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First published online 13 December 2005
doi: 10.1242/jcs.02722

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ß2-chimaerin provides a diacylglycerol-dependent mechanism for regulation of adhesion and chemotaxis of T cells

¹ Department of Immunology and Oncology, Centro Nacional de Biotecnología/CSIC, Cantoblanco, E-28049 Madrid, Spain
² Department of Immunology, Centro de Investigaciones Biológicas/CSIC, Ramiro de Maeztu 9, E-28040 Madrid, Spain
³ Department of Molecular Pharmacology, University of Pennsylvania Medical School, 3620 Hamilton Walk, Philadelphia, PA 19104, USA

^* Author for correspondence (e-mail: imerida{at}cnb.uam.es)

Accepted 3 October 2005

	Summary

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The small GTPase Rac contributes to regulation of cytoskeletal rearrangement during chemokine-induced lymphocyte adhesion and migration in a multi-step process that is very precisely coordinated. Chimaerins are Rac1-specific GTPase-activating proteins of unknown biological function, which have a canonical diacylglycerol C1-binding domain. Here we demonstrate endogenous expression of ß2-chimaerin in T lymphocytes and study the functional role of this protein in phorbol ester and chemokine (CXCL12)-regulated T-cell responses. We used green fluorescent protein-tagged ß2-chimaerin and phorbol ester stimulation to investigate changes in protein localization in living lymphocytes. Our results demonstrate that active Rac cooperates with C1-dependent phorbol ester binding to induce sustained GFP-ß2-chimaerin localization to the membrane. Subcellular distribution of GFP ß2-chimaerin in living cells showed no major changes following CXCL12 stimulation. Nonetheless Rac1-GTP levels were severely inhibited in GFP-ß2-chimaerin-expressing cells, which displayed reduced CXCL12-induced integrin-dependent adhesion and spreading. This effect was dependent on chimaerin GTPase-activating protein function and required diacylglycerol generation. Whereas ß2-chimaerin overexpression decreased static adhesion, it enhanced CXCL12-dependent migration via receptor-dependent diacylglycerol production. These studies demonstrate that ß2-chimaerin provides a novel, diacylglycerol-dependent mechanism for Rac regulation in T cells and suggest a functional role for this protein in Rac-mediated cytoskeletal remodeling.

Key words: Diacylglycerol, C1 domain, Chemokines, Immune response, Rac GTPases

	Introduction

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The chemokines are a family of low-molecular-weight chemotactic proteins that can induce directional migration of cells. CXCL12 (stromal-cell-derived factor 1; SDF-1) is a CXC chemokine initially identified as a growth-stimulating factor for B-cell progenitors (Nagasawa et al., 1994). It is expressed in a broad range of tissues and has numerous biological activities in diverse cell types including T lymphocytes, monocytes, pre-B lymphocytes and dendritic cells (Kucia et al., 2004). CXCL12-induced chemotactic activity is mediated by the seven-transmembrane G protein-coupled receptor (GPCR) CXCR4, also a known co-receptor for T-cell-tropic human immunodeficiency virus strains (Bleul et al., 1996; Oberlin et al., 1996). CXCR4 is the only chemokine receptor with ubiquitous expression in precursor and mature leukocytes, suggesting a general function in leukocyte homeostasis. During lymphocyte transendothelial migration, CXCL12 binding to its receptor results in rapid activation of T-cell integrins, including 4ß1 (Grabovsky et al., 2000). Interaction of integrins with their vascular ligands leads to arrest and firm adhesion of the circulating lymphocytes, and subsequent transmigration through the endothelium. Targeted disruption of Cxcl12 or Cxcr4 in mice showed their crucial functions for fetal development of the hematopoietic, cardiovascular and cerebellar systems (Nagasawa et al., 1996; Nagasawa et al., 1998).

As is the case for other chemokines, CXCL12 signals through heptahelical GPCR known to activate phospholipase C (PLC), which induce Ca²⁺ mobilization from intracellular stores and increase membrane diacylglycerol (DAG) levels. This lipid messenger, as well as its related analogs the phorbol esters (PE), exerts its actions through binding to specific C1 domains, first described in the classical and novel protein kinase C family (Kazanietz, 2002). DAG/PE-dependent binding leads to membrane translocation/activation of the C1-containing target proteins (Brose et al., 2004; Hurley and Meyer, 2001). The role of PLC-dependent DAG generation in the regulation of chemotaxis is debated. PLCß2 and ß3 are proposed to exert a negative role in leukocyte migration (Li et al., 2000), whereas other reports suggest that chemokine-dependent PLC activation is necessary for T-cell chemotaxis (Smit et al., 2003; Spiegel et al., 2004). In any case, the identity of the downstream signal targets of DAG generation and their exact role in the regulation of chemokine-dependent responses remains to be determined.

View larger version (40K): [in this window] [in a new window]   Fig. 1. ß2-chimaerin is expressed in lymphoid cells. (A) Expression of ß2-chimaerin mRNA was determined in Jurkat T cells (JK) by RT-PCR using specific oligonucleotides; 1, 2 and ß2-expressing plasmids were used as controls. (B) ß2-chimaerin protein expression was analyzed in total lysates of the indicated lymphoid cell lines by western blotting with anti-ß2-chimaerin antibody. Membranes were re-probed with anti-tubulin antibody as a protein loading control. H, HT; HD, HDLM2; J, Jurkat; K, Karpas; M, Molt-4; N, Namalwa; SU, SU-DHL1; T, Toledo. (C) Subcellular localization of GFP-ß2-chimaerin in Jurkat T cells was examined by confocal microscopy (green, left), the Golgi was stained with the BODIPY TR fluorescent marker ceramide (red, center); merged fluorescence is shown in the right panel. Bar, 3 µm.

