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Anchorage Dependence of Mitogen-Induced G₁ to S Transition in Primary T Lymphocytes¹

J. Geginat^*,, G. Bossi^*, J. R. Bender^§ and R. Pardi^2,*,

^* Scientific Institute San Raffaele-DIBIT, and University of Milan School of Medicine, Milan, Italy; Department of Biochemistry, Free University of Berlin, Berlin, Germany; and ^§ Molecular Cardiobiology, Boyer Center for Molecular Medicine, Cardiovascular Medicine, and the Raymond and Beverly Sackler Cardiobiology Laboratory, Yale University School of Medicine, New Haven, CT 06536

	Abstract

Top Abstract Introduction Materials and Methods Results Discussion References

 
Anchorage dependence defines the cellular requirement for integrin-mediated adhesion to substrate to initiate DNA replication in response to growth factors. In this study we investigated whether normal T cells, which spend extended periods in a nonadherent state, show similar requirements for cell cycle progression in response to TCR stimulation. Resting primary T lymphocytes were induced to enter the cell cycle by TCR triggering, and leukocyte integrins were either engaged using purified ICAM-1 or inhibited with function-blocking mAbs. Our data indicate that leukocyte integrins complement TCR-driven mitogenic signals not as a result of their direct clustering but, rather, via integrin-dependent organization of the actin cytoskeleton. Leukocyte integrin-dependent reorganization of the actin cytoskeleton cooperates with the TCR to effect mitogen-activated protein kinase activation, but also represents a required late (4–8 h poststimulation) component in the mitogenic response of normal T cells. Prolonged leukocyte integrin-dependent spreading, in the context of intercellular contact, is a requisite for the production of the mitogenic cytokine IL-2, which, in turn, is involved in the induction of D3 cyclin and is primarily responsible for the decrease in the cyclin-dependent kinase inhibitor p27^kip, resulting in retinoblastoma protein inactivation and S phase entry. Thus, T lymphocytes represent a peculiar case of anchorage dependence, in which signals conveyed by integrins act sequentially with the activating stimulus to effect a sustained production of the essential mitogenic cytokine.

	Introduction

Top Abstract Introduction Materials and Methods Results Discussion References

 
Most somatic cells require adhesion to substrate for progression through the cell cycle and the onset of DNA replication in response to growth factors (1). This phenomenon is defined as anchorage dependence and is thought to be largely dependent on active signaling by surface integrins, a highly conserved family of adhesion receptors involved in cell-extracellular matrix as well as cell-cell adhesion (2). The mechanistic basis of integrin-dependent signal transduction, and how it relates to mitogenic signals transduced by growth factor receptors are only partially understood (3). Signals delivered by growth factor receptors and integrins have been proposed to act synergistically in promoting cell proliferation, but whether they lie on independent pathways or exert a mutual permissive action on the same signaling pathway is presently unclear (4). Early events that are critical in mediating the effects of growth factors on cell cycle progression, such as mitogen-activated protein (MAP)³ kinase activation, have been shown to be strongly dependent on cell adhesion and spreading (5). Accordingly, constitutively active mutants of molecular intermediates in the MAP kinase pathway, such as Ras, Raf, or extracellular signal-regulated kinase (ERK), lead to anchorage-independent growth (6). However, most of these studies have been conducted in immortalized cell lines, whose proliferative response to growth factors is variably dysregulated, as are the constraints imposed by adhesive interactions with other cells or with the extracellular matrix.

Quiescent T lymphocytes typically require two extracellular signals to progress into the cell cycle in response to mitogenic stimulation. Signal 1 is delivered by the TCR and alone is capable of promoting second messenger generation, Ras activation, and immediate early gene expression (7). Signal 2 is provided by a fairly heterogeneous group of costimulatory receptors, which includes the Ig superfamily members CD2, CTLA-4, and CD28, as well as ß₁ and ß₂ integrins (8, 9, 10). Costimulatory receptors are operationally defined as capable of acting synergistically with the TCR in driving T cell proliferation. Indeed, selected members of the MAP kinase family, such as Jun N-terminal kinase (JNK), have been shown to be fully activated only in response to costimulation in transformed T cell lines and to be essential for IL-2 promoter activation in this model (11). However, in several models using nontransformed T lymphocytes, the engagement of costimulatory molecules does not lower the threshold number of TCRs required for mitogenesis (12, 13).

