ARTHRITIS & RHEUMATISMVol. 54, No. 6, June 2006, pp 2022–2030© 2006, American College of Rheumatology

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DOI 10.1002/art.21909

Temporal expression pattern of Duffy antigen inrheumatoid arthritis: up-regulation in early disease

The Duffy antigen is a chemokine-binding protein (orinterceptor) with broad specificity, binding inflammatory che-mokines of both CXC and CC classes with high affinity, but notconstitutive (homeostatic) chemokines (1–3). Although it isexpressed by red blood cells and some cells of other types, insynovial tissue the Duffy antigen protein shows a highlyselective distribution, localizing to the endothelial cells ofvenules (4–6). Recent evidence suggests that Duffy antigen inthe endothelium could play a role in leukocyte migration atsites of inflammation. The protein facilitates the transport ofchemokines across endothelial cells, resulting in enhancedleukocyte transendothelial migration (7,8). Consistent with thisrole, the Duffy antigen localizes to caveolae (plasmalemmalvesicles), which are involved in transcellular transport ofmolecules (9,10). In addition, mice lacking Duffy antigen showaltered recruitment of leukocytes to sites of inflammation(11–13). The aim of the present study was to examine thetemporal pattern of expression of the Duffy antigen from earlyrheumatoid arthritis (RA) (symptoms of �1 year) to long-standing disease (duration 9 years and 18 years).

Synovia were obtained by arthroscopic needle biopsyof the knee joints of 6 patients with early RA, diagnosedaccording to the criteria of the American College of Rheuma-tology (formerly, the American Rheumatism Association)(14). The duration of disease in these patients was �7 months(mean � SEM 4.2 � 0.9 months) (Table 1). The patients weretreated with nonsteroidal antiinflammatory drugs (NSAIDs)only, or received no medication. Synovia from patients withlongstanding RA (disease duration �5 years) were obtained atthe time of knee replacement surgery. Two groups of patientswith longstanding RA were studied, one with a mean � SEMdisease duration of 8.8 � 1.6 years (n � 6) and the other witha disease duration of 18.3 � 2.5 years (n � 10). Almost allpatients in both groups were receiving disease-modifying anti-rheumatic drugs (DMARDs) or steroids (Table 1).

Synovial biopsy specimens were obtained arthrosco-pically from the knees of 10 non-RA patients who had jointsymptoms suggestive of articular cartilage, meniscal, or ante-rior cruciate ligament damage. In all RA and non-RA patients,samples were obtained from the suprapatellar pouch andmedial gutter, which are reported to provide representativesampling of synovial membrane pathology (15). Synovia frompatients with longstanding RA showed classic RA histopathol-ogy, including perivascular mononuclear aggregates in thesubintima and intimal layer thickening. Synovia from patientswith early RA were infiltrated, exhibiting a diffuse pattern withless development of perivascular mononuclear aggregatescompared with samples from patients with longstanding RA.Non-RA patients showed markedly less mononuclear infiltra-tion and intimal layer thickening compared with RA patients.

Cryostat sections (10 �m thick) were fixed in formalinfor 10 minutes and washed in phosphate buffered saline.Immunohistochemistry analysis for the Duffy antigen wasperformed as described by Patterson et al (5), using ananti-human Duffy monoclonal antibody (mAb) (2C3; 20 �g/ml) (16), kindly donated by Pr. D. Blanchard (University of

Nantes, Nantes, France). An immunoperoxidase staining sys-tem (Vector, Burlingame, CA) with diaminobenzidine (Sigma,Poole, UK) as substrate, was used for detection. Duffy stainingwas quantified by counting the total number of positive andnegative blood vessels in 4 randomly selected fields of view ata magnification of 400�. Staining in 2 sections from eachsynovial sample was quantified. From this the percentage ofpositive vessels was obtained, and mean � SEM values deter-mined. The total number of vessels counted per patient was�50. Student’s t-test was used to determine the significance ofdifferences between groups.

