Glucocorticoids Suppress CCR9-Mediated Chemotaxis, Calcium Flux, and Adhesion to MAdCAM-1 in Human T Cells

The chemokine CCL25 is constitutively expressed in the small intestine and interacts with a single signaling receptor, CCR9 (1, 2). In humans, CCR9 is expressed on most (58–97%) T cells in the small intestine, ∼25% in the colon, and a small minority (3–5%) in peripheral blood (2, 3). CCR9⁺ T cells coexpress integrin α₄β₇, which binds to mucosal addressin cell adhesion molecule-1 (MAdCAM-1) (4). MAdCAM-1 is constitutively expressed on vascular endothelial cells in the small and large intestine, and together CCR9 and α₄β₇ preferentially mediate the adherence and migration of effector T cells into the intestine (5, 6).

Inflammatory bowel disease (IBD) is characterized by leukocyte infiltration, increased levels of proinflammatory cytokines in the intestine, and increased expression of MAdCAM-1 (7-10). In the peripheral blood, the frequency of circulating CCR9⁺ T cells is elevated during small intestinal Crohn’s disease, suggesting a role of CCR9⁺ T cells in small intestinal inflammation (11). CCR9, CCL25, α₄β₇, and MAdCAM-1 are attractive therapeutic targets in IBD because of their tissue and cell specificity, and clinical studies show positive effects on blocking these proteins in subsets of IBD patients (12–15).

Glucocorticoids (GCs) have broad-ranging anti-inflammatory actions, are highly effective in many chronic inflammatory disorders, and have been the mainstay of IBD treatment since the 1950s (16, 17). The anti-inflammatory effects of GCs are mainly mediated through interaction with cytosolic GC receptors (GCRs) to modify the transcription of proinflammatory genes (18, 19). Previously characterized effects of GCs on T cells include inhibiting TCR signaling, suppressing cell proliferation, inducing IκB synthesis, and subsequently reducing NF-κB–mediated transcription of proinflammatory cytokines (19–21). However, little is known about the effects of GCs on intestine-specific cell recruitment. We investigated the effect of GCs on CCR9 expression and function, as well as downstream α₄β₇ activation in primary human T cells, in vitro.

In this study we demonstrate that CCR9-mediated chemotaxis, intracellular calcium flux, and adhesion to MAdCAM-1 are significantly suppressed following corticosteroid treatment, and this is mediated through activation of GCRs. As a control, we compared the effects of GCs on CXCR4, which is coexpressed on CCR9⁺ T cells. CXCR4 is stimulated by a single ligand, CXCL12, and consistent with previous reports, GCs increase CXCR4 expression and enhance CXCL12-mediated functions (22, 23). These results identify novel effects of GCs on the function of primary human T cells that are likely to be physiologically and clinically relevant in intestinal inflammation.

PBMCs were depleted of monocytes by CD14⁺ bead selection (Miltenyi Biotec, Surrey, U.K.). Remaining leukocytes were activated by culturing on plate-bound anti-CD3 (3 μg/ml) (BioLegend, Cambridge, U.K., clone OKT3) and anti-CD28 (3 μg/ml) (BioLegend, clone CD28.2) in complete culture medium (RPMI 1640 supplemented with 10% FCS, penicillin-streptomycin) with 100 U IL-2 (PeproTech, Rocky Hill, NJ) and 100 nM all-trans retinoic acid (ATRA). After 72 h, cells were removed from CD3/CD28 stimulation and resuspended in complete culture medium with IL-2 and ATRA. Cells were used for assays between days 6 and 9 from initial isolation.

Cells were resuspended in RPMI 1640 supplemented with 10% FCS, 50 U/ml penicillin, and 50 μg/ml streptomycin. Cells were treated with 250 nM dexamethasone (DEX), 1 μM prednisolone (PRED), or 1 μM deoxycorticosterone acetate (DOC) and incubated for 16 h at 37°C with 5% CO₂. For mifepristone (MIF) treatment, cells were treated for 1 h with 5 μM MIF prior to the addition of DEX.

