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The Journal of Immunology, 2004, 172: 7821-7831.
Copyright © 2004 by The American Association of Immunologists

Alterations in Lipid Raft Composition and Dynamics Contribute to Abnormal T Cell Responses in Systemic Lupus Erythematosus1,2

Sandeep Krishnan*, Madhusoodana P. Nambiar*,{dagger}, Vishal G. Warke*, Carolyn U. Fisher*, Jeanne Mitchell{ddagger}, Nancy Delaney{ddagger} and George C. Tsokos3,*,{dagger},{ddagger}

* Department of Cellular Injury, Walter Reed Army Institute of Research, Silver Spring, MD 20910; {dagger} Department of Medicine, Uniformed Services University of the Health Sciences, Bethesda, MD 20814; and {ddagger} Walter Reed Army Medical Center, Washington, DC 20307


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
In response to appropriate stimulation, T lymphocytes from systemic lupus erythematosus (SLE) patients exhibit increased and faster intracellular tyrosine phosphorylation and free calcium responses. We have explored whether the composition and dynamics of lipid rafts are responsible for the abnormal T cell responses in SLE. SLE T cells generate and possess higher amounts of ganglioside-containing lipid rafts and, unlike normal T cells, SLE T cell lipid rafts include FcR{gamma} and activated Syk kinase. IgM anti-CD3 Ab-mediated capping of TCR complexes occurs more rapidly in SLE T cells and concomitant with dramatic acceleration of actin polymerization kinetics. The significance of these findings is evident from the observation that cross-linking of lipid rafts evokes earlier and higher calcium responses in SLE T cells. Thus, we propose that alterations in the lipid raft signaling machinery represent an important mechanism that is responsible for the heightened and accelerated T cell responses in SLE.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Systemic lupus erythematosus (SLE)4 is a complex autoimmune disease of unknown etiology. Recent studies have reported several functional abnormalities of T cells in mediating pathogenesis of SLE. However, the biochemical processes underlying these abnormal T cell responses are only beginning to be understood (1, 2). Previous studies have demonstrated abnormalities at multiple levels in the TCR signaling cascade in SLE T cells. SLE T cells display marked reduction in the expression of the CD3{zeta} chain and the kinase LCK (3, 4, 5, 6), increased activity of mitogen-activated protein kinase (7), and a concomitant up-regulation of CD3{zeta} homologue FcR{gamma}. FcR{gamma} associates with TCR and utilizes Syk kinase instead of ZAP-70 to mediate downstream signaling (8). This "rewiring" of T cells in SLE assumes significance considering that in its context, biochemical signaling events such as intracellular protein tyrosine phosphorylation and calcium responses (intracellular Ca2 concentration ([Ca2+]i)) become augmented (3). While attempting to identify mechanisms that mediate heightened T cell responses in SLE, we found that transfection of the FcR{gamma} chain into normal T cells amplifies TCR signaling events such as protein tyrosine phosphorylation and calcium signaling in a Syk kinase-dependent manner and can thus mimic SLE T cell signaling abnormalities (9). Conversely, we detected that heightened TCR signaling responses observed in SLE T cells can be dampened by replenishing CD3{zeta} into SLE T cells (10). These observations suggest the presence of mechanisms responsible for the aberrant amplification of signals in SLE T cells by the "rewired TCR" that involve close interactions between disparate sets of signaling proteins (11).

Lipid rafts are highly ordered cholesterol and ganglioside-rich platforms that can facilitate and coordinate close interactions between critical signaling molecules to amplify downstream signaling (12, 13). In normal T cells, ligation of the TCR induces rapid lipid raft clustering that leads to concentration of signaling proteins at the area of contact between APCs and T cells known as the immunological synapse (14, 15, 16). Efficient formation of the immunological synapse is critical to amplification of signals downstream of the TCR and subsequent T cell activation (15). Conversely, loss of integrity of lipid raft processes have been shown to play a major role in the pathogenesis of several diseases such as infections, allergies, and neoplasms (17).

We hypothesized a prominent role for lipid rafts in augmenting TCR-mediated responses in SLE T cells for several reasons. First, both CD3{zeta} and FcR{gamma} can signal through lipid rafts in different cellular contexts (18, 19). Second, increased tyrosine phosphorylation of cytoplasmic proteins, such as that observed in SLE T cells, has been shown to induce their recruitment to lipid rafts (20). Third, we observed that in SLE T cells, CD3{epsilon} is localized within discrete surface membrane clusters that are also enriched for linker for activation of T cells (LAT), an important resident of lipid rafts, suggesting that the bulk of CD3{epsilon} resides within lipid rafts in SLE (21). Fourth, we observed that in SLE T cells, the bulk of the residual CD3{zeta} is associated with the detergent-insoluble fraction and could be solubilized by methyl-{beta}-cyclodextrin (MbCD), a lipid raft disrupting agent, thus suggesting an increased localization of CD3{zeta} within lipid rafts in these cells (21).

Several factors can influence the strength of signals controlled by lipid rafts such as, lipid raft pool size, membrane distribution pattern, protein content, and the kinetics of cytoskeletal rearrangements following T cell stimulation. Thus, we assessed the contribution of lipid rafts to abnormal T cell responses in SLE by comparing these parameters between normal and SLE T cells. We demonstrate that the lipid raft composition and dynamics are altered in SLE T cells and that these changes may contribute to amplification of TCR-mediated signals in SLE.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Patient population

All patients studied fulfilled the American College of Rheumatology classification criteria for SLE (22). Our study included 20 inactive SLE patients, 32 active patients with SLE disease activity index (SLEDAI) ranging from 1 to 16, 5 disease controls with Sjogren’s syndrome and rheumatoid arthritis, and 48 healthy volunteers (Table I). All SLE patients, disease controls, and healthy volunteers were from the age group of 19–71 years. Patients who were being treated with prednisone were asked not to take this medication for at least 24 h before blood was drawn. Because of paucity of cell numbers obtained from the disease group, many of whom were leukopenic, different sets of experiments were performed with different patient groups but care was taken to include patients of SLEDAI scores ranging from low (0–5 range) to high (6–16 range) in each group. Within each group, patient samples were matched with normal samples of similar ages and gender. Monocyte-depleted lymphocytes from healthy controls were kindly provided by Dr. D. Hoover (Walter Reed Army Institute of Research (WRAIR), Silver Spring, MD). The study protocol was approved by the Health Use Committees of WRAIR and Walter Reed Army Medical Center. Written informed consent was obtained from all participating patients and volunteers.


