Sle Review

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Aberrant Phenotype and Function of Myeloid Dendritic Cells

in Systemic Lupus Erythematosus1

Dacheng Ding, Hemal Mehta, W. Joseph McCune, and Mariana J. Kaplan2

Systemic lupus erythematosus (SLE) is characterized by a systemic autoimmune response with profound and diverse T cell

changes. Dendritic cells (DCs) are important orchestrators of immune responses and have an important role in the regulation of

T cell function. The objective of this study was to determine whether myeloid DCs from individuals with SLE display abnormal-

ities in phenotype and promote abnormal T cell function. Monocyte-derived DCs and freshly isolated peripheral blood myeloid

DCs from lupus patients displayed an abnormal phenotype characterized by accelerated differentiation, maturation, and secretion

of proinflammatory cytokines. These abnormalities were characterized by higher expression of the DC differentiation marker

CD1a, the maturation markers CD86, CD80, and HLA-DR, and the proinflammatory cytokine IL-8. In addition, SLE patients

displayed selective down-regulation of the maturation marker CD83 and had abnormal responses to maturation stimuli. These

abnormalities have functional relevance, as SLE DCs were able to significantly increase proliferation and activation of allogeneic

T cells when compared with control DCs. We conclude that myeloid DCs from SLE patients display significant changes in

phenotype which promote aberrant T cell function and could contribute to the pathogenesis of SLE and organ damage. The

Journal of Immunology, 2006, 177: 58785889.

Systemic lupus erythematosus is an autoimmune disease of

unclear etiology that affects multiple organs. Individuals

with systemic lupus erythematosus (SLE)3 present with

numerous and diverse immune system abnormalities and display a

prominent imbalance in different T cell subset functions (13).

Indeed, individuals with SLE have autoreactive T cells that are

specific for ubiquitous self-peptides and provide pathogenic B cell

help (4, 5). A pan T cell dysfunction in SLE is characterized by

exaggerated CD4 T cell responses (6, 7) and diminished CD8 T

cell activities (8). Other alterations include changes in proliferative

T cell responses as well as increased expression of early and late

T cell activation markers (3, 7, 911). In addition, alterations inTh1 and/or Th2 lymphocyte function have been described (1216),

resulting in production of cytokines that up-regulate autoantibody

production by B cells, promote immune complex formation, and

increase the apoptotic load (17, 18). The exact role that accessory

cells might have in inducing abnormal T cell responses in SLE is

unclear; however, different groups have proposed that T cell dis-

turbances in SLE could be induced or promoted, at least in part, by

alterations in dendritic cell (DC) phenotype and function, because

these are key regulators of the immune system (1922). DCs com-

prise several subsets with different stages of maturation (20). Im-

mature, Ag-capturing DCs sense diverse signals that induce them

to undergo a maturation process, whose hallmarks are the up-reg-

ulation of the molecule CD83, as well as of a number of costimu-

latory and adhesion molecules, migration into lymphoid organs

(23), and acquisition of the capacity to activate quiescent, naive,

and memory lymphocytes (20, 2430). DCs can induce either T

cell tolerance or strong innate and adaptive immunity to specific

Ag. In general, tolerance is initiated when DCs are immature,

whereas the initiation of immunity requires an effective DC mat-

uration signal. After reaching the lymph nodes, DCs present pro-

cessed Ag to T cells via both classical (class I and II MHC) andnonclassical (CD1 family) pathways (24, 31, 32). In addition to

their maturation status, DCs can be subdivided into a CD11c

(plasmacytoid DC) and a CD11c (myeloid DC) subset (20, 33).

The CD11c subset follows a myeloid differentiation pathway in

which monocytes serve as precursors. These DC subsets have dif-

ferent migration programs and very likely have divergent roles in

the induction and regulation of the immune response (33, 34)

In the past few years, significant progress has been made in

elucidating the important role that IFN--producing plasmacytoid

DCs have in promoting autoimmune responses in SLE (22, 35, 36).

Much less is known about the role that myeloid DCs play in con-

tributing to the pathogenesis of SLE. Indeed, it is unclear whether

myeloid DCs present phenotypic and functional changes that may

promote the autoimmune features of lupus and whether such dif-

ferences might potentiate aberrant cognate interactions with T

cells. We now report that myeloid DCs of individuals with SLE

display evidence of aberrant phenotype and function which may

contribute to the T cell abnormalities described in this disease.

Materials and MethodsPatient selection

Patients with SLE and rheumatoid arthritis (RA) fulfilled the AmericanCollege of Rheumatology criteria for these diseases (3739) and were re-cruited from the outpatient rheumatology clinic and inpatient services atthe University of Michigan, and from the Michigan Lupus Cohort (AnnArbor, MI). Healthy controls were obtained by advertisement. SLE activity

Department of Internal Medicine, Division of Rheumatology, University of Michigan,Ann Arbor, MI 48109

Received for publication June 15, 2005. Accepted for publication August 12, 2006.

The costs of publication of this article were defrayed in part by the payment of pagecharges. This article must therefore be hereby marked advertisement in accordancewith 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported by Public Health Service Grants AR050554 andAR048235 and by the Anthony S. Gramer Fund in Inflammation Research (all toM.J.K.). D.D. was supported by Training Grant T32 AR 07080. W.J.M. was sup-ported by the Herb and Carol Amster Lupus Research Fund and the Klein LupusResearch Fund. This research was also supported (in part) by the National Institutesof Health through University of Michigans Cancer Center Support Grant (P30CA46592) and the Rheumatic Diseases Core Center Grant (P30 AR48310).

2 Address correspondence and reprint requests to Dr. Mariana J. Kaplan, Division ofRheumatology, University of Michigan, 5520 MSRBI, 1150 West Medical CenterDrive, Ann Arbor, MI 48109-0680. E-mail address: [email protected]

3 Abbreviations used in this paper: SLE, systemic lupus erythematosus; DC, dendriticcell; RA, rheumatoid arthritis; SLEDAI, SLE disease activity index; 6-MP, 6-mer-captopurine; MMF, mycophenolate-mofetil.

The Journal of Immunology

Copyright 2006 by The American Association of Immunologists, Inc. 0022-1767/06/$02.00

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was assessed by the SLE disease activity index (SLEDAI) (40). Patient andcontrol cells were paired and studied in parallel. Overall, 146 SLE patients, 76healthy controls and 35 RA patients were studied. Information regarding thedemographics, disease activity, and use of medications is provided in Table I.

