Stromal Cell–Derived Factor-1/CXCL12 Stimulates Chemorepulsion of NOD/LtJ T-Cell Adhesion to Islet Microvascular Endothelium

Mice used in this study were bred and housed at the Association for Assessment and Accreditation of Laboratory Animal Care, international-accredited Louisiana State University Health Sciences Center-Shreveport animal resource facility and maintained according to the National Research Council Guide for Care and Use of Laboratory Animals. Twelve-week-old female NOD and C57BL/6 mice were used for cell isolations.

All tissue culture media and reagents were purchased from Sigma (St. Louis, MO). Purified recombinant murine SDF-1α and tumor necrosis factor-α (TNF-α) proteins were purchased from R&D Systems (Minneapolis, MN). The MS1 mouse pancreatic islet microvascular endothelial cell line was cultured and maintained as we have previously reported (20). Endothelial cell cultures were routinely cultivated in T-75 flasks. For parallel plate flow chamber studies, endothelial cells were seeded into 35-mm culture dishes and grown to confluency. Confluent endothelial cell monolayers were stimulated with 10 ng/ml TNF-α 4 h before use in flow cytometric analysis or parallel plate flow chamber adhesion assays. Anti-CXCR4 antibody (10 μg/ml; R&D Systems) was used in some experiments to block T-cell CXCR4 binding to SDF-1.

Whole splenocytes from female NOD/LtJ or C57BL/6 mice were obtained from spleens that were ground between two frosted slides into a Petri dish containing RPMI as we have previously reported (21). The cell suspension was filtered twice through 70-μm cell strainers before lysis of erythrocytes. Target T-cell subpopulations were obtained by purification with CD3, CD4, or CD8 SpinSep Mouse T Cell Enrichment kits from StemCell Technologies (Vancouver, BC, Canada).

Hydrodynamic parallel plate flow chamber studies were performed as we have previously reported (20,22). Briefly, mouse leukocytes were labeled with a fluorescent dye by 30-min incubation at 37°C with 200 nmol/l Cell Tracker Green purchased from Molecular Probes. The labeled cells were resuspended in Hanks' balanced salt solution (HBSS) at 2 × 10⁵ cells/ml in a 200-ml beaker kept at 37°C and stirred at 60 rpm. A Glycotech flow chamber insert and gasket were used to form a laminar plate flow chamber that could be viewed on a microscope. The labeled cells were drawn from the beaker into the flow chamber across endothelial cell monolayers at a physiological shear rate of 1.5 dynes/cm² using a programmable digital syringe pump. All endothelial cell monolayers were washed three times with HBSS to remove remaining cytokines/chemokines before flow chamber assembly. Fluorescently labeled cells were viewed using a Nikon Eclipse TE-2000 epifluorescent microscope equipped with a Hamamatsu digital camera, and real-time digital video was captured using SIMPLE PCI software from Compix. The motion-tracking analysis feature of the software enabled calculation of individual cell-rolling velocity. Firmly adherent cells were defined as those that did not move one cell diameter over a 5-s period as determined by automated tracking and manual review of individual cells in each experimental field of view.

CD3⁺ cells were isolated as described above. Cells were rinsed in Tris-buffered saline (TBS) and spun for 5 min at 1,500 rpm. The resulting pellet was lysed in radioimmunoprecipitation assay buffer (50 mmol/l Tris-HCL, pH 8.0, 150 mmol/l NaCl, 1% Nonidet-40, 0.5% deoxycholate, and 0.1% SDS) supplemented with 0.1 μmol/l leupeptin, 0.3 μmol/l aprotinin, and 1 μmol/l phenylmethylsulfonyl fluoride (PMSF). Samples were sonicated for three 5-s intervals on ice. Protein determination was preformed using a Bio-Rad DC Protein kit (Bio-Rad, Hercules, CA) according to the manufacturer's instructions. Whole-cell protein homogenates (25 μg total protein) were loaded on 12% polyacrylamide SDS gels, and electrophoresis was performed as we have previously reported (23). Gels were transferred overnight to Immobilon-P7 (Bio-Rad), and subsequent membranes were blocked with 5% BSA in TBS for 2 h. Anti-CXCR4 antibody (E-Bioscience, San Diego, CA) was incubated overnight at 1:1,000 dilution at 4°C in blocking buffer supplemented with 0.1% polyoxyethylenesorbitan monolaurate (Tween-20). The remaining washes and incubations were performed at room temperature in TBS containing 0.1% milk and 0.1% Tween-20. Membranes were washed three times for 5 min and allowed to incubate with the peroxidase secondary anti-rabbit antibody for 2 h. After three 10-min washes, membranes were rinsed for 10 min in TBS alone. Chemiluminesence was preformed using ECL detection reagents (Amersham, Piscataway, NJ) according to the manufacturer's directions. Various exposures to Hyblot film (Denville Scientific, Metuchen, NJ) were performed to insure exposure linearity. Films were scanned and quantified using Image J (National Institutes of Health [NIH], Bethesda, MD). Densitometric values were reported as means ± SE. Three isolations from each mouse strain were performed, and samples were run in triplicate.

