INTRODUCTION

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Cell migration is a required behavior in the development and maintenance of multicellular organisms. During embryonic development, epithelial cell migration occurs at the gastrulation stage, and neural crest cell migration is required to form the developing nervous system. The axonal growth cones of elongating nerves detect and migrate in response to various chemoattractants and chemorepellents in the surrounding environment as they are directed toward their precise target. In the adult, movement of fibroblasts into areas of injury plays an important role in wound repair, and the migration of endothelial cells performs a paramount role during angiogenesis. Leukocytes are recruited to sites of inflammation, and, during normal immune surveillance, lymphocytes migrate from the blood into various microenvironments of lymphatic organs whereupon they encounter antigen presenting cells that stimulate the lymphocytes' effector functions and promote a more potent immune response. Although much is known concerning the motor that drives cell migration (actin assembly and disassembly) (1, 2), the cellular signal transduction pathways that coordinate these processes in the context of cell movement are largely undefined.

In addition to pro-migratory stimuli (chemokinesis), chemoattractants also provide a directional signal to cells leading to migration toward the source of the chemoattractant (chemotaxis). In the case of leukocytes, chemoattractants direct migration from the bloodstream into organized lymphoid organs and into inflamed tissues (3-6). Subsequently, they may target lymphocytes into specialized microenvironmental niches within lymphoid organs (4). Similarly, myeloid cells in inflamed tissues can be directed toward the invading organism via the interaction of host-derived and target-derived chemoattractants (7-9). Chemoattractants also can direct the rapid, integrin-dependent adhesion of leukocytes to various cell-associated or extracellular proteins if the corresponding chemoattractant receptor is expressed at high levels (10). The number of identified leukocyte chemoattractants has exploded in the past years with the addition of many newly cloned chemokines (chemoattractant cytokines) to the list of "classical" chemoattractants such as formyl peptides, C5a, and leukotriene B₄.

Most leukocyte chemoattractants mediate their activity by binding and stimulating specific G_i-coupled receptors (11, 12). Chemoattractant binding to receptor facilitates the receptor-coupled G_i subunit exchange of bound GDP for GTP, and subsequent dissociation of G_i from the receptor and the heterodimer. The active GTP-bound G_i subunit and the released heterodimer can then stimulate their respective effector pathways. Recent studies suggest that -mediated signaling is critical to the chemotactic response (13, 14). The G_i subunit has an intrinsic GTPase activity that converts the active GTP-bound form back to the inactive GDP-bound form. GDP-bound G_i reassociates with the heterodimer to complete the cycle and turn off the G-protein-stimulated signal(s) (15). Importantly, little is known as yet about the duration of the stimulated G-protein response to chemoattractants (16), let alone about the importance of this to proadhesive versus migratory behaviors of the cell.

Sophisticated mechanisms exist, however, to modify cellular responses through G-protein coupled receptors. Treatment with a chemoattractant can render the cell less responsive to subsequent stimulation by the same agonist, a process to which homologous receptor phosphorylation and desensitization can contribute (17). Treatment with some attractants can also render a cell less responsive to subsequent treatment with agonists that bind different receptors, a process that can be mediated by heterologous receptor desensitization (17), heterologous receptor sequestration (18), or, potentially, inhibition of downstream machinery. Both homologous and heterologous cross-talk between agonists and their receptors can help control and target leukocyte migration to chemoattractants (7-9).

Recently, a distinct molecular mechanism with the potential to modulate G-protein responses has been discovered. Genetic evidence from mutants in Saccharomyces cerevisiae (19), Caenorhabditis elegans (20), and the filamentous fungus Aspergillus nidulans (21) suggested that a group of proteins with a common domain, the RGS¹ (for regulator of G-protein signaling) domain (20, 22), can directly control aspects of G-protein-stimulated signaling pathways. RGS proteins appear to enhance the endogenous GTPase activity of G-proteins, thus decreasing the half-life of the active GTP-bound state and limiting the duration of G_i signaling. At least 19 mammalian RGS members have been identified (23). Many RGS members have been shown to bind to G_i proteins and activate the G_i subunit's intrinsic GTPase activity in in vitro biochemical assays. Although only few in vivo studies have been performed in mammalian cells, these studies have suggested that RGS proteins can regulate G-mediated events such as G-protein-gated inward rectifier K⁺ (GIRK) channels (24, 25) as well as G_i-stimulated responses. Inhibition of signaling by RGS proteins may reflect GDP-bound G_i reassociation with and hence sequestration of free active heterodimers.