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  		Table I. Details of the patients involved in this study

 
T lymphocyte isolation

PBMC were obtained by density gradient centrifugation over Ficoll and T cells were isolated by positive depletion of non-T cells by magnetic separation (Miltenyi Biotec, Auburn CA) as described previously (23) or using Rosette Sep (StemCell Technology, Vancouver, British Columbia, Canada) following the manufacturer’s instructions. In all cases, the percentage of T cells in the isolated subpopulation was >98% as determined by anti-CD3{epsilon} staining and flow cytometry (FACS) using FACSCalibur (BD Biosciences, Mountain View, CA).

Cell stimulation, CD3{epsilon} capping, and confocal microscopy

Cells (0.5 x 106) were adhered on polylysine-coated glass slides for 1 h at room temperature. For capping experiments, cells were stimulated with 1/100 IgM anti-CD3 Ab for various time points at 37°C and immediately fixed with 4% paraformaldehyde solution. For intracellular staining, cells were permeabilized with a buffer containing 0.05% (w/v) saponin in RPMI 1640 medium. Cells were stained with matching pairs of primary and fluorochrome-labeled secondary Abs (Jackson Immunoresearch Laboratories, West Grove, PA) for 1 h and 30 min, respectively, at room temperature. Cells were washed, air dried, and mounted using Gel/Mount (Biomeda, Foster City, CA) and coverslips applied. Samples were analyzed with a laser scanning confocal fluorescence microscope (1X70; Olympus, Lake Success, NY) with Lasersharp-2000 software (Bio-Rad, Richmond, CA). For some images where comparison was made between cytoplasmic and membrane distribution of proteins, the Z-stack feature of the software was used to obtain a library of images of various sections of cells, and sections showing both cytoplasmic and membrane portions clearly were selected. All images were processed to obtain the final images using Adobe Photoshop 7.0 Adobe Systems, Mountain View, CA).

GM1 staining and flow cytometric analysis

T cells, resting or activated with anti-CD3 (2 µg/ml, clone OKT3; Ortho Biotech, Raritan, NJ) and anti-CD28 (1 µg/ml; BD PharMingen, San Diego, CA) immobilized on plastic for 72 h as described before (24), were fixed with 0.75% paraformaldehyde and permeabilized with a buffer containing 0.05% (w/v) saponin in PBS and stained with cholera toxin B (CT-B)-FITC for 1 h. Cells were washed twice and fluorescence was detected by FACS.

Actin polymerization assay

The procedure was done as described before (25), with minor changes as follows: cells were washed twice with PBS and stimulated with IgM anti-CD3 (1/100) as required and treated with 3.7% paraformaldehyde for 8 min to stop the reaction. Cells were washed twice with PBS and permeabilized and stained with staining buffer containing 0.1% Triton X-100 and 0.2 µM phalloidin-FITC (Sigma-Aldrich, St. Louis, MO) for 40 min. Cells were washed, resuspended in PBS and F-actin content was analyzed by FACS. In each experiment, normal and SLE T cells were activated and analyzed together.

Preparation of detergent-resistant membrane fractions

We modified the protocols described before (20, 26) as follows: T cells (25 x 106) from healthy volunteers and lupus patients were used, either resting or following activation with IgM anti-CD3 Ab for 2 min. Cells were washed twice with PBS and resuspended in 1 ml of ice-cold lysis buffer (1% Brij 58, 25 mM MES, 150 mM NaCl, supplemented with 1 mM sodium orthovanadate, 2 mM EDTA, 1 mM PMSF, and 1 µg/ml aprotinin). Cells were homogenized with 12 strokes of Dounce homogenizer and incubated on ice for 30 min, mixed with an equal volume of 85% w/v sucrose (prepared in 25 mM MES, 150 mM NaCl, pH 6.5) and placed at the bottom of a 12-ml ultracentrifuge tube. A step gradient consisting of 6 ml of 35% sucrose and 4 ml of 5% sucrose was layered on top of it and the tubes were centrifuged for 18–20 h at 200,000 x g (39,000 rpm) at 4°C in a SW41 rotor (Beckman Instruments, Fullerton, CA). Twelve 1-ml fractions were collected starting from the top. Fractions 4 and 5 were indicative of rafts and were pooled for each experiment while fraction 12 was used as the cytosolic fraction.

Immunoblotting

Proteins from raft and cytosolic fractions were concentrated using a standard TCA precipitation protocol and then suspended in denaturing sample buffer (27). Following resolution on a 4–12% bis-Tris NuPage (Invitrogen, San Diego, CA), proteins were transferred onto polyvinylidene difluoride (PVDF) membranes and then incubated with specific Abs. The following Abs were used: anti-FcR{gamma} (Upstate Biotechnology, Charlottesville, VA), and anti-Lck (2102), anti-CD3{zeta} (clone 6B10.2), anti-phosphorylated Syk (p-Syk; clone 4D10), anti-LAT (FL-233), and anti-actin (I-19) from Santa Cruz Biotechnology (Santa Cruz, CA). HRP-coupled Abs (Santa Cruz Biotechnology) were used as secondary Abs and detection was performed with ECL (Amersham Pharmacia Biotech, Piscataway, NJ). For Vav1 association experiments, collection of actin cytoskeletal fractions has been described before (28). Briefly, the detergent-insoluble pellets obtained by lysis of 5 x 106 cells as described above were treated with cytochalasin B (0.1 mg/ml; Sigma-Aldrich) for 1 h at 37°C, samples were spun, and supernatants were collected. Samples were boiled for 2 min and reduced with 1 M DTT and immunoblotting experiments were performed with anti-Vav (Upstate Biotechnology) or anti-phospho-Vav (Santa Cruz Biotechnology), as discussed above.