Rap1 activation was measured using a kit from Stressgen (Ann Arbor, MI) according to the manufacturer's directions. SDF-1 was added to C57BL/6J or NOD/LtJ T-cells at a concentration of 100 ng/ml and allowed to react for 5 or 15 min. Controls were performed according to the manufacturer's directions. The Rap1 activation kit used to detect active GTP-Rap1 uses a glutathione S-transferase (GST)-fusion protein containing the Rap1 binding domain to affinity purify active Rap1 from cell lysates. The fusion protein is incubated with the cell lysate and captured using a glutathione-conjugated filter disc. The affinity-purified product is detected by Western blot analysis using anti-Rap1 antibody. Affinity purification of active Rap1 was performed using 500 μg total protein from each sample. Approximately 15 × 10⁶ CD3⁺ cells from either C57BL/6J or NOD/LtJ mice for each sample were rinsed in TBS and lysed at 4°C using the Tris-based kit buffer supplemented with 0.1 μmol/l leupeptin, 0.3 μmol/l aprotinin, and 1 μmol/l PMSF. Samples were passed over resin discs containing the GST-fusion capture peptide; bound protein was eluted using sample buffer (0.060 mol/l Tris-HCL, pH 6.8, 0.1% SDS, 2% glycerol, 5% β-mercaptoethanol, and 0.05% bromphenol blue) by heating to 95°C for 3 min and loaded on 12% polyacrylamide SDS gels; and Western analysis was performed. Gels were transferred to nitrocellulose membranes and blocked for 2 h in 5% BSA in TBS. Rap1 antibody (1:1,000) was incubated overnight at 4°C in 5% BSA and 0.1% Tween-20 in TBS and then washed three times for 5 min in TBS containing 0.1% milk and 0.1% Tween. A peroxidase-conjugated secondary antibody, anti-rabbit IgG (1:2000), was allowed to react for 1 h at room temperature in wash buffer. Membranes were again washed three times for 5 min in the same buffer followed by a 10-min wash in TBS alone. Chemiluminesence was preformed using Amersham ECL detection reagents according to manufacturer's directions. Various exposures to Hyblot film (Denville Scientific) were performed to insure exposure linearity. Films were scanned and quantified using Image J (NIH). Densitometric values were reported as means ± SE.

Measurement of T-cell and endothelial cell surface adhesion molecule expression was performed by flow cytometry as we have previously reported (20,22,24). Antibodies and the dilutions used against these molecules are as follows: CD18 (1:320), CD29 (1:320), CD11a (1:80), CD49d (1:80), intracellular adhesion molecule-1 (ICAM-1) (1:80), vascular cell adhesion molecule (VCAM-1) (1:80), E-selectin (1:320), and P-selectin (1:320) and isotype controls at the respective dilutions. Cultured MS1 cell lines were harvested and washed in 10 ml fluorescence-activated cell sorting (FACS) buffer (PBS plus 1% fetal bovine serum). An aliquot of 5 × 10⁵ cells was used for adhesion molecule analysis. All cells were preincubated on ice for 20 min with 50 μl 1:100 anti–FC-receptor antibody to block nonspecific binding. The above-mentioned diluted antibodies were then added to the cells and incubated on ice for 20 min. The cells were washed twice with 1 ml FACS buffer and resuspended in 300 μl FACS buffer. Immunofluorescence-stained samples were analyzed on a FACS Calibur flow cytometer (Becton Dickinson) made available through the Research Core Facility at Louisiana State University Health Sciences Center. Data analysis was performed using CELL Quest software (Becton Dickinson). The cells were gated based on forward versus side scatter, and 10,000 events were collected.