Reagents

Human rIL-4, TNF-, and IL-2 were purchased from PeproTech. HumanGM-CSF was a gift from Berlex. rIFN-2b was obtained from Schering.X-vivo 10 serum-free medium was from BioWhittaker. LPS and PHA werepurchased from Sigma-Aldrich. The following anti-human mAbs conju-

gated to FITC, PE, CyChrome, and allophycocyanin were used: anti-CD1a,CD3, CD4, CD8, CD11c, CD14, CD25, CD40L, CD69, CD80, CD83,CD86, HLA-DR, and isotype controls (all obtained from BD Biosciences).The pan T cell isolation kit was obtained from Miltenyi Biotec. CFSE wasobtained from Molecular Probes.

Generation of monocyte-derived DCs

Myeloid DCs were generated from human peripheral monocytes as previ-ously described (41). In brief, human PBMC were separated by standarddensity gradient centrifugation on Ficoll-Hypaque Plus (Amersham Bio-sciences) and resuspended at 6 106 cells/ml in RPMI 1640 with L-glutamine and 10% FBS. The cells were transferred to tissue culture platesand allowed to adhere to the plastic surface for 1 h at 37C. Nonadherentcells were then removed by washing with PBS. Monocyte recovery rateafter adherence was 89% in both healthy controls and SLE patients. The

adhered monocytes were further cultured for 57 days in DC inductionmedium (serum-free X-vivo-10 containing 20 ng/ml GM-CSF and 510ng/ml IL-4). In some experiments, cells were further purified using metri-zamide gradient (Sigma-Aldrich). The purity of myeloid DCs obtainedunder our experimental conditions was 90%, as confirmed by flow cy-tometric analysis (data not shown). At days 57, cells were harvested foranalysis or stimulated with DC maturation stimuli (41) (0.52 g/ml LPSand/or 10100 ng/ml TNF-) for 48 h. In additional experiments, freshlyisolated monocytes were cultured for 7 days with GM-CSF, with or withoutIL-4, in the presence or absence of 100 or 1000 U of rIFN-.

Immunofluorescence staining, FACS, and fluorescent microscopy

analysis

DCs were washed with PBS/0.2% BSA and FcRs were blocked by incu-bating cells for 20 min in PBS with 40% control human sera or with anti-FcAb (Miltenyi Biotec). Cells were then incubated for 30 min at 4C with

0.060.15 g/ml fluorochrome-conjugated mAb following the manufac-turers directions. Cells were washed three times with PBS/0.2% BSA,fixed in 1% paraformaldehyde and analyzed in a FACSCalibur flow cy-tometer (BD Biosciences), using previously described protocols. Data anal-ysis was performed using Lysys II software (BD Biosciences) and Win-MDI 2.8 software ( http://facs.scripps.edu ). Stained cells were gated byside scatter and forward scatter characteristics and further identified bysurface markers. The results were expressed as the percentage of cellsstaining positive for different markers as well as by mean channel fluores-cence. The cutoff point for positive staining was above the level of thecontrol isotype Abs.

For fluorescent microscopy, DCs were derived from monocytes as statedabove. At day 7, cells were harvested, washed, and stained with the specificmAbs mentioned above to characterize DC phenotype and morphology.Cells were then fixed in 1% formalin in PBS for 30 min, and transferred toa microscope slide using ProLong Antifade kit (Molecular Probes). Slideswere analyzed using a Zeiss LSM 510 META Laser Scanning Microscope(Carl Zeiss Advanced Imaging Microscopy).

Isolation and characterization of myeloid DCs from peripheral

blood

PBMCs were isolated from peripheral blood of SLE patients and controlsby Ficoll gradient as described above. PBMCs were incubated with twodifferent mixtures of Abs: 1) CD11c-FITC, CD1a-allophycocyanin, CD83-PE, and CD86-PE/Cy5, and 2) CD11c-FITC, CD14-allophycocyanin/Cy5,CD86-PE/Cy5 and HLA-DR-allophycocyanin. After incubation for 30 minwith these Abs at 4C, cells were washed, fixed in PBS/1% paraformal-dehyde, and analyzed by FACS by gating the CD11c population andexcluding CD14 cells.

Drug treatment

Monocytes were cultured as stated above to induce DC differentiation, inthe presence or absence of graded concentrations of indomethacin (0.011g/ml), hydroxychloroquine (0.022 g/ml), hydrocortisone (0.011 M),6-mercaptopurine (6-MP) (0.011 M) and mycophenolate-mofetil(MMF) (0.044 g/ml; all obtained from Sigma-Aldrich) or vehicle (42

46). A stock solution of 6-MP was prepared in dimethylformamide at aconcentration of 10 mg/ml. The stock solution was diluted in assay diluent(80% culture medium/20% ethanol) to yield a 6-MP working solution of 80g/ml or less as indicated. The working solution of 6-MP as well as theother materials prepared in assay diluent were then further diluted 1/40 intothe cell cultures for the tests. The final concentrations of ethanol (0.1%) ordimethylformamide (0.02%) do not yield significant effects on the cellcultures (11, 47). Indomethacin was prepared in a concentration of 500 mMin absolute ethanol, then diluted to final concentrations in the cell culturemedium (11). Control cells were treated with an equal volume of the sol-vent. MMF was prepared as previously described (43). At day 7, DCs werewashed and analyzed by flow cytometry for expression of differentiation(CD1a) and maturation (CD83, CD86, HLA-DR) markers.

RNA extraction and quantitative real-time RT-PCR

Total RNA was isolated from myeloid DCs using the RNeasy kit with

DNase I digestion (Qiagen) to remove possible genomic DNA contamina-tion, and reverse transcribed to cDNA using the SuperScript III first-strandsynthesis system (Invitrogen Life Technologies) with Oligo(dT)30 primer.For real-time detection of target and reference gene expression, eight pairprimers and probes were designed as follows: 83 forward (F): 5-CTGCTCCTGAGCTGCGCCTACA; CD83 reverse (R): 5-CACCACCCTCCAATAACTTGAC; CD83 probe: 5-ATCCGCAGGTTCCCTACACGGTCTCC; CD80 F: 5-CTTCAACTGGAATACAACCAAGCA; CD80 R: 5-TGCATCTTGGGGCAAAGCAGTA; CD80 probe: 5-CTCCCATCCTGGGCCATTACCTTAATC; CD1a F: 5-GTCCTCTACTGGGAGCATCACA; CD1a R: 5-GTCTTAACAGAAACAGCGTTTCC; CD1a probe:

Table I. Demographic and clinical characteristics of human subjects studied

Variable Lupus Control RA

Number studied 146 76 35Age(mean range) 44.5 11.5 37 13.5 51 14.4Females n (%) 130 (89) 55 (72.3) 18 (51.5)Males n (%) 16 (11) 21 (27.6) 17 (48.5)