Chimaerins are Rac1-specific GAPs of unknown biological function, characterized by the presence of a high-affinity DAG-binding C1 domain (Diekmann et al., 1991; Hall et al., 1990). The chimaerin family is composed of two isoforms, and ß, encoded by two different genes, with two alternative splice variants each (1 and 2). All chimaerins have a Rac-GAP domain together with a C1 module; in addition, type 2 isoforms exhibit an extended N-terminal region encompassing an SH2-type domain (Hall et al., 1993). Chimaerins are expressed abundantly in brain, where developmentally regulated expression has been shown for and ß2 isoforms (Hall et al., 2001; Lim et al., 1992). ß1-chimaerin, expressed specifically in testis, is also developmentally regulated during acrosomal assembly (Leung et al., 1993). ß2-chimaerin was first identified as a 46 kDa protein in rat cerebellum (Leung et al., 1994). When overexpressed in COS cells, ß2-chimaerin relocates to the plasma and Golgi membranes in response to PE and other DAG analogs (Kazanietz, 2002). PE-dependent ß2-chimaerin translocation requires the C1 domain (Caloca et al., 1999), and is mediated by PE binding as well as by direct interaction with Tmp21, a Golgi transmembrane protein (Wang and Kazanietz, 2002).

In this study we demonstrate endogenous expression of ß2-chimaerin in T lymphocytes and investigate its role in the modulation of CXCL12-dependent functions. Integrin-dependent adhesion, actin polymerization, and cell spreading can be induced by CXCL12, and are essential for CXCL12-induced polarized T-cell migration. We demonstrate that overexpression of green fluorescent protein (GFP)-ß2-chimaerin alters all three parameters, showing a functional role for this protein in CXCL12-regulated responses. ß2-chimaerin function requires its Rac-GAP activity and a functional C1 domain. While exerting a negative role on chemokine-induced static adhesion, ß2-chimaerin potentiates the chemotactic response to CXCL12 in a DAG-dependent fashion. This study provides new evidence for the role of non-kinase DAG receptors in the regulation of T-cell responses, and suggests a functional role for ß2-chimaerin in the regulation of CXCL12-dependent responses.

	Results

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Jurkat T cells express endogenous chimaerin
We assessed ß2-chimaerin expression in the Jurkat acute T-cell leukemia-derived cell line by reverse transcriptase polymerase chain reaction (RT-PCR), using specific primers and plasmids encoding different chimaerin isoforms as controls (Fig. 1A). Expression of endogenous protein was confirmed in these cells, as well as in different human leukemia and lymphoma cell lines, using an antibody to the ß2-chimaerin catalytic domain. Western blot analysis of cell lysates showed expression of a protein of appropriate molecular mass (Fig. 1B). High protein expression was observed in Jurkat and in Molt-4 acute lymphoblastic leukemia cells, as well as in cell lines derived from anaplastic large cell and Hodgkin's lymphomas. No protein expression was detected in lymphoma cell lines of other origins.

To study ß2-chimaerin function in T lymphocytes, we generated a GFP-tagged protein and examined its intracellular distribution. Western blot analysis indicated that GFP-ß2-chimaerin was expressed and migrated according to the predicted molecular size of the fusion protein (not shown). Confocal images showed that GFP-ß2-chimaerin localized in cytosol, and fluorescent staining indicated partial co-localization of GFP-ß2-chimaerin with the Golgi apparatus (Fig. 1C).

PMA-dependent regulation of GFP-ß2-chimaerin in Jurkat cells
ß2-chimaerin has a canonical C1-type domain, suggestive of DAG/PE-dependent regulation. To analyze its functional role, we examined the intracellular distribution of GFP-tagged ß2-chimaerin in response to the PE phorbol 12-myristate 13-acetate (PMA), and compared it with that of the isolated ß2-chimaerin C1 domain. Confocal analysis of living Jurkat T cells showed that the GFP-ß2-chimaerin C1 domain (GFP-C1), which showed a diffuse localization in untreated cells, was rapidly relocated to membranes following PMA addition (Fig. 2A and Movie 1 in supplementary material). In these cells, addition of the same PMA concentration nonetheless did not induce translocation of the full-length ß2-chimaerin (Fig. 2A and Movie 2 in supplementary material). The recently reported crystal structure of ß2-chimaerin (Canagarajah et al., 2004) suggests a conformation for the native protein in which the C1 domain is hidden and not readily available for DAG/PMA binding. We examined PMA-dependent translocation of two GFP-ß2-chimaerin constructs bearing mutations of glutamine in position 32 or isoleucine in position 130, two residues suggested to maintain a closed protein conformation through intramolecular interactions (Canagarajah et al., 2004). PMA-dependent translocation of GFP-ß2-chimaerin Q32A or I130A mutants fully mimicked that of the isolated C1 domain (Fig. 2A and Movie 3 in supplementary material). These results further confirm that the lack of PMA-dependent GFP-ß2-chimaerin translocation correlates with a buried C1 domain in the native protein conformation.

View larger version (41K): [in this window] [in a new window]   Fig. 2. PMA-dependent regulation of GFP-ß2-chimaerin in intact Jurkat cells. (A) Jurkat T cells were transfected with vectors encoding GFP-ß2-chimaerin C1 domain (GFP-C1), GFP-ß2-chimaerin, GFP-ß2-chimaerin Q32A and GFP-ß2-chimaerin I130A. At 24 hours post-transfection, cells were suspended in HBSS and plated on poly-D,L-lysine-coated chamber slides. Slides were mounted on a 37°C plate and analyzed by confocal microscopy. PMA was added at 100 ng/ml after the first frame, and images captured every 10 seconds. Time-lapse confocal recording is shown in Movies 1-3 in the supplementary material. Images are shown at the indicated times to illustrate subcellular protein localization. (B) Jurkat T cells were transfected with GFP-ß2-chimaerin or GFP-ß2-chimaerin C1 domain mutant (GFP-F215G). At 24 hours post-transfection, cells were treated as in A. PMA (800 ng/ml) was added after the first frame, and images captured every 10 seconds. Time-lapse confocal recording is shown in Movie 4 in supplementary material. Images are shown at the indicated times to illustrate subcellular protein localization. (C) Jurkat T cells were co-transfected with a EE-tagged, constitutive active (RacV12) Rac mutant and either GFP-ß2-chimaerin or GFP-F215G mutant. At 24 hours post-transfection, cells were stimulated with PMA (100 ng/ml) for 3 minutes, fixed, stained with anti-EE antibodies (red) and images acquired by confocal microscopy. GFP-ß2-chimaerin localization (green) is shown in the left column, RacV12 expression (red) is shown in the middle column, merge images are shown on the right. Time-lapse confocal recording of a similar experiment performed in living cells is shown. Bars, 5 µm.

