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The Journal of Immunology, 2003, 171: 2225-2232.
Copyright © 2003 by The American Association of Immunologists

CD28 Engagement Promotes Actin Polymerization Through the Activation of the Small Rho GTPase Cdc42 in Human T Cells

Laura Inés Salazar-Fontana1,*, Valarie Barr{dagger}, Lawrence E. Samelson{dagger} and Barbara E. Bierer2,*

Laboratories of * Lymphocyte Biology, National Heart, Lung, and Blood Institute, and {dagger} Cellular and Molecular Biology, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, MD 20892.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Engagement of the costimulatory molecule CD28 is an important step in the optimal activation of T cells. Nevertheless, the specific role of CD28 in the formation of the immunological synapse and cytoskeletal changes that occur upon TCR/CD3 complex engagement is still poorly understood. Using Ab-coated surfaces, we show that CD28 engagement in the absence of any other signal induced the formation of cytoplasmic elongations enriched in filamentous actin (F-actin), in this work called filopodia or microspikes. Such structures were specific for engagement of CD28 on mAb-coated surfaces because they could not be observed in surfaces coated with either poly(L-lysine) or anti-CD3 mAb. The signaling pathway coupling CD28 to cytoskeletal rearrangements required Src-related kinase activity and promoted Vav phosphorylation and Cdc42 activation independently of the {zeta}-chain-associated kinase (ZAP-70). CD28-induced filopodia required Cdc42 GTPase activity, but not the related Rho GTPase Rac1. Moreover, Cdc42 colocalized to areas of increased F-actin. Our results support a specific role for the activation of the small Rho GTPase Cdc42 in the actin reorganization mediated by CD28 in human T cells.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
T cell activation requires the relocalization and clustering of Ag and costimulatory receptors at the area of contact between the T cell and the APC (1). The T cell/APC conjugate needs to be stabilized and sustained for a limited period of time to initiate and direct T cell effector functions (2). Once the T cell contacts the APC, a reorganization of different receptors and signaling molecules takes place at the area of cell-cell contact. The assembly of T cell signaling molecules is intimately related to the reorganization of the actin/myosin and microtubule cytoskeleton. The signaling pathway relating the TCR/CD3 complex to actin reorganization is dependent on the actin-regulatory molecules Vav and Wiskott-Aldrich syndrome protein (WASp), 3 which in turn control the activation of Rho GTPases (3).

It is now generally accepted that CD28 is an important costimulator for the TCR/CD3 complex promoting a sustained signaling cascade that will allow the T cell to proliferate and function. CD28 is a 44-kDa protein present on the surface of T cells; its engagement, either with Abs or its ligands, CD80 (B7.1) or CD86 (B7.2), allows optimal cytokine production, survival, and expansion of naive T cells, and is critical to the prevention of T cell unresponsiveness or anergy (4). The biochemical pathways affected by CD28 engagement have been studied extensively, although unique and specific signals mediated by CD28 and not by the TCR/CD3 complex have been difficult to identify. CD28 triggering induces tyrosine phosphorylation of its cytoplasmic domain by the Src kinase Lck and the subsequent binding of the regulatory subunit of the phosphoinositide 3-kinase (PI3K) (5). This initial event is followed by activation of Tec kinases and tyrosine phosphorylation of the adaptor molecule linker for activation of T cells, Grb2, Vav, SLP-76, p62dok, the serine-threonine phosphorylation of p21-activated kinase (PAK), and protein kinase B or Akt (6, 7, 8). These biochemical events are not unique to CD28 because they also occur following engagement of the TCR/CD3 complex alone. Recent reports have shown that ZAP-70 activity is not required for CD28-mediated phosphorylation of linker for activation of T cells, Vav, and SLP-76 (6, 7, 9, 10), suggesting a distinct and separable signaling pathway for CD28 from the TCR/CD3 complex. These CD28-mediated biochemical events have been correlated with cytokine expression (e.g., IL-2); however, their role in modifying the actin cytoskeleton and in mediating initial CD28-mediated adhesion contacts in human T cells remains poorly understood.

Small Rho GTPases, namely Cdc42, Rac, and Rho, form a subgroup of the Ras superfamily of 20- to 30-kDa GTP-binding proteins that have been associated with the remodeling of the actin cytoskeleton in many cell types, including T lymphocytes (11). In the hemopoietic system, Rho and Rac activities are essential during T cell development (12), in chemotaxis (13) and integrin adhesion (14). Cdc42 in T cells has been related to polarization toward the APC (15), but its role in other aspects of T cell biology remains unclear.