For double-label immunofluorescence staining ofDuffy antigen and �v�3 integrin, cryosections were fixed for 10minutes in ice-cold acetone and incubated with anti-DuffymAb as above, followed by incubation with goat anti-mouseAlexa Fluor 488 (1:200; Molecular Probes, Eugene, OR) in10% human serum. Sections were blocked in mouse serum(1:20) and then incubated with biotin-conjugated anti-human�v�3 mAb (10 �g/ml; Chemicon, Southampton, UK) followedby streptavidin–Alexa Fluor 594 (1:1,000; Molecular Probes)containing 10% human serum.

Immunohistochemistry analysis revealed Duffy antigenexpression in the endothelial cells of postcapillary venules inall RA and non-RA synovia (Figures 1A–D); arterioles werenegative for Duffy antigen. Staining was negative when sec-tions were treated with isotype-matched control mouse IgGinstead of primary antibody. Synovial sections treated withantibodies to other endothelial markers, including von Wille-brand factor, CD31, CD34, CD105, and CD146, showed adifferent distribution pattern compared with that demon-strated for Duffy antigen, with labeling of endothelial cells inall vessels, i.e., postcapillary venules, capillaries, or arterioles(6). The percentage of vessels staining positive for Duffyantigen (Figure 1E) was 63.1 � 3.3% (mean � SEM) innon-RA tissue. This proportion was significantly higher insynovia from patients with early RA (79.8 � 2.3%) (P �0.013). With increased disease duration the proportion ofpositive vessels decreased, such that at 9 years it was 62.7 �8.8%, which approached statistical significance compared withearly RA (P � 0.051). At 18 years’ disease duration, Duffyantigen expression had further declined, with a mean percent-age positive vessels of 45.7 � 3.9%; this value was highlysignificantly lower than that in samples from patients with earlyRA (P � 0.00005) and significantly lower than that in non-RAsamples (P � 0.02).

These results show a temporal and phasic pattern ofDuffy antigen expression. In the early stage of RA (within thefirst few months), there is an up-regulation of the Duffyantigen in the venules of the synovium. Following this, withincreasing disease duration, there is a down-regulation ofDuffy antigen expression such that at 18 years, levels are belowthose in non-RA controls. Our earlier study (5) showed adown-regulation of synovial Duffy antigen expression in pa-tients with longstanding RA (mean disease duration 18 years),compared with non-RA controls. This finding was difficult toreconcile with a proposed proinflammatory role of Duffyantigen and with findings of other studies showing up-regulated expression of this protein in inflamed lungs duringsuppurative pneumonia and in kidneys with a variety ofinflammatory diseases (17–19). The current results are inaccordance with those described in these other reports and

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extend our earlier findings by showing that Duffy antigenexpression is increased, but only in early RA, and not afterprolonged disease.

It was possible that the enhanced Duffy antigen expres-sion in early RA was occurring on newly angiogenic bloodvessels. To test this we performed double-label immunofluo-rescence microscopy using antibodies to Duffy and �v�3integrin, which is a marker for angiogenic vessels (20). Themean � SEM percentage of Duffy-positive blood vessels thatcolocalized with �v�3 was 32.7 � 6.3% in non-RA controls,

19.4 � 4.0% in patients with early RA, and 14.5 � 5.2% inpatients with longstanding RA (mean disease duration �18years) (6 patients per group; 30 Duffy-positive vessels sampledper patient). Thus, in early RA, the amount of �v�3 colocal-ization with Duffy antigen was relatively low and did notcorrespond to the increased expression pattern of Duffy anti-gen shown in Figure 1. Therefore, it is unlikely that Duffyantigen is preferentially expressed on newly angiogenic bloodvessels in early RA.

To our knowledge this is the first study to demonstrate

Table 1. Clinical characteristics of the non-RA controls and RA patients*

Patient Diagnosis/pathologyDisease

duration† Treatment

Non-RA1 Articular cartilage damage 14 Analgesic, steroid, NSAID2 Articular cartilage degeneration 10 None3 Meniscal tear 1 None4 ACL damage �1 None5 Articular cartilage and meniscal

degeneration4 None

6 Articular cartilage damage 5 None7 ACL and articular cartilage

damage�5 None

8 Meniscal tear, articularcartilage degeneration

�1 None

9 Meniscal tear, articularcartilage degeneration

2 NSAID

10 ACL damage 1 NoneEarly RA

1 RA 4 None2 RA 1 NSAID3 RA 7 None4 RA 3 NSAID5 RA 4 NSAID6 RA 6 None

Longstanding RA(mean 9 years)

