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Regulator of G Protein Signaling 1 (RGS1) Markedly Impairs G_i Signaling Responses of B Lymphocytes

Chantal Moratz^*, Veronica H. Kang^*, Kirk M. Druey^1,*, Chong-Shan Shi^*, Astrid Scheschonka^2,*, Philip M. Murphy, Tohru Kozasa and John H. Kehrl^3,*

^* B Cell Molecular Immunology Section, Laboratory of Immunoregulation, and Laboratory of Host Defenses, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD 20892; and Department of Pharmacology, University of Texas Southwestern Medical Center, Dallas, TX 75235

	Abstract

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Regulator of G protein signaling (RGS) proteins modulate signaling through pathways that use heterotrimeric G proteins as transducing elements. RGS1 is expressed at high levels in certain B cell lines and can be induced in normal B cells by treatment with TNF-. To determine the signaling pathways that RGS1 may regulate, we examined the specificity of RGS1 for various G subunits and assessed its effect on chemokine signaling. G protein binding and GTPase assays revealed that RGS1 is a G_i and G_q GTPase-activating protein and a potential G₁₂ effector antagonist. Functional studies demonstrated that RGS1 impairs platelet activating factor-mediated increases in intracellular Ca⁺², stromal-derived factor-1-induced cell migration, and the induction of downstream signaling by a constitutively active form of G₁₂. Furthermore, germinal center B lymphocytes, which are refractory to stromal-derived factor-1-triggered migration, express high levels of RGS1. These results indicate that RGS proteins can profoundly effect the directed migration of lymphoid cells.

	Introduction

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Germinal centers within lymphoid tissue form specialized microenvironments essential for the induction of thymus-dependent B cell Ab production, the affinity maturation of the humoral immune response, and the induction of B cell memory. Although many of the genetic and biochemical mechanisms that underlie these processes remain poorly understood, some of the receptor-ligand interactions controlling these events activate heterotrimeric G proteins. For example, chemokine-chemokine receptor interactions regulate lymphocyte migration patterns as well as the establishment of germinal centers (1, 2, 3, 4). Furthermore, pretreatment of B lymphocytes with pertussis toxin, which blocks G_i-mediated signaling, inhibits chemoattractant-induced B cell migration (5, 6, 7, 8).

Heterotrimeric G proteins, which couple heptahelical receptors to effectors in signal transduction pathways, consist of three subunits: , ß and . Each subunit has multiple isoforms; the subunit isoforms are grouped into four families, G_i, G_s, G_q, and G_12/13 (reviewed in Ref. 9). Upon receptor activation, the subunit complex exchanges GTP for GDP and dissociates from the ß subunit. Both GTP-bound G and the released ß subunit can activate downstream effectors. Several mechanisms regulate the duration and magnitude of G protein signaling (reviewed in Ref. 10). Protein kinases can phosphorylate G protein-coupled receptors (GPCR)⁴, allowing members of the arrestin family to bind the phosphorylated receptor, precluding subsequent G protein activation. The levels of GPCR expression often decline following ligand simulation, thereby reducing the number of available receptors for subsequent restimulation. G protein signaling can also be inhibited by proteins that accelerate the intrinsic GTPase activity of G subunits. GTPase activators (GAPs), were first discovered for the small GTPase proteins Ras and EF-tu (reviewed in Refs. 11 and 12). Recently, G subunit GAPs have also been identified and termed regulator of G protein signaling proteins (RGS; reviewed in Refs. 13 and 14).

Insights into the function of the RGS family members arose from the identification of evolutionarily conserved homologues in Saccharomyces cerevisiae (Sst2) and Caenorhabditis elegans (Egl-10 and C05B5.7) (15, 16). Sst2p contains a split RGS domain and in yeast is a key negative regulator of the mating response to pheromone. In C. elegans, the Egl-10 protein contains an RGS domain and functions in signal transduction pathways that regulate egg laying and movement (15). Mammalian RGS proteins also regulate G protein-linked signal transduction pathways. Introduction of RGS family members into yeast blunted the responses to pheromone and partially complemented an sst2 mutation while the same RGS proteins expressed in HEK 293 cells stably transfected with CXCR1 blunted the activation of mitogen activated protein kinase (MAPK) following IL-8 stimulation (16). RGS proteins inhibit signaling pathways that utilize either G_i or G_q as signal transducers (reviewed in Refs. 13 and 14). Recently, p115 RhoGEF has been shown to have an RGS-like domain that has GAP activity for G_12/13 (17).

RGS proteins bind G subunits and do so most efficiently in a form that mimics a transition state in GTP hydrolysis (G treated with GDP and AlF₄^-) and possess GAP activity for G_i and G_q subfamily members (18, 19, 20, 21). In the crystal structure of RGS4 complexed with G_i1-GDP-AlF₄^-, the RGS domain forms a four-helix bundle that directly contacts G_i at the three so-called "switch regions" (22), which undergo the greatest conformational change during the GTPase cycle. Mutagenesis studies of RGS4 revealed that altering the contact residues identified in the crystal structure resulted in a loss of G binding and an inability to inhibit G signaling (23, 24). Overall these studies indicate that RGS proteins stabilize G_i in its transition state for GTP hydrolysis. Besides acting as GAPs for G_i and G_q, certain RGS proteins also act as effector antagonists, i.e., compete with effectors for binding to G_q (25). Despite these advances in our understanding of their mechanisms of actions, the physiologic roles of most RGS proteins remain poorly defined.

Chemokine receptor signaling is a prime arena for regulation by RGS proteins. This type of regulation may help target cells to particular sites and keep them localized there despite the continued exposure to chemokines and chemoattractants. The initial studies of RGS1 indicated high levels of expression in tonsil germinal centers. Here we show the specificity of RGS1 for various G subtypes, discern some of the signals that induce RGS1 expression in B lymphocytes, and provide evidence that RGS1 down-regulates signaling initiated by the platelet-activating factor (PAF) receptor and CXCR4 in stable RGS1-transfected lines. We also confirm that RGS1 protein is specifically expressed in germinal center B cells that, although they express the CXCR4 receptor, are refractive to stromal-derived factor (SDF)-1-induced migration.