All GTPases exist in an inactive GDP-bound and an active GTP-bound conformation, regulated by GTPase-activating proteins and guanine nucleotide exchange factors (GEFs), respectively (16). GTP binding to Rho proteins induces a conformational change that increases the affinity for effector molecules containing a GTPase binding domain (16). Vav is a GEF for both Cdc42 and Rac1 (17) and is tyrosine phosphorylated following both TCR/CD3 complex and CD28 engagement (5). Studies in Vav knockout mice have shown that T cells from these animals suffer defects in actin polymerization and capping formation upon TCR/CD3 engagement that can be corrected by transfection of an activated form of Rac1 (18).

In the present study, we show that CD28 engagement promoted the formation of filopodial extensions enriched in filamentous actin (F-actin) in human CD4+ T cells. We show further that the signaling pathway leading to the formation of CD28-induced filopodia and Cdc42 activation required the activation of the Src kinase p56Lck and/or p59Fyn, but was independent of the Syk kinase family member, ZAP-70. Transfection experiments demonstrated that CD28-induced microspike formation was mimicked by expression of a constitutively active Cdc42, but not Rac1, molecule. CD28 appears to mediate actin polymerization in human T cells through the activation of the Rho GTPase Cdc42. The role of CD28 in T cell adhesion and in secretion of cytokines is further discussed.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell lines and reagents

Human T cell lines were cultured at 37°C in 5% CO2 in RPMI 1640 (MediaTech, Herndon, VA) supplemented with 10% heat-inactivated FCS (Life Technologies, Gaithersburg, MD), 2 mM L-glutamine, 10 mM HEPES, pH 7.2, 100 U/ml penicillin (MediaTech), 100 µg/ml streptomycin (MediaTech), and 50 µM 2-ME (Bio-Rad, Hercules, CA), termed complete RPMI. The Jurkat T cell leukemia cell line (clone J77) was the gift of K. Smith (Cornell University, New York, NY). The ZAP-70/Syk-deficient clone, P116 (ZAP-70-), was the gift of R. Abraham (Duke University, Durham, NC); the Jurkat variant JCaM1.6 (American Type Culture Collection, Manassas, VA) is Lck deficient, but retains low expression of the Src kinase p59Fyn. These cell lines, as well as the JcaM1 cells reconstituted with either wild-type (WT) Lck (JcaM1/Lck) or WT Fyn (JcaM1/Fyn), both gifts from D. Straus (University of Chicago, Chicago, IL), were grown in complete RPMI 1640. All cell lines were mycoplasma negative, verified by routine screening with the mycoplasma set primer (Stratagene, La Jolla, CA).

Resting human PBLs were isolated from adult healthy donors by density-gradient centrifugation on Ficoll-Paque PLUS (Amersham Pharmacia Biotech, Uppsala, Sweden), resuspended in complete RPMI, and used within 24 h of isolation. T cells blasts were generated by incubating 5 x 106 PBLs for 3 days in the presence of soluble anti-CD3 (25 µg/ml) and anti-CD28 (40 ng/ml) mAbs. After this time, cells were washed and resuspended in complete RPMI and rested overnight before use. Human CD4+ T cells were obtained by using human CD4 enrichment columns from R&D Systems (Minneapolis, MN). Purity was evaluated by flow cytometry, and the proportion of CD4+ cell was always greater than 95%.

The anti-human CD28 mAb 9.3, the gift of C. June (University of Pennsylvania, Philadelphia, PA), was used as purified ascites fluid. The anti-Vav (H-211), anti-Cdc42 (P1), anti-Lck (3A5), and anti-Fyn (15) were all purchased from Santa Cruz Biotechnology (Santa Cruz, CA); the anti-phosphotyrosine mAb 4G10 was purchased from Upstate Biotechnology (Lake Placid, NY). The anti-GFP mouse mAb was obtained from Clontech, BD Biosciences (Palo Alto, CA).