1 RA 6 Steroid2 RA 8 SSZ, NSAID3 RA 16 NSAID, steroid, analgesic4 RA 10 NSAID5 RA 8 NA6 RA 5 MTX, steroid, analgesic

Longstanding RA(mean 18 years)

1 RA 15 D-Pen, NSAID, analgesic2 RA 7 Steroid, NSAID3 RA 20 Steroid4 RA 25 NA5 RA 26 Steroid, NSAID6 RA 15 Gold, NSAID7 RA 5 Steroid, MTX8 RA 28 Steroid, MTX9 RA 20 HCQ, steroid, MTX, gold10 RA 22 Steroid

* RA � rheumatoid arthritis; NSAID � nonsteroidal antiinflammatory drug; ACL � anterior cruciateligament; SSZ � sulfasalazine; NA � information not available; MTX � methotrexate; D-Pen �D-penicillamine; HCQ � hydroxychloroquine.† In months for early RA; in years for non-RA and longstanding RA.

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Figure 1. Temporal pattern of expression of Duffy antigen in rheumatoid arthritis (RA). A and B, Immunohistochemistry analysis, showingexpression of Duffy antigen (brown staining) in the venules of synovia from a patient with early RA (A) and a patient with longstanding RA (B).C and D, Detail of Duffy-stained vessels (dark brown/black staining) in synovia from a patient with early RA (C) and a patient with longstandingRA (D) (different patients from those in A and B). Vessels that were Duffy negative were also observed (arrows). (Diaminobenzidine [DAB] stainingin A and B, DAB/nickel staining in C and D; original magnification � 100 in A, � 50 in B, � 400 in C and D.) E, Mean and SEM percentage ofDuffy antigen expression in vessels from non-RA controls and from patients with early RA or longstanding RA. Duffy antigen expression wasincreased in patients with early RA compared with that in non-RA controls (� � P � 0.013). Expression then decreased after disease duration of9 years and 18 years (�� � P � 0.00005 versus early RA and P � 0.02 versus non-RA).

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the expression of a chemokine receptor in early human RA,together with changes in its temporal expression pattern.Other studies have shown increased levels of chemokines,including CCL2, CCL5, and CXCL8, in joints of patients withearly RA (disease duration �3 months or �12 months)(21,22). Since these chemokines show high affinity to Duffyantigen (2,23), the up-regulation of Duffy expression in earlyRA may be functionally important in enhancing leukocytemigration into the synovium. It is interesting that in the synoviaof patients with early RA, the leukocyte infiltration pattern wasdiffuse. Therefore, the increased levels of Duffy antigen inearly RA may be involved in the enhanced leukocyte migrationassociated with this early diffuse pattern of infiltration. Duffyantigen may be less involved in leukocyte migration in morelong-standing disease, when Duffy levels are reduced and theperivascular follicles of mononuclear cells develop.

It is unlikely that treatment could be responsible forthe up-regulation of Duffy antigen in early RA, since both thenon-RA patients and the patients with early RA were takingNSAIDS only, or no medication (Table 1). It is possible thatsteroids or DMARDs may affect Duffy antigen expression inlongstanding RA. However, this does not appear to be the casefor steroids since there was no obvious difference in thepercentage of Duffy-positive vessels in patients with longstand-ing RA who were taking steroids (mean � SEM 52.7 � 5.4%,n � 10) compared with those taking other medications (53.4 �9.0%, n � 4). Future investigation into the effects of treatmenton Duffy antigen expression would, however, be of interest.

Supported by grants from the Biotechnology and BiologicalSciences Research Council, AstraZeneca, the Arthritis Research Cam-paign, the Wellcome Trust, and the Droitwich Medical Trust.

Lucy Gardner, BScCatherine Wilson, BScAngela M. Patterson, PhDKeele University at Robert Jones

and Agnes Hunt Orthopaedic HospitalOswestry, UKBarry Bresnihan, MD, FRCPOliver FitzGerald, MD, FRCPSt Vincents HospitalDublin, IrelandMichael A. Stone, MResAstraZenecaMacclesfield, UKBrian A. Ashton, PhDJim Middleton, PhDKeele University at Robert Jones

and Agnes Hunt Orthopaedic HospitalOswestry, UK

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2. Gardner L, Patterson AM, Ashton BA, Stone MA, Middleton J.The human Duffy antigen binds selected inflammatory but nothomeostatic chemokines. Biochem Biophys Res Commun 2004;321:306–12.