Based on these observations, we hypothesized that RGS proteins might also regulate cellular migratory and proadhesive responses to chemoattractants. It is not apparent, however, whether to expect that RGS proteins would stimulate or inhibit chemotaxis and adhesion responses. RGS4 and RGS8 have unexpectedly been shown to stimulate rather than inhibit the opening of GIRK channels in Xenopus oocytes following the addition of acetylcholine or dopamine. Subsequent to the increased conductance through these channels, RGS4 and RGS8 also stimulated the rapid closure of the same channels following agonist removal (24, 25). In addition, recent studies of light-stimulated rhodopsin/transducin signaling clearly indicate that certain cellular responses (e.g. the amplitude of light-induced membrane depolarization) are regulated not by the duration of the G-protein signal, but rather by the rate of receptor desensitization, a G-protein-independent process (26). Thus, if and how RGS proteins regulate proadhesive and migratory responses must depend on the dynamics of the signaling pathways involved, and must be assessed experimentally.

This study was undertaken to address this issue using lymphoid cell responses to classical chemoattractants and chemokines as a model. The results suggest that regulation of the GTPase activity can significantly influence the migratory and adhesive response to chemoattractants and demonstrate the potential for RGS control of chemoattractant-directed cell motility.

	EXPERIMENTAL PROCEDURES

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Materials-- The cDNAs for human FLAGged formyl peptide receptor (fPR) (27), human FLAGged CXCR1 (CXCR1) (28), human FLAGged C5a receptor (C5aR) (29, 30) and human FLAGged CCR-2B (31) encoded within pRC-CMV (Invitrogen, Carlsbad, CA) have been described previously (10). "FLAG" epitope addition to the N terminus of these receptors does not affect the binding of their corresponding agonist nor their subsequent responses (10). The L1/2.2 line was derived by transient transfection of the parental L1-2 cells with a chemoattractant receptor and selecting for the cells that migrated to ligand. The migrated cells were expanded (losing receptor expression in the process) and the cycle repeated once more to generate the L1/2.2 line. The L1/2*CXCR1/CCR-2B cell line was developed as follows. L1-2 cells were stably transfected with the FLAGged CXCR1 as described in Ref. 10, FACS sorted for high expression using the anti-FLAG monoclonal antibody M1, and selected for functional expression of the receptor by chemotaxis to interleukin-8 (IL-8) (32, 33). Subsequently, the CXCR1-positive cells were stably transfected with FLAGged CCR-2B and cells with functional levels of receptor obtained by chemotaxis of the double transfectants to the CCR-2B ligand, monocyte chemoattractant protein (MCP-1) (34). The L1/2*fPR/CXCR1 cell line was developed in an analogous way as the L1/2*CXCR1/CCR-2B cell line. The cDNAs for RGS1, RGS2, and RGS3 encoded within pRC-CMV and RGS4 encoded within pCR3 (Invitrogen) have been described previously (22). The RGS4N88S mutant was made by annealing primers containing the appropriate mutation (AAT to AGT) to the pCR3 vector containing RGS4, performing 12 cycles of PCR and then digesting the starting vector with DpnI according to manufacturer's instructions enclosed in the QuikChange site-directed mutagenesis kit (Stratagene, La Jolla, CA). Clones were isolated that had lost the unique SspI site in RGS4 following mutagenesis and were sequenced to confirm that there were no additional mutations. pGreen Lantern was obtained from Life Technologies, Inc. and contains a "humanized" GFP gene with a S65T mutation. FMLP and recombinant human C5a was purchased from Sigma. IL-8 was a generous gift from Sandoz Forschungsinstitut (Vienna, Austria), and MCP-1 was obtained from R&D Systems (Minneapolis, MN).