Cross-linking of lipid rafts and measurement of Ca2+ influx

Five million cells were washed with RPMI 1640 and incubated with 1 µg/ml Indo-AM (Molecular Probes, Eugene, OR) for 30 min at 37°C, washed with RPMI 1640, and kept on ice. Cells were analyzed using an Epics Altra (Coulter, Hialeah, FL) flow cytometer equipped with a high-power dual wavelength (365- and 488-nm argon laser). CT-B subunit patching of rafts was adopted as described before (29), with changes as follows: cells were stained with CT-B (10 µg/ml; Calbiochem, San Diego, CA) for 10 min followed by anti-CT-B (1/250; Calbiochem) for 4 min on ice and then warmed up to 37°C in 1 min. For some experiments, cells were treated with varying doses of MbCD for 30 min at 37°C as reported before (20, 30). Samples were run and at 40 s, either OKT3 (10 µg/ml) or the isotype control mIgG2a was added followed by goat anti-mouse cross-linker at 1 min and the ratio of the fluorescence, which is directly proportional to free cytosolic Ca2+, was recorded for a period of 10 min as described before (3, 10). In some cases, MbCD-treated cells were treated with 1 µM thapsigargin (Sigma-Aldrich) and calcium response was analyzed immediately.

Densitometry and statistical analysis

Densitometric analysis of the autoradiograms was performed with the software program GelPro (Media Cybernetics, Silver Spring, MD). Statistical analysis of the data was done by paired t test using the software MINITAB, version 13 (Minitab, State College, PA). A value of p ≤ 0.05 was considered to be significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
SLE T cells display robust capacity to generate lipid rafts

To understand the role of lipid rafts in altered signal transduction in SLE T cells, we first asked whether SLE T cells contained larger pools of lipid rafts compared with normal T cells by analyzing the surface and total (surface and intracellular) GM1 expression in these cells. GM1 ganglioside has been traditionally used as a lipid raft marker (27, 29, 31, 32, 33) and can be detected by flow cytometry by virtue of its ability to bind to CT-B (32, 34, 35). Normal and SLE T cells, either freshly isolated or stimulated with anti-CD3 and anti-CD28 for 72 h, were stained with CT-B-FITC and analyzed by flow cytometry. Although similar surface levels of GM1 were found in freshly isolated normal and SLE T cells (Fig. 1A), SLE T cells have a significantly larger intracellular pool of GM1 than normal T cells (Fig. 1B; normals = 1-fold, SLE = 1.43-fold, p = 0.009). Anti-CD3- and anti-CD28-mediated activation resulted in dramatic increases in both surface and total GM1 expression in both cell types. However, unlike freshly isolated T cells, activated SLE T cells generated higher amounts of GM1 ganglioside than activated normal T cells detected both in the total and membrane pool (surface (Fig. 1A), normals = 15.96-fold, SLE = 22.06-fold, p = 0.036; total (Fig. 1B), normals = 2.84-fold, SLE = 4.16-fold, p = 0.008). These observations were unaffected by the SLEDAI scores of the patients or the treatment regimen, suggesting that higher levels of GM1 expression occurred independent of SLE disease activity. To determine whether the higher levels of GM1 expression following activation were due to increased production of GM1 by SLE T cells, we repeated the activation studies in T cells isolated from normal and SLE patients in the presence or absence of sphingosine N acetyltransferase inhibitor, fumonisin B1 (32, 36). The dose of fumonisin B1 used (25 µM) was monitored carefully to ensure that it did not affect the function of T cells, as suggested by optimum production of IL-2 by normal T cells (Ref.32 and data not shown). Activation of cells in the presence of the vehicle DMSO alone did not limit the production of GM1 in SLE or normal T cells (Fig. 1C; normals = 3.17-fold, SLE = 4.57-fold, p = 0.04). However, in the presence of fumonisin B1, both subsets displayed markedly lower levels of total GM1 (normals = 1.32-fold, SLE = 2.91-fold, p = 0.015; Fig. 1C), whereas unstimulated normal and SLE T cells cultured in the presence of fumonisin B1 did not demonstrate appreciable differences in the levels of GM1 expression (data not shown). These findings were observed at both 48 h (data not shown) and 72 h (Fig. 1C). This experiment confirms that the observed increase in GM1 levels in these cells was due to increased production induced by T cell activation. We conclude that SLE T cells possess more extensive basal levels of lipid rafts and greater capacity to generate lipid rafts than normal T cells in response to stimulation.



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  		FIGURE 1. SLE T cells have larger lipid raft compartments and produce lipid rafts more robustly than normal T cells. T cells from normal subjects and SLE patients were cultured in 24-well plates coated with anti-CD3 and anti-CD28 for 72 h as described in Materials and Methods. Cells were harvested, fixed with 0.75% paraformaldehyde, and stained directly or following permeabilization with CT-B-FITC and GM1 staining detected by FACS. Values on the y-axis represent the ratio of fold increases in mean fluorescence intensity (MFI) of each time point over MFI of normal T cells at 0 h. Demonstration of (A) surface GM1 expression (n = 5, *, p < 0.05, (B) total GM1 expression (n = 5, *, p < 0.05, and (C) inhibition of GM1 production by fumonisin B1 (n = 3, *, p < 0.05.