Cell stimulation and immunoprecipitation

Before stimulation, cells were washed twice with RPMI 1640 without additives and then incubated for 10 min at 37°C before stimulation. In brief, 1 x 107 cells were left unstimulated or stimulated for the indicated times with 1 µg of anti-CD28 mAb 9.3. Cells were then pelleted and lysed by addition of 1 ml of ice-cold lysis buffer (1% Nonidet P-40, 150 mM NaCl, 25 mM Tris-HCl, pH 7.5, 1 mM EDTA, 1 mM sodium orthovanadate, 10 µg/ml leupeptin, 10 µg/ml aprotinin, and 1 mM PMSF) and kept on ice for 20 min. Cell lysates were clarified by centrifugation at 14,000 x g for 10 min at 4°C. For immunoprecipitation, precleared cell lysates were incubated for 2 h at 4°C with the appropriate antisera and 20 µl of protein A-agarose. The resin was then washed four times with ice-cold washing buffer (0.1% Nonidet P-40, 150 mM NaCl, 25 mM Tris-HCl, pH 7.5, 1 mM sodium orthovanadate, 10 µg/ml leupeptin, 10 µg/ml aprotinin, and 1 mM PMSF). Proteins were boiled in SDS sample buffer, separated by SDS-PAGE, transferred to polyvinylidene difluoride (Millipore, Bedford, MA), and probed with primary Abs, followed by HRP-conjugated secondary Abs (Amersham Pharmacia Biotech). Polypeptides recognized in the Western blot were detected using ECL, according to manufacturer’s instructions (Amersham Pharmacia Biotech).

Cdc42 activation assay

Immunoprecipitation of Cdc42-GTP and Rac1-GTP was performed using Cdc42 activation kit from Upstate Biotechnology. Briefly, 1 x 107 cells were stimulated for the indicated times and immediately lysed in Mg2+ lysis buffer containing 25 mM HEPES, pH 7.5, 150 mM NaCl, 1% Igepal CA-630, 10 mM MgCl2, 1 mM EDTA, 10% glycerol, 1 mM sodium orthovanadate, 0.2 mM PMSF, 10 µg/ml aprotinin, and 10 µg/ml leupeptin. After 20 min on ice, the lysates were clarified in a microfuge for 10 min at 14,000 rpm. Postnuclear lysates were incubated with 8 µg of GST-PAK-PBD for 1 h at 4°C with rotation. The pellets were then washed three times with 200 µl of Mg2+ lysis buffer and resuspended in 60 µl of SDS sample buffer. GST-PAK-PBD-associated Cdc42 was identified by 12% SDS-PAGE and Western blotting with specific antisera.

Immunofluorescence, confocal microscopy, and time-lapse imaging

Glass coverslips (Fisher Scientific, Pittsburgh, PA) were coated overnight at 37°C with either 1 mg/ml of poly(L-lysine) (PLL) (Sigma-Aldrich, St. Louis, MO), or 10 µg/ml anti-CD28 mAb, or anti-CD3 mAb in 0.1 M phosphate buffer, pH 9.0, and then washed three times with Dulbecco’s PBS (DPBS) without Ca2+/Mg2+ (Life Technologies). A total of 2 x 105 cells was washed and resuspended in 200 µl of DPBS without Ca2+/Mg2+ and seeded on the coverslips for the indicated times. Cells were then fixed with 3.7% formaldehyde for 10 min, washed three times with DPBS without Ca2+/Mg2+, and permeabilized with 0.1% Triton X-100 for 4 min. Cells were washed three times in 0.22 µm filtered DPBS containing 2% BSA (Sigma-Aldrich) and blocked with the same buffer for 30 min. Actin was visualized by incubating fixed permeabilized cells with 1/500 dilution of phalloidin-Texas Red in 2% BSA-DPBS (Molecular Probes, Eugene, OR) for 30 min at room temperature (RT). Cells were mounted with Prolong Anti-Fade (Molecular Probes, Eugene, OR) mounting medium and visualized using a Zeiss epifluorescence microscope coupled to a Spot camera system (Zeiss, Oberkochen, Germany). Images were edited using Adobe Photoshop software (Adobe, San Jose, CA).