Lupus disease activity (mean SEM) 4.5 3SLEDAI 2 (%) 77SLEDAI 2 (%) 69

MedicationsAntimalarials (%) 68.5 40.5Azathioprine (%) 5.47 0Cyclophosphamide (%) 1.36 0Methotrexate (%) 1.36 51.4Mycophenolate mofetil (%) 24.65 0Prednisone (0.5 mg/kg/day) (%) 34.2 51.4Prednisone (0.51 mg/kg/day) (%) 9.5 5.7Prednisone (1 mg/kg/day) (%) 4.1 8.5

5879The Journal of Immunology

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5-CTTGGCGGTGATAGTGCCTTTACTTCTT; CD86 F: 5-GACAGGCATTTGTGACAGCACTA; CD86 R: 5-TCT GCA GTC TCA TTGAAA TAA GC; CD86 probe: 5-TTCCTGCTCTCTGGTGCTGCTCCTCT;CD14 F: 5-GGTGCCGCTGTGTAGAAAGAAGC; CD14 R: 5-GGTTCTGGCGTGGTCGCAGAGAC; CD14 probe: 5-TTATCGACCATGGAGCGCGCGT; DRa F: 5-TCAAGGTGCATTGGCCAACATAG; DRaR: 5-CTCTCAGTTCCACAGGGCTGTTC; DRa probe: 5-CGATCACCAATGTACCTCCAGAG; PBGD-F 5-GGCAATGCGGCTGCAA-3 ;PBGD-R 5-GGGTACCCACGCGAATCAC-3 ; PBGD probe 5-CTCATCTTTGGGCTGTTTTCTTCCGCC; -actin F: 5-AGCCTTCCTTC

CTGGGCATGGA; -actin R: 5-CTCAGGAGGAGCAATGATCTTGA;-actin probe: 5-ACATCCGCAAAGACCTGTACGCCAACA.Probes were labeled with 5-6-FAM/3-TAMRA (Integrated DNA

Technologies). HotStar Taq polymerase (Qiagen) was used. PCR was per-formed using MyCycler Thermal Cycler (Bio-Rad) in a total reaction mix-ture of 20 l containing 50 ng of cDNA, 1 HotStar TaqPCR buffer, and400 nM probe/primers mixture. After denaturation at 95C for 15 min, 55cycles were performed at 95C for 15 s, followed by 60C for 1 min.Comparative cycle threshold method with PCR efficiency correction, alsoknown as Pfaffls method, was used for quantification, as previously de-scribed (48). The expression levels of the target genes were adjusted to theexpression levels of the housekeeping genes PBGD and -actin.

Cytokine determination

Cytokines were measured using two different methods. The human cyto-kines IL-1, IL-2, IL-4, IL-5, IL-6, IL-7, IL-8, IL-10, IL-12p70, IL-13,

IFN-, GM-CSF, and TNF- were simultaneously quantified in duplicateusing the human cytokine multiplex kit (Linco Research). Supernatantsobtained from day 7 (unstimulated DCs) or day 9 (DCs treated with mat-uration stimuli for 48 h) cultures were aliquoted and stored at 80C untilused. Sera from the same patients were also stored. Cytokine concentra-tions were determined following manufacturers instructions. In brief, stan-dards and samples were added into the appropriate wells. Mixed beadswere added to each well and the plate was incubated with agitation for 1 hat room temperature. Plate was washed and a detection Ab mixture wasadded to each well and incubated for 30 min at room temperature. Strepta-vidin-PE was added to each well containing the detection Ab mixture. Theplate was then incubated with agitation on a plate shaker for 30 min atroom temperature, washed, and sheath fluid was added for 5 min. Plate wasread on Luminex 100 (Luminex) and the concentration reported as pico-grams per milliliter.

In addition, the BD CBA Human Inflammation kit (BD Biosciences)

was used to quantitatively measure IL-8, IL-1, IL-6, IL-10, TNF-, andIL-12p70 protein levels in sera and supernatant samples, following man-ufacturers instructions. In brief, mixed capture beads were added to assaytubes. Human inflammation standard dilutions and test samples were addedto the assay tubes. Samples were incubated for 1.5 h at room temperatureand protected from direct exposure to light. Wash buffer was added andtubes were centrifuged at 200 g for 5 min. Supernatants were discardedleaving 100 l of liquid in each assay tube. Human inflammation PE de-tection reagent was added to the tubes and samples were incubated for 1.5 hat room temperature, washed, and analyzed by FACS using BD CytometricBead Array Software.

T cell proliferation and determination of T cell activation

T cells were isolated by negative selection using magnetic beads and in-structions provided by the manufacturer (pan T cell isolation kit; MiltenyiBiotec). Purity was 95%, as assessed by CD3 expression. T cell prolif-

eration was analyzed using the intracellular fluorescent dye CFSE. Witheach cell division, the CFSE fluorescence intensity of the cells is reducedby half (4951). T cells from healthy controls were labeled with 2 MCFSE in RPMI 1640/10% FBS for 2 min at room temperature. After wash-ing to remove unbound fluorescent dye, 4 106 T cells were coculturedwith allogeneic DCs from SLE or healthy controls. Conditions includedunstimulated DCs or DCs stimulated with LPS and TNF- for 48 h, asdescribed above. Cells were cocultured at a T cell:DC ratio of 5:1 to 10:1.After 1 and 5 days, cells were harvested and stained with mouse anti-human CD3-allophycocyanin/Cy7, CD11c-PE, and CD25-CyChrome, thenfixed with 2% formaldehyde/PBS and analyzed by FACSCalibur, usingCellQuest software (BD Biosciences) and WinMDI. In additional experi-ments, expression of additional activation markers (CD69 and CD40L) wasmeasured on these T cells. Expression was measured 24 h and 5 days aftercocultures were started. Proliferation of T cells was analyzed by the log-arithmic reduction of CFSE staining of CD3-positive cells. The percentage

of T cells that had undergone specific numbers of cell divisions (G1G5) orno cell divisions (G0) was calculated (52, 53). Controls included PHA-stimulated T cells and unstimulated T cells.

Statistical analysis

The difference between means was analyzed using paired t test or one-wayANOVA with post hoc analysis and Bonferroni correction. Spearman andPearsons correlation were used to assess correlation between differentvariables. Analyses were performed with SPSS version 11.5. A value ofp 0.05 was considered to be statistically significant.