As described for COS cells (Caloca et al., 2001), at higher PMA concentration ß2-chimaerin relocated rapidly to both internal and plasma membrane sites (Fig. 2B and Movie 4 in supplementary material). To assess the contribution of the C1 domain to PMA-dependent translocation, we generated a GFP-ß2-chimaerin construct in which the conserved phenylalanine in C1 domain position 3 was mutated to glycine (F215G). Studies with PKC showed that this residue is crucial for PMA binding when the ligand is presented in micelles, as indicated by the lack of binding in point mutants at this position (Kazanietz et al., 1995). The F215G mutation fully impaired PMA-dependent translocation of the mutant, confirming that C1-mediated binding to PMA is essential for membrane translocation in living cells (Fig. 2B). Lack of PMA-dependent GFP-ß2-chimaerin F215G translocation mirrored the behavior of a C246A mutant, shown to prevent DAG/PMA binding to ß2-chimaerin (Caloca et al., 2001).

At high PMA doses, the cells displayed lamellipodium-like extensions, a phenotype suggestive of Rac activation (Ridley et al., 1992). We considered that Rac-GTP levels might determine the extent of GFP-ß2-chimaerin membrane translocation. We thus analyzed PMA-dependent GFP-ß2-chimaerin translocation in cells expressing a constitutive active Rac mutant (RacV12). In the absence of additional stimulation, GFP-ß2-chimaerin localization did not change in RacV12-expressing cells, indicating that high levels of active Rac are not sufficient to induce membrane translocation (Fig. 2C, upper panel). Nonetheless, in RacV12-expressing cells, GFP-ß2-chimaerin relocated to the plasma membrane in response to the same, lower PMA dose that triggered relocation of the isolated C1 domain (Fig. 2C, middle panel). In RacV12-expressing cells, PMA-dependent translocation was not observed for the GFP-ß2-chimaerin F215G mutant, confirming the essential role of the C1 domain for membrane translocation in living cells (Fig. 2C, lower panel). All together, these results suggest that levels of active Rac cooperate with C1-dependent PMA binding to induce sustained GFP-ß2-chimaerin localization to the membrane.

View larger version (37K): [in this window] [in a new window]   Fig. 3. GFP-ß2-chimaerin-expressing Jurkat T cells show reduced PMA-induced F-actin polymerization. Jurkat T cells transfected with (A) GFP-ß2-chimaerin or (B) a GFP-coupled dominant-negative (RacN17) Rac mutant were stimulated with PMA (800 ng/ml, 5 minutes) or unstimulated (-), fixed and processed to determine GFP-ß2-chimaerin (A) or GFP-RacN17 (B) localization (green) and actin polymerization by Rhodamine-phalloidin staining (red). Control cells not expressing GFP constructs are indicated with an asterisk. Phase-contrast micrographs show the cells analyzed (left). Images are representative of the field observed for each condition. (C) Quantification of F-actin polymerization of experiments described in A and B was determined by analyzing intensity of fluorescence of Rhodamine-phalloidin staining. Values are the mean ± s.d. fluorescence intensity (arbitrary units) corresponding to 20 cells for each condition. Bars, 5 µm.

GFP-ß2-chimaerin inhibits CXCL12-induced cell spreading
Cell spreading is a flattening of the cell that increases the surface area of the contact region with the extracellular matrix, to allow stronger adhesion. Integrin-dependent spreading is observed following addition of CXCL12, a potent regulator of cytoskeletal remodeling which, through binding to a GPCR, induces PLCß-mediated DAG generation. To get further insight into the functional relevance of ß2-chimaerin activity, we next examined ß2-chimaerin-dependent regulation of integrin 4ß1-dependent spreading onto vascular cell adhesion molecule-1 (VCAM-1) in response to CXCL12. Addition of CXCL12 to control GFP-expressing cells induced characteristic spreading, with rapid lamellipodia-like extension, indicative of Rac activation (Fig. 4 and Movie 5 in supplementary material). Cell spreading was abolished in cells co-expressing GFP and the dominant-negative Rac mutant RacN17 (not shown) confirming previous data showing that Rac regulates integrin-mediated T-lymphocyte spreading and adhesion (D'Souza-Schorey et al., 1998). When analyzed, most GFP-ß2-chimaerin-expressing cells remained rounded after CXCL12 addition, and only a few cell projections were observed (Fig. 4 and Movie 5 in supplementary material). Spreading was observed in cells co-transfected with GFP-ß2-chimaerin and RacV12, indicating that the effect of this DAG receptor is due to modification of CXCL12-dependent Rac activation (Fig. 4).

View larger version (60K): [in this window] [in a new window]   Fig. 4. GFP-ß2-chimaerin-expressing Jurkat T cells show reduced CXCL12-dependent cell spreading to immobilized VCAM-1. Jurkat T cells were transiently transfected with vectors encoding GFP, GFP-ß2-chimaerin or GFP-ß2-chimaerin plus RacV12. After 24 hours, cells were suspended in HBSS and plated onto VCAM-1-coated slides. Slides were mounted on a 37°C plate and analyzed by confocal microscopy. CXCL12 (200 ng/ml) was added after the first frame and images captured every 10 seconds. Time-lapse confocal recording is shown in Movie 5 in supplementary material. Images are shown at the indicated times to illustrate cell spreading. Bar, 10 µm.