Transfection of L1/2 Cells-- L1/2.2, L1/2*CXCR1/CCR-2B, and L1/2*fPR/CXCR1 cells grown in RPMI 1640 containing 10% iron-supplemented bovine calf serum, 1 mM sodium pyruvate, penicillin-streptomycin, 2 mM L-glutamine, and 500 µM -mercaptoethanol (Growth Medium) were harvested by centrifugation after reaching >3 × 10⁶ cells/ml (post-logarithmic growth). Cells were resuspended in RPMI 1640 at 1.0-1.5 × 10⁸/ml and 0.8 ml of cells transferred to an Electro Chamber (0.4-cm gap, Life Technologies, Inc.). The DNAs to be co-transfected were mixed in a sterile microcentrifuge tube, brought to 137 mM NaCl, and 500 µg of yeast tRNA (Boehringer Mannheim) in phosphate-buffered saline added. The DNA mixture was added to cuvette, mixed by inversion, and allowed to stand at room temperature for 10 min. The cells were electroporated at 310 V/1180 microfarads using a Cell-PORATOR (Life Technologies, Inc.). After electroporation, cells were allowed to stand at room temperature for 10 min and then diluted into 5-7 ml of Growth Medium and incubated at 37 °C and 8% CO₂ for 5-6 h. Cells were harvested by centrifugation, resuspended in 0.7 ml of Growth Medium, and layered on top of 0.7 ml of Histopaque 1119 (Sigma) in a microcentrifuge tube. The cells were centrifuged at 400 × g for 30 min at room temperature. Viable cells were harvested from the interface, washed in Growth Medium, and incubated overnight in Growth Medium containing 2 mM sodium butyrate (Sigma) to maximize the expression of proteins transcribed from the cytomegalovirus promoter (35) and to increase the cells' intrinsic migratory capacity if they were to be used for most chemotaxis assays. Cells were not incubated with sodium butyrate if they were transfected with any of the GFP-RGS chimeras since the GFP intensity was a direct measure of RGS expression and there was no need to maximize the expression of the GFP-RGS chimeras.

FACS Analysis of Transfected Cells-- 5 × 10⁵ cells were stained with the mouse anti-FLAG M1 monoclonal antibody (IBI, New Haven, CT) or isotype control antibody (MOPC-141, Sigma) at 3 µg/ml for 1 h on ice in 1× phosphate-buffered saline containing 2% bovine serum albumin (Fraction V, Sigma), 1 mM CaCl₂, 1 mM MgCl₂, and 0.1% NaN₃. Cells were washed and stained with phycoerythrin-conjugated goat anti-mouse IgG (Jackson ImmunoResearch Laboratories, West Grove, PA) for 1 h on ice at the concentrations indicated in the figure legends. Cells were analyzed on a FACScan using Cellquest software (Becton Dickinson, San Jose, CA).

Agonist-stimulated Chemotaxis of Receptor-transfected L1/2 Cells-- RPMI 1640 containing 0.5% BSA (Fraction V) was CO₂-equilibrated overnight. The following day, agonists were resuspended in RPMI/BSA to the indicated concentration and 0.6 ml were aliquoted into a 24-well plate (Corning Costar Corp., Cambridge, MA). The plates were returned to the CO₂ incubator to re-equilibrate for >1 h. L1/2 cells transiently transfected the previous day were harvested and washed once in RPMI/BSA. The cells were resuspended in RPMI/BSA at 1.0 × 10⁷/ml. Transwell cell culture chamber inserts (polycarbonate 5-µm pore size, Costar Corp., Cambridge, MA) were placed in the wells containing agonist, and 100 µl of cells (1.0 × 10⁶) were added to each transwell. The cells were allowed to migrate through the porous bottom for 4 h at 37 °C. The transwell inserts were removed, and the number of migrated cells was counted by mixing with a known number of polystyrene beads (15 µm, Polysciences, Inc., Warrington, PA) and analyzing the ratio of cells to beads on the FACScan as described previously (10). Dead cells, displaying decreased forward scatter and propidium iodide uptake, were eliminated from all analysis.

Quantitative and Qualitative Western Blot Analysis-- For quantitative analysis, 15 µg of whole cell lysate and increasing amounts of His₆-RGS2 (kindly provided by David Siderovski, Amgen Institute, Toronto, Canada) or glutathione S-transferase-RGS3 were separated on an 11% SDS-PAGE gel, transferred to polyvinylidene difluoride membranes, and blocked overnight in WB buffer (25 mM Tris, pH 7.4, 300 mM NaCl, 1 mM CaCl₂) containing 5% nonfat dried milk. The blots were washed twice for 10 min each with WB buffer containing 0.05% Tween 20 (WBT), incubated with rabbit polyclonal anti-RGS2 (1:400) or anti-RGS3 (1:800) antiserum (22) for 1 h in WBT, washed twice with WBT for 10 min each, and incubated with alkaline phosphatase-conjugated goat anti-rabbit IgG (Zymed Laboratories Inc., South San Francisco, CA) at a 1:500 dilution for 1 h in WBT. The blots were then washed extensively in WBT three to four times for 10 min each, then washed twice with 25 mM diethanolamine, pH 9.5, 300 mM NaCl, 1 mM CaCl₂, 1 mM MgCl₂ for 10 min each. Bound antibodies were detected using nitro blue tetrazolium/5-bromo-4-chloro-3-indolyl phosphate or the chemiluminescent substrate CDP-Star (NEN Life Science Products) and visualized with Hyperfilm-ECL (Amersham Pharmacia Biotech). Quantification of immunoreactive band intensity was performed by comparison to known quantities of His₆-RGS2 and glutathione S-transferase-RGS3 probed side by side. The quantification was performed on a LKB Bromma UltroScan XL enhanced laser densiometer using GelScan XL software (Pharmacia LKB Biotechnology, Uppsala, Sweden).