3. Nibbs R, Graham G, Rot A. Chemokines on the move: control bythe chemokine “interceptors” Duffy blood group antigen and D6.Semin Immunol 2003;15:287–94.

4. Hadley TJ, Peiper SC. From malaria to chemokine receptor: theemerging physiologic role of the Duffy blood group antigen. Blood1997;89:3077–91.

5. Patterson AM, Chamberlain G, Siddall H, Gardner L, MiddletonJ. Expression of the Duffy antigen/receptor for chemokines(DARC) by the inflamed synovial endothelium. J Pathol 2002;197:108–16.

6. Middleton J, Americh L, Gayon R, Julien D, Mansat M, Mansat P,et al. A comparative study of endothelial cell markers expressed inchronically inflamed human tissues: MECA-79, DARC (Duffyantigen receptor for chemokines), von Willebrand factor, CD31,CD34, CD105 and CD146. J Pathol 2005;206:260–8.

7. Lee JS, Frevert CW, Wurfel MM, Peiper SC, Wong VA, BallmanKK, et al. Duffy antigen facilitates movement of chemokine acrossthe endothelium in vitro and promotes neutrophil transmigrationin vitro and in vivo. J Immunol 2003;170:5244–51.

8. Rot A. Contribution of Duffy antigen to chemokine function.Cytokine Growth Factor Rev 2005;16:687–94.

9. Chaudhuri A, Nielsen S, Elkjaer M, Zbrzezna V, Fang F, PogoAO. Detection of Duffy antigen in the plasma membranes andcaveolae of vascular endothelial and epithelial cells of noneryth-roid organs. Blood 1997;89:701–12.

10. Middleton J, Neil S, Wintle J, Clark-Lewis I, Moore H, Lam C, etal. Transcytosis and surface presentation of IL-8 by venularendothelial cells. Cell 1997;91:385–95.

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13. Luo H, Chaudhuri A, Zbrzezna V, He Y, Pogo AO. Deletion ofthe murine Duffy gene (Dfy) reveals that the Duffy receptor isfunctionally redundant. Mol Cell Biol 2003;23:3030.

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15. Kirkham B, Portek I, Lee CS, Stavros B, Lenarczyk A, Lassere M,et al. Intraarticular variability of synovial membrane histology,immunohistology, and cytokine mRNA expression in patients withrheumatoid arthritis. J Rheumatol 1999;26:777–84.

16. Blanchard D, Tournamille C, Petit-Le Roux Y, Wasiowska K,LeVanKim C, Loirat MJ, et al. A new murine monoclonalantibody directed against FY6 antigen [abstract]. Vox Sang 1998;74 Suppl 1:71.

17. Liu X, Hadley TJ, Xu L, Peiper SC, Ray PE. Up-regulation ofDuffy antigen receptor expression in children with renal disease.Kidney Int 1999;55:1491–500.

18. Segerer S, Regele H, Mack M, Kain R, Cartron JP, Colin Y, et al.The Duffy antigen receptor for chemokines is up-regulated duringacute renal transplant rejection and crescentic glomerulonephritis.Kidney Int 2000;58:1546–56.

19. Lee JS, Frevert CW, Thorning DR, Segerer S, Alpers CE, CartronJP, et al. Enhanced expression of Duffy antigen in the lungs duringsuppurative pneumonia. J Histochem Cytochem 2003;51:159–66.

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DOI 10.1002/art.21906

Failure to confirm coxsackievirus infection in primarySjogren’s syndrome

An extensive debate has developed concerning thepathogenic role of various viral strains in the development ofprimary Sjogren’s syndrome (SS) (1–4). Recently, Triantafyl-lopoulou et al presented interesting data concerning theassociation between coxsackievirus infections and primary SS(5). Differential display revealed the presence of a 94-bpfragment of coxsackievirus B4 gene in salivary glands of 3patients with primary SS. Coxsackievirus RNA was detected in7 of 8 patients with primary SS, but in no patients withsecondary SS or controls, using seminested reversetranscriptase–polymerase chain reaction (RT-PCR) with spe-cific primers for the 5�-noncoding region (5�-NCR) of theenteroviral genome. We tried to replicate these findings byusing seminested RT-PCR with specific primers for the 5�-NCR of the enteroviral genome on high-quality RNA suitablefor gene expression studies of salivary glands.