Cells and chemokines were resuspended in chemotaxis buffer (RPMI 1640 supplemented with 0.1% BSA and 10 mM HEPES [pH 7.40]). Transwell inserts (Corning Life Sciences, Corning, NY) containing 4 × 10⁵ live cells in 75 μl were placed in the wells of 96-well plates containing 235 μl chemotaxis buffer with or without chemokine (250 nM CCL25 or 10 nM CXCL12). All chemokines were sourced from PeproTech, and CCL25 was a gift from ChemoCentryx (Mountain View, CA). Cells and chemokines were incubated together for 2 h at 37°C in 5% CO₂. All assays were performed in a minimum of triplicate data points. Live migrated cells in the lower chamber were quantified by CellTiter-Glo (Promega, Southampton, U.K.) according to the manufacturer’s instructions, and verified by trypan blue staining. To ensure that equivalent numbers of live cells were loaded into chemotaxis assay inserts, cells were adjusted by trypan blue staining and by CellTiter-Glo measurements.

Cells were washed and labeled at 4°C for 30 min with Ab diluted in PBS with 0.1% BSA and 0.01% sodium azide. The following Abs were used: anti-CCR9 (R&D Systems, Abingdon, U.K., clone 248621), anti-CXCR4 PE (eBioscience, Hatfield, U.K., clone 12G5), anti-CD3 Alexa Fluor 700 or allophycocyanin-Cy7 (eBioscience, clone UCHT1), anti-CD4 FITC (eBioscience, clone OKT4), anti-CD8 eFluor 605NC (eBioscience, clone RPA-T8), and SYTOX Green (Life Technologies, Carlsbad, CA). To detect unconjugated anti-CCR9, labeled cells were washed and incubated with allophycocyanin-labeled anti-mouse IgG (BioLegend). Flow cytometry was performed with an LSR II or Fortessa (BD Biosciences, Franklin Lakes, NJ) and data were analyzed using FlowJo software version 9.3.3 (Tree Star, Ashland, OR).

Human PBMCs were isolated by Ficoll (GE Healthcare, Little Chalfont, U.K.) density gradient centrifugation and T cells were isolated by negative bead selection (pan T cell isolation kit, Miltenyi Biotec). PBMCs or ATRA-cultured cells were labeled with monoclonal anti-CCR9 Ab (R&D Systems, clone 248621) followed by anti-mouse allophycocyanin secondary Ab (BioLegend). Cells were washed and labeled with anti-allophycocyanin beads (Miltenyi Biotec) and isolated by positive selection according to the manufacturer’s instructions.

RNA was isolated using the RNeasy mini kit and reversed transcribed to cDNA using SuperScript III and oligo(dt) primers (Invitrogen). Quantitative PCR was performed with primers specific for CCR9 (forward, 5′-GCTGCCTGCTCAGAACCCACA-3′, reverse, 5′-GTGGGTGTCATGGTGGGTCAG-3′), CXCR4 (forward, 5′-CGCTACCTGGCCATCGTCC-3′, reverse, 5′-GAGGGCAGGGATCCAGACG-3′), CCRL1 (RT² qPCR Primer Assay for human CCRL1; Qiagen), and GAPDH (forward, 5′-GGGGCTGGCATTGCCCTCAA-3′, reverse, 5′-TTGCTGGGGCTGGTGGTCCA-3′) using SYBR Green JumpStart Taq ReadyMix (Sigma-Aldrich, Gillingham, U.K.). Samples were run in triplicate on a Rotor-Gene 3000 (Qiagen) with the following conditions: 95°C for 15 min and second phase of 40 cycles at 94°C for 15 s, 60°C for 30 s, and 72°C for 30 s. Melt curve analysis was performed on all PCR products. Gene expression for each sample was normalized to GAPDH.