Agonist-activated Adhesion of Receptor-transfected L1/2 Cells to VCAM-1-- Agonist-activated adhesion was performed as described in Ref. 10 with the following modifications. 16 µl of cells were allowed to settle onto the VCAM-1-coated well for 7 min. Following addition of 4 µl of 500 nM FMLP and incubation for 4 min, the slides were inverted into a coupling jar with ice-cold HBSS, 10 mM Hepes, pH 7.4. After allowing the non-adherent cells to fall off the slide for 3 min, the slide was removed and re-inverted onto a cold metal block to prevent drying while other slides in the group were being handled. 20 µl of HBSS (without divalent cations) ,10 mM Hepes, pH 7.4, 5 mM EDTA, 5 mM EGTA, 0.5% BSA (Fraction V) (Recovery Medium) containing approximately 10,000 polystyrene beads (15 µm, Polysciences, Inc., Warrington, PA) were aliquoted into each well containing adherent cells. The slide was placed in a humidified chamber at room temperature to prevent drying until all adhesion assays were performed. The adherent cells and beads were resuspended by repetitive pipetting and transferred to a polypropylene FACS tube containing 200 µl of Growth Medium. The well was washed with 20 µl of Recovery Medium without beads and combined with the first wash. A "100% adhesion" control was made by transferring 16 µl of cells and 20 µl of Recovery Medium containing beads directly into a FACS tube. The number of adherent cells and their GFP intensity was counted by analyzing the ratio of cells to beads on the FACScan as described previously (10). PMA-stimulated adhesion was performed as described above except 4 µl of 1 µM PMA was added to the cells and incubated for 30 min before the slide was inverted into ice-cold HBSS, 10 mM Hepes, pH 7.4.

Analysis of Subcellular Distribution of RGS Family Members by Confocal Microscopy-- RGS family members were tagged by fusing them by their N termini with an enhanced GFP mutant. Fusion of sequences to the N termini of RGS members have previously been shown not to inhibit RGS function (36-46). The full-length sequence of different RGS members were PCR-amplified using Pfu DNA polymerase (Stratagene, La Jolla, CA) and primers containing a KpnI site. The resulting PCR products were cut using KpnI and cloned non-directionally into the KpnI site of pEGFP-C1 (CLONTECH) by standard techniques. Vector clones with the correct orientation were tested for their ability to regulate L1/2 migration to chemoattractants. Selected clones were linearized with ApaLI, transfected into L1/2*fPR (47) using the transfection method detailed above, and placed under G418 selection to obtain stable transfectants. Individual clones of L1/2 cells stably transfected with GFP, GFP-RGS1, GFP-RGS2, and GFP-RGS3 were obtained by limiting dilution. Clones with similar GFP expression levels were selected for further confocal microscopy analysis.

	RESULTS

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RGS3 Inhibits Chemotaxis Signaled through Co-transfected Chemoattractant Receptors-- To examine the effects of RGS proteins, we developed a procedure for efficient transient transfection of the pre-B L1-2 cell, an Abelson virus transformed C57L/J lymphoma (48, 49), as described under "Experimental Procedures." This method reproducibly yielded ~50% transfection efficiency of L1/2.2 cells. Importantly, co-transfected vectors are taken up together and coordinately expressed in this system. For example, Fig. 1 illustrates correlated expression of transfected FLAGged CXCR1 (the experimental gene in this example) and GFP (the marker gene). As shown in Fig. 1d, the cells that express CXCR1 also express GFP, demonstrating efficient co-transfection of the two vector-encoded genes (most transfected cells are in upper right quadrant, i.e. double expressors). Moreover, except at the lowest levels of expression, there is a positive correlation between the expression of the two transfected genes; thus, the level of GFP expression can be used as an indicator of the relative expression from a co-transfected vector.