 
Qualitative alterations in the composition of lipid rafts in SLE

Next, we asked whether the protein composition of lipid rafts differed between SLE and normal T cells. We had previously demonstrated that in SLE T cells, residual CD3{zeta} was limited to detergent-insoluble fractions, thus suggesting that CD3{zeta} might be localized to lipid rafts of SLE T cells (21). However recent evidence has accumulated suggesting that detergent-insoluble fractions may not necessarily depict lipid rafts (37, 38). Therefore, to clarify this point and to identify possible alterations in the protein composition of lipid rafts in SLE, we performed immunofluorescence confocal microscopy to compare the location of critical proteins such as LAT, CD3{zeta}, the CD3{zeta} homologue FcR{gamma}, and phospholipase C{gamma}1 (PLC{gamma}1) in normal and SLE T cells, relative to lipid raft marker GM1. Several interesting findings emerged from these experiments. We noted that consistent with our previous results (21), both LAT and CD3{zeta} were limited to discrete clusters on the membrane of SLE T cells that also colocalized with GM1, whereas they were uniformly distributed on the normal T cell membrane (Fig. 2 and data not shown). Additionally, we noted that FcR{gamma} chain was also present within GM1-enriched clusters on the membrane of SLE T cells. These observations were in contrast to those in normal T cells in which FcR{gamma} chain expression was not detected (Fig. 2 and Refs. 8 and 39). In addition, while a majority of normal T cells displayed intracellular stores of PLC{gamma}1, in a majority of SLE T cells, PLC{gamma}1 was restricted to the cell membrane (percentage of cells displaying intracellular stores of PLC{gamma}1: normals = 67 ± 15%, SLE = 28 ± 14.9%, p = 0.046). Activation of normal and SLE T cells with anti-CD3 induced a shift in the localization of PLC{gamma}1 to the membrane (percentage of cells displaying intracellular stores of PLC{gamma}1: normal cells = 9 ± 5.03%, SLE = 17 ± 7.06%, p = 0.16), consistent with the findings reported by others in Jurkat cells (Ref.40 and Fig. 2). The above findings suggest that the majority of T cells in SLE resemble activated T cells. Our observations that the pattern of distribution of these crucial proteins did not vary between T cells from normal subjects and individuals with Sjogren’s disease suggest that the observed alterations in the lipid raft composition are unique to SLE T cells (Fig. 2).



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  		FIGURE 2. Altered composition of lipid rafts in SLE T cells demonstrated by confocal microscopy. Normal and SLE T cells were adhered to polylysine-coated slides (Sigma-Aldrich) and stained directly or following activation with IgM anti-CD3 for 2 min. Cells were fixed and stained with CT-B-FITC (green), followed by permeabilization and staining with anti-LAT (1/50), anti-FcR{gamma} (1/50), or anti-PLC{gamma}1 (1/50) followed by corresponding tetramethylrhodamine isothiocyanate (TRITC)-conjugated secondary Abs (red), and analyzed by a laser scanning confocal microscope as described in Materials and Methods. The merged images (yellow) indicate colocalization of respective proteins with GM1. Twenty-five cells were counted per field and the percentage of positive cells was calculated. A representative cell is shown for each sample. Magnification used was x100 oil objective. These results are representative of six experiments.

 
To confirm our observations from confocal microscopy and to detect possible alterations in the distribution of other crucial signaling molecules in the lipid raft fractions of SLE T cells, we isolated lipid rafts from normal and SLE T cells by performing sucrose density gradient ultracentrifugation. Lipid raft fractions were identified as fractions 4 and 5 (Fig. 3A) by the presence of GM1 expression and enrichment for LAT protein, that is constitutively associated with lipid rafts (41), and fraction 12 was considered as the cytosolic fraction (31). Because such experiments required large numbers of T cells and we were limited by the low yield of cells from SLE patients, the proteins were concentrated by the standard TCA precipitation method for further analysis (27). Comparison between raft and cytosolic fractions of normal and SLE T cells revealed that CD3{zeta} was present in low amounts in lipid rafts of SLE T cells and in even lower levels in the cytosolic fractions (Fig. 3B). Additionally, we detected FcR{gamma} chain exclusively in SLE T cells in both raft and cytosolic fractions, while the expression and distribution of LAT was similar between normal and SLE T cells (Fig. 3C). Our results showing that LAT levels remain unaltered despite increases in GM1-containing lipid rafts in SLE are consistent with observations made by others in effector CD8 T cells that bore higher levels of lipid rafts and yet displayed similar levels of LAT compared with resting naive and memory cells (33). Next, we compared the levels of LCK within raft and non-raft fractions of normal and SLE T cells. In normal T cells, substantial amounts of LCK are found within lipid rafts (42, 43). Consistent with previous reports (6), levels of LCK were markedly lower in both lipid raft and cytosolic fractions of SLE T cells compared with respective fractions of normal T cells (Fig. 3D). In addition, we noted that detectable levels of active Syk kinase (phosphorylated Syk) were consistently observed only in the lipid raft fractions of SLE cells (Fig. 3D), while it was undetected in raft and non-raft fractions of normal cells. Activation of SLE T cells by anti-CD3 induced association of activated Syk with lipid rafts of SLE but not normal T cells. We could not detect ZAP-70 in the lipid rafts of SLE T cells (data not shown). These results complement our previous observations that SLE T cells preferentially utilize Syk kinase whereas normal T cells utilize ZAP-70 (8, 44). The preferential usage of Syk kinase by SLE T cells also suggests that SLE T cells may be in a state of activation. Previously, we have shown that activated effector T cells generated from normal T cells utilize Syk instead of ZAP-70 (39). We could not detect PLC{gamma}1 in either raft or cytosolic fractions of both cell types by Western blotting, possibly due to low levels of expression of this protein or because of the limitations imposed by a low number of T cells in the density gradient experiments (data not shown). There was also no detectable difference in the phosphorylation pattern of LAT in SLE and normal T cells (data not shown). We also observed that the above qualitative changes in the lipid raft composition were unaffected by disease activity (SLEDAI scores).



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  		FIGURE 3. Altered composition of lipid rafts in SLE T cells demonstrated by sucrose density gradient analysis. Normal and SLE T cells (25 x 106) were lysed with a lysis buffer containing 1% Brij 58 as such or following activation with IgM-anti-CD3 (1/100) for 2 min and raft and cytosolic fractions were extracted by discontinuous sucrose density gradient ultracentrifugation. Proteins of each fraction were concentrated by standard TCA precipitation and resolved on a 4–12% bis-Tris-NuPage and transferred to PVDF membranes. The blot was probed with anti-LAT (1/1000), anti-CD3{zeta} (1/1000), anti-actin (1/1000), anti-FcR{gamma} (1/1000), anti-p-Syk (1/1000), or anti-LCK (1/1000). A, Identification of raft fractions as fractions 4 and 5 in SLE T cells by probing blot with CT-B-HRP (Sigma-Aldrich). Various fractions depict portions of the cell with varying density, fraction 12 being the heaviest and depicting cytosolic portion (B) nonreduced gel showing CD3{zeta} association with raft and cytosolic fractions of normal (n = 2) and SLE (n = 3), (C) nonreduced gel showing FcR{gamma} and LAT association with raft and cytosolic fractions of normal (n = 6) and SLE (n = 6), (D) reduced gel (1 M DTT) comparing localization of p-Syk and LCK in normal (n = 6) and SLE (n = 6). Densitometric ratios of various proteins with LAT or {beta}-actin are shown below the gels. R, Raft fraction; C, cytosolic fraction; Stim., stimulus.