ResultsMyeloid lupus DCs display abnormal levels of differentiation

and maturation markers and have abnormal responses tomaturation stimuli

Confirming a previous study (54), no morphologic differences in

the capacity of monocytes to differentiate into DCs were found

between SLE and controls, using light and fluorescent microscopy

(Fig. 1). After 57 days of culture, DCs from SLE patients and

healthy controls were found to be increased in size and developed

typical dendrites. To assess whether DCs from SLE individuals

and controls differ in their differentiation and maturation poten-

tials, monocyte-derived DCs from 31 individuals with SLE, 8 pa-

tients with RA, and 20 healthy controls obtained at day 7 were

stained with anti-human Abs to CD1a (to evaluate DC differenti-

ation) (31) and CD83 (a marker of DC maturation) (55), and ex-

pression of these markers was measured by FACS. As shown inFig. 2, AC, SLE patients, have increased numbers of cells ex-

pressing the differentiation marker CD1a, and decreased numbers

of cells expressing the maturation marker CD83 (p 0.05). No

FIGURE 1. Monocyte-derived DCs from SLE patients display normal

morphology. A, Images represent different magnifications of cells from one

individual with SLE, after culture in X-vivo medium with IL-4 and GM-

CSF for 7 days. Cells display the characteristic membrane and nuclear

features of DCs. Arrows point at the cells that are displayed at higher

magnification on the left panels. B, Fluorescent microscopy image of onecell from a patient with SLE that displays the classic morphology DCs, at

day 7 in culture. The cell expresses both CD1a and CD83.

5880 ABNORMAL PHENOTYPE AND FUNCTION OF LUPUS DCs

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significant differences were observed in mean fluorescence inten-

sity (data not shown). No significant differences were detected be-

tween RA patients and controls in CD1a expression (Fig. 2A) or

CD83 expression (Fig. 2B). The differences between SLE and con-

trols were confirmed at the mRNA level by real-time RT-PCR

(Fig. 2D). Indeed, lupus patients (but not RA patients) displayed

significantly decreased mRNA levels of the monocyte marker

CD14 and higher levels of CD1a mRNA at day 7, suggesting an

acceleration of the differentiation from the monocyte stage to the

myeloid DC stage (Fig. 2D). Baseline CD14 levels in monocytes

did no differ between controls and lupus patients (data not shown),

suggesting that the differences seen on day 7 DCs were the con-

sequence of accelerated differentiation from the monocyte to the

DC stage and not due to baseline down-regulation of CD14 on

lupus monocytes. Interestingly, while CD83 protein and mRNA

levels were significantly lower in lupus DCs at day 7 (Fig. 2, BD),

FIGURE 2. Aberrant expression of differentiation and maturation markers in monocyte-derived DCs in SLE. Bar graph represents A, CD1a, and B, CD83

percent expression, in 20 healthy controls, 8 RA patients, and 31 SLE patients. , p 0.05, results represent mean SEM of independent experiments.

C, A representative density plot of CD83 and CD1a expression from one control and one SLE patient. D, Bar graphs represent CD1a, CD14, and CD83

mRNA expression by real-time RT-PCR relative to housekeeping genes in day 7 DCs from 7 healthy controls, 22 SLE, and 7 RA patients. Results are

reported as the fold change compared with healthy controls. , p 0.05, results represent mean SEM of independent experiments.


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mRNA levels of other specific maturation and differentiation markers

(CD86, CD80, and HLA-DR) were significantly higher in SLE than

in healthy controls in the absence of exogenous maturation stimuli

(Fig. 3A). These findings were confirmed at the protein level (Fig. 3,

B and C), both as numbers of cells expressing these markers (data not

shown) and as mean fluorescent intensity in each cell. When com-

pared with healthy controls, there were no differences in the expres-

sion of maturation markers in RA patients (Fig. 3). These results in-

dicate that differentiation and maturation are enhanced in monocyte-

derived DCs from SLE patients, but that the maturation marker CD83

is selectively down-regulated in lupus DCs.

We considered the possibility that an in vivo serum effect could

induce early responses in monocytes that would lead/promote the

observed differences in maturation. However, when we treated lu-

pus and healthy control monocytes with autologous sera, no dif-

ferences were found between control and SLE sera on the induc-

tion of expression of specific maturation markers (CD80: 11.5

4.6% for untreated control DCs; 10.5 4.1% for control DCs

treated with autologous sera; 32.6 11.2% for untreated lupus

DCs and 34.2 12% for lupus DCs treated with autologous sera.

CD83: 28.2 11.1% for untreated control DCs; 5.35 0.8% for

control DCs treated with sera; 14.5 2% for untreated lupus DCs

and 2 0.1 for lupus DCs treated with autologous serum, p NS

when comparing lupus vs control sera; results represent mean

SEM of three SLE patients and four healthy controls). These re-

sults suggest that the differences found on maturation markers

were intrinsic to lupus monocyte derived-DCs, rather than an ex-

ogenous effect secondary to a serum factor or to serum withdrawal.

Furthermore, when we cultured lupus monocytes in GM-CSF and

IL-4 in the presence or absence of IFN-, there were no significant

differences in the expression of CD83 between IFN-treated or un-

treated cells, suggesting that adding this cytokine did not alter the

phenotypic abnormalities seen in SLE DCs (CD83: 39 10% in

IL-4 GM-CSF; 30 5% in IL-4 GM-CSF 100 U of

IFN-; 30 4.4% in IL-4 GM-CSF 1000 U IFN-; results

represent mean SEM of eight independent experiments, p 0.05).

Similarly, when lupus monocytes were cultured in GM-CSF without

IL-4 but in the presence of IFN-, this combination of cytokines did

not result in changes in CD83 expression (10 4% in GM-CSF

alone; 15 5% in GM-CSF 100 U of IFN-; 19 5% in GM-

CSF 1000 U of IFN-; results represent mean SEM of eight

independent experiments; p 0.05 between all conditions).To exclude the possibility that medications could account for the

differences in expression of differentiation and maturation markers

between controls and SLE patients, monocytes were treated with

graded concentrations of drugs commonly used in the management of

SLE, including hydrocortisone (for steroids), indomethacin (for non-

steroidal anti-inflammatory drugs), chloroquine (for antimalarials),

6-MP (for azathioprine), and MMF, while these cells were being dif-

ferentiated into DCs in vitro. At doses equivalent to the ones used to

treat SLE patients, we did not find any significant changes in the

expression of differentiation and maturation markers induced by these

drugs (Table II). Further, there were no significant correlations be-

tween the use of specific immunosuppressive drugs by SLE patients

and the phenotypic differences found in SLE DCs (Table III).Abnormal differentiation and maturation correlated with specific

lupus clinical and serologic manifestations. Indeed, decreased

CD83 expression in DCs correlated significantly with current or

previous evidence of lupus nephritis, and with levels of anti-

dsDNA Abs. Increased CD1a expression in lupus DCs correlated

with decreased levels of complement (defined as C3 levels 83

mg/dl and/or C4 levels 13 mg/dl) (Fig. 4B). These correlations

were seen both at the protein (Fig. 4) and mRNA level (p 0.05

for same variables). Increased levels of CD80 at the protein level

correlated with hemologic manifestations of SLE (leucopenia and

immune-mediated thrombocytopenia) (r 0.49, p 0.02) and

levels of anti-dsDNA Abs (r 0.46, p 0.03), and negatively

correlated with skin manifestations of SLE (r 0.59, p 0.004). Increased levels of CD86 at the protein level strongly cor-

related with disease activity (SLEDAI) (r 0.74, p 0.001) and

with previous or current lupus nephritis (r 0.45, p 0.04). The

correlations between CD80 and CD86 with these specific clinical

variables were also confirmed at the mRNA level (data not shown).