	Materials and Methods

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Production of recombinant RGS1

A bacterial expression vector for RGS1 and RGS4 were constructed using Vent polymerase and the coding region of RGS1 or RGS4 to generate a PCR fragment flanked by XhoI and BamHI restriction sites, which were subcloned into the corresponding sites of the His-tag fusion vector pET14b (Novagen, Madison, WI). The resulting constructs pET14b-RGS1 and pET14b-RGS4 were transformed into the bacterial strain BL21 (DE3) pLysS and induced with 1 mM isopropyl ß-D-thiogalactoside (IPTG) for 3 h at 37°C. The His-tagged RGS proteins were purified from a 100-ml culture by metal chelation chromatography as outlined by the manufacturer (Novagen). The purified proteins were dialyzed against HED buffer (50 mM Na-HEPES (pH 8.0), 1 mM EDTA, 1 mM DTT, and 10% glycerol).

G protein-binding assays

HS-Sultan cells (5 x 10⁶) were lysed in a 0.5 ml of buffer that consisted of 20 mM HEPES (pH 8.0), 1 mM EDTA, 6 mM MgCl₂, 3 mM DTT, 380 mM NaCl, and 1% Triton X-100 for 20 min at 4°C. The lysates were centrifuged at 14,000 x g for 20 min to remove particulates and transferred to fresh tubes before activation with 20 µM GDP, 20 µM GDP plus 30 µM AlF₄^-, or 30 µM GTP-S for 30 min at 30°C. RGS1 attached to Ni + 2NTA beads was added, and the mixture was incubated for another 90 min at 4°C. The beads were washed four times with lysis buffer that included the appropriate nucleotide, the bound G proteins were eluted with SDS-sample buffer, and the samples were fractionated on SDS-PAGE before transfer and analysis by immunoblotting for various G subunits (see below).

G proteins and GAP assays

G_s, G_i1, and G_o were expressed in and purified from Escherichia coli (26). G_z, G₁₂, and G_qR183C were expressed in Sf9 cells and purified as described (27). G_i1, G_o, G_s, G_z, and G₁₂ (50 pM) were loaded with 5–10 µM [-³²P]GTP (5000 Ci/mmol) at 20°C (for G_s) or 30°C (for G_i1, G_o, G_z, and G₁₂) for 20 to 30 min in the presence of 5 mM EDTA. Samples were then gel-filtered at 4°C through a Sephadex G-50 spin column equilibrated with buffer A (50 mM HEPES (pH 8.0), 1 mM DTT, 5 mM EDTA,and 0.05% of the detergent C₁₂E₁₀) to remove free [-³²P]GTP and ³²P_i. Hydrolysis of GTP was initiated by adding G loaded with [-³²P]GTP in buffer A containing 8 mM MgSO₄ and 1 mM GTP with the indicated amount of RGS proteins. The reaction mixture was incubated at 4°C (for G_i1, G_o, and G_s) or 15°C (for G_z and G₁₂). Aliquots (50 µl) were removed at the indicated times and mixed with 750 µl of 5% (w/v) Norit in 50 mM NaH₂PO₄. The mixture was centrifuged at 2000 rpm for 5 min, and 400 µl of supernatant containing ³²P_i was counted by liquid scintillation spectrometry. Direct measurement of the k_cat for GTPase activity of G_q was assayed with the use of mutant G_qR183C (28). An analogous mutant of G_i, G_iR178C, has markedly reduced GTPase activity but still responds to RGS proteins. The slow GTPase activity of G_qR183C made it possible to load [-³²P]GTP on G_q without accelerating GDP-GTP exchange by agonist bound receptor. G_qR183C was loaded with 10 µM [-³²P]GTP in the presence of 50 mM HEPES (pH 7.4), 0.1 mg/ml BSA, 1 mM DTT, 1 mM EDTA, 0.9 mM MgSO₄, 30 mM (NH₄)₂SO₄, 4% glycerol, and 5.5 mM CHAPS at 20°C for 2 h. The reaction mixture was gel filtered through a Sephadex G50 spin column equilibrated with 50 mM HEPES (pH 7.4), 1 mM DTT, 1 mM EDTA, 0.9 mM MgSO₄, 0.1 mg/ml BSA, and 1 mM CHAPS. GTPase assays were initiated by addition of 1 mM GTP and the indicated amount of RGS proteins and incubation at 20°C. Aliquots (50 µl) were removed and processed as described above.

Cell lines

The HS-Sultan (a B lineage human plasmacytoma), Molt-4 (a human T lineage leukemia), Jurkat (a human acute T cell leukemia line), COS-7 (an SV40-transformed fibroblast like kidney line), PC12 (a rat adrenal pheochromocytoma), Ramos (a human Burkitt’s B cell chronic lymphoma), and K562 (a human chronic myelogenous leukemia) cell lines were obtained from the American Type Culture Collection (ATCC, Manassas, VA). The Burkitt lymphoma cell lines MC116 and CA46 were kind gifts of Dr. Ian Magrath (National Cancer Institute (NCI), National Institutes of Health (NIH), Bethesda, MD). The SuDHL5 and SuDHL6 B lymphoma cell lines were kind gifts of Dr. Lazlo Krenacs (NCI, NIH, Bethesda, MD). The human pre-B cell line, NALM-6, was a kind gift of Dr. Thomas Tedder (Duke University, Durham, NC). All the lymphoid cell lines were maintained in RPMI 1640 supplemented with 5% to 10% FCS whereas the nonlymphoid cells were maintained in DMEM plus 10% FCS. HS-Sultan cells overexpressing RGS1 were generated using the retroviral vector LXSN-RGS1 as previously described (16). Twelve separate pools of neomycin-resistant cells were isolated and tested for RGS1 expression; three pools that expressed RGS1 at high levels, termed TF7, TF8, and TF12, were used in these studies. A pool of LXSN vector-transfected and neomycin-resistant cells served as a control.