Shear-mediated detachment assays were performed to determine changes in T-cell integrin affinity and avidity necessary for T-cell firm adhesion (25). Briefly, NOD/LtJ T-cells were introduced into the flow chamber and allowed to adhere to endothelial monolayers for 15 min under static conditions. After the incubation period, flow was resumed at 0.15 dyne/cm² using cell-free perfusate, and the shear stress was doubled every 30 s using a programmable syringe pump (0.15–38.4 dynes/cm²). The percentage of cells remaining adherent was determined by counting the number of adherent cells remaining on completion of the respective shear stress interval and dividing that number by the total number of initially adherent cells. The number of cells remaining adherent during a given shear stress interval reflects the degree of adhesion molecule affinity and/or avidity.

Data were statistically compared using Prism 4.0 software (GraphPad). The number of firmly adherent cells was compared using a one-way ANOVA with Bonferroni's post hoc test to determine statistical differences between experimental groups. Firm adhesion data are reported as the mean and SE. Rolling velocity data from 1,200 cells per treatment group were compared using a Kruskal-Wallis nonparametric ANOVA with a Dunn's post hoc test to determine statistical differences between experimental groups. Rolling velocity data are reported as a bar graph illustrating the mean rolling velocity and SE and a relative frequency histogram distribution identifying cell populations rolling at various velocity intervals. The percentage of remaining firmly adherent cells during the detachment assay was compared by one-way ANOVA with Bonferroni's post hoc test between experimental groups at each shear interval. Densitometric values for CXCR4 protein expression between C57BL/6J and NOD/LtJ mice were compared using an unpaired Student's t test. Rap1 activity assays were compared using a one-way ANOVA with Bonferroni's post hoc test to determine differences between time points. A P value of <0.05 was required to achieve statistical significance for all experimental procedures.

Previous reports demonstrate that SDF-1 facilitates firm adhesion of normal T-cells under hydrodynamic flow conditions (16,17). However, the effect of SDF-1 on diabetogenic or autoimmune T-cell firm adhesion is not known. Figure 1A illustrates the experimental design used to investigate the effect of SDF-1 on NOD/LtJ T-cell firm adhesion to TNF-α–activated mouse islet microvascular endothelial cells under hydrodynamic flow. Mouse T-cells were drawn across unstimulated, 10 ng/ml TNF-α–stimulated, or 10 ng/ml TNF-α–stimulated and SDF-1–coated islet microvascular endothelial cell monolayers using a hydrodynamic parallel plate flow chamber to measure T-cell–endothelial cell biophysical interactions (22). Figure 1B shows biophysical data of diabetogenic NOD/LtJ CD3 T-cell firm adhesion to control, TNF-α–stimulated, or TNF-α–stimulated plus SDF-1–coated (1, 10, 50, and 100 ng/ml) islet endothelial cells in a dose-response fashion. NOD/LtJ CD3 T-cell firm adhesion to TNF-α–stimulated islet endothelium was significantly increased under hydrodynamic flow conditions compared with unstimulated islet endothelial monolayers. Surprisingly, NOD/LtJ T-cell firm adhesion to TNF-α–activated endothelium was significantly inhibited by surface coating of SDF-1 in a dose-dependent manner, demonstrating chemorepulsion of T-cell firm adhesion. Complete inhibition of TNF-α–dependent firm adhesion was accomplished using a dose of 100 ng/ml SDF-1. Importantly, 100 ng/ml SDF-1 had no effect on NOD/LtJ T-cell firm adhesion to unstimulated endothelium.