View larger version (30K): [in this window] [in a new window]   Fig. 5. GFP-ß2-chimaerin-expressing Jurkat T cells show reduced CXCL12/PMA-induced Rac activation and adhesion to immobilized VCAM-1. (A) Control Jurkat cells (JK) or Jurkat cells stably transfected with GFP-ß2-chimaerin (JKß2) were plated onto fibronectin and stimulated with CXCL12 (200 ng/ml, 60 minutes) or PMA (100 ng/ml, 30 minutes). Cells were lysed and Rac-GTP determined by pull-down assays with PAK-RBD-GST (see Materials and Methods). Samples were separated by SDS-PAGE and Rac-GTP levels determined by western blotting with anti-Rac antibody. One-tenth of each sample was analyzed for total Rac level and ß2-chimaerin expression. (B) Jurkat cells were transfected with GFP or GFP-ß2-chimaerin-expressing vectors; GFP-positive cells were sorted after 24 hours. Sorted cells were added to wells coated with BSA or sVCAM-1, either alone (medium) or co-immobilized with CXCL12. Where indicated, cells were stimulated with PMA (100 ng/ml, 5 minutes) before addition to sVCAM-1-coated wells. Cells were allowed to adhere at 37°C for 2 minutes. Unbound cells were washed and the extent of adhesion determined as described (see Materials and Methods). Data represent the mean ± s.d. of at least three independent experiments performed in triplicate.

The 4ß1 integrin has an important role in T-cell migration to peripheral lymph nodes and infected tissue, mediating the capture and subsequent firm adhesion of the migrating lymphocytes to the vascular wall. We next examined ß2-chimaerin-dependent regulation of integrin 4ß1-dependent adhesion to VCAM-1. We observed no apparent differences between basal integrin 4-dependent adhesion in cells expressing GFP-ß2-chimaerin or control GFP. As reported for Jurkat cells (Feigelson et al., 2001), CXCL12 markedly increased adhesion of control GFP-expressing cells to VCAM-1, whereas the HP1/2 anti-4 antibody completely blocked the adhesion (not shown). ß2-chimaerin expression abolished any CXCL12-stimulated adhesion to VCAM (Fig. 5B). This was also the case for PMA-induced cell adhesion, which suggests an important role for DAG/PMA-dependent ß2-chimaerin regulation in integrin-induced adhesion.

View larger version (27K): [in this window] [in a new window]   Fig. 6. Effect of a GFP-ß2-chimaerin GAP-defective mutant on CXCL12/PMA-induced adhesion. (A) Vectors encoding GFP, GFP-ß2-chimaerin or a ß2-chimaerin GAP-defective mutant bearing a deletion of the EIE sequence (GFP-EIE) were transfected into HEK293 cells. At 24 hours post-transfection, cells were lysed and Rac-GTP levels determined by pull-down assays with PAK-RBD-GST. One-tenth of each sample was analyzed for total Rac level and ß2-chimaerin expression. (B) Jurkat T cells were transfected with the indicated plasmids; cells were sorted 24 hours post-transfection and GFP-positive cells tested for adhesion as in Fig. 5. Data represent the mean ± s.d. of at least three independent experiments performed in triplicate.

View larger version (26K): [in this window] [in a new window]   Fig. 7. Effect of GFP-ß2-chimaerin on CXCL12/PMA-induced adhesion of primary blood T lymphocytes. Human peripheral blood mononuclear cells were prepared, and T lymphocytes were isolated and nucleofected as described in the Materials and Methods. After 6 hours, cells were added to wells coated with sVCAM-1 alone (medium), or co-immobilized with CXCL12 and allowed to adhere (37°C, 2 minutes). Where indicated, cells were stimulated with PMA (100 ng/ml, 5 minutes) before addition to the wells. The inset shows endogenous ß2-chimaerin expression in cell lysates of Jurkat and hPBLs, determined by western blotting. Tubulin expression was measured as a control of total protein levels. Data represent the mean ± s.d. of two independent experiments performed in triplicate.

GFP-ß2-chimaerin expression in primary T cells decreases CXCL12- and PMA-mediated adhesion to VCAM
We next evaluated the functional effect of ß2-chimaerin and the GAP-inactive mutant in primary human peripheral blood T lymphocytes (PBLs). Primary PBLs expressed endogenous chimaerin, although expression was lower than that observed in Jurkat T cells (Fig. 7, inset). As for Jurkat cells, GFP-ß2-chimaerin expression in primary T cells severely impaired CXCL12-induced adhesion to VCAM-1 (Fig. 7), confirming a role for ß2-chimaerin as a negative modulator of chemokine-induced integrin-dependent adhesion. Again, and as described above for Jurkat cells, the GAP-inactive mutant did not affect chemokine-regulated adhesion. Prevention of adhesion by the wild-type protein and lack of effect of the GAP mutant were also observed following PMA stimulation, suggesting that regulation of the CXCL12-dependent effect is mediated by DAG generation.

Role of the C1 domain in CXCL12/PMA-dependent regulation of ß2-chimaerin
The previous results demonstrated that, via its Rac-GAP activity, GFP-ß2-chimaerin acts as a potent attenuator of CXCL12-induced cell spreading and adhesion. Videomicroscopy analysis of GFP-ß2-chimaerin and the mutants (see Fig. 2A) suggest that, as indicated by structural studies (Canagarajah et al., 2004), the ß2-chimaerin C1 domain is not exposed in the native conformation. This renders a protein that requires a considerable conformational change to bind DAG, raising a question about the functional relevance of this particular domain on CXCL12-regulated cell adhesion and spreading.

View larger version (19K): [in this window] [in a new window]   Fig. 8. CXCL12- and PMA-dependent regulation of ß2-chimaerin requires a functional C1 domain. (A) Jurkat T cells were transfected with a GFP-ß2-chimaerin C1 domain mutant (GFP-F215G). At 24 hours post-transfection, cells were sorted and GFP-positive cells tested for PMA-induced adhesion to sVCAM-1 as in Fig. 5. Data represent the mean ± s.d. of two independent experiments performed in triplicate. (B) Jurkat cells were transfected with GFP-ß2-chimaerin F215G and plated onto VCAM-1-coated slides, mounted on a 37°C plate and analyzed by confocal microscopy. CXCL12 (200 ng/ml) was added after the first frame and images are shown at the indicated times to illustrate cell spreading. Time-lapse confocal recording is shown in Movie 6 in supplementary material. (C) Jurkat T cells transfected with the indicated plasmids were sorted and GFP-positive cells were tested for CXCL12-induced adhesion to sVCAM-1 as in Fig. 5. Data represent the mean ± s.d. of two independent experiments performed in triplicate. Bar, 5 µm.