Isolation of tonsil B cells and tonsil B cell subsets

Tonsillar B cells were isolated as described in Current Protocols in Immunology. First, lymphocytes were isolated by Ficoll-Hypaque (Amersham Pharmacia Biotech, Uppsala Sweden) density gradient centrifugation (29). T cells were then depleted by rosetting with neuraminidase (Life Technologies, Gaithersburg, MD)-treated sheep RBC (NIH Media Unit, Frederick, MD) and subsequent Ficoll-Hypaque density gradient centrifugation. The purity of tonsillar B cells was routinely greater than 95% using this method as defined by post separation immunofluorescense staining with anti-CD4, anti-CD8, and anti-CD19 (PharMingen, San Diego, CA). Subsequently, the cells were washed and analysis was performed on a FACScalibur flow cytometer with CELLQuest software (Becton Dickinson, Mountain View, CA). The purified B cells were stimulated with various reagents for 12 h in RPMI 1640 and 10% FCS. IL-8, IL-10, and TNF- were purchased from R&D Systems (Minneapolis, MN), anti-CD40 from PharMingen, and PAF, PMA, and sphingosine-1-phosphate were purchased from Sigma (St. Louis, MO). B cell subsets were obtained from the B cell-enriched fraction by cell sorting. In brief, separation of the naive, memory, and germinal center B cells were isolated according to a schematic developed by Yong-Jun Liu (30, 31). The cells were incubated with mAbs to CD19, IgD, and CD38 (PharMingen) for 15 min and washed in FACS staining buffer (PBS without Ca²⁺/Mg²⁺ plus 1% BSA). Using a FACStar^Plus (Becton Dickinson), the cells in the CD19-positive gate were sorted, based on their differential expression of CD38 and IgD, into naive, memory, and germinal center B cells. Reanalysis of each sorted population by flow cytometry indicated a population purity of greater than 97% for each subset sorted. Lysates of the sorted fractions were analyzed by immunoblotting (see below).

Flow cytometry and migration assays

HS-Sultan-transfected cell lines were harvested and counted, and 1 x 10⁶ cells per well (96-well round-bottom plate, Costar, Cambridge, MA) were used for staining. The cells were washed in staining buffer (1% BSA fraction V (ICN Biomedicals, Aurora, OH) in PBS, blocked with 10% normal rat serum (Cedarlane Laboratories, Acccurate Chemical & Scientific, Westbury, NY) in staining buffer for 10 min. After washing, the cells were stained with mAb, anti-CD4, anti-CXCR4, and anti-CD19 (PharMingen). Subsequently, the cells were washed, and analysis was performed on a FACScalibur flow cytometer with CellQuest software (Becton Dickinson). Cell migration was assessed in triplicate in 24-well 5-µm pore size polycarbonate membrane filter transwell insert culture plates (Costar). Transfected cell lines were harvested and resuspended at 1 x 10⁷ cells per ml in culture medium (RPMI 1640, penicillin (100 U/ml), and streptomycin (100 µg/ml); all from Life Technologies), 10% FBS (Tissue Culture Biologicals, Tulare, CA). Transwell culture plates were set up. Six hundred microliters of medium or medium plus chemoattractant was placed in the bottom of the well; then 100 µl of cells (1 x 10^-6 cells) were placed in the upper chamber of the well. The chemokine SDF-1 (R&D Systems) was used at 10 ng/ml, 100 ng/ml, or 1000 ng/ml concentrations. The cultures were incubated at 37°C, 5% CO₂ for 4 h. Migrated cells were analyzed on a FACScalibur. Results are shown as percentage of total input. Chemotaxis studies with tonsil B cells were performed as described above except that migrated cells were harvested and stained with mAbs anti-CD19, anti-IgD, and anti-CD38 for 15 min at 4°C, washed, and analyzed on a FACScalibur. Additionally, the studies used a dose of 50 ng/ml of SDF-1, which was determined to be optimal to induce maximal migration in a series of titration experiments. Using Cellquest software, the percentage migration of each subpopulation of B cells was determined.

Determination of intracellular calcium levels

Cultured cells were harvested, washed in HBSS buffer (HBSS (Biofluids, Rockville, MD), 10 mM HEPES, and 1% FBS), and resuspended at 1 x 10⁷ cells/ml in HBSS buffer plus the fluorescent calcium probe Indo-1 (indo-1/acetocxymethylester) (Sigma or Molecular Probes, Eugene OR) was added at a final concentration of 2 µg/ml plus Pleuronic detergent (Molecular Probes) at a final concentration of 300 µg/ml (32). The cells were incubated 30 min at 30°C while protected from light. The cells were washed with HBSS buffer and resuspended to 1 x 10⁶ in HBSS buffer. Cells were warmed at 37°C for 3 min before stimulation. To stimulate, cells were loaded into the Time Zero module (Cyteck, Fremont, CA) and run at 1000 cells/s. A baseline was collected for 30 s, and then a sham of 50 µl of HBSS buffer was injected. Finally at 60 s the stimulant was injected. The measurement for calcium flux was performed on a FACSVantage flow cytometer (Becton Dickinson Immunocytometry Systems, San Jose, Ca) equipped with an argon laser tuned to 488 nm and a krypton laser tuned to 360 nm. Indo-1 fluorescence was analyzed at 390/20 and 530/20 for bound and free probe, respectively. The data were analyzed using the FlowJo software (Tree Star, San Carlos, CA). Results are shown as ratio fluorescence (violet/blue).