View larger version (45K): [in this window] [in a new window]   Fig. 1.   Coordinate expression of marker and experimental plasmid-encoded proteins into L1/2 cells. L1/2.2 cells were co-transfected with 25 µg of pRC-CMV*FLAGged CXCR1 and either 12.5 µg of empty pRC-CMV (panels a and b) or 12.5 µg of pGreen Lantern (panels c and d) and grown as described under "Experimental Procedures." The cells were stained with 3 µg/ml of either isotype control MOPC-141 (panels a and c) or M1 monoclonal antibody (panels b and d) followed by phycoerythrin-conjugated goat anti-mouse IgG (1:800 dilution). The M1 antibody recognizes the FLAG epitope engineered onto the N terminus of CXCR1 and can used to visualize receptor expression on the y axis. GFP's intrinsic fluorescence is visualized on the x axis. Similar results were obtained in three separate experiments.

Effects of RGS Proteins on Chemotaxis of Cells Stably Expressing CXCR1 and CCR-2B-- To study the effects of different RGS proteins and the importance of relative expression levels in an otherwise constant cellular environment, we next transfected RGS-encoding or control vectors along with a vector encoding GFP into L1/2 cells stably transfected with CXCR1 and CCR-2B (L1/2*CXCR1/CCR-2B). A typical histogram of GFP expression by each of the four RGS transfectants is depicted in Fig. 2a. The L1/2*CXCR1/CCR-2B cell line averages 30% transient transfection efficiency and there is little or no difference in the GFP expression among the various transfectants (Fig. 2a). In this model, transfected cells express homogeneous levels of CXCR1 and CCR-2B independent of the co-transfected RGS member (as assessed by immunofluorescence staining and flow cytometric analyses); thus, all cells have the potential to respond to chemoattractant. However, the RGS super-transfected cells can be distinguished since they co-express GFP.

View larger version (26K): [in this window] [in a new window]   Fig. 2.   GFP expression levels before and after migration of transiently transfected L1/2 cells to chemoattractant. Panel a, L1/2*CXCR1/CCR-2B cells were transiently transfected with 12.5 µg of pGreen Lantern and 25 µg of pRC-CMV (thin line), pRC-CMV*RGS1 (dashed line), pRC-CMV*RGS2 (dotted line), or pRC-CMV*RGS3 (dark line). Cells were grown as described under "Experimental Procedures," and their GFP expression levels quantified by FACS analysis. The presented histograms are an average of six independent transfections. Panel b, L1/2*CXCR1/CCR-2B cells transiently transfected with pGreen Lantern and either pRC-CMV (thin line) or pRC-CMV*RGS3 (dark line) were subjected to a chemotaxis assay to 1 nM IL-8. The migrated cells were harvested and their GFP expression pattern quantified by FACS analysis. The presented histograms are an average of six independent transfections. The percent inhibition of chemotaxis by RGS3 at each of the different expression levels is denoted. ND, not determined.

View larger version (38K): [in this window] [in a new window]   Fig. 3.   RGS inhibition of L1/2*CXCR1/CCR-2B chemotaxis to IL-8 and MCP-1. L1/2*CXCR1/CCR-2B were transiently transfected with 12.5 µg of pGreen Lantern and 25 µg of pRC-CMV (open squares), pRC-CMV*RGS1 (filled squares), pRC-CMV*RGS2 (filled circles), or pRC-CMV*RGS3 (filled triangles). The transiently transfected populations were subjected to a chemotaxis assay to the indicated concentrations of IL-8 (panels a-c) or MCP-1 (panels d-f). Migrated cells were harvested and quantified for the number of cells and the GFP expression levels of those cells by FACS analysis. The migration was calculated by comparing the number of input cells that were GFPnegative (panels a and d), GFPintermediate (panels b and e) ,and GFPhigh (panels c and f) to the number of cells, expressing the respective GFP levels as defined in Fig. 2b, that migrated toward the chemoattractants. These data are the mean ± S.E. from four to five separate experiments.

View larger version (34K): [in this window] [in a new window]   Fig. 4.   Comparison of wild type and mutant RGS4 inhibition of L1/2*CXCR1/CCR-2B chemotaxis to IL-8. L1/2*CXCR1/CCR-2B were transiently transfected with 12.5 µg of pGreen Lantern and 25 µg of pRC-CMV (open squares), pCR3*RGS4wt (filled squares), or pCR3*RGS4N88S (filled triangles). The transiently transfected populations were subjected to a chemotaxis assay to 1 nM IL-8. Migrated cells were harvested and quantified for the number of cells and the GFP expression levels of those cells by FACS analysis. The migration was calculated by comparing the number of input cells that were GFPnegative, GFPintermediate, and GFPhigh to the number of cells, expressing the respective GFP levels as defined in Fig. 2b, that migrated toward the chemoattractant. Similar data were obtained in two independent experiments.