 
Increased lateral membrane mobility in SLE T cells

While exploring alterations in the lipid raft composition in SLE T cells by confocal microscopy, we unexpectedly noted that activation of SLE T cells with the strong CD3 cross-linker, IgM anti-CD3{epsilon} for 2 min, resulted in the formation of CD3 caps in a majority of cells, whereas T cells from normal and Sjogren’s syndrome demonstrated only receptor clustering (Fig. 2). We suspected an increase in lateral membrane mobility of proteins and lipids in SLE T cells as a possible mechanism behind this phenomenon. To address this issue, we compared the kinetics of anti-CD3-induced capping of TCR-CD3 complexes in normal and SLE T cells by fluorescent microscopy. Cells were activated with IgM anti-CD3{epsilon} for various time points as depicted in Fig. 4, and the number of cells demonstrating capping of CD3{epsilon} were counted. Similar to our earlier observations in CD4 cells (39) and as observed in Fig. 2, in normal T cells, TCR-CD3 complexes were uniformly distributed on the surface membrane. Cross-linking of CD3{epsilon} induced TCR-CD3 clustering within 2 min (8 ± 3.27% capped, n = 6, p = 0.014) and, within 10 min, a majority of cells capped (75 ± 11.9% capped, n = 6, p = 0.001; Fig. 4, top panel, boxes 1a, 2a, and 3a, and bottom right panel). By contrast, freshly isolated SLE T cells consistently exhibited clustering of TCR-CD3 complexes on the surface (12 ± 3% capped) and demonstrated receptor capping in a majority of T cells within 2 min of CD3{epsilon} cross-linking (79 ± 5.26% capped, n = 6, p = 0.002; Fig. 4, top panel, boxes 1d, 2d, and 3d, and bottom right panel), demonstrating that the kinetics of receptor capping or lateral membrane mobility is accelerated in SLE T cells. The caps were maintained up to 30 min of CD3 cross-linking without internalization of the receptor complexes in all three cell types (Fig. 4, top panel, column 4). All of these observations occurred independent of SLE disease activity and treatment regimen. Our observations that disease control cells from patients with Sjogren’s syndrome displayed a pattern similar to that of normal T cells (cf Fig. 4, top panel, boxes 1a, 2a, and 3a and 1g, 2g, and 3g, and bottom right panel) suggest that faster lateral membrane mobility of proteins is a feature specific to SLE T cells. Because the CD3 clusters and caps also colocalized GM1 (Fig. 4, lower left panel), these results suggest that SLE T cells are characterized by faster kinetics of lipid raft clustering and polarization.



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  		FIGURE 4. IgM anti-CD3-induced receptor capping is accelerated in SLE T cells. Freshly isolated T cells from normal, SLE, and Sjogren’s patients were adhered to polylysine-coated slides, directly fixed or activated with IgM-anti-CD3{epsilon} Ab at 37°C at 2, 10, and 30 min to induce CD3{epsilon} cap formation before fixation with 4% paraformaldehyde. Cells were then surface stained with CT-B FITC (red) and anti-CD3{epsilon} Ab (clone UCHT1) and counterstained with anti-mouse-TRITC (red) and analyzed by a laser scanning confocal microscope. The merged images (yellow) represent colocalization of respective proteins with GM1. Twenty-five cells were counted per field and the percentage of positive cells was calculated. A representative cell is shown for each sample. Magnification used was x100 oil objective. These results are representative of six experiments. Bottom left panel, Single-field image of clusters of cells representative of 2-min activation. Bottom right panel, Statistical representation of the capping data showing percentages of cells that demonstrate CD3 capping (*, p < 0.05).

 
Alterations in actin dynamics in SLE T cells

Remodeling of cytoskeleton is an essential component of receptor cap formation and integrity of the immunological synapse. Because actin polymerization is a critical event in this process (45, 46, 47), we asked whether faster kinetics of receptor capping observed in SLE T cells was associated with accelerated kinetics of actin polymerization in these cells. Comparison of F-actin polymerization levels (detected by binding of polymerized F-actin to phalloidin) in normal and SLE T cells activated for various time points with IgM anti-CD3 (Fig. 5A, top panel) demonstrated that while in normal T cells, peak levels of actin polymerization occurred at 1 min of activation and in SLE T cells it occurred between 30 s and 1 min (Fig. 5A, bottom panels). Another notable difference was that, while in normal T cells, high levels of polymerized F-actin were maintained up to 15 min following activation; in SLE T cells, F-actin levels dropped rapidly to values less than those observed in normal T cells by 15 min following activation (Fig. 5A, bottom right panel). These observations demonstrate that compared with normal T cells, SLE T cells display faster kinetics of actin polymerization and depolymerization. We used IgM anti-CD3 Ab for these experiments because of its ability to cross-link CD3 without addition of a cross-linker Ab as would have been required with OKT3 (39), thus useful in short time point analysis of actin polymerization.