Stress-activated protein kinase (SAPK) and serum response element (SRE) reporter assays

For the SAPK assays, COS cells were transfected via a DEAE-dextran method with the following plasmids: MT3-HA-SAPK-p46 (1 µg, provided by Dr. J. Kyriakis, Boston, MA), pcDNAG₁₂-Q229L (2 µg, provided by Dr. S. Gutkind, NIH), or pcDNAG₁₃-Q226L (provided by Dr. S. Gutkind), and the presence or absence of varying concentrations of FLAGpCMV2-RGS1 or pCR3-FLAG-RGS4. Transfected DNA levels were normalized with control plasmids. Seventy-two hours following the transfection, HA-immunoprecipitates were subjected to in vitro kinase assays using c-Jun (1-79) as a substrate, and the samples were size fractionated by SDS-PAGE. Following autoradiography, the c-Jun (1-79) bands were quantitated using NIH Image. For the SRE reporter assays, COS cells were transfected with pSRE-LUC (0.35 µg, Stratagene, San Diego, CA), pCMV-ßgal (0.35 µg), and pcDNAG₁₂-Q229L (0.5 µg) or pcDNAG₁₃-Q226L in the presence or absence of FLAGpCMV2RGS1 using SuperFect (Qiagen, Valencia, CA). Twenty-four hours later, the DMEM plus 5% FCS was removed and replaced with DMEM plus 0.5% serum. The following day, the cells were harvested, and lysates were prepared in 100 µl reporter lysis buffer (Promega, Madison, WI). Using a luminometer (Analytical Luminescence Laboratory, San Diego, CA), 10 µl of the supernatant was tested for luciferase activity with a luciferase substrate (Promega), and 10 µl was tested for ß-galactosidase activity with a galactan chemiluminescent substrate (Tropix, Bedford, MA). The amount of luciferase activity was normalized to amount of ß-galactosidase activity in each sample. Each transfection was done in either duplicate or triplicate.

Immunoprecipitations and immunoblots

RGS1 Abs were prepared by immunizing rabbits with recombinant RGS1, an N-terminal peptide (MPGMFFSANPK) coupled to keyhole limpet hemocyanin (KLH), a C-terminal peptide (NDLNANSLK) coupled to KLH, or an internal peptide (DDKMNKRRPK) coupled to KLH. The antiserum generated against the N-terminal peptide (referred to as 2247) proved to be the most efficacious and, since it was raised against a peptide outside of the RGS domain, does not cross-react with other RGS proteins. For immunoblotting, the samples (25–100 µg protein, Bio-Rad Protein Assay, Bio-Rad, Richmond, CA) were fractionated by SDS-PAGE and transferred to pure nitrocellulose. The membranes were blocked with 10% milk in TTBS for 1 h, and then incubated with the appropriate dilution of Ab in 5% milk and 0.05% sodium azide in TTBS overnight (0.1% Tween 20, 100 mM Tris Cl (pH 75), 0.9% NaCl). The blots were washed twice with TTBS before the addition of a biotinylated goat anti-rabbit Ig (DAKO, Carpinteria, CA) diluted 1:10,000 in TTBS containing 10% FCS. Following a 1-h incubation, the blot was washed twice with TTBS and then incubated with streptavidin conjugated to HRP (DAKO). The signal was detected by enhanced chemiluminescence (ECL) following the recommendations of the manufacturer (Amersham). The anti-RGS1 antiserum (no. 2247) was used at 1:200 dilution; the rabbit Abs reactive with G_i1, G_i2, G₁₂, or G_i3 were used as recommended (Santa Cruz Biotechnology, Santa Cruz, CA); and the affinity purified anti-peptide Ab reactive with G_i1 and G_i2 (AS/7) was used at a 1:1000 dilution (a kind gift of Dr. P. Goldsmith, NIDDK, NIH).

	Results

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RGS1 is expressed in B cell lines and normal B cells

To examine RGS1 protein expression in B lymphocytes, we generated RGS1-specific antisera for immunoblotting. Recombinant RGS1 was produced in bacteria with a histidine tag to facilitate purification via metal chelation chromatography. The antisera were screened by immunoblotting recombinant RGS1 and by the detection of RGS1 in cell lysates from HS-Sultan cells stimulated with phorbol esters. These cells are representative of mature B cells partially differentiated toward plasma cells. Phorbol esters had been previously shown to markedly increase RGS1 (BL34) mRNA expression in HS-Sultan and tonsillar B cells (33). The RGS1 antisera recognized recombinant RGS1 via immunoblotting, and both the anti-recombinant RGS1 and the antiserum no. 2247 immunoprecipitated a 26-kDa protein in lysates prepared from HS-Sultan cells stimulated with PMA (data not shown). This molecular mass agrees well with the predicted molecular mass of RGS1, and the endogenous RGS1 protein comigrated with recombinant RGS1.

Next, we analyzed RGS1 expression in human B cell lines representative of various stages of B cell development as well as other hemopoietic cell lines. Most of the cell lines had low or undetectable levels of RGS1, with the exception of a follicular B cell lymphoma cell line SuDHL5, which expressed high levels (Fig. 1). Two pre-B cell lines, NALM-6 and PB-697, failed to express RGS1 (Fig. 1 and data not shown). One T cell line, MOLT-4, contained low levels of RGS1 whereas the other T cell line, Jurkat, was negative. Previously described HS-Sultan cell lines stably transfected with RGS1 (TF8 andTF12) and recombinant RGS1 served as positive control (16). Further immunoblotting analysis of a panel of Burkitt lymphoma cell lines revealed that several of them also constitutively expressed high levels of RGS1 (data not shown). We also examined the kinetics of RGS1 induction in HS-Sultan cells following exposure to phorbol esters. PMA induced RGS1 expression within 4 h of stimulating the cells (Fig. 1). These analyses of protein expression confirm that RGS1 expression is restricted and that it is not expressed in T cell, fibroblast cell, or most B and pre-B cell lines. Additionally, they confirm that HS-Sultan cells lack RGS1 unless stimulated by PMA, corroborating data suggesting that B cells must be activated to express RGS1.

View larger version (45K): [in this window] [in a new window]   FIGURE 1. Expression of RGS1 in various B cell lines. Cell lysates from various cells lines indicated below were analyzed by immunoblotting for RGS1 expression. In addition, 200 ng of recombinant RGS1 (lane 1) and cell lysates prepared from two of the HS-Sultan RGS1 transfectants (TF no. 8 and TF no. 12, lanes 2 and 3) served as positive controls. On a separate immunoblot (lanes 13–17), lysates (75 µg each) from HS-Sultan cells stimulated with PMA (50 ng/ml) for various duration were analyzed for RGS1 expression. The immunoblots were detected by ECL. Representative of one of three experiments performed.