SDF-1 binds to its primary receptor CXCR4 and signals through a pertussis toxin–sensitive G_iα-protein–coupled pathway (26,27). Therefore, we examined whether pertussis toxin pretreatment of NOD/LtJ CD3 T-cells could prevent SDF-1–mediated chemorepulsion of T-cell adhesion to TNF-α–activated endothelial monolayers. Figure 1B shows that pretreatment of NOD/LtJ CD3 T-cells with 1 μg/ml pertussis toxin for 30 min completely restores adhesion to TNF-α–activated endothelium coated with 100 ng/ml SDF-1.

By way of comparison, we then performed C57BL/6J CD3 T-cell firm adhesion studies to various islet endothelial monolayers. Figure 1C demonstrates that TNF-α increases C57BL/6J CD3 T-cell firm adhesion and that 50 ng/ml SDF-1 further enhances firm adhesion consistent with previous reports (15,17,28). However, it has been shown that low doses (5–10 nmol/l) of SDF-1 stimulate T-cell chemotaxis, whereas a high dose of SDF-1 (100 nmol/l) stimulates chemorepulsion of T-cell migration (26). Therefore, we examined whether 100 nmol/l (800 ng/ml) SDF-1 might also stimulate chemorepulsion of C57BL/6J CD3 T-cell firm adhesion to TNF-α–activated monolayers. Figure 1C illustrates that 800 ng/ml SDF-1 significantly prevented C57BL/6J CD3 T-cell firm adhesion to TNF-α–activated islet endothelium, which was reversed by pretreating C57BL/6J T-cells with pertussis toxin.

We further examined whether SDF-1 stimulated chemorepulsion of different NOD/LtJ T-cell populations. Figure 1D and E illustrates that SDF-1 treatment resulted in chemorepulsion of firm adhesion of both CD4 and CD8 T-cells from NOD/LtJ mice. Importantly, pretreatment of these T-cell populations with pertussis toxin prevented SDF-1 chemorepulsion of firm adhesion, reiterating a role for G_iα-protein–coupled signaling.

The process of leukocyte recruitment is a multistep event in which leukocyte capture and rolling are necessary to initiate firm adhesion. Moreover, SDF-1 has been reported to destabilize L-selectin–dependent T-cell rolling independent of shedding, suggesting that SDF-1 could alter rolling behavior (29). Therefore, we examined whether SDF-1 chemorepulsion of firm adhesion involved defective T-cell rolling. Figure 2 reports biophysical rolling data comparing C57BL/6J and NOD/LtJ CD3 T-cell–rolling interactions under hydrodynamic flow conditions with unstimulated, TNF-α–stimulated, or TNF-α–stimulated plus 100 ng/ml SDF-1–coated mouse pancreatic islet endothelium. Figure 2A illustrates the number of T-cells rolling on the various islet microvascular endothelial cell cultures. TNF-α stimulation significantly increased the number of rolling C57BL/6J and NOD/LtJ CD3 T-cells. Interestingly, SDF-1 treatment further augmented the number of rolling NOD/LtJ CD3 T-cells but not C57BL/6J T-cells. Figure 2B illustrates the average rolling velocity of CD3 T-cells. Both C57BL/6J and NOD/LtJ CD3 T-cells rolled significantly slower on TNF-α–stimulated or TNF-α–stimulated plus SDF-1–treated monolayers compared with unstimulated monolayers. These data clearly demonstrate that SDF-1 does not antagonize NOD/LtJ CD3 T-cell rolling.

The process of leukocyte slow rolling is important in facilitating cell signaling, which results in cellular activation and conversion to firm adhesion (22,30). Thus we determined the population frequency of CD3 T-cells rolling at various velocity intervals. Figure 2C and D report rolling velocity histograms for C57BL/6J and NOD/LtJ CD3 T-cells, respectively. A similar fraction of CD3 T-cells from both C57BL/6J and NOD/LtJ mice rolled at slow velocities (<50 μm/s), which are necessary for conversion to firm adhesion. In all of the rolling parameters analyzed, NOD/LtJ CD3 T-cells showed an equivalent response to C57BL/6J CD3 T-cells, indicating that rolling responses are not defective in response to SDF-1.