Confocal imaging and time-lapse microscopy conditions have been described elsewhere (19). Spreading assays were initiated by injecting 20 µl of a concentrated cell suspension (2 x 106 cells/ml) into the bottom of treated chambers containing RPMI 1640 supplemented with 25 mM HEPES. For time-lapse imaging, chambers were mounted at 37°C on a temperature-controlled inverted microscope (Zeiss Axiovert 135TV) equipped with a x63 oil-immersion Plan-Apochromat objective (NA 1.4; Zeiss). Confocal images were collected using a LSM 410 confocal microscope (Zeiss) equipped with an internal 633-nm laser and an external krypton/argon laser (Spectra-Physics, Mountain View, CA). Time-lapse sequences were recorded using a custom macro program that periodically focuses on the coverslip surface, captures confocal interference reflexion microscopy and fluorescence images at the coverslip, and then captures a fluorescence image 5 µm above the coverslip.

Constructs and transient transfection

DNAs encoding human WT, dominant-negative (DN or T17N), or constitutively active (CA or Q61L) Cdc42 and Rac1 (a gift of S. Gutkind, National Institutes of Health, Bethesda, MD) were subcloned into the BamHI and EcoRI restriction sites of pBKSII and subsequently cloned into the SacI and EcoRI restriction sites of pEGFP-C3 (Clontech, BD Biosciences) to produce a fusion protein encoding enhanced green fluorescent protein (EGFP), followed in frame by the respective Rho GTPase.

Jurkat cells (1 x 107) were removed from culture, washed, resuspended in 0.5 ml RPMI 1640 to which 10% FCS had been added (termed 10% FCS-RPMI), and incubated with 10 µg of one of the following constructs: pEGFP-C3 (pEGFP), pEGFP-C3-Cdc42WT (Cdc42-WT), pEGFP-C3-Cdc42T17N (Ccd42-DN), pEGFP-C3-Cdc42Q61L (Cdc42-CA), pEGFP-C3-Rac1WT (Rac1-WT), pEGFP-C3-Rac1T17N (Rac-DN), or pEGFP-C3-Rac1Q61L (Rac-constitutively active (CA)) for 15 min at RT. Jurkat cells were then electroporated at 260 V, 800 microfarads, low {Omega} (Life Technologies); incubated for 15 min at RT; and then transferred to complete RPMI medium and incubated at 37°C. Human CD4+ T cells were tranfected using the Amaxa method following manufacturer’s instructions (Amaxa Biosystems, Koeln, Germany).

Twelve to 18 h after the transfection, 0.5 x 106 cells were evaluated for GFP expression by flow cytometry (Coulter Epics flow cytometer; Beckman Coulter, Fullerton, CA) and confirmed by Western blotting of postnuclear lysates. Cells were then washed and used either for immunofluorescence or confocal imaging, as described earlier.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
CD28 engagement induces F-actin-enriched filopodial extensions or microspikes in human T cells

To understand the effect of CD28 on the actin cytoskeleton in human T cells in the absence of other receptor or coreceptor engagement, we used surfaces coated with a mAb against the costimulatory molecule CD28. This method has been previously used to assess the effect of TCR/CD3 complex engagement in the reorganization of actin in T cells (19, 20, 21). To evaluate the capacity of CD28 to induce actin reorganization, we used Jurkat cells, human resting CD4+ T cells, and human CD4+ blasts on coverslips previously coated with either anti-CD28 mAb, anti-CD3 mAb, or an irrelevant substrate such as PLL. Cells were fixed, permeabilized, and stained for F-actin content using phalloidin-Texas Red. Following CD3 engagement, Jurkat cells spread in a concentric and uniform fashion, forming a prominent peripheral ring enriched in F-actin (Fig. 1A), as expected (19, 20, 21). Jurkat cells demonstrated prominent filopodial extensions enriched in F-actin content when seeded on anti-CD28 mAb-coated coverslips (Fig. 1A). These extensions were specific for anti-CD28-coated surfaces because they did not occur on PLL (Fig. 1A) or an isotype control or irrelevant mAb (data not shown). The same filopodial extensions could be detected in resting CD4+ T cells seeded in surfaces coated with an anti-CD28 mAb, structures that became more dramatic in human CD4+ T cells blasts (Fig. 1A). The formation of these filopodial extensions could not be ascribed to fibronectin/integrin-like interactions because the experiments were always performed in the absence of serum or calcium/magnesium.