View larger version (32K): [in this window] [in a new window]   FIGURE 2. Induction of RGS1 in normal tonsil B cells. Purified tonsil B cells were stimulated for 12 h with media, anti-CD40 (5 µg/ml), anti-CD40 plus IL-10 (50 ng/ml), IL-8 (50 ng/ml), PAF (10-6 M), PMA (25 ng/ml), sphingosine-1-P (50 ng/ml), or TNF- (50 ng/ml). Cell lysates were prepared and analyzed for RGS1 expression by immunoblotting with the RGS1 anti-peptide Ab no. 2247. This experiment is one of several similar experiments that were performed.

RGS1 Binds G_i, G_q, and G₁₂, but not G_s from B lymphocytes

To determine which G subunits RGS1 may regulate in B cells, we reacted recombinant RGS1 immobilized on N_i²⁺ NTA beads with cell lysates from HS-Sultan cells that had been treated with GDP or with GDP with AlF₄^-. The bound proteins were solubilized and size fractionated by SDS-PAGE, and their identity was determined by immunoblotting with antisera specific for different G subunits (Fig. 3A). In each case, an HS-Sultan cell lysate was simultaneously analyzed to verify that it contained the G subunit being studied. RGS1 readily extracted G_i1/2, G_i3, and G_q from the cell lysate treated with GDP and AlF₄^-, but not from the lysate treated with GDP alone. We failed to detect any G_s associated with RGS1 under either condition. We also examined whether RGS1 could extract G₁₂ from the HS-Sultan cell lysates (Fig. 3B). In contrast to the results with G_i, which exhibited an AlF₄^--dependent extraction by RGS1, G₁₂ was extracted whether the lysate was GDP, GTP-S, or GDP and AlF₄^- treated. Since the N_i²⁺ NTA beads alone did not extract G₁₂, the extraction depended upon the presence of RGS1. In contrast to RGS1, recombinant RGS4 did not extract G₁₂ from the HS-Sultan cell lysates (data not shown). Based on these results, we predicted that RGS1 would act as a GAP for G_i and G_q subunits, but not G_s subunits. Although RGS1 extracted G₁₂ from cell lysates, it failed to do so in an AlF₄^--dependent manner, suggesting that it was unlikely to be a G₁₂ GAP.

View larger version (25K): [in this window] [in a new window]   FIGURE 3. The specificity of RGS1 for G subunits in B cells. A, RGS1 binds Gi and Gq in a GDP plus AlF4--dependent manner. HS-Sultan cell lysates were treated with GDP or GDP plus AlF4- and reacted with His-tagged recombinant RGS1 bound to beads. After washing, the bound proteins were eluted and analyzed by immunoblotting with Abs against various G subunits. A cell lysate was simultaneously analyzed to verify the presence of the G subunit in the lysate. The immunoblot was detected by ECL. B, RGS1 binds G12 independent of its nucleotide loading. HS-Sultan cell lysates were treated with GDP (lane 3), GDP plus AlF4- (lane 4), or GTP-S (lane 5) and reacted with His-tagged recombinant RGS1 bound to beads. After washing, the bound proteins were eluted and analyzed by immunoblotting for G12 as above. The presence of RGS1 in the eluted fractions was verified by immunoblotting using a mAb reactive with the His-tag (bottom panel). Each experiment was performed twice.

RGS1 acts as GAP for G_i and G_q, but not for G_s or G₁₂

To directly test the effects of RGS1 on the GTPase activity of various G subunits, we measured the catalytic activity of purified recombinant G subunits during a single GTPase cycle in the presence or absence of recombinant RGS1 (Fig. 4). RGS4 served as a positive control since its GAP activity in these assays is well documented (19). RGS1 proved nearly as effective G_i GAP as RGS4, although approximately 3-fold more RGS1 than RGS4 was needed to achieve a similar level of GAP activity. However, when tested against two other G_i subfamily members, G_z and G_o, RGS1 proved less efficient than did RGS4. To measure the GAP activity of RGS1 for G_q, we employed a mutant G_q, G_qR183C. The analogous mutant in G_i, G_iR178C, has a markedly reduced GTPase activity, but still responds to RGS proteins (19). The slow GTPase activity of G_qR183C made it a suitable target for testing potential G_q GAPs. Both RGS1 and RGS4 showed good GAP activity for G_q, although at equal molar concentrations RGS4 was slightly superior to RGS1 (Fig. 4). As expected, neither RGS4 nor RGS1 enhanced the intrinsic GTPase activity of G_s. Furthermore, despite the ability of RGS1 to bind G₁₂, it lacked GAP activity. In contrast, p115 RhoGEF, a recently discovered G12 GAP (17), significantly increased the rate of G₁₂ GTP hydrolysis.

View larger version (34K): [in this window] [in a new window]   FIGURE 4. GAP activity of RGS1 for specific G subunits. Different concentrations of recombinant RGS1 or recombinant RGS4 were tested for their ability to accelerate the GTPase activity of Gi1, Gq R183C, Gz, Go, Gs, and G12. Hydrolysis of GTP was initiated by adding the G subunit loaded with [-32P]GTP to a buffer containing MgSO4 with the indicated amount of RGS protein. The reaction mixture was incubated, and aliquots were removed at the indicated times and processed, and the amount of 32Pi was counted by liquid scintillation spectrometry. HED is buffer without RGS protein. The GAP assays were performed twice with similar results.