We next examined the role of CXCR4 in modulating SDF-1 chemorepulsive effects on NOD/LtJ CD3 T-cells. Figure 3A shows CXCR4 protein expression between C57BL/6J and NOD/LtJ T-cells. Interestingly, NOD/LtJ CD3 T-cells showed a significant 60% reduction of CXCR4 protein expression compared with C57BL/6J T-cells. Figure 3B reports the effect of anti-CXCR4 antibody (10 μg/ml) blockade on SDF-1 chemorepulsion. Inhibition of SDF-1/CXCR4 binding conferred a significant yet partial 43 ± 2% reversal of SDF-1 chemorepulsion, suggesting involvement of additional pathways. Given the significantly reduced CXCR4 expression and its partial involvement in mediating SDF-1 chemorepulsion, we next examined whether SDF-1–dependent Rap1 activation was also altered between C57BL/6J versus NOD/LtJ T-cells. SDF-1/CXCR4 interactions result in activation of Rap1 kinase activity, which alters adhesion molecule functions (31,32,33). Figure 3C reports Rap1 kinase activity from C57BL/6J or NOD/LtJ T-cells stimulated with 100 ng/ml SDF-1 at 0, 5, and 15 min. SDF-1 treatment stimulated a rapid, transient increase in C57BL/6J Rap1 activity that is consistent with previous studies (32,33). Conversely, Rap1 activation by SDF-1 was significantly delayed in NOD/LtJ T-cells, and basal Rap1 activity was distinctly absent in NOD/LtJ T-cells compared with C57BL/6J. Together, these data highlight a significant difference in NOD/LtJ CXCR4 expression and function in response to SDF-1, which suggests that other signaling pathways likely contribute to SDF-1 chemorepulsion of NOD/LtJ T-cells.

Proper surface expression of endothelial cell adhesion molecules is an obligatory requirement for T-cell firm adhesion (34). Therefore, we next examined whether 100 ng/ml SDF-1 treatments altered endothelial cell surface adhesion molecule expression. Flow cytometric analysis was performed on nonstimulated, TNF-α–stimulated, and TNF-α–stimulated plus SDF-1–coated mouse pancreatic islet endothelial cells to measure the surface expression of endothelial cell adhesion molecules P-selectin, E-selectin, ICAM-1, and VCAM-1 (Fig. 4). Figure 4 shows that low basal levels of constitutive P-selectin, ICAM-1, and VCAM-1 expression were observed on unstimulated endothelium. Expression of P-selectin, E-selectin, ICAM-1, and VCAM-1 were all significantly upregulated on stimulation with 10 ng/ml TNF-α. Importantly, treatment with SDF-1 did not significantly alter TNF-α induction of increased adhesion molecule expression, and SDF-1 treatment alone did not affect resting adhesion molecule expression.

VLA-4 (CD49d/CD29) and LFA-1 (CD11a/CD18) are the primary T-cell integrin adhesion molecules that mediate firm adhesion to endothelial cells (35). Therefore, we next examined whether 100 ng/ml SDF-1 altered the surface expression of these adhesion molecules. Figure 5 reports surface expression staining by flow cytometry analysis for CD29 and CD49d on either C57BL/6J or NOD/LtJ CD3 T-cells. Stimulation with 100 ng/ml SDF-1 for either 5 or 15 min did not significantly change C57BL/6J or NOD/LtJ CD49d/CD29 surface expression. Likewise, Fig. 6 demonstrates that SDF-1 treatment also did not alter C57BL/6J or NOD/LtJ CD3 T-cell surface expression of CD11a/CD18 over the 15-min experimental period.