To establish whether DCs from individuals with SLE normally

up-regulate maturation markers in response to maturation stimuli,

we proceeded to treat day 7 monocyte-derived DCs with LPS

and/or TNF- for 48 h. As show in Fig. 5C, lupus DCs signifi-

cantly up-regulate mRNA of the maturation markers CD86, CD80,

and CD83. However, when compared with healthy controls, the

degree of up-regulation for CD80 and CD86 was blunted in the

lupus group both at the protein and mRNA level ( p 0.05) (Fig.

5), using either LPS and/or TNF- as maturation stimuli. These

experiments suggest that lupus DCs have the capacity to respond

FIGURE 3. Increased expression of maturation markers in lupus DCs.

A, Bar graphs represent mRNA expression of different maturation markers

relative to housekeeping genes by real-time RT-PCR in day 7 DCs from 7

healthy controls, 22 SLE patients, and 7 RA patients. Results are reported

as the fold change compared with healthy controls. , p 0.05, results

represent mean SEM of independent experiments. B, Bar graphs compare

CD86 mean fluorescent intensity in day 7 DCs from 8 healthy controls, 5 RA

patients, and 24 SLE patients. Representative histogram displays expression of

CD86 on day 7 myeloid DCs from one healthy control, one RA patient, and

one SLE patient. C, Bar graphs compare CD80 mean fluorescent intensity in

day 7 DCs from 8 healthy controls, 5 RA patients, and 24 SLE patients.

Representative histogram displays expression of CD80 on day 7 myeloid DCsfrom 1 healthy control, 1 RA patient, and 1 SLE patient. , p 0.05, results

represent mean SEM of independent experiments.


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to maturation stimuli, as shown by up-regulation of maturation

markers; however, the level of up-regulation of these markers isdecreased when compared with healthy controls. When compared

with healthy controls, there were no statistical differences in the

capacity of RA DCs to up-regulate maturation markers after ex-

posure to exogenous maturation stimuli (Fig. 5, A and B).

To exclude the possibility that abnormal differentiation and mat-

uration observed in SLE DCs were consequences of the in vitro

conditions used, the expression of maturation and differentiation

markers in myeloid DCs directly obtained from peripheral blood

was examined. As shown in Fig. 6A, the myeloid DC population

obtained directly from lupus blood is also characterized by higher

numbers of CD1a cells, lower numbers of CD83 cells and

higher expression of CD86 cells when compared with healthy

controls, further confirming our in vitro findings. In addition, high

levels of CD1a cells significantly correlated with levels of anti-

dsDNA Abs (Fig. 6B) and with hypocomplementemia (p

0.004). Furthermore, similar to what we found in DCs generated in

vitro, there were no significant correlations between the use of

specific immunosuppressive drugs and phenotypic abnormalities

seen in lupus DCs isolated from peripheral blood (data not shown).

Lupus DCs secrete higher levels of IL-8

To evaluate whether SLE DCs show differential secretion of cy-

tokines, we measured the secretion of a number of cytokines using

cytometric bead array and a cytokine multiplex kit array. Control,

RA, and SLE DCs secrete detectable amounts of IL-6, IL-7, IL-8,

IL-10, IL-13, IFN-, and TNF-. The cytokine secreted at the

highest levels was the proinflammatory cytokine IL-8, and lupusDCs secreted significantly higher levels of IL-8 than control DCs

both before (5459.2 813.7 vs 2137.9 759.3 pg/ml, respec-

tively, mean SEM, p 0.05) and after TNF- stimulation

(8760.6 1048.6 vs 4817.2 689.4, respectively, mean SEM,

p 0.05) (Fig. 7A). Levels of IL-8 secreted by lupus DCs corre-lated with expression of the maturation marker CD80 (r 0.56,

p 0.01, Pearsons correlation), and patients with higher produc-

tion of IL-8 had higher anti-dsDNA (Fig. 7B) and anti-cardiolipin

levels in sera (r 0.53, p 0.007 and r 0.45, p 0.02,

respectively, Pearsons correlation). When compared with healthy

controls, there were no significant increases in IL-8 secretion by

DCs from RA patients (Fig. 7B). No significant differences be-

tween control, SLE, and RA patients were observed when the other

cytokines were measured in DC supernatants (data not shown). Of

interest, IL-8 was also the most abundant cytokine detected in

plasma from SLE patients and serum levels were higher than in

healthy controls, although no statistical significance was found

(57.9 vs 36.3 pg/ml, P NS).

Lupus DCs up-regulate T cell proliferation and activation

Unstimulated and stimulated DCs from lupus patients induced a

significant increase in allogeneic T cell proliferation when com-

pared with DCs from healthy controls. As expected, unstimulated

DCs from healthy controls were poor stimulators of allogeneic T

cell proliferation and, after stimulation with maturation stimuli,

their capacity to induce allogeneic T cell proliferation increased

significantly. When cocultured with lupus DCs, allogeneic T cells

showed a significantly higher percentage of proliferating cells than

what was observed with control allogeneic DCs, and the effect was

more pronounced in DCs treated with maturation stimuli (LPS and

TNF-) (Fig. 8, A and B). This increase in proliferation required

cell-cell interactions, as supernatants from lupus DCs did not in-duce similar increases in T cell proliferation (data not shown).

Similarly, there was a statistically significant increase in the ex-

pression of the activation marker CD40L in allogeneic T cells

Table III. Correlation analysis of specific immunosuppressive/immunomodulator use and expression of differentiation and maturation markers inlupus DCsa

Corticosteroids Antimalarials Azathioprine Mycophenolate

CD1a p 0.24 (0.28) p 0.50 (0.16) p 0.63 (0.121) p 0.46 (0.15)CD83 p 0.77 (0.07) p 0.12 (0.37) p 0.29 (0.262) p 0.16 (0.24)CD86 p 0.14 (0.31) p 0.99 (0.0002) p 0.71 (0.08) p 0.47 (0.17)CD80 p 0.65 (0.11) p 0.96 (0.009) p 0.47 (0.18) p 0.24 (0.27)

IL-8 p 0.42 (0.20) p 0.30 (0.26) p 0.30 (0.18) p 0.52 (0.16)

a Results represent p value (r ), using Spearman correlation. Negative numbers indicate a negative correlation. All p values were 0.05.