We therefore analyzed the presence of the coxsackievi-rus genome in salivary glands of 9 patients with primary SS and9 controls who attended 2 French specialty centers (therheumatology departments of Bicetre and Strasbourg Hospi-tals). All the 9 patients had primary SS (according toAmerican-European consensus group criteria [6]) and a focusscore �1. Seven patients had anti-SSA antibodies, 4 patientshad anti-SSB antibodies, and 2 patients had no autoantibodies.The controls were subjects who had experienced subjectivesymptoms of oral or ocular dryness but who met none of theobjective criteria for SS (no autoantibodies, no lymphocyticinfiltrate on minor salivary gland biopsy). Only women wereincluded in both groups. The mean � SD ages of patients andcontrols were 59 � 10 years (range 40–80 years) and 56 � 10years (range 37–70 years), respectively.

To limit RNA degradation, minor salivary gland sam-ples were immediately stored in RNA Later (Qiagen, Chats-worth, CA) after biopsy, and RNA was extracted the sameweek and stored at �80°C. The integrity and loading of RNAwere analyzed using the Agilent 2100 Bioanalyzer (AgilentTechnologies, Palo Alto, CA); all samples reached a 28S:18Sribosomal RNA ratio of �1.7 (Figure 1). Moreover, RT-PCRamplification of the �-actin housekeeping gene was performedin all primary SS and control samples to verify the integrity ofRNA. One microgram of RNA of each minor salivary glandwas submitted to seminested RT-PCR with specific primers forthe conserved part of the 5�-NCR of the enteroviral genome,validated by the Centers for Disease Control and Prevention

(CDC), Atlanta, GA (7). The primer sequences used for thefirst PCR were as follows: for forward primer MB-EV1,5�-CTCCGGCCCCTGAATGCG-3�; for reverse primer MB-EV2, 5�-ATTGTCACCATAAGCAGCCA-3�. One microliterof the first PCR was submitted to seminested amplification(the second PCR) with the same forward primer MB-EV1 andthe nested reverse primer CDC-EV (5�-ACACGGACA-CCCAAAGTAGTCGGTTCC-3�).

The other reaction components used in the PCRs werethe same as previously reported, as were the thermocyclerconditions (incubation at 95°C for 5 minutes, followed by 40cycles at 95°C for 30 seconds, 56°C for 30 seconds, and 72°C for40 seconds) (8). Total RNA from poliovirus 1 Sabin straincultures was included as a positive control in each amplifica-tion reaction. The assays included 1 negative control (water)for each tested sample (the same water sample was submittedto the 2 PCRs). The greatest precautions were taken to limitthe risk of contamination. PCRs were performed in a labora-tory in which no enteroviral strains had ever been studiedbefore, the positive control tubes were opened in a roomseparate from that in which the salivary gland samples wereprocessed, and negative controls were intercalated betweeneach studied RNA sample.

No enteroviral DNA could be detected in any of the 18salivary gland specimens studied. Thus, no amplification prod-uct was observed after the first PCR (Figure 2). No amplifica-tion product was obtained by seminested amplification (thesecond PCR) of 15 salivary gland specimens. Seminestedamplification yielded a smear in 2 negative controls (water)and in specimens from 1 control subject and 1 patient withprimary SS, as well as amplification products of different sizes(not including the expected size of 115 bp, consistently found

Figure 1. Electrophoregram of total RNA extracted from a labialsalivary gland of 1 patient with primary Sjogren’s syndrome using theAgilent 2100 Bioanalyzer. This characterization of total RNA, per-formed in each sample, demonstrates the absence of RNA degrada-tion, given the conserved 28S:18S ribosomal RNA ratio, the absence ofadditional peaks below ribosomal bands, and the absence of shifttoward shorter fragments.

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