Jurkat T cells stably transfected with GFP-ß2-chimaerin show enhanced CXCL12-dependent chemotaxis
CXCL12 is a potent chemotactic agent for T lymphocytes. To study the role of ß2-chimaerin in chemokine-regulated cell migration, we generated stable Jurkat cell transfectants expressing GFP-ß2-chimaerin or GFP as controls, and analyzed CXCL12-induced chemotaxis in a Transwell assay. GFP-ß2-chimaerin levels were readily detected in western blot using anti-chimaerin antibodies, and CXCR4 expression levels were comparable in transfected and parental cell lines, as measured by flow cytometry (not shown). As in transiently transfected cells, CXCL12-induced cell adhesion to VCAM was severely impaired (not shown). When CXCL12-induced chemotaxis was examined at different times, GFP-ß2-chimaerin expressing cells showed increased migration compared to parental Jurkat cells (Fig. 9A). A CXCL12 dose-response curve showed enhanced migration of GFP-ß2-chimaerin cells compared with parental cells from the lowest CXCL12 concentrations tested (Fig. 9B).

View larger version (26K): [in this window] [in a new window]   Fig. 9. GFP-ß2-chimaerin expression enhances CXCL12-dependent migration of Jurkat T cells. (A) The chemotactic response of parental (control) and GFP-ß2-chimaerin cells towards CXCL12 (200 ng/ml) was determined in a 5-µm-pore chemotaxis chamber (see Materials and Methods). Data are expressed as a percentage of input cells, and are shown as the mean ± s.d. of at least three independent experiments. (B) Chemotactic responses of parental (control) and GFP-ß2-chimaerin-expressing cells towards CXCL12 at the concentrations indicated were determined at 1 hour post-stimulation. Data are expressed as in A, showing the mean ± s.d. of at least three independent experiments. (C) The chemotactic response of parental, GFP- and GFP-ß2-chimaerin-expressing cells towards CXCL12 (200 ng/ml) was determined 1 hour post-stimulation. Before seeding into the chamber, cells were incubated with the indicated concentrations of U73122 or EGTA (30 minutes) or with PTx (100 ng/ml overnight) at 37°C. Data are expressed as in (A), and are shown as the mean ± s.d. of at least three independent experiments.

	Discussion

Top Summary Introduction Results Discussion Materials and Methods References

 
The identification of non-PKC types of DAG-binding proteins has provided new functions for this lipid messenger in the regulation of cell responses. In mammals, there are five alternative non-PKC families of DAG/PE target proteins: protein kinase D (PKD), diacylglycerol kinases (DGK) ß and , Ras guanyl nucleotide-releasing proteins (RasGRP), chimaerins and mammalian uncoordinated 13 (Munc13). Non-PKC DAG-binding proteins are reported to be essential for T-cell functions: RasGRP1, a DAG-modulated GEF for Ras, is required for positive selection in the thymus and correct homeostasis of mature T cells (Ebinu et al., 1988; Priatel et al., 2002); recent experiments in genetically modified animals also revealed that PKD exerts important and divergent effects in T-cell differentiation (Marklund et al., 2003). The existence of distinct families of DAG-receptors in T lymphocytes reinforces the essential role of PLC-dependent DAG generation in the regulation of T-cell responses, increasing the complex network of DAG-modulated signals in this particular cell type. Chimaerins were the first high-affinity non-PKC DAG-binding proteins discovered, although endogenous mRNA expression had been reported only in the nervous system. In the present study we demonstrate for the first time endogenous chimaerin expression in T cells, and provide evidence of its role as a direct modulator of CXCL12-dependent adhesion and migration of T lymphocytes.

Here we show that ß2-chimaerin expression in T cells reduces CXCL12-induced Rac activation and concomitantly diminishes CXCL12-induced 4ß1-mediated cell spreading and adhesion to immobilized VCAM-1. This effect is lost in cells expressing ß2-chimaerin GAP-inactive form. These experiments demonstrate that GAP activity is essential for ß2-chimaerin biological activity in T lymphocytes, at difference from the suggested requirements for 2-chimaerin function (Hall et al., 2001). Whereas adhesion in most cell types depends strictly on Rac activation, the exact mechanism that governs chemokine-induced adhesion of T lymphocytes is a question of active debate. Several reports have shown that chemokines upregulate integrin-dependent adhesion by PI3K-independent mechanisms, questioning the relevance of Rac activation in regulating this process. Our results showing the potent effect of ß2 chimaerin expression on CXCL12-dependent cell spreading and adhesion to VCAM-1 throw new light on the essential role of Rac in the regulation of this process. Similar effects to those described for Jurkat T cells are observed in primary lymphocytes, confirming that they are not specific for this particular cell line. Moreover, the lack of effect in cells expressing a ß2-chimaerin mutant defective for DAG binding indicates the essential role of this lipid for correct ß2-chimaerin function. DAG-dependent modulation of Rac-GTP levels thus represents another example of the broad regulatory role of DAG in the regulation of lymphocyte responses.