View larger version (27K): [in this window] [in a new window]   Fig. 5.   Western blot analysis of transiently transfected RGS2 and RGS3. L1/2.2 cells were transiently transfected with 5 µg each of pRC-CMV*FLAGged fPR, pRC-CMV*FLAGged C5aR, and pRC-CMV*CXCR1 and 20 µg of empty pRC-CMV (lanes a and c), pRC-CMV*RGS2 (lane b), or pRC-CMV*RGS3 (lane d). Cells were harvested by centrifugation, disrupted by homogenization and boiled in sample buffer. 15 µg of whole cell lysate was subjected to SDS-PAGE and probed with appropriate polyclonal antibodies as described under "Experimental Procedures."

View larger version (30K): [in this window] [in a new window]   Fig. 6.   Western blot analysis and migration of L1/2 cells transiently transfected with GFP-RGS fusion constructs. Panel A, L1/2*fPR/CXCR1 cells were transiently transfected with 12.5 µg of empty pEGFP (lane a), pEGFP-RGS1 (lane b), pEGFP-RGS2 (lane c), or pEGFP-RGS3 (lane d). Cells were harvested by centrifugation, disrupted by homogenization and boiled in sample buffer. 5 µg (lanes a-c) or 1 µg (lane d) of whole cell lysate was subjected to SDS-PAGE and probed with anti-GFP antibody (1:2000 dilution) as described under "Experimental Procedures." Panel B, the above cells were subjected to a chemotaxis assay to 1 nM IL-8. Migrated cells were harvested and quantified for the number of cells and the GFP expression levels of those cells by FACS analysis. The migration was calculated by comparing the number of input cells that were GFPnegative, GFPdim, and GFPbright to the number of cells that migrated toward the chemoattractant. The GFPnegative, GFPdim, and GFPbright gates were set in the first, second, and third decade of fluorescent intensity as assessed by flow cytometry, respectively. There were no GFP-RGS2 cells in the GFPvery bright gate (fourth decade), which precluded calculating their migration in relation to cells expressing GFP, GFP-RGS1 and GFP-RGS3 at this level. These data are the mean ± S.E. from six to eight separate experiments.

Effects of RGSs on Adhesion Triggered through Co-transfected Chemoattractant Receptors-- Signals in the high endothelial venules of lymph nodes and on inflamed endothelium cause the rapid shear-resistant integrin-dependent adhesion of leukocytes (reviewed in Ref. 4). The signal(s) that promote integrin-dependent adhesion in vivo are pertussis toxin-inhibitable, suggesting that G_i-coupled receptors are involved (52), and this rapid adhesion event can be modeled by expressing chemoattractant receptors and stimulating the transfected receptor in the presence of the appropriate cell adhesion molecules (e.g. VCAM-1) (10). Therefore, L1/2*fPR/CXCR1 cells transfected with GFP or any of the "active" GFP-RGS chimeras were stimulated by FMLP addition to bind VCAM-1-coated wells via their endogenous ₄₁ integrins. In the absence of agonist, there is low background adhesion to VCAM-1 (2% of cells bind spontaneously), but after FMLP addition there is a rapid increase in the total cells bound (>20% of input cells bind after 4 min of stimulation with 100 nM FMLP). As seen in Fig. 7, expression of bright levels of GFP-RGS1, GFP-RGS3, and GFP-RGS4 inhibited the adhesion signal and decreased the recovery of those cells from the VCAM-1 coated wells. In contrast, none of the GFP-RGS chimeras inhibited PMA-stimulated L1/2 cell adhesion to VCAM-1, a G_i -independent event (data not shown).

View larger version (38K): [in this window] [in a new window]   Fig. 7.   Chemoattractant-stimulated integrin-dependent adhesion of L1/2 cells transiently transfected with RGS members. L1/2*fPR/CXCR1 cells were transiently transfected with 12.5 µg of pEGFP, pEGFP-RGS1, pEGFP-RGS3, or pEGFP-RGS4. Cells were grown and subjected to an adhesion assay as described under "Experimental Procedures." The cells were stimulated with 100 nM FMLP for 4 min and adherent cells harvested for analysis by flow cytometry. The data presented are the mean ± S.D. from triplicate wells normalized to the adhesion of GFPnegative cells in the same wells. The gates used to separate the different GFP expression levels were the same gates defined in Fig. 6B. These data are the mean ± S.E. from three to five separate experiments.