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  		FIGURE 1. CD28 engagement promotes the formation of filopodial extensions enriched in F-actin human T cells. A, F-actin staining in Jurkat and human CD4+ T cells after 25-min incubation on coverslips coated with either 1 mg/ml PLL (top), 10 µg/ml anti-CD28 mAb (middle), or 10 µg/ml anti-CD3 (bottom). After fixation and permeabilization, F-actin was visualized by staining with phalloidin-Texas Red. Images were taken using an epifluorescence microscope under a x100 oil objective. B, Real-time formation of CD28-induced filopodia in Jurkat cells. A total of 0.4 x 106 cells was applied to anti-CD28 mAb-coated chambers, and confocal interference reflexion microscopy images were collected under a x63/1.3 oil objective using a LSM 410 confocal microscope. Images were captured for a total period of time of 28.5 min, at an interval of 1.5 min each.

 
To define the time course for such morphological changes, Jurkat T cells were filmed over time using confocal microscopy. Images at the level of the area of contact between the cell and the coverslip were taken periodically for a total period of 28.5 min. The presence of CD28 cross-linking prompted the cells to adhere and to begin a dynamic phenomenon of cytoplasmic projections over time that resemble filopodia (Fig. 1B). These structures could not be observed when coverslips were coated either with PLL alone or anti-CD3 (data not shown).

Taken together, our data demonstrate that engagement of CD28 in human T cells induces filopodial extensions enriched in F-actin content in human T cells. These CD28-induced microspikes were immediately observed at the area of contact between the cell and the anti-CD28 mAb-coated surface, suggesting the existence of an early CD28-mediated activation pathway leading to actin reorganization.

The Src family tyrosine kinases Lck and Fyn, but not ZAP-70, are required for CD28-induced filopodial extensions

In T cells, triggering of the costimulatory molecule CD28 induces tyrosine phosphorylation of its cytoplasmic domain through the participation of the Src family kinases Lck and Fyn (9). The requirement for ZAP-70 kinase activity in CD28 signaling pathway remains unclear. To address the role of Src family tyrosine kinases and Syk kinases in the formation of CD28-induced filopodial extensions, we used a panel of Jurkat cell lines deficient in the different enzymatic activities. As shown in Fig. 2A (left panel), the absence of p56Lck completely abrogated the formation of F-actin-enriched filopodial elongations upon CD28 engagement. Reconstitution of p56Lck restored the formation of CD28-induced filopodia. Similar results were obtained in cells overexpressing p59Fyn, suggesting that high levels of Fyn can compensate for Lck in mediating CD28 actin reorganization. Surprisingly, the lack of ZAP-70 kinase did not affect the formation of CD28-induced microspikes. Differences in the magnitude of CD28-induced microspikes could not be ascribed to different surface expression of CD28 among cell lines (Fig. 2A, right panel) or expression of Lck among the different clones of Jurkats (Fig. 2B). Moreover, this CD28-promoted phenotype could be observed in 80% of cells plated on anti-CD28-coated coverslips, as quantitated in Fig. 2C. Taken together, we conclude that CD28-induced microspike formation required the presence of Src kinases, but was independent of ZAP-70 kinase activity.



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  		FIGURE 2. CD28-induced filopodia requires the activation of Src family kinases, Lck and Fyn, but is independent of ZAP-70 kinase. A, F-actin staining in Jurkats (WT), JcaM1, JcaM1/Lck, JcaM1/Fyn, and P116 (ZAP-70-) cell lines after 25-min incubation on coverslips coated with either 1 mg/ml PLL or 10 µg/ml anti-CD28 mAb. After fixation and permeabilization, F-actin was visualized by staining with phalloidin-Texas Red. Images were taken using an epifluorescence microscope under a x100 oil objective. B, Jurkats (WT), JCaM1, JCaM1/Lck, JCaM1/Fyn, and P116 (ZAP-70-) were equivalent for CD28 surface expression. Briefly, 2 x 105 cells were stained with 20 µl of anti-CD28-PE Ab and adquired in a Coulter cytometer. C, Postnuclear lysates from 0.5 x 106 cells were resolved in a 10% SDS-PAGE, followed by blotting with antisera against p56Lck or p59Fyn, as shown. D, Quantitation of CD28-induced microspike-forming cells as a fraction of total cells among the different cell lines. Results are representative of three different experiments in which the number of positive cells was estimated over a total number of 30. Bars represent average ± SD.