Table II. Expression of maturation and differentiation markers on day 7 monocyte-derived DCs, after in vitro drug treatmenta

CD1a CD83 CD86 CD80

Untreated 18.08 6.5 24.86 0.27 34 1.67 89.2 0.4HC 0.01 M 19.08 1.76 39.9 9.6 36.7 0.07 84 0.3HC 0.1 M 24 5.01 27.1 4.2 28 10.47 82.5 0.5HC 1 M 31 9 26.06 6.7 26.5 3.2 83.2 26-MP 0.01 M 10.25 0.2 33.5 2.7 46.7 3.45 84.4 0.96-MP 0.1 M 13.3 0.25 27.2 4.2 36.6 0.075 82.2 0.8

6-MP 1 M 14.8 0.9 35.4 4.9 47 3.5 84.1 3INDO 0.01 g/ml 10.6 5.6 34.4 0.08 33.5 1.7 83 9INDO 0.1 g/ml 14.6 0.52 35.2 1 35 16.22 77.8 5INDO 1 g/ml 15.5 1.4 35.6 6.5 32.3 16.2 77 6MMF 0.04 g/ml 14 8.5 30.9 8.3 51 0.5 92 5MMF 0.4 g/ml 10.7 5.7 32 11.5 51 11 93 2.8MMF 4 g/ml 18 3.5 37 18.4 41 2.05 94 3HCQ 0.02 g/ml 13.5 7.1 28 10.5 51 .3 89 2HCQ 0.2 g/ml 6.9 2.85 31 8.3 57 11.2 93 3HCQ 2 g/ml 3.44 1.4 28.7 11.6 51 6 91 1.1

a HC, Hydrocortisone; 6-MP, 6-mercaptopurine; INDO, Indomethacin; MMF, mycophenolate mofetil; HCQ, hydroxychloroquine. Vehicle results did not differ from resultsfrom untreated cells. Values represent mean SEM of three independent experiments and all values are p 0.05 when compared to untreated cells.


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when they were cocultured for 24 h with either unstimulated or

stimulated lupus DCs, as compared with DCs from healthy con-

trols (Fig. 8C). An increase in the percentage of cells that coex-

press CD25 and CD40L in allogeneic T cells was also observed

after these cells were cocultured with either unstimulated or stim-

ulated lupus DCs for 5 days, as compared with unstimulated or

stimulated DCs from healthy controls (percent of CD25

CD40L

T cells: 2.4 1% with unstimulated control DCs were added;

5.8 2.1% when unstimulated lupus DCs were added; 4.8 1.2%

when stimulated control DCs were added;. 10.8 3% when stim-

ulated lupus DCs were added; p 0.05 between control and lupus

cells for both conditions; results represent mean SEM of inde-

pendent experiments using five controls and 11 SLE patients). No

significant differences were seen for the early T cell activation

marker CD69 between control and lupus DCs (data not shown). As

expected, DCs treated with maturation stimuli were more effective

at inducing allogeneic T cell activation (Fig. 8).

Discussion

This study establishes that myeloid DCs from individuals with SLEdisplay distinct phenotypic and functional differences characteristic of

proinflammatory DCs and promote abnormal T cell responses. These

phenotypic changes are characterized by accelerated differentiation

from the monocyte to the myeloid DC stage (as measured by in-

creased expression of the DC differentiation marker CD1a), up-regu-

lation of different costimulatory molecules which are involved in T

cell priming, even in the absence of exogenous maturation stimuli;

and increased production of the proinflammatory cytokine IL-8. In

addition, myeloid DCs from SLE patients display a selective down-

regulation of the maturation marker CD83. These abnormalities were

seen not only on in vitro-derived DCs, but also in DCs obtained from

peripheral blood, indicating that this phenomenon is not the conse-

quence of added cytokines in vitro or a serum withdrawal effect but

likely represent true abnormalities on the differentiation/maturation

pathways of myeloid DCs in vivo.

These phenotypic differences correlate with specific clinical and

serologic manifestations of the disease. Because the population

studied was fairly well-controlled, with an SLEDAI of 4.5 3

(mean SEM), the results of this study suggest that the abnor-

malities observed in SLE DCs are not merely the result of active

disease but might have pathogenic significance. Further, these phe-

notypic differences in lupus DCs likely have functional relevance,as both unstimulated and stimulated myeloid DCs from lupus pa-

tients induce significant increases in allogeneic T cell proliferation

and activation, when compared with healthy controls. Immature

DCs are known to have a low T cell activation potential (20) and

this promotion of T cell proliferation and activation is likely the

consequence of an increased mature phenotype. Accelerated dif-

ferentiation from the monocyte (or potentially other myeloid pre-

cursors) to the DC stage, and up-regulation of maturation markers

could promote and enhance the capabilities of lupus DCs to prime

and activate T cells in the spleen and other lymphoid organs, fur-

ther contributing to the T cell hyperresponsiveness and enhanced

activation described in SLE. Supporting this idea, a recent study

reported up-regulation of CD11b

CD11c

DCs in the thymus andspleen in aged BWF1 lupus-prone mice (56), suggesting acceler-

ated migration to these organs. Studies in humans have also shown

that myeloid and plasmacytoid DCs are markedly decreased in

SLE blood and it has been speculated, although not proven, that

this might reflect an accelerated migration of these cells from the

blood into tissues (22, 57). Certainly, the up-regulation of CD86

and other maturation markers in DCs would support the possibility

that a mature DC phenotype would be associated to increased

migration to lymphoid organs of SLE DCs and stimulation and

priming of T cells in these organs. DCs can induce either T cell

tolerance or strong innate and adaptive immunity to specific Ag. In

general, tolerance is initiated when DCs are immature, whereas the

initiation of immunity requires an effective DC maturation signal.

Therefore, accelerated DC maturation in SLE in the absence of

extrinsic danger signals suggests that these cells can become very

FIGURE 4. Aberrant differentia-

tion and maturation in SLE correlatewith clinical and serologic features of

lupus. A, Correlation of a low percent

of CD83-positive day 7 DCs, with

past or current lupus nephritis and

with levels of anti-dsDNA Abs. B,

Correlation of an increased percent of

CD1a-positive day 7 DCs with serum

complement levels; , p 0.05,

mean SEM of independent exper-

iments. Similar correlations were

seen at the mRNA level.


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efficient autoantigen-presenting cells and drive autoimmune re-

sponses. In addition, the role of costimulatory molecules is well

documented in murine models of lupus, including the observation

that treatment with a combination of blocking mAbs to CD80 and

CD86, before the onset of murine lupus, significantly improves

survival and severity of the disease (58, 59).

