints tested simultaneously are in agreement with this view. Identical expanded T cell clones have also been reported in simultaneous joints (39). The differences between the fraction of CD45RO+ memory T cells that express CCR4+ or CCR5+ in the synovial fluid is of interest. In contrast to CCR4, CCR5+ T cell accumulation into the joint occurs even within the CD45RO+ memory subset. Although nearly 100% of the CCR4+ cells express CD45RO, the proportion of the CD4+/CD45RO+ cells expressing CCR4+ is not consistently increased for individual patients. A general increase in CCR4+/CD4+/CD45RO+ T cells, which are not joint-specific, would be consistent with the view that the TARC/CCR4 receptor/ligand pair is involved in systemic immunity and not specific local immunity (40). Alternately, the observation of decreased CCR4+ cells in the blood of JRA patients and the accompanying TARC expression may be due to the serum TARC causing a down-regulation of the CCR4 receptor expression on the PBLs.

In conclusion, we have demonstrated the selective accumulation of the CCR4+ T cells in the joint fluid of JRA patients. Our finding of differences in cytokine mRNA in the sorted populations also suggests that these cells play anti-inflammatory roles especially in the early years of disease. The reported differences for CCR4 and its ligand TARC between JRA and RA, suggest that different therapeutic strategies will be necessary to effectively treat pediatric patients.

We thank Drs. A. Grom, M. Scola, C. Karp, and J. Rottman for discussion and critical review of the manuscript, Drs. J. Rottman and L. Wu (Millenium Pharmaceuticals) for providing the CCR4 and CCR5 mAbs, and Drs. R. Colbert, T. Griffin, A. Grom, R. Hirsch, D. Lovell, and L. Houk for providing the patient samples.

1

This work was supported in part by the Children’s Hospital Research Foundation of Cincinnati, the Schmidlapp Foundation, and National Institutes of Health Grants AR39979, AR47363, and AR44059.

3

Abbreviations used in this paper: JRA, juvenile rheumatoid arthritis; RA, adult rheumatoid arthritis; TARC, thymus- and activation-regulated chemokine; CF, competitive fragment; MDC, macrophage-derived chemokine; CXCR, CXC chemokine receptor; PBS-BSA, PBS containing 1% BSA.

4

M. P. Scola, S. D. Thompson, H. I. Brunner, M. K. Tsoras, D. Witte, A. A. Grom, M. A. van Dijk, M. H. Passo, J. B. Rottman, and D. N. Glass. Increased expression of IL-12p35, IL-15 and IL-18 in juvenile rheumatoid arthritis synovial tissue. Submitted for publication.

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