The aim of the present study was to investigate the molecular basis of leukocyte integrin-mediated costimulation of TCR-driven proliferation in primary T lymphocytes. The existing evidence concerning this issue is rather controversial; some reports, based on the premise that integrins alone do not transduce signals required for cell proliferation, suggest that leukocyte integrins merely provide the adhesive contact needed by the TCR to engage its cognate ligand on the apposing cell (14, 15, 16). However, other groups, which make use of primary cells from leukocyte integrin knockout animals, show selected defects in the lymphocyte’s mitogenic response under experimental conditions not requiring intercellular adhesion, indicating that an integrin-dependent signaling component is indeed required in the process (17, 18). Since mitogenic signals in nontransformed, quiescent cells ultimately lead to increased expression and/or activation of cyclin-dependent kinases (Cdk), which trigger the biochemical events allowing cells to progress from G₁ to S in the cell cycle (1, 19), we explored the role of leukocyte integrins in promoting critical steps in this pathway, from the early activation of MAP kinases to the expression and function of G₁ cyclins, Cdks, and their inhibitors. Our results suggest that leukocyte integrins complement TCR-driven mitogenic signals not as a result of their direct clustering, but as an indirect response to integrin-dependent organization of the actin cytoskeleton. The integrity of the actin cytoskeleton and a sustained "spread" cell shape not only allow efficient MAP kinase activation, but also represent required late (4–8 h poststimulation) components in the mitogenic response of normal T cells by affecting the expression levels of genes, such as IL-2, that are critically involved in the process.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion References

 
Cells

Buffy coats from normal donors were obtained from the local blood bank. PBMC isolation was described previously (20). B cells and monocytes were depleted by plastic adherence and passage through a nylon wool column. NK cells and residual APCs were depleted by a panning technique using anti-CD16 and anti-MHC class II Abs, respectively. The T cells obtained were >95% CD3⁺ and did not contain APCs as detected by flow cytometry using anti-CD14 and anti-human Ig Abs as markers for monocytes and B cells, respectively.

Abs and reagents

mAb Anti-CD3 (OKT3; IgG2a), anti-CD16 (KD1; IgG1), anti-CD18 (TS1.18; IgG1), anti-CD11a (TS1.22; IgG1), and anti-MHC class II (CA141; IgG1) were affinity purified in our laboratory. mAb anti-CD14 (3C10) was a gift from M. Alessio (Dibit, Milan, Italy). Polyclonal goat anti-lactate dehydrogenase was a gift from M. Bianchi (Dibit). Polyclonal anti-human Ig Abs were purchased from Dako (Copenhagen, Denmark). Recombinant ICAM-1 was provided by Anna Randi (Glaxo Wellcome, Stevenage, U.K.). Cytochalasin D (CCD), digitonin, PMA, poly-L-lysine, myelin basic protein, and neutralizing polyclonal goat anti-IL-2 Abs were purchased from Sigma (St. Louis, MO). The MEK inhibitor PD98059, Pansorbin cells, and ionomycin were purchased from Calbiochem (La Jolla, CA). The GST-c-Jun_1–79 fusion protein construct was a gift from Michael Karin (University of California, La Jolla, CA). mAb anti-JNK (clone G151-333) was purchased from Upstate Biotechnology (Lake Placid, NY), mAb anti-cyclin D2 (G132-43) was purchased from Pierce (Rockford, IL). mAb anti-cyclin D3 (C-16), anti-c-Jun (KM-1), and anti-ERK-2 (D-2) and polyclonal rabbit anti-Cdk4 (C-22), anti-Cdk6 (C-21), anti-Cdk2 (M2), anti-Rb (C-15), anti-p38 (C-20) anti-Lck (2102), anti-c-Fos (4), and anti-p27 (C-19) Abs were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Polyclonal goat anti-mouse and horseradish peroxidase-conjugated goat anti-mouse and anti-rabbit Abs were purchased from Zymed (South San Francisco, CA).