Discrimination of DAG-dependent signals by distinct types of DAG-binding proteins represents a complex biological problem. Our studies using GFP-tagged proteins expressed in living lymphocytes provide some clues for the specific regulation of this particular DAG receptor. PMA addition to intact lymphocytes induces translocation to the membrane of the GFP-fused ß2-chimaerin C1 domain: a domain shown to bind PMA with high affinity in vitro (Caloca et al., 1997). Membrane localization of the full-length GFP-ß2-chimaerin in living T cells, nonetheless, requires higher PMA doses than those needed for membrane localization of other C1-containing proteins such as PKC, RasGRP1 or even its own isolated C1 domain. The crystal structure of ß2-chimaerin reveals that, in the inactive state, the N-terminal domain maintains a `closed' conformation through intramolecular interactions, sterically blocking Rac binding and concealing the C1 domain in the absence of additional stimuli (Canagarajah et al., 2004). Accordingly, mutations in residues reported as responsible for maintaining the C1 domain hidden and not readily available to DAG/PMA-binding (Q32 and I130) render a protein that responds to PMA stimulation as efficiently as the isolated ß2-chimaerin C1 domain. These results in living T lymphocytes fully support the structural data and suggest a model where ß2-chimaerin would exist in equilibrium between an open/membrane-associated and closed/cytosolic conformation. Membrane localization would be favored by RacGAP-domain-dependent binding to Rac and C1-mediated DAG/PMA interaction. Accordingly, our experiments demonstrate that PMA-dependent translocation of GFP-ß2-chimaerin is more effective in cells expressing constitutive active Rac. In addition, GFP-ß2-chimaerin translocation, observed at higher PMA concentration, could be the result of PMA-dependent displacement of this equilibrium to the ß2-chimaerin membrane-bound form. This cooperation between DAG/PMA binding and active Rac, as a result of the domain combination present only in the chimaerin family, would provide a mechanism for discriminating among distinct DAG-dependent cellular signals. This type of mechanism would also ensure a feedback regulation mode in which ß2-chimaerin relocates to membranes, stimulating Rac-GTP hydrolysis, in response to Rac-GTP levels.

Although the ß2-chimaerin-occluded C1 domain is relatively inefficient for protein translocation, our results demonstrate that a mutation that impairs its membrane binding, fully prevents the ß2-chimaerin effect on CXCL12-dependent responses. These data, again in agreement with the crystal structure, suggest that the C1 domain is essential for ß2-chimaerin Rac-GAP function, probably disrupting the steric inhibition on the Rac-binding site.

Further experiments should help to clarify the sequence of events responsible for ß2-chimaerin transition from a closed to an open/active conformation. Type 2 chimaerins contain an `atypical' N-terminal SH2 domain, highly conserved in and ß isoforms, which includes a Glu residue replacement of the invariant Trp in the N-terminus of the `classical' SH2 domains (Hall et al., 1993; Leung et al., 1994). This suggests that the ß2-chimaerin SH2 domain may not bind to tyrosine-phosphorylated residues in other proteins. Accordingly, our preliminary studies did not detect association of ß2-chimaerin with tyrosine-phosphorylated proteins following CXCL12 stimulation. A role for the SH2 domain maintaining the inactive conformation of chimaerins would explain previous studies in neuroblastoma cells showing the distinct functional effects and subcellular localization of 1 and 2 chimaerin isoforms (Hall et al., 2001).

Whereas ß2-chimaerin-expressing cells showed a marked decreased in CXCL12-induced spreading and adhesion to immobilized integrins, they migrated more efficiently in response to the chemokine. CXCL12-mediated migration of ß2-chimaerin-expressing cells was PTx-sensitive, as was the case for parental cells, indicating CXCR4/G_i protein-mediated signaling. The migratory advantage of ß2-chimaerin-expressing cells was very sensitive to inhibition of PLC activity, even at inhibitor concentrations that did not affect parental cell migration. As modulation of Ca²⁺ elevation did not affect cell migration, our results show that the ß2-chimaerin-enhanced effect on cell migration depends directly on CXCL12-induced DAG elevation by a G_i-coupled PLC-dependent mechanism.

Chemokine binding to its receptor triggers a series of signaling events that result in directional cell migration toward the chemoattractant source. The migrating cell is characterized by an actin-rich membrane protrusion at the leading edge and a tail structure at the trailing edge. This morphologic polarization involves dynamic actin polymerization at the cell front and actin disassembly behind the leading edge, as well as cell body contraction and retraction of the trailing edge (Sanchez-Madrid and del Pozo, 1999). The signaling events that regulate directional migration have been studied thoroughly. The front edge requires high Rac activity that promotes assembly of F-actin networks, whereas Rho activity and actomyosin array assembly are crucial at the trailing edge (Xu et al., 2003). The activities of Rho and Rac negatively regulate one another; Rho activity at the trailing edge inhibits protrusive lamellipodium formation at this site, promoting the development of a single leading edge (Worthylake and Burridge, 2003). In addition, during migration, actin cytoskeletal remodeling must be coordinated with cycles of integrin-mediated cell-substratum attachment at the leading edge and detachment at the trailing edge. RhoA and Rho-kinase promote migration in Jurkat T cells and neutrophils by inhibiting actin cytoskeleton-dependent cell adhesion and integrin activation outside the leading edge (De La Roche et al., 2002; Worthylake and Burridge, 2003). This dynamic antagonism between Rac and Rho signaling pathways maintains cell polarization and allows directional responsiveness to the chemoattractant. The spatial and temporal activation of these two proteins, and therefore their GEFs and GAPs, requires precise regulation. Our results suggest a model in which ß2-chimaerin-dependent downmodulation of Rac in response to DAG elevation could be implicated in maintaining the dynamic equilibrium between `front' and `back' signals, thus enhancing the migratory response. PLCß is proposed to be stimulated by active Rac (Snyder et al., 2003.); this suggests a negative-feedback loop in which Rac-dependent PLC activation induces elevation of DAG levels, leading to ß2-chimaerin activation and subsequent downregulation of Rac activity behind the leading edge.

In conclusion, our findings demonstrate for the first time a functional role for ectopically expressed ß2-chimaerin in the regulation of chemokine-modulated responses in T lymphocytes. The spatial restriction imposed by the C1 domain allows strict regulation of the ß2-chimaerin Rac-GTPase-activating function, which acts as a positive modulator of polarized migration while reducing chemokine-induced integrin adhesion. Additional experiments aimed to knock down endogenous protein will help to provide further insight into the physiological relevance of ß2-chimaerin in the regulation of T-cell functions.