Subcellular Distribution of RGS Proteins Assessed by Confocal Scanning Microscopy-- GFP and the different GFP-RGS chimeras were expressed stably in L1/2*fPR cells and observed by confocal scanning microscopy. GFP is expressed in L1/2 cells at a fairly uniform concentration throughout the cell (Fig. 8a) as is GFP-RGS1 (Fig. 8b) and GFP-RGS4 (data not shown). However, a majority of the GFP-RGS2 is localized in a subcellular compartment away from the plasma membrane where RGS proteins would be expected to regulate receptor-coupled G-GTP levels (Fig. 8c). Staining with the cell-permeable DNA dye SYTO-59 identified this region of RGS2 localization as the nucleus (data not shown). Most of the remaining non-nuclear GFP-RGS2 is diffusely associated with the plasma membrane or a periplasma membrane region, with little in the cytoplasm in general. The same predominately nuclear localization was also observed when GFP-RGS2 was transfected into the adherent, non-lymphoid Balb/c 3T3 cell line (data not shown). Strikingly, the large GFP-RGS3 was cytosolic and totally excluded from the nucleus (Fig. 8d). Membrane association of RGS1, RGS3, and RGS4 (data not shown) was difficult to assess in the context of the intense diffuse cytoplasmic fluorescence of these proteins, but the GFP-RGS2 chimera was apparently on or near the plasma membrane due to the contrast with the largely negative cytoplasm. At the level of resolution available, no change in GFP-RGS distribution was apparent following FMLP stimulation (data not shown).

View larger version (132K): [in this window] [in a new window]   Fig. 8.   Laser scanning confocal images of the intracellular distribution of GFP-RGS fusion proteins. L1/2*fPR cells were stably transfected with empty pEGFP (panel a), pEGFP-RGS1 (panel b), pEGFP-RGS2 (panel c), or pEGFP-RGS3 (panel d), and clones that expressed similar GFP intensity were isolated. Confocal slices through the middle of the cells were obtained. The bar in the lower left corner of each panel denotes a length of 5 µm.

	DISCUSSION

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The RGS proteins are a recently identified family of molecules with the potential to regulate diverse heterotrimeric G-protein-mediated signaling events in vivo. In yeast, pheromones bind a G-protein-coupled receptor that signals downstream events that lead to mating preparation (19). These downstream events include changes in gene transcription, morphological and cytoskeletal changes, and growth arrest. If mating does not occur, however, the yeast becomes desensitized to the pheromone signal and recovers from the growth arrest (19). Mutational screens were used to isolate a protein (Sst2p) that, when expressed normally, promoted the recovery from pheromone-stimulated growth arrest. Genetic analysis suggested that Sst2p regulated the yeast G subunit (GPA1) and in an unidentified manner inhibited G-protein-mediated signaling (19). In independent studies, mutational screens to detect egg-laying defects in C. elegans revealed a protein (EGL-10) that modulates the G-protein-regulated egg laying behavior (20). Additionally, the flbA gene in A. nidulans antagonized a Gprotein-dependent developmental change (21). All three genes modulated G-protein-mediated events, and all three contained a common domain. This domain has been named the RGS domain (for regulator of G-protein signaling (20, 22).

Recently, mammalian genes containing RGS domains have been isolated by a variety of techniques (22, 36, 38, 42, 53-58). Analysis of previously cloned mammalian genes revealed two human proteins containing RGS domains: BL34/1R20 (renamed RGS1), an early response/activation gene up-regulated during PMA-induced plasmacytoid differentiation (53) and enriched among B-cell specific genes (54); and GOS8 (renamed RGS2), an activation gene of mononuclear cells (55). RGS3 was subsequently isolated by homology from a B-cell cDNA library (22). RGS4 was obtained using a brain cDNA library to screen for genes that compensated for the yeast RGS mutant (22).

The RGS domain appears to mediate direct physical interaction with G G-proteins (36). Genetic data from flbA action suggested that the intrinsic GTPase activity of the G-protein was required for RGS-mediated inhibition of function (21), and in vitro assays using purified RGS members and G-proteins have confirmed these observations. RGS proteins had no effect on the G-protein's ability to bind GTP or to release GDP, actions akin to GDS and GDI proteins on small GTP-binding proteins (59). Instead, RGSs enhance the intrinsic GTPase activity of the G-protein acting as GTPase-activating proteins (GAPs). Most RGS members tested modulate the activity of G_i and/or G_q G-proteins but do not modulate G_s G-proteins (37-39, 60, 61). No selectivity toward the different members of the G_i subfamily thought to mediate both the chemotatic and proadhesive activities of chemoattractant receptors has been reported.