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  		FIGURE 5. Accelerated actin cytoskeleton rearrangement events in SLE T cells. A, FACS analysis of phalloidin-FITC staining of normal and SLE T cells following activation with IgM-anti-CD3 (1/100) for indicated time periods. The reaction was stopped after each time point by treatment with 3.7% paraformaldehyde. The top panel represents overlay of histograms representing each time point (open) with that of 0 s (shaded). The bottom left panel is a representative histogram overlay comparing phalloidin staining at selected time points (open) with that at 0 s (shaded). The bottom right panel is a statistical comparison of phalloidin staining at various time points. Values on the y-axis represent ratios of fold increases in MFI of each sample over MFI at 0 h (*, p < 0.05). Data are representative of five separate experiments. B, Whole-cell lysates from normal and SLE T cells, nonactivated or activated with IgM-anti-CD3 (1/100) for 2 min, were loaded (20 µg/lane) and resolved on 4–12% bis-Tris NuPage gels, blotted onto PVDF membranes, and probed with anti-Vav1 (1/1000), anti-phospho-Vav1 (p-Vav1; 1/1000), and anti-actin (1/1000). A representative gel of six experiments is shown. The right panel shows a statistical analysis of Vav1 expression in 13 normal subjects and 21 SLE patients. The values on the y-axis are represented as the mean of ratios of densitometric values of Vav1:actin of each sample. C, Comparison of phospho-Vav1 expression in cytochalasin-B treated Triton X-100-insoluble fractions from 5 x 106 normal or SLE T cells. Data are representative of six separate experiments.

 
To understand the mechanisms that alter the rate of actin polymerization in SLE, we explored the expression and activity of key proteins involved in actin polymerization such as WASP, Vav1, and Rac-1 (48, 49, 50). Immunoblot analysis of lysates obtained from normal and SLE T cells demonstrated that expression of WASP and Rac-1 remained unchanged in SLE T cells (data not shown). However, we observed that in up to 55% of SLE T cells, there was a significant increase in the expression of Vav1 (normal subjects, n = 13; SLE, n = 21, p = 0.014; Fig. 5B). This increase in Vav1 expression was unrelated to SLE disease activity and was observed at random in patients with SLEDAI scores ranging from 0 to 16. Of the 45% patients that did not demonstrate an increase in a Vav1:actin ratio, only one patient demonstrated a decrease, whereas all other patients displayed similar levels of Vav1:actin ratio as normal subjects (data not shown). Vav1 has been shown to be important for reorganization of actin cytoskeleton, and in Vav1-deficient mice, Ab-mediated TCR capping is abrogated (50, 51). Therefore, we explored the possibility that Vav1 may be involved in mediating rapid kinetics of actin polymerization in SLE. Phosphorylation of tyrosine residues is essential for the activation of Vav1 (52, 53), and Vav1 induces actin polymerization primarily by its association with actin-binding proteins such as Rac-1, Rac-2, and Rho G proteins (49, 50, 54). Thus, we compared the amounts of phosphorylated Vav1 that associates with actin-bound proteins in normal and SLE T cells following treatment of 1% Triton X-100-insoluble fractions with cytochalasin B, which is known to release proteins from the cytoskeletal network (28). Analysis of actin cytoskeletal fractions of normal and SLE T cells revealed similar amounts of Vav1 in normal and SLE T cells (data not shown). However, the levels of phosphorylated Vav1 were higher within the cytoskeletal fractions of freshly isolated SLE T cells than within normal T cells (Fig. 5C). Following anti-CD3 Ab stimulation, the levels of phosphorylated Vav1 remained high in SLE T cells with 2.7 ± 0.23 (p = 0.034)- and 3.1 ± 0.5, (p = 0.04)-fold increases observed at 15 and 30 s, respectively (Fig. 5C). Taken together, the increase in Vav1 expression in SLE T cells and the presence of higher amounts of phosphorylated Vav1 within actin cytoskeletal fractions strongly suggest a role for Vav1 in mediating accelerated kinetics of actin polymerization in SLE T cells.

Enhanced calcium signaling in SLE T cells is mediated by lipid rafts

Calcium flux has been shown to be lipid raft dependent (20, 29). Thus, we asked whether previous cross-linking of lipid rafts evoked different levels of TCR-CD3-mediated calcium responses in normal and SLE T cells. Cross-linking of lipid rafts by treating cells with CT-B followed by anti-CT-B has been described before (29, 55) and has been shown to recruit additional phosphoproteins to lipid rafts (29). Consistent with this view, we observed in normal T cells that recruitment of PLC{gamma}1 to membrane patches, an event critical for TCR-induced calcium flux, could be induced by cross-linking either TCR-CD3 receptor complex by anti-CD3{epsilon} or lipid rafts by CT-B/anti-CT-B (Fig. 6A). We observed that resting T cells demonstrated PLC{gamma}1 in both membrane and cytoplasm, with a majority of cells demonstrating cytoplasmic stores of PLC{gamma}1 (Fig. 6A), a result consistent with that reported previously for Jurkat cells (20). Following anti-CD3 and CT-B/anti-CT-B treatment, a majority of cells demonstrated a shift in PLC{gamma}1 location to the membrane fractions (Fig. 6A, right panel). Analysis of calcium responses of normal and SLE T cells to anti-CD3 revealed that although previous cross-linking of lipid rafts induced calcium fluxes in both normal and SLE T cells, calcium responses observed in SLE were more rapid and higher than those observed in normal cells (Fig. 6B). Our results are consistent with our previous findings that in SLE T cells, there is higher and sustained calcium responses than in normal T cells following stimulation with anti-CD3 (56). We also reasoned that if the augmented calcium responses observed in SLE T cells were mediated through lipid rafts, disruption of lipid rafts by MbCD should abrogate this heightened response. MbCD disrupts lipid rafts by depleting cholesterol and can alter signaling by altering associations of signaling proteins with the lipid raft compartment (30). We noted that although we could achieve near total abrogation of anti-CD3-induced calcium responses by treating both normal and SLE T cells with MbCD, SLE T cells required higher doses of MbCD (12 mM) to achieve the same decrease observed in normal T cells (10 mM; Fig. 6C), thus arguing for a direct role for lipid rafts in mediating the enhanced calcium response in SLE T cells. These results were consistently observed in SLE patients of varying levels of disease activity (SLEDAI scores 0, 4, 8, 12, and 12) as shown in the right panel of Fig. 6C. We observed that decreases in calcium responses following treatment with 12 mM MbCD could be reversed following resting both normal and SLE T cells for 10 h, thus showing that at the doses used for the above experiments, MbCD is not irreversibly cytotoxic (Fig. 6C). We also observed that prior treatment of these cells with MbCD did not abolish calcium response to thapsigargin (Fig. 6E), thus ruling out the possibility that the observed reductions in calcium response were due to depletion of calcium reserves caused by MbCD (57). Taken together, these results demonstrate that calcium signaling is augmented in SLE T cells, at least in part, via a lipid raft-dependent mechanism.