Data presented thus far demonstrate that SDF-1 treatment is chemorepulsive for NOD/LtJ CD3 T-cell firm adhesion to activated islet endothelium that does not involve alterations in cell capture, rolling, or surface adhesion molecule expression. An additional layer of regulation of firm adhesion is accomplished by altering integrin affinity and/or avidity for its counter-ligand. SDF-1 has been reported to stimulate changes in VLA-4 and LFA-1 affinity and avidity, which are important for firm adhesion by mediating resistance against increasing shear-mediated detachment (17,28,36). Therefore, we performed shear-mediated detachment assays to examine whether SDF-1 stimulates chemorepulsion of firm adhesion by altering changes in integrin affinity or avidity. NOD/LtJ CD3 T-cells were perfused onto various endothelial cell monolayers and allowed to adhere under static conditions for 15 min. Hydrodynamic flow was then reestablished and doubled every 30 s to determine the resistance of NOD/LtJ CD3 T-cell adhesion to increasing shear, which is directly proportional to degree of integrin affinity and avidity. Figure 7A illustrates that NOD/LtJ CD3 T-cells readily detach from unstimulated islet endothelial cells, with <10% remaining adherent at a shear stress of 1.5 dynes/cm². Conversely, NOD/LtJ CD3 T-cells adherent to TNF-α–stimulated islet endothelium were significantly resistant to shear-mediated detachment, with ∼40% of the population still adherent at a shear stress of 38.4 dynes/cm². SDF-1 treatment (100 ng/ml) completely inhibited NOD/LtJ CD3 T-cell resistance to shear-mediated detachment on TNF-α–activated islet endothelium, with <10% remaining adherent at 0.6 dyne/cm². Importantly, pretreatment of NOD/LtJ CD3 T-cells with pertussis toxin completely reversed SDF-1–dependent loss of shear resistance.

The above results indicate that NOD/LtJ integrin affinity or avidity is diminished by SDF-1, which suggests that integrin activity could be constitutively greater in NOD/LtJ CD3 T-cells. Therefore, we measured the absolute numbers of C57BL/6J or NOD/LtJ firmly adherent CD3 T-cells to control or TNF-α–stimulated islet endothelial monolayers. Figure 7B illustrates that NOD/LtJ CD3 T-cells show a significantly greater number of firmly adherent cells under both control and TNF-α–stimulated conditions compared with C57BL/6J CD3 T-cells. These results demonstrate that integrin affinity and avidity is significantly greater in NOD/LtJ CD3 T-cells, consistent with the results from shear-mediated detachment experiments.

Recent reports have shown that SDF-1 genetic polymorphisms are associated with early onset of autoimmune diabetes (18,19). Interestingly, some studies suggest that this may not be true for all ethnic backgrounds or in disease conditions involving other autoimmune disorders (37,38). Nonetheless, a recent study reported that genetic polymorphisms of SDF-1 result in decreased protein expression in lymphoblasts on activation using allele-specific transcript quantification methods (39). Consistent with these findings, a recent study suggested that SDF-1/CXCR4 signaling protects against autoimmune diabetes in NOD mice as inhibition of CXCR4 activity exacerbates adoptive transfer of diabetes (40). Together, these observations clearly implicate SDF-1 in regulating the development of autoimmune diabetes; however, the manner in which this could occur is not known. Our data corroborate the importance of SDF-1 in controlling diabetogenic T-cell function and provide a potential mechanistic explanation for how SDF-1 may be critical in regulating diabetogenic T-cell entry into pancreatic islets.

The process of leukocyte recruitment is essential for initiating and sustaining inflammatory states within tissues. As such, the ability to recruit a leukocyte into a particular tissue niche is regulated at several levels, with chemokines serving important roles in directing cell-cell adhesion and chemotaxis responses (12). It has been reported that genetic mutations or increased expression of various chemokines facilitate the development of autoimmune diabetes in NOD mice and patients (41–45). However, the specific cellular responses and mechanisms of chemokine regulation of diabetogenic T-cell recruitment are completely unknown. Moreover, nothing is known regarding the manner in which SDF-1 affects diabetogenic versus normal T-cell homing and adhesion, highlighting a key deficiency in our understanding of autoimmune T-cell recruitment. In this study, we have revealed that SDF-1 mediates a novel chemorepulsive effect on diabetogenic T-cell firm adhesion compared with enhancing normal T-cell firm adhesion. This finding is in stark contrast to all previous studies investigating the effect of SDF-1 on T-cell recruitment, which show that SDF-1 augments T-cell adhesion to recombinant adhesion molecules and endothelial monolayers (16,17,46).