Cell stimulation and propidium iodide staining

Dishes were coated with 0.01% poly-L-lysine in distilled H₂O or with the indicated mAbs or ICAM-1 at 10 µg/ml in 50 mM Tris-Cl, pH 9.5, at room temperature for 3 h or overnight at 4°C. Dishes were then washed with PBS and blocked for 1 h at room temperature with 1% BSA in PBS. Cells (2 x 10⁶/ml) were plated in prewarmed RPMI medium containing 10% FCS. To promote binding of resting T cells to ICAM-coated dishes the incubation was performed in the presence of 1 mM EGTA and 5 mM Mg²⁺ (21). In some experiments requiring long term incubations, the anti-CD11a mAb TS1.22 was used as a substitute for ICAM-1. For inhibition experiments, cells were pretreated with various inhibitors for 1 h at 37°C. The MEK inhibitor PD98059 was used at 50 µM; at this concentration it completely blocked ERK activation without significantly inhibiting JNK or p38 (not shown) as determined by band-shift assay and in vitro kinase assay, respectively (see below). Cells stimulated with soluble anti-CD3 mAb were first incubated on ice for 30 min with saturating concentrations of OKT3 (1 µg/10⁶ cells), excess Ab was removed with ice-cold PBS, and cells were resuspended in prewarmed RPMI medium containing 10% FCS and 10 µg/ml goat anti-mouse Ig polyclonal Ab. After 72 h cells were collected, washed with PBS, and fixed for 30 min in 75% ethanol. Cells were then centrifuged, resuspended in 50 µg/ml propidium iodide in 0.1% sodium citrate and 0.05% Nonidet P-40 containing 50 µg/ml RNase A and incubated for at least 1 h at room temperature. Cell cycle analysis was performed by flow cytometry (FACScan, Becton Dickinson, Milan, Italy).

5-Bromo-2'-deoxyuridine (BrdU) incorporation assay

T lymphocytes stimulated as indicated by solid phase bound ligands were pulsed with 40 µM BrdU (Sigma) for 2 h at 16 and 40 h poststimulation, followed by fixation in 75% ethanol, partial DNA denaturation in 3 M HCl, and direct immunofluorescence using a 1/50 dilution of a FITC-conjugated anti-BrdU mAb (Becton Dickinson, Milan, Italy). Fluorescent cells were visualized using a Zeiss Axiophot fluorescence microscope (Zeiss, Jena, Germany).

Immunoblotting and electrophoretic mobility shift assay

Before lysis, cells were washed twice with ice-cold PBS. Lysis buffer (50 mM Tris-Cl (pH 7.4), 0.5% Nonidet P-40, 0.15 M NaCl, 2 mM EDTA, 10 mM NaF, 10 mM P₂O₇, 0.5 mM Na₃VO₄, 100 µg/ml PMSF, and 1 µg/ml aprotinin and leupeptin; Sigma) was added, and samples were incubated for 15 min on ice. The soluble fraction was cleared by centrifugation, and the protein concentration was determined (DC Protein Assay, Bio-Rad, Hercules, CA). Sample buffer (2% SDS, 10% glycerol, 0.1 M Tris (pH 6.8), 0.001% bromophenol blue, and 100 mM DTT; Sigma) was added, samples were boiled for 5 min, and equal amounts of protein were loaded on a standard SDS-PAGE. For electrophoretic mobility shift assay, which allows the quantification of phosphorylated, activated ERK (22), the separating gel contained 10% acrylamide and 0.073% bisacrylamide. Proteins were transferred on a nitrocellulose membrane (Amersham, Aylesbury, U.K.). Blocking of nonspecific binding and all incubations with Abs were performed in blocking buffer (5% dry nonfat milk in Tris-buffered saline (pH 7.4) and 0.05% Tween-20). Blots were developed with an enhanced chemiluminescence kit (Amersham).

Preparation of subcellular fractions

To assess nuclear translocation of MAP kinases in T cells exposed to various stimuli we prepared cellular fractions by a sequential extraction procedure as follows. Cytosolic proteins were extracted with low concentrations of digitonin in extraction buffer (0.025% in 25 mM HEPES (pH 7.6), 90 mM potassium acetate, 2.5 mM MgCl₂, 2 mM EGTA, 0.2 mM CaCl₂, 12 mM glucose, and phosphatase and protease inhibitors) for 10 min on ice, followed by supernatant collection and further washes of the insoluble pellet. Nuclear proteins were extracted by osmotic disruption of nuclei (0.3 M NaCl in extraction buffer) for at least 10 min on ice, and the supernatant was collected. Finally, after additional washings, membrane proteins were extracted with extraction buffer containing 1% Nonidet P-40. The purity of the fractions obtained was assessed by immunoblotting with anti-lactate dehydrogenase, anti-Rb, and anti-Lck Abs as markers for cytosolic, nuclear, and membrane-associated proteins, respectively.