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  		FIGURE 6. Role of lipid rafts in mediating increased calcium responses in SLE T cells. A, PLC{gamma}1 distribution in freshly isolated normal T cells or after cross-linking CD3{epsilon} with anti-CD3 (OKT3) and goat anti-mouse IgG Abs or after cross-linking surface GM1 with CT-B followed by anti-CT-B as detailed in Materials and Methods. Cells were fixed with paraformaldehyde, permeabilized, and stained with anti-PLC{gamma}1 (1/1000) and secondary anti-rabbit-TRITC (red) and analyzed by confocal microscopy. Twenty-five cells were counted per field. Data are representative of cells positively staining for PLC{gamma}1. Data are representative of three similar experiments. Magnification used was x100 oil objective. Right panel shows statistical representation of PLC{gamma}1 localization. B, [Ca2+]i measurement was performed in normal and SLE T cells loaded with Indo-AM either without or after cross-linking lipid rafts with CT-B/anti-CT-B. Cells were stimulated with OKT3 and anti-mouse IgG and [Ca2+]i was measured with EPICS Altra flow cytometer. A representative of five separate experiments is shown. [Ca2+]i was also measured in normal and SLE T cells upon stimulation with OKT3 and goat anti-mouse IgG (C) following incubation for 30 min with varying concentrations of MbCD in the left panel, and statistical analysis in the right panel (D) following rest for 10 h after treatment with 12 mM MbCD (only SLE T cells) or (E) following incubation with 12 mM MbCD for 30 min followed by treatment with 1 µM thapsigargin. Mean ratio:mean fluorescence ratio corresponding to [Ca2+]i, *, not determined.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In this study, we provide evidence of significant differences in the lipid raft composition and dynamics between normal and SLE T cells. First, SLE T cells have a more extensive lipid raft pool and are more robust in their capacity to generate lipid rafts. Second, SLE T cells qualitatively differ in their lipid raft composition. Third, actin cytoskeletal dynamics are accelerated in SLE. Fourth, calcium responses are augmented in SLE T cells, at least in part, via a lipid raft-dependent mechanism. These findings were observed independent of the disease activity or treatment. Our results shed new light on molecular mechanisms that contribute to increased and accelerated TCR-mediated signaling in SLE T cells.

Among mechanisms that control lipid raft-mediated signaling, the size and distribution of lipid rafts are considered to be significant. The small size of individual rafts, for example, is believed to be important for maintaining the signaling proteins present in them in an inactive state, whereas clustering of multiple rafts induces activation of these proteins (17, 20). Thus, preclustering of lipid rafts observed herein in SLE T cells might tilt the balance in favor of preactivation of signaling proteins contained within lipid rafts and may mediate faster downstream signaling. This view is also supported by our observations that SLE T cells bear phosphoproteins within lipid rafts such as Syk and Vav1 and by previous observations that effector CD4 T cells also display clustering of surface receptors (39). Similarly, a larger pool of lipid rafts can also contribute to accelerated signaling. Thus, memory CD4 and CD8 T cells that have lower thresholds of activation possess larger lipid raft pools than naive T cells (32, 33).

Although it is not known presently what drives preclustering of lipid rafts in SLE T cells, our observations that the lateral membrane mobility of proteins (e.g., CD3 complex) and lipids (e.g., GM1) is significantly enhanced in SLE T cells suggest its role in facilitating preclustering of lipid rafts. It must be mentioned that our results differ from those reported 20 years ago and suggested a defect in receptor capping in SLE T cells (58). We believe that this difference is due to the fact that IgM anti-CD3 Ab used in our studies is a stronger cross-linker of CD3{epsilon} than OKT3, which requires an additional cross-linker such as goat anti-mouse IgG to exert its cross-linking effect, as observed by us previously (G. C. Tsokos, D. L. Farber, and S. Krishnan, unpublished observation and Ref.24). Therefore, IgM anti-CD3 Ab may overcome the limitations of weaker responses to cross-linkers such as OKT3 used in the previous study. In support of this view is our observation that peak receptor capping occurred in normal T cells within 10 min using IgM anti-CD3, whereas OKT3-induced maximum capping occurred within 30 min (our unpublished observation). Additionally, the previous study used lower concentrations of OKT3 (625 ng/ml) than us. When we used higher concentrations of OKT3 (10 µl/ml), we could demonstrate receptor capping in SLE T cells, albeit at a slower rate (between 2 and 5 min, data not shown). Other processes such as dysregulation of glycosylation of proteins in SLE may also be responsible in mediating lipid raft clustering. It has been shown that dysregulation of {beta}1,6-N-acetylglucosaminyltransferase V (Mgat5), an enzyme in the N-glycosylation pathway, can induce T cell activation by directly enhancing TCR clustering and thus lowering the threshold for T cell activation (59). We had previously observed that defects in glycosylation of the transcription factor Elf-1 might contribute to defective CD3{zeta} chain expression in SLE T cells (60, 61). Thus, it is likely that other defects in protein glycosylation may also coexist in SLE T cells that mediate lipid raft clustering.

The role of actin cytoskeletal rearrangements in mediating lipid raft clustering is unclear. However, it has been shown that lipid raft polarization to the immunological synapse is dependent upon actin cytoskeleton reorganization in T cells (49). Thus, during interactions between SLE T cells and APC, faster kinetics of actin polymerization may lead to rapid formation of the immunological synapse and thus accelerate TCR signaling. Because polarization of lipid rafts to immunological synapse is Vav1 dependent and Ab-mediated TCR capping is affected in the absence of Vav1 (49, 50, 51), our observation of higher levels of Vav1 expression, combined with higher levels of activated Vav1 association with actin-binding proteins in SLE T cells, strongly suggests that Vav1 plays an important role in mediating faster kinetics of receptor capping and actin polymerization and immunological synapse formation in SLE.