How then might SDF-1 stimulate chemorepulsion of NOD/LtJ T-cell firm adhesion? One explanation could be related to the doses of SDF-1 previously used because we report here for the first time that SDF-1 can prevent normal T-cell firm adhesion at a concentration known to stimulate chemorepulsion of T-cell migration (800 ng/ml) (26). However, several lines of data support the hypothesis that SDF-1 may be working through alternative receptor-signaling pathways besides CXCR4. First, CXCR4 protein expression is significantly decreased in NOD/LtJ T-cells. Second, immuno-neutralization of CXCR4 partially attenuates SDF-1 chemorepulsion, whereas G_iα-protein inhibition by pertussis toxin completely reverses chemorepulsion. As such, SDF-1 signaling responses between normal and diabetogenic T-cell populations appear to have a common point of origin, G_iα-protein, but show divergent activation of downstream Rap1 pathways. This alternative receptor could be RDC1/CXCR7, which avidly binds SDF-1 and signals through a G-protein–coupled receptor pathway (47,48). Thus, it is possible that downstream SDF-1 signaling pathways mediated by CXCR4 versus RDC1/CXCR7 may be different between diabetogenic versus normal T-cells and that the activity between the two may differentially control for chemoattraction versus chemorepulsion. It is also possible that a co-repressor signal from the endothelium may also be involved. Interestingly, preliminary experiments examining the effect of NOD T-cell detachment on recombinant VCAM-1 ± SDF-1 coating showed that SDF-1 augments NOD T-cell resistance to shear-mediated detachment compared with islet microvascular endothelial cell monolayers (M. Hueng, C.G. Kevil, unpublished observations). Such a co-repressor function of the endothelium is possible because the Slit-2/Robo system has been shown to counteract SDF-1/CXCR4 function and to regulate leukocyte chemotaxis (49,50). Future experiments are necessary to determine whether RDC1/CXCR7 signaling or Slit/Robo interactions participate in SDF-1–mediated chemorepulsion of NOD/LtJ T-cell firm adhesion.

The remaining question is how does SDF-1 mediate chemorepulsion of diabetogenic versus normal T-cell firm adhesion? We have presented data demonstrating that SDF-1 treatment renders diabetogenic T-cells vulnerable to shear-mediated detachment that is due to changes in integrin affinity and avidity. This result suggests that integrin affinity or avidity is likely to be constitutively greater on diabetogenic versus normal T-cells. Support for this idea is strong because the absolute numbers of adherent diabetogenic T-cells was greater on either unstimulated or TNF-α–stimulated islet endothelial cells compared with normal T-cells. These findings are striking for two reasons. First, these data demonstrate that NOD/LtJ T-cell surface integrins are activated to a greater degree, thereby increasing the likelihood that these cells will readily adhere and migrate into islets. Second, our data identify a previously unknown response that integrin affinity and avidity is reversible by SDF-1. Figure 7C illustrates our working hypothesis in which integrin activation is constitutively greater in NOD/LtJ T-cells, which can be reversed by SDF-1 treatment. It is possible that other factors could influence integrin activation, such as differences in binding pocket structure or subunit assembly, which cannot be dismissed. However, specific questions that need to be answered in the future are 1) whether SDF-1 treatment specifically affects integrin affinity or avidity or both and 2) which of the NOD/LtJ T-cell integrin proteins are primarily affected by SDF-1 treatment.

In conclusion, we provide novel evidence that SDF-1 exerts a diverse response in regulating diabetogenic immune cell recruitment versus nonautoimmune cells. Here, we show that diabetogenic T-cell integrin activity is constitutively elevated in NOD/LtJ T-cells, which can be inactivated by SDF-1 that could serve to diminish the autoimmune cell recruitment. This may appear counterintuitive because several reports suggest that several chemokines and their receptors (e.g., IP-10, RANTES, CXCR3, CCR7, CCR5, etc.) may contribute to the disease process of type 1 diabetes (41,42). However, our understanding of chemokine regulation of immune cell trafficking during normal and inflammatory states is constantly evolving with the identification of novel functions and new molecules (47,48,51). Future studies are clearly needed to better understand the unique findings above and how chemokines may differentially regulate immune cell trafficking during autoimmune diabetes.

C.G.K. has received American Diabetes Association Award 1-05-JF-26.