Vehicle (H₂O) and DEX-treated cells were separately labeled with anti-CCR9 (R&D Systems, clone 248621) followed by anti-mouse IgG conjugated to allophycocyanin in PBS with 0.1% BSA. Vehicle-treated cells were labeled with anti-CD3 Alexa Fluor 700 and DEX-treated cells with anti-CD3 allophycocyanin–eFluor 780 (eBioscience, clone UCHT1). An equivalent number of vehicle- and DEX-treated cells were combined in HBSS with 0.02% Pluronic F-127, 1 μg/ml Fura Red AM (Life Technologies), and incubated for 30 min at 37°C. Cells were resuspended in HBSS with 0.1% BSA, 1 mM CaCl₂, 0.5 mM MgCl₂, and 10 mM HEPES with Live/Dead cell marker, SYTOX Green (Life Technologies), and incubated at 37°C until flow cytometry was performed. Data were acquired on an LSR II SORP and analysis was performed in FlowJo. The gating strategy was as follows: live cells (SYTOX Green⁻), excluding cell doublets (determined by forward light scatter area versus forward light scatter height), CCR9-allophycocyanin⁺ cells, and, finally, vehicle- and DEX-treated cells, distinguished by their distinct anti-CD3 fluorochrome. Fura Red was excited by both violet (406 nm, 25 mW) and green (532 nm, 150 mW) lasers, and the emission was detected using 630LP and 660/20BP and 685LP and 710/50BP filters sets, respectively. The ratiometric “Fura Red ratio” was calculated as the increasing signal stimulated by the violet laser over the decreasing signal stimulated by the green laser using the kinetics application in FlowJo. Baseline calcium flux was recorded for 25 s before cells (180 μl) were transferred to an aliquot of stimulant (20 μl chemokine or ionomycin at 10-fold the desired final concentration). The following final concentrations were tested: 125 nM CCL25, 10 nM CXCL12, and 5 μg/ml ionomycin. Recording was continuous and 8,000–10,000 total events/s were recorded for 120 s in total.

Lyophilized recombinant human MAdCAM-1 and VCAM-1 (R&D Systems) were resuspended in PBS and stored in aliquots at −80°C. Polystyrene 96-well, flat-bottom, tissue culture–treated plates (Corning, Sigma-Aldrich) were coated with adhesion molecules by incubating overnight at 4°C with 0.15 μg/ml MAdCAM-1 or VCAM-1 diluted in PBS (50 μl/well). Wells were washed twice with PBS, and nonspecific binding sites were blocked with 1% BSA in PBS for 1 h at 37°C followed by an additional three washes with room temperature assay buffer (RPMI 1640 supplemented with 0.1% BSA and 10 mM HEPES). Chemokine was diluted in assay buffer to 2-fold the final desired concentration and added to appropriate wells (30 μl). The following final concentrations were tested: 500 nM CCL25, 10 nM CXCL12, and 10 mM MnCl₂. Cells were resuspended in room temperature assay buffer to a density of 2 × 10⁶ cells/ml and added (30 μl) gently but quickly to the plate using a multichannel pipette. The plate was centrifuged for 15 s and immediately transferred to a 37°C incubator for 2 min. Nonadherent cells were removed by washing five times with room temperature assay buffer. Adherent cells were quantified using CellTiter-Glo (Promega) and visualized by phase-contrast microscopy (×10 magnification, captured with a Nikon Eclipse TS100 microscope, and photographed with a Nikon Coolpix 995).

Chemicals and reagents were supplied by Sigma-Aldrich unless otherwise specified, and reconstituted according to the manufacturer’s recommendations. Aliquots of DEX, PRED, DOC, and MIF were stored in frozen aliquots at −20°C.

Statistical analysis was performed using the Prism program (GraphPad Software, San Diego, CA).

CCR9⁺ T cells in the peripheral blood are a minority population (2–5% of CD3⁺ cells). To obtain a sufficient number of CCR9⁺ cells while avoiding the use of immortalized cell lines, we cultured PBLs in the presence of ATRA, which induces CCR9 and β₇ integrin expression (6, 24, 25). This increased the proportion of CCR9⁺ T cells to 25–70% of the total. In addition to surface CCR9, most ATRA-cultured cells expressed CXCR3, CXCR4, and β₇ integrin, a minority expressed CCR2 or CCR6 (<20%), and the cells were uniformly negative for CCR3, CCR5, and CCR10 (Supplemental Fig. 1A).