 
CD28 cross-linking promotes the phosphorylation of Vav and activation of the Rho GTPase Cdc42 independently of ZAP-70 kinase

We and others have previously observed the ability of CD28 to induce Vav phosphorylation upon triggering either with mAb or CD80/CD86 (5). CD28-mediated Vav phosphorylation has been reported to be independent of ZAP-70 activity (22). Vav phosphorylation results in an augmentation of its GDP/GTP exchange activity (GEF) for the small Rho family GTPases Rac1 and Cdc42, proteins implicated in actin cytoskeletal changes in other cell types (23). We evaluated the requirement of ZAP-70 in CD28-mediated phosphorylation of Vav and activation of Cdc42 and Rac1 in T cells. Engagement of CD28 was able to induce the rapid phosphorylation of Vav even in the absence of ZAP-70 (Fig. 3A), as reported (22). In agreement with these results, CD28 triggering also promoted Cdc42 activation. Cdc42 activation occurred after 5 min in ZAP-70-expressing (WT) Jurkats, but occurred even earlier in ZAP-70-deficient P116 Jurkats, suggesting that CD28 ligation promotes Cdc42 activation and Vav phosphorylation in a ZAP-70-independent manner. No changes in Rac1 activation could be observed between WT Jurkats and P116 (ZAP-70-) cell line upon CD28 cross-linking potentially secondary to the basal level of Rac1-GTP in both cell lines (Fig. 3B).



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  		FIGURE 3. CD28-induced Vav phosphorylation and Cdc42 activation are independent on ZAP-70 kinase. A, A total of 1 x 107 Jurkat or P116 (ZAP-70-) cells was stimulated for the indicated periods of time with 1 µg of anti-CD28 mAb. Vav was immunoprecipitated using 1 µg of the specific rabbit antisera, followed by 20 µl of protein A-agarose. The presence of tyrosine-phosphorylated Vav was assayed by blotting the membrane with the antiphosphotyrosine Ab 4G10. The membranes were stripped and reblotted for Vav to confirm equal amount of protein. Results are representative of two different experiments. B, A total of 1 x 107 WT Jurkat T cells or P116 (ZAP-70-) was stimulated for the indicated periods of time with anti-CD28 mAb (1 µg) to evaluate the activation of endogenous Cdc42. GTP-bound forms of Cdc42 were immunoprecipitated using GST-PAK-PBD and resolved in a 12% SDS-PAGE, followed by Western blot for the specific GTPases. Results are representative of three different experiments.

 
Cdc42 is responsible for CD28-induced actin reorganization and microspike formation in human T cells

To compare the roles of Cdc42 and Rac1 in the formation of CD28-induced microspikes, we created GFP-tagged forms of either WT, DN (T17N), or CA (Q61L) human Cdc42 and Rac1. Jurkat cells were transiently transfected, and GFP-positive cells (40–50% of the cells) were assessed for CD28-induced filopodia formation.

We first evaluated the effect of the different Cdc42 and Rac1 constructs in Jurkat cells (Fig. 4A). In cells seeded on PLL coverslips, Cdc42-CA localized to cytoplasmic elongations and promoted a spiky phenotype reminiscent of CD28 activation. Furthermore, an inactive Cdc42 mutant (Cdc42-DN) completely abolished the formation of CD28-induced filopodia even in the presence of CD28 engagement. Intact GTPase activity was required to localize Cdc42 to the cytoplasmic membrane and F-actin-enriched areas, including filopodia. No filopodial extensions were observed when a CA form of Rac1 was used, and no CD28-induced filopodia abrogation was observed when the DN form was used (Fig. 4A). To further confirm the role of Cdc42 in CD28-induced filopodia, we transfected resting CD4+ T cells with either control vector or the DN or CA forms of Cdc42. As shown in Fig. 4C, a comparable effect was reproduced in freshly isolated human T cells, confirming the observation that CD28-mediated actin reorganization and microspike formation in human T cells require the activation of the small Rho GTPase Cdc42, but not Rac1.