The present study was undertaken to assess the ability of RGS proteins to influence chemoattractant-stimulated cell migration and adhesion. We have demonstrated that some RGS members modulate the ability of transfected lymphoid cells to chemotax and adhere. Some but not all cellular responses to G-protein-linked receptors are regulated by the duration of G-protein signaling (26), and our results suggest strongly that chemoattractant-directed leukocyte migration and adhesion falls into this category. RGS1, RGS3, and RGS4 were potent inhibitors of L1/2 cell chemotaxis to each of the chemoattractants tested, including FMLP, C5a, IL-8, and MCP-1. RGS3 and RGS4 also inhibited chemotaxis to MIP-1 by L1/2*CXCR1/CCR-1 cells (not shown). RGS1 and RGS3 inhibited chemotaxis at all concentrations of chemoattractant with no apparent shift in the EC₅₀ of the response; this is consistent with its action being downstream of the receptor. A decrease in response without a change in the EC₅₀ of the agonist has been observed previously for RGS4 and RGS8 inhibition of G-activated GIRK channels stimulated by acetylcholine or dopamine (24, 25). The ability of these RGSs to modulate chemotaxis was dependent on their GAP activity since a mutant RGS4, shown previously to be impaired in its G_i binding activity and subsequent GAP activity toward G_i, was much less effective at regulating chemotaxis than wild type RGS4. Although less dramatic than their effects on chemotaxis, RGS1, RGS3, and RGS4 also reduced the rapid, integrin-dependent adhesion stimulated through chemoattractant receptors as well, suggesting that the half-life of the activated G-protein subunit also plays a role in the intensity of the proadhesive signal, albeit quantitatively less significant than for chemotaxis.

RGS2 had no effect, under any experimental conditions, on L1/2 cell chemotaxis. Even when comparing populations normalized (by quantitative Western blot analysis or equal GFP fusion protein expression) for RGS levels at which RGS3 demonstrated dramatic inhibition of chemotaxis, no inhibition by RGS2 was detected. Although the unique, predominantly nuclear intracellular localization of GFP-RGS2 is of uncertain significance, it raises another possible correlation with the lack of RGS2 effect on chemotaxis. We conclude that RGS2 is relatively impotent as a modulator of the chemotactic response in lymphoid cells. An inability of RGS2 to regulate certain biological responses (IP₃ generation and K⁺ channels) that other RGSs modulate has been observed in other systems, as well (24, 62). RGS2 also differs from other RGS family members in its interaction with G proteins. RGS4, RGS5, and RGS16 but not RGS2 bound well in vitro to G_i2 (the predominant G_i protein in leukocytes (63, 64)), G_i3, and G_o (42, 61). This observation has been extended very recently to show that RGS2 binds G_q and can modulate G_q-stimulated IP₃ generation in vitro (61). Thus, it is possible that in L1/2 cells, RGS2 can preferentially associate with G_q versus G_i proteins. The inability of RGS2 to inhibit chemotaxis may reflect this selectivity of G interaction. However, these and our results do not rule out that RGS2 can act similar to RGS1, RGS3, and RGS4 in other cells; for example, all four RGSs inhibited IL-8-stimulated MAP kinase activity in 293T cells transfected with CXCR1 (22). If RGS2 itself localizes primarily to the nucleus in L1/2 cells (like the GFP-RGS2 chimera), it may be partially sequestered from the required site of action.

Our results confirm that RGS proteins can modify chemoattractant and proadhesive responses, and suggest that differential and/or regulated expression of RGS members may allow fine tuning of leukocyte motility and adhesion in response to chemoattractant stimuli. Since RGS1 and RGS2 were first described as activation genes in leukocytes, and RGS3 has been demonstrated in PMA-stimulated tonsillar B and T cells,² it is reasonable to hypothesize that RGS activity is modulated during lymphocyte activation and/or development, and it will be important to explore their differential regulation during leukocyte development and in response to proinflammatory or antigenic stimuli.

In conclusion, RGS1, RGS3, and RGS4 modulate the ability of motile lymphoid cells to migrate in a gradient of chemoattractant, presumably by decreasing the half-life of the activated G-protein. The results suggest that the duration of the G_i active state is a major determinant of the chemotactic response, which requires coordination and integration of multiple downstream responses over a prolonged time period. In contrast, RGS2 was unable to regulate migration to any chemoattractant when it was expressed at any level achievable in our system. RGS1, RGS3, and RGS4 significantly inhibited rapid, chemoattractant triggered adhesion as well, although to a lesser extent than chemotaxis. We postulate that, within the intracellular environment, the regulated expression of RGS molecules in conjunction with G_i specificities, regulatory interactions, and subcellular localization of RGS proteins may help control leukocyte migratory responses to chemoattractant stimuli, thus regulating the microenvironmental trafficking of leukocytes in vivo.