By Transwell chemotaxis, ATRA-cultured cells migrated to increasing concentrations of soluble, recombinant CCL25 (via CCR9), CXCL11 (via CXCR3), and CXCL12 (via CXCR4) at concentrations comparable to previous reports (Supplemental Fig. 1B) (1, 26, 27). Consistent migration to CCL20 (via CCR6) or to CCL5 (via CCR5) could not be measured (data not shown), likely owing to low receptor expression.

To study the effect of corticosteroids on CCR9 function, ATRA-cultured cells were treated for 16 h in the presence of vehicle (H₂O) or 250 nM DEX, a synthetic corticosteroid with pure GCR activity. The following day, Transwell chemotaxis to CCL25 (via CCR9) was measured. For comparison, chemotaxis to CXCL11 (via CXCR3) and CXCL12 (via CXCR4) was also measured. A consistent concentration of chemokine to induce cell migration between 75 and 90% of the maximal signal was used: 250 nM CCL25, 90 nM CXCL11, and 10 nM CXCL12 (refer to Supplemental Fig. 1B). DEX treatment reduced the proportion of viable cells (32 ± 3% reduction, n = 3), as measured by trypan blue and CellTiter-Glo, and this was corrected for by adjusting the chemotaxis assay so that equal numbers of live cells were assayed for each condition.

Between independent experiments, the proportion of cells that migrated was variable (Fig. 1A); however, DEX treatment consistently reduced CCL25-mediated chemotaxis (48 ± 8% reduction, p < 0.01), reduced CXCL11-mediated chemotaxis (24 ± 6% reduction, p < 0.05), and increased CXCL12-mediated chemotaxis (53 ± 21% increase, p < 0.05) (Fig. 1A, 1B).

To determine whether changes in directed cell migration reflected changes to chemokine receptor expression, levels of surface protein were measured by flow cytometry. Following 16 h of DEX treatment, surface expression of CCR9, as measured by mean fluorescence intensity (MFI), was equivalent between vehicle- and DEX-treated cells (Fig. 1C, 1D). In contrast, surface expression of CXCR3 was reduced whereas surface expression of CXCR4 increased following DEX treatment (Fig. 1C, 1D).

The effects of DEX on CXCR4 surface expression and cell migration have been previously described in other cell types; however, to our knowledge, these are the first reports of suppressive effects on CCR9- and CXCR3-mediated cell migration (22, 28). Although the effects of DEX on CXCR3 were novel, our main interest was intestinal-specific homing, and therefore we continued to study CCR9 using CXCR4 as a positive control.

In human T cells and eosinophils, DEX-enhanced migration through CXCR4 is dose-dependent at concentrations >10 nM (22, 29). To determine whether differences in CCR9 and CXCR4 functions were due to the effect of GC concentrations, two lower doses of DEX were tested in our system. Following 25 and 125 nM DEX, CCR9-mediated migration was equivalent or suppressed, respectively, whereas CXCR4 function was either equivalent or increased relative to untreated cells (Supplemental Fig. 2A–C). These data demonstrate that the effects of DEX on CCR9 and CXCR4 are dose-dependent.

CCR9 is the single identified signaling receptor for CCL25; CCL25 also binds the nonsignaling, decoy receptor, CCRL1 (30). To determine whether suppressed CCR9 function was the consequence of sequestering of ligand by CCRL1, mRNA levels were measured (CCRL1-specific Abs for detection by flow cytometry were not available at the time of this work). CCRL1 mRNA was detected in CCR9-expressing ATRA cells (p < 0.001) and in small intestinal tissue homogenate, which has been previously described (Supplemental Fig. 2D) (31). By quantitative PCR, DEX increased CXCR4 mRNA expression (p < 0.05) and had no effect on levels of CCR9 or CCRL1 mRNA (Supplemental Fig. 2E). Therefore, the suppressive effects of DEX on CCR9 function do not correspond to changes in CCR9 expression or to changes in mRNA levels of CCRL1.