At the biochemical level, interactions between disparate signaling proteins present within lipid rafts impose another level of regulation upon TCR signaling. Thus, lipid rafts of naive, effector, and memory CD8 T cells display distinct protein profiles with distinct functional outcomes (33), and Th1 and Th2 cells differentially recruit TCR complexes to lipid rafts and display different functions (62). Previously, we had reported "rewiring of TCR" in SLE T cells in which CD3{zeta} is replaced by the FcR{gamma} chain in the TCR-CD3 complex and that this modified receptor complex preferentially utilizes Syk instead of ZAP-70 kinase (11, 44). Furthermore, we demonstrated that the excitation threshold of normal T cells could be lowered akin to SLE T cells by overexpressing FcR{gamma} that also increased the expression and activation of Syk kinase (9). Thus, our observations that FcR{gamma} and p-Syk are recruited to lipid rafts in SLE T cells suggest a prominent role for lipid rafts in mediating heightened T cell responses in SLE. Although the precise mechanisms mediating increased calcium responses in SLE are not known, there are important leads in this direction. There is growing support for a connection between Vav1 and PLC{gamma}1 in inducing calcium flux (54). Previous studies in Vav1 knockout mice have revealed that Vav1 is important for maintaining sustained calcium levels following T cell activation (63). Our observations of higher Vav1 activity in SLE, greater association of PLC{gamma}1 with lipid rafts, and higher calcium flux in SLE T cells following cross-linking lipid rafts strongly support this theory and suggest that lipid rafts act as a platform for mediating these events, although we cannot rule out contribution from other important factors. It must be mentioned here that although we could not determine the phosphorylation status of PLC{gamma}1, previous studies have demonstrated that localization of PLC{gamma}1 within lipid rafts would be sufficient to induce its activation (40). It is presently unclear why FcR{gamma} is up-regulated in SLE. However, the dependence of CD3{zeta} on Src kinases for phosphorylation and association with microfilament cytoskeleton (64, 65), the reduced expression of Src kinase Lck in SLE (6), and the ability of FcR{gamma} to associate with lipid rafts, combined with the increased association of Syk with lipid rafts (Fig. 3D) that has been shown to phosphorylate FcR{gamma} (66), suggest FcR{gamma} to be an ideal candidate to functionally associate with the TCR signaling complex in the context of reduced CD3{zeta} in SLE (8).

Similar to our findings in human SLE, previous studies have noted lowering of activation threshold for T cells from lupus-prone mice that are also hyperresponsive to TCR stimulation (67). Our present findings are significant in this regard and alterations in lipid raft dynamics could be an important contributor of enhanced T cell functions in murine SLE models as well. Currently it is not known what drives T cells in SLE to display activation phenotypes. We have drawn significant parallels between TCR signaling in SLE and effector T cells and raised the possibility that these activation phenotypes displayed by SLE T cells may be due to the presence of effector cells perpetuated via chronic stimulation by autoantigens (11, 44). This view is supported by 1) our previous observations that, similar to SLE T cells, effector CD4 T cells also display membrane clustering of TCR complexes (11, 39); 2) observations reported herein that higher levels of PLC{gamma}1 localize to cell membrane of SLE T cells similar to activated Jurkat T cells (40); and 3) association of active Syk, a kinase used preferentially in effector T cells (39), is present within lipid rafts of SLE T cells. In addition, these findings lend support to the view that biochemical markers of T cell markers are more reliable than phenotypic markers such as CD25 and CD69 in determining the long-term activation status of T cells (39, 68). The T cell population analyzed by us included various T cell subsets such as CD4 and CD8, and within CD4 T cells, probably Th1 and Th2 cells. Presently, it is unclear whether there exist differences in lipid raft-mediated signaling in these different subsets.

In conclusion, we demonstrate that alterations in the lipid raft distribution, composition, and biochemical associations contribute to the increased and accelerated TCR-mediated responses in SLE T cells. In addition, our studies add SLE to the list of diseases in which abnormal lipid raft processes are used to generate disease-specific immune abnormalities (17). Elucidating the precise signaling pathways regulated by lipid rafts in SLE may serve to effectively target these pathways and dampen increased T cell responses.


   Acknowledgments
 
We gratefully acknowledge Dr. Ellis Reinherz (Dana-Farber Cancer Institute, Boston, MA) for the kind gift of IgM anti-CD3 Ab, Dr. Michael Koenig (WRAIR, Silver Spring, MD) for help with confocal microscopy, Drs. Ashima Saxena and Ramchandra Naik (WRAIR) for help with sucrose gradient experiments, Dr. Juliann Kiang (WRAIR) for help with statistical analysis, and Leah Perkins (WRAIR) for her excellent technical assistance. We also thank Drs. Donna Farber (University of Maryland, Baltimore, MD) and Mojgan Ahmadzadeh (National Institutes of Health, Bethesda, MD) for critical reading of this manuscript and Dr. Gary Kammer (Wake Forest University School of Medicine, Winston Salem, NC) for extensive critical discussions.


   Footnotes
 
1 This work was supported by research grants from the National Institutes of Health (RO1-AI42269 and RO1-AI49954) awarded to G.C.T. Back

2 The opinions expressed herein are the private ones of the authors and they do not represent those of the Department of Defense or the Department of the Army. Back

3 Address correspondence and reprint requests to Dr. George C. Tsokos, Department of Cellular Injury, Walter Reed Army Institute of Research, Silver Spring, MD 20910-7500. E-mail address: gtsokos{at}usuhs.mil Back

4 Abbreviations used in this paper: SLE, systemic lupus erythematosus; ZAP-70, {zeta}-associated protein 70; [Ca2+]i, intracellular Ca2+ concentration; LAT, linker for activation of T cells; MbCD, methyl-{beta}-cyclodextrin; SLEDAI, SLE disease activity index; F-actin, filamentous actin; PVDF, polyvinylidene difluoride; CT-B, cholera toxin B; PLC{gamma}1, phospholipase C{gamma}1; MFI, mean fluorescence intensity; TRITC, tetramethylrhodamine isothiocyanate; p-Syk, phosphorylated Syk. Back

Received for publication February 5, 2004. Accepted for publication April 13, 2004.


   References
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 Introduction
 Materials and Methods
 Results
 Discussion
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