In vitro kinase assays

JNK kinase assay was performed as described previously (23). Briefly, 10 µg of purified, bacterially expressed GST-c-Jun_1–79 was incubated with whole cell extracts containing equal amounts of protein for 4 h at 4°C. The pellet was washed and resuspended in kinase buffer containing 5 µCi of [-³²P]ATP. The reaction was incubated for 20 min at 37°C and was stopped by the addition of sample buffer. To measure the activity of the p38 MAP kinase, after preclearing with normal rabbit Abs prebound to Pansorbin cells the kinase was precipitated from lysates containing equal amounts of protein with 1 µg of anti-p38 Ab prebound to protein A-Sepharose. After extensive washing, the pellet was resuspended in kinase buffer containing 5 µCi of [-³²P]ATP and 10 µg of myelin basic protein. The kinase reaction was conducted for 20 min at 37°C and was stopped with sample buffer. The activities of Cdk4 and Cdk6 were assessed as previously described (24). Briefly, 20 x 10⁶ cells stimulated as indicated were washed, resuspended in Cdk lysis buffer (50 mM HEPES (pH 7.5), 150 mM NaCl, 1 mM EDTA, 2.5 mM EGTA, 1 mM DTT, 0.1% Tween-20, 0.1 mM Na₃VO₄, 1 mM NaF, 10 mM ß-glycerophosphate, and 1 mM sodium pyrophosphate plus protease inhibitors) and sonicated, and the nuclei were spun down. The kinases were immunoprecipitated for 2 h at 4°C with 1 µg of polyclonal rabbit anti-Cdk4 and anti-Cdk6 Abs (Santa Cruz Biotechnology) from the precleared lysates containing equal amounts of proteins. Samples were washed twice in Cdk lysis buffer and once in Cdk kinase buffer (50 mM HEPES (pH 7.5), 10 mM MgCl₂, 5 mM MnCl₂, and 1 mM DTT). The kinase reaction was conducted in 30 µl of Cdk kinase buffer containing 1 µg of recombinant GST-pRb protein (Santa Cruz Biotechnology) and 10 µCi [-³²P]ATP for 30 min at 37°C and was stopped by the addition of sample buffer. The samples were boiled, and a standard SDS-PAGE was performed. The gels were dried and exposed to an x-ray film (Amersham).

ELISA

IL-2 released by activated T cells was measured using a protocol previously described by Dr. H. Gallatti (INTEX, Muttenz, Switzerland). The supernatant of T cells stimulated for various time periods or a reference concentration of rIL-2 was serially diluted and incubated with the anti-human IL-2 mAbs 3D5 and 7B1, which had been immobilized to microtiter plates, in the presence of the soluble, peroxidase-coupled anti-human IL-2 mAb 13A6. The amount of bound peroxidase was enzymatically determined with tetramethylebenzidine-H₂O₂ as a substrate in potassium citrate, pH 4.1, for 10 min at room temperature. The reaction was stopped with 1 M sulfuric acid, and colorimetric analysis was performed at 450 nm with a multichannel photometer (Shimadzu, Kyoto, Japan).

	Results

Top Abstract Introduction Materials and Methods Results Discussion References

 
Entry into S phase in response to TCR cross-linking requires integrin-dependent spreading in quiescent T cells