To investigate whether the observed effects of DEX were mediated through activation of GC receptors, we studied the effects of a competitive antagonist of steroid receptors, MIF in the presence of DEX, a pure GCR agonist (32). ATRA-cultured cells were treated with DEX in the presence or absence of MIF for 16 h.

DEX did not affect CCR9 expression in ATRA-cultured cells, and neither did MIF or DEX plus MIF (Fig. 2A, 2B). In contrast, although MIF alone did not affect CXCR4 expression, it abrogated the increased expression of CXCR4 induced by DEX (Fig. 2A, 2B). By itself, MIF did not affect chemotaxis to either CCL25 or CXCL12 (Fig. 2C). MIF in combination with DEX completely reversed the effect of DEX on CCR9- and CXCR4-mediated chemotaxis (Fig. 2C). These results are in accord with reports that DEX increases CXCR4 surface expression and CXCL12-mediated chemotaxis through activation of the GC receptor (22, 23). Furthermore, these results demonstrate that the effect of DEX on CCR9-mediated chemotaxis is also GC receptor mediated.

DEX is routinely used in experimental studies, both in vitro and in vivo, because it is a pure GC with potent anti-inflammatory activity (19, 32). However, many clinically prescribed steroids have mixed steroid receptor activity; PRED, for example, is routinely prescribed in IBD and has mixed GC and mineralocorticoid receptor agonist activity. Agonists of the mineralocorticoid receptor do not produce anti-inflammatory effects although they have a critical role in sodium and water homeostasis. To test whether the effects of GC activation on CCR9-mediated migration were shared by another clinically relevant GC, the effects of PRED were assessed. As a comparison, the effects of DOC, a corticosteroid with pure mineralocorticoid activity, was also compared (32).

ATRA-cultured cells were treated with a physiologically relevant dose of DEX, PRED, or DOC for 16 h. Following steroid treatment, the proportion of cells expressing CCR9 was equivalent to vehicle-treated cells (Fig. 3A), and the amount of surface CCR9 (MFI measured by flow cytometry) was also equivalent between treatment groups (Fig. 3B). Similar to the effect of DEX on CCR9-mediated chemotaxis, PRED reduced CCL25-stimulated migration (p < 0.01; Fig. 3C). DOC, a pure mineralocorticoid, had no effect on CCL25-stimulated migration (Fig. 3C). DEX and PRED treatment both increased CXCR4 surface expression whereas DOC had no effect (Supplemental Fig. 3A, 3B). DEX and PRED enhanced CXCL12-mediated migration, whereas migration was equivalent between vehicle and DOC-treated cells, indicating that steroid-treated cells were still capable of chemotaxis through coexpressed CXCR4 (Supplemental Fig. 3C). These data further support the observation that activation of GCRs, and not mineralocorticoid receptors, impairs CCR9 function in human T cells.

GCs predominantly affect cell function through transcriptional modifications; however, rapid, transcription-independent mechanisms have also been described, such as steric modification of cell surface protein interactions (33–35). To examine rapid, nontranscriptional modifications, the effect of 1 h DEX treatment was examined. Cells treated with vehicle or DEX for 1 h had equivalent CCR9 and CXCR4 surface expression and equivalent migration to CCL25 and CXCL12 (data not shown). Therefore, the observed effect of GCs on CCR9 and CXCR4 function are unlikely to be due to steric hindrance of ligand binding.

To test whether DEX treatment reduced CCR9-mediated chemotaxis in primary human CCR9⁺ T cell, cells were sorted from PBMCs by magnetic bead separation. Approximately 3–5% of circulating T cells express CCR9 on the cell surface and, following sorting, the cell population comprised >80% CCR9⁺CD3⁺ T cells, which were predominantly CXCR4⁺ (Fig. 4A)

In primary human CCR9⁺ T cells, DEX treatment reduced the proportion of viable cells (15 ± 4% reduction, n = 3), although it did not alter the proportion of CCR9⁺ or CXCR4⁺ cells among live cells (Fig. 4A). As previously observed in ATRA-cultured cells, the density of surface CCR9 was equivalent between vehicle and DEX-treated cells, whereas CXCR4 expression was significantly elevated in the DEX treatment group (p < 0.05; Fig. 4B).