Given that mature myeloid DCs are able to break tolerance and

induce lupus autoantibodies in normal hosts (60), the increase in DC

maturation in SLE suggests that these abnormalities might be very

relevant in the induction and perpetuation of autoimmunity in SLE.

Increased secretion of IL-8 by lupus DCs could potentially con-

tribute to tissue damage, as this cytokine has been proposed to play

an important role in the development of lupus nephritis (6164)

and has been found to be elevated in lupus patients with CNS in-

volvement (65). IL-8 is produced by numerous cell types, includingmonocytes/macrophages and DCs (64, 66, 67). IL-8 is mainly active

on neutrophils, promoting their recruitment and also their strong ac-

tivation which triggers the leukotriene pathway, induces the release of

their granular content, elastase and lactoferrin and increases their ad-

herence to endothelial cells (6871). In fact, migration of neutrophils

is influenced by DCs primarily by IL-8 (64). IL-8 is also a chemoat-

tractant for other cell types including T cells (72). In addition, we

found a significant association between IL-8 secretion and serum lev-

els of anti-dsDNA. Anti-dsDNA can enhance the release of proin-

flammatory cytokines, including IL-8 and TNF- from mononuclear

cells to augment inflammatory reactions and can polarize the immune

reaction toward the Th2 pathway (7274). Therefore, an increase in

the production of this cytokine by DCs might enhance the ability torecruit cells in the glomerulus and enhance neutrophil adhesion to

vascular endothelium which in turn may contribute to renal and vas-

cular damage. Interestingly, a recent study has shown that the IL-8

gene and its receptor CXCR-2 are up-regulated in PBMCs from SLE

patients using microarrays (75).

The functional relevance of decreased levels of CD83 in SLE is

unclear at this point. CD83 is a cell surface membrane glycopro-

tein whose surface expression is largely restricted to DCs (55). The

precise functions of this molecule remain unknown (76, 77), but

CD83 may serve important roles during intercellular interactions

(7779), as membrane-bound CD83 increases the stimulatory ca-

pacity of DCs (79). Further, previous studies suggest that CD83

mediates adhesion to monocytes and CD8 T cells (80). CD83-Ig

enhances T cell proliferation and increases the proportion of CD8

T cells (80), and engagement of CD83 delivers a significant signal

FIGURE 5. Responses of monocyte-derived DCs to maturation stimuli

are abnormal in lupus DCs. A, Bar graphs display mean fluorescence in-

tensity of CD86 in DCs from 10 healthy controls, 5 RA patients, and 20

SLE patients before and after stimulation with TNF-. B, Representative

histograms of CD86 expression on DCs from 1 healthy control, 1 patient

with SLE, and 1 patient with RA before and after stimulation with LPS and

TNF-. C, Bar graphs display mRNA expression by real-time RT-PCR of

differentiation and maturation markers normalized to housekeeping genes

in DCs, after stimulation with LPS and TNF- in 6 healthy controls and 17

SLE patients. Results are reported as the fold change when compared with

unstimulated day 7 DCs. , p 0.05 when compared before and after

stimulation; results are presented as mean SEM.

FIGURE 6. Aberrant expression of differentiation and maturation mark-

ers in lupus myeloid DCs obtained from peripheral blood. A, Bar graphs

display expression of DC differentiation and maturation markers in freshly

isolated CD11c cells from peripheral blood, in 11 healthy controls and 24

SLE patients. B, Bar graphs display correlation of anti-dsDNA Abs with

CD1aDCs in peripheral blood. , p 0.05, results represent the mean

SEM of independent experiments.


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specifically supporting the expansion, survival and function of

newly primed naive CD8 T cells (81). CD8 T cells in lupus-

prone mice are impaired in expansion, acquisition of memory, se-

cretion of cytokine, and suppression of autoimmunity (8, 82) and

because CD83 appears to have a role in CD8 function, it is pos-

sible that down-regulation of the former could contribute to ab-

normal CD8 function in SLE. In addition, CD4 T cells that

develop in a CD83 mutant animal fail to respond normally fol-lowing allogeneic stimulation (83), at least in part due to an altered

cytokine expression pattern characterized by an increased produc-

tion of IL-4 and IL-10 and diminished IL-2 production, findings

typically seen in SLE. Thus, absence or decrease of CD83 in SLE

DCs may result in the generation of T cells with an altered acti-

vation and cytokine profile. Future studies will address these pos-

sibilities. The factors inducing down-regulation of CD83 in lupus

DCs remain to be determined. Although we detected that autolo-

gous serum down-regulates the percentage of DCs that express

CD83, this phenomenon was observed for both lupus and control

serum, and therefore cannot explain the differences in expression

of this molecule observed between the two groups. Similarly, treat-

ment with IFN- did not alter CD83 expression.

DCs from SLE patients responded to extrinsic maturation stimuli

by up-regulating the expression of maturation markers, further in-

creasing IL-8 secretion and enhancing T cell proliferative and activa-

tion responses. However, the up-regulation of some of the membrane-

bound maturation markers was blunted when compared with the

degree of up-regulation seen in healthy controls. It is possible that the

preactivation status of lupus DCs, as confirmed by the overexpression

of maturation markers even before maturation stimuli, makes the lu-

pus cells more refractory to further up-regulation of cell surface mat-

uration markers. Lupus DCs treated with maturation stimuli did be-

come more efficient at inducing allogeneic T cell proliferation, T cell

activation and cytokine secretion, than untreated lupus DCs.

Although we cannot entirely exclude the possibility that immuno-

suppressive drugs could play a role in the phenotypic and functional

differences of lupus myeloid DCs, correlation analysis did not show

an association between expression of specific differentiation and mat-

uration markers and secreted cytokines with different medications

used by SLE patients. Further, when we treated monocytes with drugs

commonly used to treat SLE and differentiated them into DCs, we

found no differences in the maturation and differentiation patterns

seen in untreated or treated cells. Also, while RA patients are also

exposed to immunosuppressive medications, DCs from patients with

this disease did not show any significant differences in the expression

of maturation and differentiation markers and cytokine profile when

compared with healthy controls. These observations suggest that ourfindings are not related to the use of medications in these patients. In

addition, an effect of corticosteroid treatment on DC function can

reasonably be excluded, as these agents have a short biological half-

life of 1236 h (84) and therefore can only have a marginal effect on

maturation of DCs after a 7-day culture. Further supporting that our

results are not related to medication use, a recent report in a novel

lupus-prone mouse strain, B6.Sle3 (not subject to any type of immu-

nosuppressive treatment), indicates that its DCs are hyperstimulatory,

more mature and proinflammatory, overexpress CD80 and CD86

among other costimulatory molecules, and induce T cell hyperactivity

(85), similar to our findings in human lupus DCs.