RGS1 inhibits G12-Q229L signaling in COS cells

Although there is no information about the role of G₁₂ proteins in lymphocytes, they may be involved in chemokine-induced cytoskeletal changes that occur as lymphocytes switch from a spherical to a polarized motile morphology. Although RGS1 failed to act as a G₁₂ GAP, the RGS1 G protein binding data indicated that HS-Sultan cells contained significant levels of G₁₂ and that RGS1 could bind GTP-G₁₂ and, thus, potentially act as an effector antagonist. To test that possibility, we transiently expressed in COS cells a GTPase-deficient form of G₁₂, G_12-Q229, which is largely GTP bound. Since the transient expression of G₁₂-Q229L in COS cells activates the SAPK pathway (33), we could test the effects of RGS1 on SAPK activation by concomitantly transfecting an epitope-tagged version of SAPK and assessing its activity in an in vitro kinase assay (Fig. 5). Expression of G₁₂-Q229L increased SAPK activity 3- to 10-fold depending upon the experiment, and in each of five experiments the coexpression of RGS1 inhibited the activation of SAPK. In contrast, RGS4 at similar expression levels had either no effect or actually augmented SAPK activity. Although RGS1 inhibited G_12-Q229L-induced SAPK activation, it did not inhibit G₁₃-Q226L-induced SAPK activation in COS cells. Since G₁₂-Q229L is also a potent activator of serum response element (SRE)-dependent transcription (34, 35, 36), we could examine whether RGS1 expression impaired the activation of a SRE reporter construct. Cotransfection of COS cells with an SRE-responsive reporter construct and G₁₂-Q229L resulted in a 27-fold increase in luciferase activity compared with a control plasmid (Fig. 5). The concomitant expression of RGS1 resulted in a dose-dependent reduction in reporter gene activity. Higher levels of RGS1 expression than that shown resulted in further inhibition of SRE reporter activity. RGS1 failed to significantly impair G₁₃-Q226L-mediated SRE activation, a result consistent with the data from the SAPK assay. These results indicate that RGS1 can impair signal transduction via those receptors that utilize G₁₂ as a signal transducer.

View larger version (23K): [in this window] [in a new window]   FIGURE 5. RGS1 inhibits activated G12-mediated signal transduction. A, RGS1 impairs G12-Q229L-induced SAPK activation. COS cells were transiently transfected with expression constructs for HA-SAPK, G12-Q229L, and varying amounts of FLAG-RGS1 (1, 2, or 4 µg) or FLAG-RGS4 (4 µg). HA-immunoprecipitates were performed 36 h after the transfection and subjected to an in vitro kinase assay using a GST-N-terminal fragment of c-Jun fusion protein. The amount of kinase activity was quantitated by autoradiography and NIH Image, and expressed as fold induction. The expression of RGS1 and RGS4 was verified by immunoblotting with an anti-FLAG mAB. This experiment is one of five performed. B, RGS1 does not impair G13-Q226L-mediated SAPK activation. Similar experiment as in A, except G13-Q226L was used. C, RGS1 impairs G12-Q229L-induced activation of an SRE reporter. COS cells were transfected with expression constructs for CMV-ßGal, SRE-luc, G12-Q229L, or G13-Q226L, and varying amounts of RGS1 (the amount of plasmid transfected is indicated below in micrograms). The amount of luciferase and ß-galactosidase activity in each sample was measured, and the luciferase activity was normalized to the ß-galactosidase activity. The data are shown as the fold induction (activated G12 or G13 vs control). Data are from one of three experiments performed.

Next, we tested whether RGS1 modulates G protein signaling in B cells. Because of difficulties in manipulating primary B lymphocytes, we used as a model system the B cell line HS-Sultan, since it lacks RGS1 unless stimulated. To identify suitable agonists, we exposed HS-Sultan cells to ligands known to bind heptahelical receptors that activate heterotrimeric G proteins and monitored intracellular calcium levels. Of the ligands tested (SDF-1, macrophage inflammatory protein (MIP)-1, RANTES, IL-8, ATP, isoproterenol, histamine, oxotremorine, sphingosine-1-phosphate, lysophosphatidic acid (LPA), and PAF) only the lipids PAF and LPA triggered a significant increase in intracellular calcium. The chemokine SDF-1 triggered an almost negligible increase in intracellular calcium. Many B lymphocyte cell lines express PAF receptors, and PAF has been shown to increase intracellular calcium levels and activate mitogen-activated protein kinase (MAPK) (37, 38). Although little is known about the effects of LPA on B lymphocyte function, LPA increases intracellular calcium levels and activates MAPK in a variety of other cell types (39).

We transduced HS-Sultan cells with a retroviral vector that encodes RGS1 and isolated separate pools of transduced cells by neomycin selection. HS-Sultan cells transduced with an empty retroviral vector served as a control. An immunoblot of cellular lysates demonstrated RGS1 expression in the RGS1-transduced cell lines (an immunoblot of lysates from RGS1-transfected cell lines no. 8 and no. 12 is shown in Fig. 1). We exposed the HS-Sultan vector control and RGS1-transfected cells to concentrations of PAF that had triggered the strongest increase in intracellular calcium. As expected, PAF triggered a sharp increase in the intracellular calcium levels of the control cells; however, a similar concentration of PAF raised the intracellular calcium levels in the RGS1-transfected cells very modestly. Furthermore, the intracellular calcium levels rapidly returned to baseline in the transfected cells (Fig. 6A). Similarly, the control cell line increased its intracellular calcium level following LPA stimulation, whereas the RGS1 transfectants again responded poorly (data not shown).

View larger version (19K): [in this window] [in a new window]   FIGURE 6. RGS1 impairs GPCR signaling in B cells. A, Changes in free intracellular calcium in RGS1-transfected lines (TF7, 8, 12 (dashed lines)) and vector alone transfected control line (solid line) in response to PAF signaling as measured by the change in the ratio of free vs bound intercellular Ca2+. In each stimulation, buffer was added at 30 s, and PAF (10-5 M) was injected at 60 s as indicated on the x-axis denoting time. The y-axis labeled as fluorescence intensity represents the ratio fluorescence (violet/blue) of the calcium probe indo-1. Shown are representative responses of each line from at least three experiments done in duplicate in each experiment. B, RGS1 impairs SDF-1 directed cell migration. RGS1-transfected lines and the vector-transfected control line were assessed for their ability to migrate in response to increasing concentrations of SDF-1 in a standard chemotaxis assay. The data are shown as the percentage of total input migrating on the y-axis and the concentration of SDF-1 on the x-axis. The graph represents mean and SD of two independent experiments in which each concentration point was done in triplicate for each line.