In the Transwell chemotaxis system, sorted CCR9⁺ T cells migrated efficiently to increasing concentrations of CCL25 with a characteristic bell-shaped dose-response curve, with maximum migration at 250 nM CCL25 (data not shown). These cells also displayed a robust migration response to 10 nM CXCL12. DEX treatment reduced migration of CCR9⁺ cells to CCL25 (p < 0.01), whereas migration to CXCL12 increased (p < 0.05; Fig. 4C). When corrected for background migration and normalized to vehicle-treated cells, DEX treatment reduced CCR9-mediated chemotaxis by 36 ± 4% (p < 0.05) and increased CXCR4-mediated chemotaxis by 38 ± 21% (Fig. 4D).

To determine whether DEX specifically affected chemotaxis of either CD4⁺ or CD8⁺ T cells, the proportion of both populations was examined before and after migration to chemokine. CCR9⁺-sorted T cells included both single-positive CD4 and CD8 T cells, and the ratio of CD4 to CD8 was unchanged following DEX treatment (Fig. 4E). Following chemokine-mediated cell migration, the ratio of CD4 to CD8 T cells was equivalent between treatment groups (Fig. 4E), demonstrating that DEX affected the migration of both CD4 and CD8 T cells. These data demonstrate that the function of CCR9, as expressed by primary human CD4 and CD8 T cells, is suppressed following activation of GCRs.

Intracellular calcium flux is a rapid and sensitive measure of chemokine receptor activation (36). Therefore, to further study the effects of DEX on CCR9, we examined this signaling event. This necessitated the design of a novel flow cytometry–based assay that allows simultaneous stimulation and monitoring of vehicle- and DEX-treated CCR9⁺ T cells, using two distinct anti-CD3 fluorochromes to differentially label each population, and gating on CCR9⁺CD3⁺ cells (Fig. 5A). Additionally, nonviable cells and doublets were excluded by appropriate gating (Fig. 5A).

CCL25 induced calcium flux in ATRA-cultured cells and this effect was confined to the CCR9⁺ population (Fig. 5B). Virtually all ATRA-cultured cells expressed CXCR4 (refer to Fig. 1C), and CXCL12 induced calcium flux in both CCR9⁺ and CCR9⁻ cells (Fig. 5B). DEX treatment reduced calcium flux in response to CCL25 by 28 ± 6% (p < 0.05; Fig. 5C). In contrast, in the same cells, DEX treatment increased CXCL12-mediated calcium mobilization by 24 ± 10% (p < 0.05; Fig. 5C). Calcium flux in response to ionomycin was equivalent between treatment groups (Fig. 5C). Representative calcium flux traces from one independent experiment are shown in Fig. 5D. Collectively, these results show that although CCR9 mediated signaling was inhibited by DEX treatment, the cells were viable and capable of mounting a calcium response through coexpressed CXCR4, or when stimulated with ionomycin.

During cell extravasation, chemokines stimulate both directed cell migration and integrin-mediated cell adhesion (37). To study the effects of DEX on chemokine-stimulated cell adhesion, we optimized a static adhesion assay of T cells to MAdCAM-1 or VCAM-1 bound to the surface of plastic tissue culture plates. Adhesion to MAdCAM-1 is mediated by α₄β₇ integrin, whereas adhesion to VCAM-1 is predominantly mediated by α₄β₁ integrin (38).