Our results on the differentiation and maturation status of my-

eloid derived DCs differ from the ones previously described byKoller et al. (54), where no significant increases in the expression

of maturation markers in lupus DCs and a decrease in T cell pro-

liferation by lupus DCs were found. We believe that these discrep-

ancies might be secondary to differences in methodology. The

number of patients studied by Koller et al. (54) was significantly

smaller and disease activity also appeared to be lower, although a

different scoring system was used. That group used positive selec-

tion to isolate monocytes, which could contribute to activation of

these cells, as has been reported for other PBMCs (86). In addition,

their cells were cultured for longer periods of time and under con-

ditions that were not serum-free. It has been demonstrated that

serum contains growth factors that could affect DC development,

differentiation and maturation in vitro (87, 88) and that serum-freeconditions like the ones used in our study lead to more optimal DC

harvest that non-serum free conditions (89). Furthermore, Koller et

al. (54) used irradiated DCs for their mixed-lymphocyte reaction

studies, while we used nonirradiated DCs. Irradiated DCs can un-

dergo apoptosis (90) and posttranslational modifications of pro-

teins during apoptosis can potentially modify T cell activation and

proliferation (90). Our results do confirm a previous observation

that CD83 expression on lupus DCs is not up-regulated after mat-

uration stimuli (91), are supported by similar ex vivo findings in

DCs isolated from peripheral blood and, as mentioned, are remi-

niscent of what has been reported in lupus animal models (85).

Our observations that DCs from RA patients do not display the

phenotypic abnormalities seen in SLE are in consensus what has

been previously been reported in the RA literature in monocyte-

derived DCs (92, 93). Our observations may not necessarily indicate

FIGURE 7. Increased IL-8 secretion by myeloid DCs from SLE pa-

tients. A, Results represent the mean concentration in supernatants (pico-

grams per milliliter) SEM in DC cultures from 12 healthy controls and

27 SLE patients. Concentrations were adjusted according to initial mono-

cyte number; , p 0.05 when comparing controls vs SLE in each culture

condition. B, Results represent the mean concentration in supernatants (pi-

cograms per milliliter) SEM in DC cultures from 7 healthy controls, 5

RA patients, 13 SLE patients with negative anti-dsDNA Abs, and 9 SLEpatients with elevated anti-dsDNA Abs. The variability in measurements

between A and B is due to different methodology (CBA for A, Luminex

array for B) and different samples measured.


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that all RA DCs have normal phenotype and function. Indeed,

monocyte-derived DCs might not reflect changes occurring in the

inflamed synovium and there is evidence that DCs in the joint of

RA patients show a distinct phenotype, with differential expression

of specific TLRs (94). DCs are abundant both in synovial tissueand in synovial fluid of RA patients (95) and it has been proposed

that RA synovitis may be a delayed-type hypersensitivity reaction

generated by the interaction of synovial DCs and T cells (96).

Furthermore, the cytokine profile in RA and SLE is known to be

distinctly different (35, 93, 97) and there is evidence that the mech-

anisms leading to autoimmunity in SLE and RA are probably quite

distinct (98, 99). Therefore, the lack of hyperactivated DCs in RA

patients does not imply that the abnormal DC phenotype observed

in SLE is not important in maintaining autoimmunity, and this

statement is supported by findings in lupus animal models (56, 85).

Immune complexes consisting of DNA and anti-dsDNA Abs

isolated from the sera of patients with SLE can induce plasmacy-

toid DCs to produce high levels of IFN- (100). Increased IFN-

production in SLE causes increased monocyte differentiation into

DCs (22). These cells are able to capture apoptotic cells and

present their Ags to autologous T cells and induce potent MLRs

(22). We propose that, in addition to the IFN-, there are endog-

enous abnormalities in myeloid DC differentiation and maturation,

as a serum effect and exogenous IFN- could not account for, or

abrogate, the differences in phenotype and function seen in lupusmonocyte-derived DCs. Purified nucleosomes directly induce in

vitro DC maturation of mouse bone marrow-derived DC, human

monocyte-derived DC and purified human myeloid DCs as ob-

served by stimulation of allogenic cells in MLR, IL-8 secretion,

and CD86 up-regulation (101), findings similar to the ones we

report in this study. Further, nucleosomes complexed with antinu-

cleosome Abs can activate DCs (102). Therefore, autoantigens and

immune complexes could play an important role in vivo by induc-

ing accelerated differentiation and maturation of lupus DCs.

Taken together, our data suggest that monocyte-derived DC dif-

ferentiation and maturation are altered in SLE and contribute to

enhanced T cell proliferation and activation and to an increase in

the secretion of proinflammatory cytokines. We hypothesize that

these events could help to initiate and maintain the autoimmune

response in lupus.

FIGURE 8. Lupus DCs promote increased al-

logeneic T cell proliferation and activation. A,

Bar graph represents percentage of allogeneic T

cells in phases G0-G5, after coculture for 5 days

with either unstimulated or stimulated (LPS

TNF-) DCs from 11 healthy controls and 19

SLE patients. Proliferation of untreated T cells

and PHA-treated T cells treated is also shown.

Results represent the mean SEM of indepen-

dent experiments. , p 0.05 when comparing

proliferation of T cells adding SLE DCs with

healthy control DCs and with untreated T cells.

, p 0.05 when comparing proliferation of T

cells stimulated with healthy control DCs vs un-

treated T cells. Both control and SLE-stimulated

DCs induced higher T cell proliferation rates

when compared with unstimulated healthy con-

trol or lupus DCs (p 0.02 for G0 in SLE; p

0.02 for G2

in control; p 0.04 for G2 in SLE

and p 0.04 for G3 in control). B, Representa-tive dot plots display CFSE and CD25 expres-

sion in allogeneic T cells after exposure to stim-

ulated DCs from 1 SLE patient or DCs from 1

healthy control. C, Bar graph represents percent-

age of CD40L allogeneic T cells after 24 h co-

culture with unstimulated or stimulated (LPS

TNF-) DCs from 4 healthy controls and 7 SLE

patients. Untreated T cells and PHA-treated T

cells treated are also shown. Results represent

the mean SEM of independent experiments. ,

p 0.01 when comparing unstimulated SLE vs

unstimulated healthy control DCs and p 0.03

when comparing stimulated lupus DCs vs stim-

ulated control DCs. There was also a statistically

significant difference (p 0.04) in up-regulationof CD40L between untreated and stimulated

DCs for both control and SLE patients.


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AcknowledgmentsWe thank Drs. David Fox, Bruce Richardson, and Michael Denny for crit-

ical review of the manuscript; Joel Joshua for help in patient recruitment,

and Berlex for providing human GM-CSF.

DisclosuresThe authors have no financial conflict of interest.

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