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  		FIGURE 4. Cdc42 is responsible for CD28-induced filopodia in human T cells. A, A total of 1 x 107 WT Jurkats was transfected by electroporation with 10 µg of either pEGFP-C3, or GFP-tagged forms of WT, DN, or CA Cdc42 or Rac1. After 12–18 h, 0.2 x 105 cells were plated on anti-CD28-coated coverslips for 25 min, fixed, permeabilized, and stained for F-actin. Localization of GFP-tagged proteins is shown in green, and F-actin in red. Immunofluorescence images were taken under a x100 oil objective. B, Postnuclear lysates from 1 x 106 transfected WT Jurkats cells were resolved in a 12% SDS-PAGE, followed by blotting with the corresponding anti-GFP Ab to confirm the expression of GFP-tagged proteins. C, A total of 5 x 106 purified human CD4+ T cells was transfected using Amaxa method with 5 µg of either pEGFP-C3, or GFP-tagged forms of DN or CA Cdc42. After 12–18 h, 0.5 x 105 cells were plated on PLL or anti-CD28-coated coverslips for 25 min, fixed, permeabilized, and stained for F-actin. Localization of GFP-tagged proteins is shown in green, and F-actin in red. Immunofluorescence images were taken under a x100 oil objective.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
CD28 ligation plays a central role in the prevention of anergy in naive T cells by promoting optimal cytokine production and survival. The role of CD28, however, in regulating the formation and stabilization of the T cell/APC contact is less well understood. The topology of CD28/B7 interactions is similar to that of many adhesion molecules, such as CD2/CD58 (CD48), and has led to the suggestion that CD28 might have a dual role as adhesion and signaling molecule (1). Within seconds of interaction between the T cell and the APC and before Ag recognition occurs, actin cytoskeletal changes can be observed at the surface of the T cell (24). This initial crawling of the T cell around the APC is likely to depend on multiple adhesion forces, of which one may be the binding of CD28 to B7. An adhesive role for CD28 is supported by analysis of T cells from mice deficient in CD28 (25) and by the observation that CD28 ligation induces actin polymerization and formation of focal adhesion-like contacts (26). Our results support and extend these data using a different methodological approach. We chose, as stimulator, anti-CD28-coated surfaces to assess the capacity of CD28 to reorganize the actin cytoskeleton in the absence of any other signal elicited from the engagement of either the TCR/CD3 complex or adhesion molecules, a method previously used to explore the signaling pathway linking the TCR/CD3 complex to the reorganization of the actin cytoskeleton (19, 20, 21).

We analyzed the molecular basis of the regulation by CD28 of actin projections. Ligation of CD28 with mAbs immobilized to a surface induced CD4+ T cells to form filopodia and microspikes that, in appearance, were reminiscent of the phenotypic alterations induced by Cdc42 activation in fibroblasts and other cell types (23). CD28 ligation induced Cdc42 activation; a role for Cdc42 was confirmed by overexpression of WT and mutated forms of this GTPase in both Jurkat cells and resting human peripheral blood CD4+ T cells. The ability of CD28 to induce Cdc42 activation and filopodia projections was shown further to be independent of the activation of the ZAP-70/Syk kinase family, but dependent on a Src kinase family member p56Lck or p59Fyn.

Much is known about the signaling pathways initiated by CD28 following its engagement with either mAb or CD80/CD86, and our current observations extend this appreciation (27). Engagement of CD28 either with mAb or CD80/CD86 results in tyrosine phosphorylation of the cytoplasmic tail of CD28 by p56Lck and subsequent recruitment and activation of PI3K (5). However, our analysis of the role of PI3K in Cdc42 activation and filopodia projections was encumbered by the fact that constitutively high levels of PI3K products are present in Jurkat T cells secondary to deficient expression of the tumor suppressor protein phosphatidylinositol-3,4,5-triphosphate 3-phosphatase (28). The facts 1) that treatment of Jurkat T cells and human CD4+ T cells with the PI3K inhibitor wortmannin inhibited CD28-induced filopodia (our unpublished results), and 2) that no filopodial elongations were observed in the absence of CD28 cross-linking are consistent with the hypothesis that CD28-mediated filopodia formation requires PI3K activity.

PI3K has been shown to recruit the GEF Vav to the plasma membrane (11). Vav in turn has been shown to be tyrosine phosphorylated by Src kinases (p56Lck and p59Fyn) upon CD28 ligation in the absence of TCR/CD3 engagement (9) and in a ZAP-70-independent manner (22). One target GTPase for Vav activity in vitro is Rac1 (3); its target(s) in vivo is not known. Our data confirmed the ZAP-70-independent, CD28-mediated tyrosine phosphorylation of Vav; Vav activation correlated with CD28-induced actin reorganization into filopodial extensions. Maximal Cdc42 activation was observed between 1 and 5 min after CD28 ligation, but Vav phosphorylation and filopodial extensions were sustained for as long as 30 min. We note that the biochemical demonstration of Cdc42 activation is insensitive and that the biochemical analysis itself does not allow for different subcellular pools to be assayed differentially (29). A small change in the pool of active Cdc42 may be sufficient to induce actin reorganization. Furthermore, the simple correlation of Vav phosphorylation with both Cdc42 activation and microspike formation does not demonstrate causality nor exclude the possibility that other GEFs, as yet unidentified, might play a role in CD28-mediated actin polymerization.