SDF-1 attracts naive and memory B cells, but not germinal center B cells despite their expression of CXCR4 (3). We used T cell-depleted human tonsil cells in chemotaxis assays to confirm the migratory responsiveness to SDF-1 of each B lymphocyte subset. The cells, which migrated in response to SDF-1, were assessed for their cell surface expression of CD19, CD38, and IgD. IgD⁺CD38-negative B cells reside in the mantle zone region and are considered to be naive B cells. Germinal center B cells (CD38⁺) can be divided into IgD⁺ and IgD^- fractions, whereas memory B cells are IgD^-/CD38^-. The majority of B cells in each fraction expressed the chemokine receptor CXCR4 (Fig. 7A); however, in a standard chemotaxis assay, only the memory cells (IgD^-/CD38^- B cells) and the naive B cells (IgD⁺/CD38^-) migrated in response to SDF-1. In contrast, the two other populations were refractory to chemokine (Fig. 7B). In preliminary experiments, germinal center B cells are refractive in migrational responses to other B cell chemoattractants (data not shown). To examine whether RGS1 expression correlates with nonresponsiveness to SDF-1, we used FACS to isolate three populations of CD19-positive lymphocytes based on their differential expression of IgD and CD38. We tested the germinal center B cell fractions (IgD^-, CD38⁺), the memory B cell fraction (IgD^-, CD38^-), and the naive B cell fraction (IgD⁺, CD38^-) for their expression of RGS1 by immunoblotting. We found that the naive and memory B cells expressed low levels of RGS1 whereas the germinal center B cells contained significant levels of RGS1 as assessed by immunoblotting, equivalent to what we observed following PMA activation (Fig. 7C). Thus the refractoriness of germinal center B cells to SDF-1-triggered chemotaxis may be because they express RGS1.

View larger version (41K): [in this window] [in a new window]   FIGURE 7. Correlation of SDF-1 migratory refractoriness and RGS1 expression. A, CXCR4 expression by human tonsil B cell subsets. Human tonsil B cells were phenotyped into subsets based on differential expression of IgD and CD38 into naive, memory, and germinal center B cells. The expression level of CXCR4 receptor was assessed on each B cell subset, with the MFI of CXCR4 staining by the majority of each subset given. B, SDF-1-induced migratory responsiveness of human tonsil B cell subsets. Human tonsillar B cells were used in a chemotaxis assay to determine the migratory responsiveness to an optimal concentration (50 ng/ml) of SDF-1. Input and transmigrated populations were analyzed and phenotyped by flow cytometry. Thus, the percentage of each subset population migrating could be determined. The data are expressed as the percentage of total for each subset; thus, the percentage of naive cells migrating is indicative of the percentage of total naive cells in the population, not the percentage of total or percentage of B cells. C, RGS1 expression in human tonsil B cell subsets. Human tonsillar B cells were sorted into naive, memory and germinal center B cells. Lysates were made from each of the subpopulations, and 30 µg of each lysate was used to fractionate by SDS-PAGE, was transferred to nitrocellulose membrane, and was used for immunoblotting with anti-RGS1 Ab. Shown is a representative blot from three blots using lysates from three independent sorts.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

A surprising result in this study is that, not only did RGS1 bind G₁₂, but also its expression impaired the activation of downstream signaling by GTP-G₁₂. The direct downstream effectors of GTP-bound G₁₂ and G₁₃ are poorly characterized although a recent study showed that activated G₁₃ directly stimulates the guanine nucleotide exchange activity of p115 RhoGEF, indicating that it is a G₁₃ effector (42). However, activated G₁₂ failed to have a similar effect, suggesting that these subunits may possess both overlapping and distinct effectors. Consistent with that possibility, G₁₂ and G₁₃ differentially activate Na⁺/H⁺ exchanger isoforms (43). We also provide evidence that G₁₂ and G₁₃-triggered signaling pathways differ. G₁₂, but not G₁₃, signaling is impaired by RGS1. Precisely how RGS1 inhibits G₁₂-mediated signaling requires clarification. We have no evidence that the GAP activity of RGS1 is important for its inhibition of G₁₂ signaling. However, since all the GAP assays in this study were done in the absence of receptors, it remains possible that RGS1 is a G₁₂ GAP in the presence of the appropriate receptor. A precedent for such a possibility is that the G specificity of RGS2 was revealed only in the presence of a receptor (28). Nevertheless, based on the G₁₂ binding data, we would predict that RGS1 inhibits G₁₂ signaling pathways by behaving as a G₁₂ effector antagonist.

We analyzed the consequence of RGS1 expression on signal transduction initiated by two chemoattractants, LPA and PAF. The presence of RGS1 impaired the increase in intracellular calcium that occurs following exposure to PAF or LPA. Both PAF and LPA are potent phospholipid agonists produced by a variety of cell types. Although there is little information on the effects of LPA on B-lymphocyte function, PAF has been extensively studied. The binding of PAF to its receptor stimulates c-Fos and c-Jun transcription; increases phospholipid turnover; results in the tyrosine phosphorylation of several proteins including p53/56^lyn and p59^fyn; and increases NF-B binding activity in nuclear extracts (44, 45, 46). Functionally, PAF augments the proliferation of B cell lines, stimulates TNF- production, and increases Ig secretion (47, 48). Both PAF and LPA increase intracellular calcium by activating phospholipase C-ß, which eventually results in calcium mobilization. Since pertussis toxin only partially inhibits the PAF-induced calcium flux in B cells, PAF likely signals through G_q in addition to G_i to mobilize calcium (38, 49). Thus, the effect of RGS1 of PAF-induced Ca⁺² mobilization is consistent with its GAP activity for G_i and G_q.