	Materials and Methods

Top Summary Introduction Results Discussion Materials and Methods References

 
Reagents and antibodies
Orthovanadate, phenylmethylsulfonyl fluoride (PMSF), poly-DL-lysine, paraformaldehyde (PFA), Igepal CA-630 (NP40), human fibronectin, phorbol 12-myristate 13-acetate (PMA), EGTA and pertussis toxin (PTx) were from Sigma (St Louis, MO). Leupeptin and aprotinin were purchased from Roche (Basel, Switzerland). CXCL12 was from Peprotech (London, UK), U73122 from Calbiochem (San Diego, CA) and recombinant human VCAM-1/Fc chimera from R&D Systems (Minneapolis, MN). BODIPY TR ceramide, Rhodamine-phalloidin and BCECF-AM [2',7'-bis(carboxyethyl)-5(6')-carboxyfluorescein-acetoxymethyl ester] were from Molecular Probes (Leiden, The Netherlands). The anti-ß2-chimaerin monoclonal antibody (mAb) was generated as described (Wang and Kazanietz, 2002). Anti -tubulin was from Sigma (St Louis, MO) and anti-Rac1 from BD Transduction Laboratories (Lexington, KY). The anti-CXCR4-01 mAb was generated as described (Vila-Coro et al., 1999). Horseradish peroxidase (HRP)-conjugated goat anti-mouse and anti-rat IgG were from Dako (Glostrup, Denmark). The HP1/2 anti-4 mAb was a gift from F. Sánchez-Madrid (Hospital de la Princesa, Madrid, Spain). Mouse monoclonal anti-EE Ig were kindly provided by P. Hawkins (Babraham Institute, Cambridge).

RT-PCR
RNA was prepared from Jurkat cells using the guanidinium-isothiocyanate acid phenol method. First-strand cDNA synthesis was done using random primers. PCR was performed with primers 5'-TGCTGAAGAATACCAGCCTC-3' and 5'-TTGGAATAGGTATCATATGTG-3'.

Recombinant plasmids
Generation of plasmid encoding EGFP-ß2-chimaerin wt and EIE mutant have been described (Caloca et al., 1999; Caloca et al., 2003). For T-lymphocyte transfection, ß2-chimaerin cDNA was excised from CMV plasmids and subcloned at the EGFP C-terminus of the pEFbosEGFP vector modified from pEGFP (Clontech) as described (Sanjuan et al., 2001). The GFP-fused construct of ß2-chimaerin C1 domain was generated as described (Carrasco and Merida, 2004). The ß2-chimaerin F215G mutant was generated by site-directed mutagenesis using the QuikChange mutagenesis kit from Stratagene (La Jolla, CA). We used pEFbosEGFP-ß2-chimaerin as a template, and generated the replacement of Phe215 by Gly with the primer 5'-GAGAAGACACACAACGGTAAGGTCCACACGTTC-3' (mutated codon underlined). The resulting construct (GFP-ß2-chimaerin-F215G) was confirmed by sequencing. EGFP-ß2-chimaerin Q32A and I130A mutants were previously described (Canagarajah et al., 2004). Plasmids encoding -chimaerins, used as RT-PCR controls, were a kind gift of C. Hall (Institute of Neurology, London, UK). Plasmids encoding constitutive active (RacV12) and dominant-negative (RacN17) plasmids in-frame with the N-terminal EE-tag epitope were a generous gift of P. Hawkins (The Babraham Institute, Cambridge, UK). Plasmid encoding RacN17 in-frame with EGFP was a gift of F. Sánchez-Madrid (Hospital de la Princesa, Madrid, Spain). GST-RBD Pak was a gift of A. Berns (The Netherlands Cancer Institute, Amsterdam, The Netherlands).

Cell culture and transfection
Human leukemia and lymphoma cell lines were obtained from the American Type Culture Collection (Manassas, VA). Jurkat and Molt-4 are cell lines derived from acute T-cell leukemias; Karpas 299 and SUDHL-1 were derived from anaplastic large cell lymphomas (ALCL), which have the t(2;5)(p23;q35) translocation, resulting in the NPM-ALK fusion gene. HDLM2 is a Hodgkin/Reed-Sternberg (H-RS) cell line of T-lymphocyte origin. Toledo was derived from a non-Hodgkin's B-cell lymphoma. HT cells were derived from a human B-cell lymphoma, and Namalwa from a Burkitt's lymphoma. Jurkat cells were cultured in DMEM (BioWhittaker, Walkersville, MD) supplemented with 10% fetal bovine serum (FBS; Sigma), 2 mM glutamine, 10 mM HEPES, and 100 U/ml each penicillin and streptomycin. The remaining cell lines were maintained in RPMI 1640 (BioWhittaker) supplemented as for Jurkat cells. Cells in logarithmic growth phase were transfected (1.2x10⁷ in 400 µl complete medium) with 20 µg plasmid DNA by electroporation in a Gene Pulser (Bio-Rad, Hercules CA; 270V, 975 µF). Cells were immediately transferred to 10 ml complete medium and assayed after 24 hours. To generate stable GFP-ß2-chimaerin transfectants, Jurkat cells were transfected and sorted to obtain GFP-positive cells (Altra Hypersort, Beckman-Coulter, Miami, FL). Stably transfected populations were maintained in G418 selection medium. HEK293 cells were maintained in DMEM medium supplemented as for Jurkat cells. Cells were transfected using jetPEI reagent (Polyplus transfection SAS, Illkirch, France) according to the manufacturer's instructions and assayed 24 hours later.

Human peripheral blood mononuclear cells were prepared from buffy coats using a Ficoll density gradient. T lymphocytes were isolated by negative selection using anti-CD14-conjugated microbeads and MACS separation columns (Miltenyi Biotec, Bergisch Gladbach, Germany) to eliminate monocytes, followed by additional negative selection with Dynabeads conjugated with anti-CD19 (Dynal ASA, Oslo, Norway) to remove B lymphocytes. Purity was >95% for each sample, as analyzed by flow cytometry using the T3b anti-CD3 mAb. For nucleofection, isolated human T lymphocytes were washed with PBS/EDTA 0.3 mM and resuspended in T-cell Nucleofector solution at 5x10⁶ cells/100 µl. Cells were nucleofected with 5 µg plasmid DNA using an Amaxa Nucleofector (Amaxa, Cologne, Germany). After transfection, cells were transferred immediately to complete medium and assayed after 6 hours. GFP expression by transfected cells was analyzed by flow cytometry.