Superficially, our results would appear to be discordant with those of Kaga et al. (8, 26), who demonstrated translocation of Rac1 to focal adhesion-like contacts and activation of PAK1 and mitogen-activated protein kinase kinase kinase, after CD28 ligation in T cells. These investigators, however, allow for the possibility of cross talk in their analyses and did not specifically investigate Cdc42 activation in their experiments (8, 26). The existence of cross talk among members of the Rho family of GTPases has been shown in other systems (23). In fibroblasts, Cdc42 activation not only triggered induction of filopodia and microspikes, but also induced downstream Rac1-mediated responses (30). We suggest that the effects observed by Kaga et al. (26) may be, at least in part, mediated by Cdc42, which in turn acts upon Rac1 as a mediator of actin reorganization. We do not eliminate a contribution of Rac1 in the effects that we observe (see below); we do endorse the hypothesis that Cdc42 may be an important and essential intermediate in the cascade.

Cross talk between Rac and Cdc42 may also explain the observed inhibition of microspike formation by DN forms of Rac, particularly if Rac activation is downstream of Cdc42 in this system. Overexpressed, mutated forms of Rac have profound effects in T cell morphology and actin reorganization. We are not surprised that there is an effect of overexpression of mutated forms of Rac1 on T cell morphology, as was observed in Fig. 4. However, only the constitutively active form of Cdc42 was able to induce the formation of filopodial extensions in the absence of any stimulation; no filopodial extensions were detected in cells transfected with the constitutively active form of Rac1. Unlike Cdc42, we were unable to demonstrate a specific role for Rac1 in filopodia formation.

A direct target/effector molecule of Cdc42 is the WASp, and WASp activation results in the recruitment of the actin-nucleating complex Arp 2/3 (31). Upon engagement of the TCR/CD3 complex, WASp and Arp 2/3 are recruited to the area of contact between the T cells and the APC or anti-CD3-coated beads (29, 32). In our hands, no active Cdc42 was precipitated in assays using the PBD domain of WASp (data not shown), possibly because of the very small pool of Cdc42-GTP required for actin reorganization upon CD28 cross-linking. We also could not detect tyrosine phosphorylation of WASp after CD28 engagement; whether WASp becomes phosphorylated in T cells and phosphorylation is important for WASp function are still unknown (31). The role of WASp and Arp 2/3 in CD28-mediated actin reorganization is currently under investigation

Recent reports have proposed that the immunological synapse is more likely to be related to the delivery of a secondary T cell signaling and effector functions, such as cytokine release, than for early TCR signaling (2). Cdc42 is required for reorientation of the T cell microtubule-organizing center after binding to an Ag-bearing APC and CD28-induced PI3K activation sustains T cell microtubule-organizing center reorientation (15, 33). Future experiments will be required to address the role of CD28-mediated activation of Cdc42 in the formation of T cell-APC conjugates and the immunological synapse.


   Footnotes
 
1 Current address: Laboratory of Lymphocyte Biology, Laboratory of Immunology, National Institutes of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD 20892. E-mail address: lfontana{at}niaid.nih.gov Back

2 Address correspondence to Dr. Barbara E. Bierer, Brigham and Women’s Hospital, 75 Francis Street, Boston, MA 02115. E-mail address: BBierer{at}partners.org Back

3 Abbreviations used in this paper: WASp, Wiskott-Aldrich syndrome protein; CA, constitutively active; DN, dominant negative; DPBS, Dulbecco’s PBS; EGFP, enhanced green fluorescent protein; F-actin, filamentous actin; GEF, guanine nucleotide exchange factor; PAK, p21-activated kinase; PI3K, phosphoinositide 3-kinase; PLL, poly(L-lysine); RT, room temperature; WT, wild type. Back

Received for publication October 29, 2002. Accepted for publication June 20, 2003.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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