CCR9⁺ ATRA-cultured cells were sorted using magnetic beads and treated for 16 h in the presence of vehicle or DEX. The great majority of sorted CCR9⁺ cells expressed CXCR4, and as previously demonstrated, DEX treatment increased the surface expression of CXCR4 without affecting CCR9 (Fig. 6A). CCL25 stimulated dose-dependent adhesion of CCR9⁺ cells to MAdCAM-1, with peak adhesion occurring at 500 nM CCL25 (data not shown). Cell adhesion to MAdCAM-1 increased following stimulation with 500 nM CCL25 or 10 nM CXCL12 by 2.8 ± 0.7-fold (p < 0.01) and 4 ± 0.4-fold (p < 0.01), respectively; CXCL12 also stimulated adhesion to plate-bound VCAM-1, by an average of 2.3 ± 0.3-fold (p < 0.05) (representative data are shown in Fig. 6B; pooled data were normalized to buffer-stimulated cell adhesion and statistical significance was determined by a paired t test). However, CCL25 did not stimulate adhesion to VCAM-1 (data not shown). These data are in accord with previous reports of CCL25 stimulated adhesion to MAdCAM-1 without adhesion to VCAM-1, and CXCL12 stimulated adhesion to VCAM-1 or MAdCAM-1, measured in vitro (4, 39). DEX treatment reduced CCL25-stimulated cell adhesion to MAdCAM-1 by 60 ± 9% (p < 0.05) and increased CXCL12-stimulated adhesion to MAdCAM-1 or VCAM-1 by 113 ± 16% (p < 0.001) and 609 ± 111% (p < 0.05), respectively (Fig. 6C). Adhesion to MAdCAM-1 stimulated by MnCl₂, which activates integrins by inducing conformational change, was unaffected by DEX treatment (Fig. 6C) (40).

In this study, we demonstrate that GCs suppress chemotaxis, intracellular calcium flux, and integrin-mediated cell adhesion stimulated by the intestine-specific chemokine receptor pair, CCL25 and CCR9. Interestingly, following corticosteroid treatment, the direction and magnitude of effects were not universally shared among three studied chemokine receptors: CXCR3 surface expression was reduced whereas CCR9 was unaffected; both CCR9- and CXCR3-mediated chemotaxis were suppressed; and coexpressed CXCR4 had increased surface expression and enhanced downstream functions. The observations relating to CCR9 and CXCR3 are novel, whereas the effect of DEX on CXCR4 has been previously described (22, 23).

Collectively, our data confirm previous reports that GCs increase CXCR4 mRNA and surface protein expression and increase CXCL12-mediated calcium flux and cell migration, effects that contrast with the reduction in CCR9-mediated signaling and function. Therefore, although CCR9-mediated signaling was impaired, the cells were viable and capable of functioning through coexpressed CXCR4. The effects of DEX on CCR9 were reproduced by PRED and abrogated by the GC receptor antagonist MIF, whereas the pure mineralocorticoid agonist DOC did not affect CCR9-mediated chemotaxis. Collectively, these results demonstrate that steroids with GC activity interfere with the function of CCR9 on human T cells, and this effect is not a general consequence of all corticosteroid activity. Supplemental Fig. 4 provides a diagrammatic summary of the observations from this study, which could be interpreted as showing that intracellular signaling, probably involving transcription, modulates CCR9 signaling. Further work is needed to elucidate specific underlying mechanisms that act to inhibit CCR9 signaling and function.

In this study, we focused on developing tools to study primary blood-derived human T cells. This was on the basis of early work demonstrating that chemotaxis of Molt-4 cells (cell line that constitutively expresses CCR9) was unaffected by DEX (data not shown). These data highlighted the importance of performing studies with primary cells rather than immortalized cell lines, which may have altered intracellular signaling pathways. Owing to the low frequency of CCR9⁺ T cells in the circulation (∼3%), we have used a previously described technique for inducing CCR9 expression. This requires that T cells are expanded in the presence of ATRA, IL-2, and anti-CD3/anti-CD28. The cells are then rested in the presence of IL-2 and ATRA. We have also observed the same effect in primary CCR9⁺ T cells sorted from peripheral blood; these cells have not undergone in vitro TCR activation. We cannot rule out the influence of other mediators that may be released from highly activated T cells over the course of 16 h. If this is the case, this would be downstream of GCR activation, as the GCR antagonist MIF ameliorated the suppressive effects of DEX to CCR9 function.