The induction of RGS1 in B lymphocytes would be predicted to impair the recruitment of B cells to inflammatory sites by PAF and LPA and inhibit their effects on B cell function. We previously showed that the transient expression of RGS1, RGS3, or RGS4 inhibited the migration of a pre-B cell line transfected with CXCR1 and CCR2 to IL-8 or MCP1 (50). In that study, the inhibitory effect of RGS3 substantially exceeded that of either RGS1 or RGS4. Furthermore, in HEK 293 cells transfected with CXCR1, RGS3, and RGS4 both proved superior to RGS1 in inhibiting IL-8-induced MAPK activation (16). In this study, RGS1 potently inhibited the migration of a B cell line triggered by the chemokine SDF-1 via its cognate receptor CXCR4. In contrast, when we tested the migratory response to SDF-1 of HS-Sultan cell lines permanently transfected with RGS3, SDF-1 attracted the RGS3-transfected cells just like it did the vector control cells (C. Moratz, unpublished observation). This is in contrast to previous experiments in which RGS3 inhibited chemotaxis in transient transfections; however, these current data are the results of RGS3 stable overexpression in a B cell line in response to signaling by an endogenous receptor. Thus, similar to the findings with G_q-coupled receptors where receptor-RGS protein specificity has been noted (51, 52), RGS proteins may differ in their abilities to inhibit signaling through specific chemokine receptors.

Both SDF-1 and CXCR4 are necessary for normal B cell development (53, 54, 55). SDF-1-induced migratory response depends upon the release of G_i-associated G_ß subunits (53, 56, 57). RGS1 does not bind G_ß (20); therefore, it is unlikely to directly influence the interaction of G_ß with its downstream effectors. However, since RGS1 is a GAP for G_i, its presence will reduce the duration that G_i remains bound to GTP in the cell. Since the GDP-bound form of G_i has a high affinity for G_ß, it will rapidly recombine with free G_ß, thereby reducing the amount of G_ß available to interact with effectors. An apt illustration of the ability of RGS proteins to indirectly inhibit G_ß signaling was provided by experiments in yeast with Sst2p and RGS4 (16). The response to pheromone is mediated by the release of ß from the yeast G subunit. Although both Sst2p and RGS4 inhibit responses triggered by pheromone exposure, they fail to inhibit when the signal transduction pathway is initiated directly by G_ß. Thus, the inhibition of HS-Sultan cells to migrate in response to SDF-1 by RGS1 is likely related to its G_i GAP activity.

What role might RGS1 have in germinal center B cells? Although its elevated expression of germinal center B cells suggests a role in regulating signaling through a GPCR or receptors, no such receptors are known to participate in the B cell proliferation or the selection processes that occurs in the germinal center. One possibility is that high levels of RGS1 inhibit signaling through a chemokine receptor that regulates egress from the germinal center. RGS1 levels may remain high in germinal center B cells until they are ready to exit the germinal center. At that point, their RGS1 levels decline, they become chemokine responsive, and they escape from the germinal center. A candidate for such a chemokine is SDF-1. The majority of the B cells that reside in germinal center are not attracted by SDF-1 despite their expression of CXCR4 receptors. Germinal center B cells are less responsive to B cell chemoattractants in general; thus it is possible that the migrational unresponsiveness reflects an overall lack of motility rather than an effect of RGS1 expression. It may also indicate that RGS1 may regulate more than just CXCR4 signaling or that other RGSs are involved in regulating B cell responses. However, we know that RGS1 antagonizes SDF-1-triggered chemotaxis based on the inhibition of SDF-1-induced migration of the HS-Sultan line once RGS1 is introduced. A prediction of this model is that the lack of RGS1 would not impair germinal center formation, but that germinal center B cells would not be retained properly in the germinal center region. Another chemokine important in germinal center formation is B cell-attracting chemokine (BCA-1), which selectively attracts B lymphocytes via the BLR1 receptor (CXCR5) (1, 58). In mice lacking CXCR5, the emigration of activated B lymphocytes into B cell follicles to form germinal centers is impaired, suggesting that BCA-1 attracts activated B cells into the germinal center (1). HS-Sultan cells migrated poorly to BCA-1; thus, we could not determine whether RGS1 has a role in regulating migration induced by signaling through the CXCR5 receptor.

As more RGS family members are subjected to G binding, GAP assays, and signaling assays, subtle differences among them are becoming evident. RGS1 has emerged as a good G_i and G_q GAP and as a potential G₁₂ effector antagonist. Based on its expression in germinal center B cells, it likely regulates signaling through GPCRs important for germinal center B cell function. Finally, its up-regulation by TNF- and its inhibitory effects on LPA and PAF signaling suggests that it may also have a role in modulating B cell function in inflammatory responses.

	Acknowledgments

 
We thank Mary Rust for her excellent editorial assistance, Gaye Lynn Wilson and Pamela C. Sternweis for their technical assistance, and Dr. Anthony S. Fauci for his support and encouragement.

	Footnotes

 
¹ Current address: Klinik und Poliklinik fuer Neurologie, Universitat Goettingen, 37075 Gottingen, Germany.

² Current address: Molecular Signal Transduction Section, Laboratory of Allergic Diseases, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD 20892.

³ Address correspondence and reprint requests to Dr. John H. Kehrl, National Institute of Allergy and Infectious Diseases, Building 10, Room 11B-13, 10 Center Drive, MSC 1876, Bethesda, MD 20892-1876. E-mail address:

⁴ Abbreviations used in this paper: GPCR, G protein-coupled receptor; RGS, regulator of G protein signaling; PAF, platelet-activating factor; BCA-1, B cell-attracting chemokine-1; MAPK, mitogen-activated protein kinase; SAPK, stress-activated protein kinase; SDF-1, stromal-derived factor-1; MFI, mean fluorescence intensity; GAP, GTPase-activating protein; SRE, serum response element; KLH, keyhole limpet hemocyanin; ECL, enhanced chemiluminescence; LPA, lysophosphatidic acid.

Received for publication August 31, 1999. Accepted for publication December 9, 1999.

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