Western blotting
Cells were lysed in lysis buffer (20 mM Tris-HCl pH 7.5, 150 mM NaCl, 5 mM EDTA, 5 mM sodium pyrophosphate, 1 mM Na₃VO₄, 1% Nonidet P-40, 1 mM PMSF and 10 µg/ml each aprotinin and leupeptin) for 15 minutes on ice. After centrifugation (15,000 g, 15 minutes, 4°C), supernatants were assayed for total protein (DC protein assay, Bio-Rad). Equivalent amounts of proteins were separated by SDS-PAGE and transferred to nitrocellulose membranes (Hybond, Amersham Pharmacia Biotech). Membranes were blocked, and blots subsequently developed with the indicated antibodies and HRP-conjugated anti-rat or anti-mouse antibodies using the ECL detection kit (ECL; Amersham Pharmacia Biotech) according to the manufacturer's protocols.

Rac activation assay
A pull-down assay was used to isolate Rac-GTP by binding to the RBD (Rac-binding domain) of PAK-1 (Benard et al., 1999). Jurkat T cells attached to fibronectin-coated plates (10 µg/ml) or HEK293 cells transfected and stimulated as indicated in the figure legends, were lysed in a buffer containing 10% glycerol, 50 mM Tris-HCl pH 7.4, 100 mM NaCl, 1% NP-40, 2 mM MgCl₂ and protein inhibitors. Lysates were incubated with a GST fusion protein of RBD-PAK-1 in the presence of glutathione-Sepharose 4B beads (1 hour, 4°C). After extensive washing, bound proteins were eluted in loading buffer. PAK-GST-associated Rac was detected with an anti-Rac1 mAb; 30 µg lysate was probed with the same antibody to detect total Rac.

Chemotaxis assay
Parental or stably transfected Jurkat T cells (5x10⁵ in 0.1 ml complete medium) were placed in the upper well of 5-µm-pore Transwells (Costar, Cambridge, MA). The lower well contained 500 µl complete medium alone or with CXCL12 (200 ng/ml or as indicated). Plates were incubated (37°C) for the times indicated, and cells that migrated to the bottom chamber were counted on a flow cytometer for 30 seconds. Cell migration was calculated as a percentage of input cells. For drug treatment, before the chemotaxis assay cells were incubated in complete medium with U73122 (1-5 µM) or EGTA (0.1-1 mM) at 37°C for 30 minutes or with PTx (100 ng/ml; 37°C, overnight).

Adhesion assays
Jurkat T cells were transfected with the GFP-fused constructs and sorted 24 hours later to recover GFP-positive cells. Plates (96-well, high binding; Costar, Cambridge MA) were coated with soluble recombinant human VCAM-1 consisting of domains 1-4 fused to the Fc portion of IgG1 (Munoz et al., 1996) (sVCAM-1, 0.7 µg/ml), alone or co-immobilized with CXCL12 (650 ng/ml). Before addition to wells, GFP-positive Jurkat cells were labeled with BCECF-AM. Cells were resuspended in adhesion medium (RPMI 1640, 0.5% BSA) and added to wells (6x10⁴ cells in 100 µl) in triplicate. After a 15 second centrifugation, plates were incubated (37°C, 2 minutes) and unbound cells were removed by washing with medium. Adhered cells were lysed with 0.1% SDS in PBS and the extent of adhesion quantified in a fluorescence analyzer (Polarstar Galaxy; BMG Labtechnologies, Offenburg, Germany). Where indicated, labeled cells were stimulated with PMA (100 µg/ml, 37°C, 5 minutes) before addition to wells. For transfected human T lymphocytes, after the adhesion step and washes, bound cells were detached, counted and analyzed by flow cytometry to determine the percentage of GFP-positive adhered cells.

Confocal microscopy
For immunofluorescent staining, at 24 hours post-transfection with indicated plasmids, cells were transferred to poly-DL-lysine-coated coverslips and allowed to attach for 15 minutes. Where indicated, attached cells were stimulated with PMA (800 ng/ml, 5 minutes). Cells were then fixed with 2% PFA (10 minutes), washed, permeabilized with PBS 0.2% Triton X-100 and stained with the indicated fluorescent marker. We used fluorescent BODIPY TR ceramide for Golgi detection and Rhodamine-phalloidin for F-actin staining according to manufacturer's protocols. For EE-RacV12 detection, cells were stained with an anti-EE IgG followed by anti-mouse IgG coupled to Cy3. Fluorescence was analyzed on a Leica confocal microscope (TCS-NT, Wetzlar, Germany) and images processed using ImageJ software.

For imaging of living cells, at 24 hours post-transfection, cells were suspended in HBSS (HEPES-balanced salt solution) and plated on poly-DL-lysine or human VCAM-1 (1 µg/ml, R&D Systems) coated chamber slides. Slides were mounted on a 37°C plate; PMA or CXCL12 were added at indicated concentrations after the first frame and images captured every 10 seconds.

	Acknowledgments

 
We thank M. Mellado, J. M. Rodríguez and J. Stein for critical assessment of the manuscript and I.M. group members for helpful discussions, C. Moreno-Ortiz and A. Moreno for flow cytometry and sorting, and C. Mark for excellent editorial assistance. This work was supported by grants BFU2004-01756 from the Spanish Ministry of Science, GR/SAL/0655/2004 from the Madrid regional government and G03/179 from the Ministry of Health. M.S. and S.C. are fellows of the Spanish Ministry of Education; E.D. is a fellow of the Spanish Ministry of Science. The DIO was founded and is supported by the Spanish Council for Scientific Research (CSIC) and by Pfizer. The authors declare that they have no competing financial interests.

	Footnotes

 
Supplementary material available online at http://jcs.biologists.org/cgi/content/full/119/1/141/DC1

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