To further elucidate changes to intracellular signaling, we developed a sensitive technique for measuring intracellular calcium flux. Intracellular calcium mobilization is a rapid and transient response following chemokine receptor activation. CCL25-stimulated calcium flux in primary peripheral blood cells has not been previously described because conventional assays are insufficiently sensitive when the proportion of CCR9⁺ cells is low. Using our more sensitive assay, where individual chemokine receptor-expressing cells can be selectively monitored, we confirm that CCL25 induces calcium flux in CCR9⁺ ATRA-cultured human T cells. Using this assay we were able to demonstrate that DEX suppressed CCR9 stimulation, whereas the same cells showed an enhanced signal when stimulated through coexpressed CXCR4. The described calcium assay allowed nonviable cells to be excluded from analysis and untreated and DEX-treated cells to be simultaneously stimulated and monitored in the same sample tube, excluding the possibility of differences between technical replicates. These data demonstrate that CXCR4- and CCR9-mediated intracellular signaling pathways are differentially affected by GC treatment in primary derived human T cells.

MAdCAM-1 is expressed in mucosal tissues, including the intestine, and in IBD, expression increases in active lesions (7). DEX inhibits inflammation-induced expression of MAdCAM-1, measured in vitro (41). To our knowledge, this is the first demonstration that DEX also inhibits CCL25-stimulated adhesion to MAdCAM-1. Interestingly, the same cells had enhanced adhesion to MAdCAM-1 or VCAM-1 when stimulated through coexpressed CXCR4, demonstrating a preferential suppression of CCR9 signaling. These results are further evidence that corticosteroids can differentially modulate intracellular signaling and functions downstream of chemokine receptor activation.

Corticosteroids are regularly used to treat IBD, and characterizing their molecular effects could reveal novel mechanisms of action. Previous studies of DEX conducted in vitro have used concentrations between 100 nM and 1 μM (22, 28, 42, 43). The concentrations of DEX (250 nM) and PRED (1 μM) used in these experiments are physiologically relevant concentrations, calculated on the basis of recommended dosing relative to blood volume, and for DEX correspond to levels measured in the serum after oral administration (44, 45). It is therefore tempting to speculate that these results identify a novel effect of GCs that could contribute to therapeutic efficacy in treating intestinal inflammation, by inhibiting CCR9-medated adhesion and migration of effector T cells. However, to define more exactly the physiological and pathophysiological relevance of these effects, in vivo experiments would be necessary. These constitute a body of work that is beyond the scope of this study; however, such work could reveal a novel approach for controlling intestinal specific inflammation.

GCs are effective in both ulcerative colitis and Crohn’s disease, and their use is limited mainly by adverse side effects, particularly with chronic administration. However, if intestine-specific actions of GCs could be maintained, while limiting systemic effects, for instance by targeting CCR9⁺ or α4β7⁺ cells, their therapeutic benefit and utility could be enhanced. One might hypothesize that delivering GCs to gut-homing leukocytes could have dual effects: suppressing gut homing and exerting additional anti-inflammatory effects, such as transcriptional repression of proinflammatory cytokine expression.

The suppressive effects of GCs on CCR9 and α₄β₇ function are relevant in human disease, as therapies targeting these surface proteins are being evaluated in clinical trials. Treatments targeting CCR9 (CCR9 antagonist, CCX282-B, vercirnon) and α₄β₇ (anti-α₄β₇, vedolizumab) have recently been tested in large, multinational phase II/III clinical trials in IBD, and a subset of enrolled patients was on concomitant steroid treatment (12, 15, 46). It is therefore important to be aware of the effects of GCs on the therapeutic target under investigation to inform the analysis of study results and the design of future clinical trials.

S.K. has provided consultancy services for a number of pharmaceutical and healthcare companies, including Abbvie, AstraZeneca, ChemoCentryx, Dr. Falk Pharma, Ferring, GSK, MSD, Mitsubishi Pharma, Pfizer, Vifor, and Warner-Chilcott, and has received research support from ChemoCentryx, MSD, and Warner-Chilcott. E.W. has received grant support and reagents from ChemoCentryx. The remaining authors have no financial conflicts of interest.