To address the role of integrin-dependent costimulation in cell cycle progression of mitogen-stimulated T cells, we used anti-CD3 Abs, in soluble or immobilized form, as a substitute for cognate recognition by the Ag receptor complex and immobilized ICAM-1 to engage the _L/ß₂ integrin (or LFA-1, hereafter termed integrin) on T cells. The requirement for costimulation by the two receptors to promote S phase entry in quiescent T cells is shown in Fig. 1. While saturating amounts of soluble anti-CD3 Abs (Fig. 1, C and C') or immobilized ICAM-1 (Fig. 1, B and B') alone did not induce S phase entry, immobilized anti-CD3 Abs in either the absence (Fig. 1, E and E') or the presence (Fig. 1, D and D') of coimmobilized ICAM-1 effectively promoted, albeit to a variable degree, the onset of DNA replication in primary T cells. Conversely, when the immobilized anti-CD3 Ab was used alone (Fig. 1, E and E'), cells underwent initial spreading, which has been shown to involve dynamic interactions between integrins and the actin-based cytoskeleton (20). Two to four hours after plating, cells detached from the dish and rapidly formed tight homotypic clusters, which were entirely integrin dependent (as shown by their complete inhibition using soluble anti-LFA-1 Ab, see Fig. 1F') and were a requirement for progression into S phase (compare Fig. 1, E to F). Costimulation was also observed when cells treated with soluble anti-CD3 Ab were plated onto immobilized ICAM-1. Under these conditions, however, the fraction of cells entering S phase as a result of costimulation was low, probably because, as previously reported (25), the two stimuli induced spreading of primary T lymphocytes only transiently when delivered in trans. These problems could be largely overcome by plating soluble anti-CD3-stimulated cells onto immobilized anti-LFA-1 Ab, which allowed efficient spreading and yielded levels of T cell proliferation comparable to those observed using coimmobilized ligands (not shown). Notably, the coengagement of other costimulatory receptors, which is certainly possible under conditions in which large homotypic clusters are formed, is unlikely to be responsible for the observed phenomena, as cells were most efficiently driven into the cell cycle by costimulation with coimmobilized anti-CD3 and ICAM-1, a condition in which they remained as individual cells for the entire stimulation period (see Fig. 1D'). To confirm this hypothesis, we analyzed S phase entry at a single cell level by BrdU incorporation, followed by direct immunofluorescence with FITC-conjugated anti-BrdU Ab, in T lymphocytes stimulated with coimmobilized anti-CD3 and ICAM-1. Fig. 2 clearly shows that a significant fraction of T cells, firmly adherent as individual cells onto the solid phase bound costimulatory ligands throughout the culture period, undergoes DNA replication starting at 40 h poststimulation.

View larger version (80K): [in this window] [in a new window]   FIGURE 1. Requirements for TCR-stimulated G1 to S transition in primary T cells. Purified T cells were incubated in poly-L-lysine-coated dishes in the absence or the presence of soluble anti-CD3 mAb plus goat anti-mouse Ig (A, A' and C, C', respectively), stimulated with ICAM-1-coated dishes in the absence or the presence of coimmobilized anti-CD3 mAb-coated dishes (B, B' and D, D', respectively), or stimulated with immobilized anti-CD3 mAb in the absence or the presence of soluble, function-blocking anti-LFA-1 Abs (E, E' and F, F', respectively). Proliferation was assessed by propidium iodide staining after 72 h of stimulation. The numbers indicate the fraction of cells in the S/G2/M phases of the cell cycle. Phase contrast microscopic images were taken 20 min (A'–D') or 8 h (E' and F') poststimulation. Results are representative of 10 separate experiments.

View larger version (91K): [in this window] [in a new window]   FIGURE 2. Intercellular contact is not required to promote S phase entry in T cells stimulated by coimmobilized anti-CD3 mAb and ICAM-1. Resting T cells were bound at low density onto anti-CD3 mAb- plus ICAM-1-coated dishes and were incubated for 16 h (a, c, and e) or 40 h (b, d, and f), following which they were pulsed with 40 µM BrdU for 4 h. After fixation and partial DNA denaturation, cells were stained with the Hoechst 33342 dye to visualize the nuclei (c and d) and with FITC-conjugated anti-BrdU Ab (e and f) to detect DNA-replicating cells. a and b represent differential interference contrast (DIC) analysis of the same fields shown in the lower panels.

Since a proposed role for costimulatory molecules is the enhancement of the strength and the duration of signaling by the TCR (16, 26, 27), we assessed whether coimmobilization of the TCR and leukocyte integrins affected the rate of internalization of the Ab-stimulated Ag receptor complex. Fig. 3 shows that contact with immobilized anti-CD3 induced a steady internalization of the receptor complex in a dose-dependent manner. When cells were stimulated with saturating amounts of anti-CD3 (10 µg/ml), approximately 90% of the entire surface pool was internalized by 90 min poststimulation; the simultaneous engagement of leukocyte integrins by immobilized ICAM-1 did not have a significant effect on the rate and the extent of TCR complex internalization. In contrast, suboptimal concentrations of anti-CD3 (5 µg/ml) led to a slower and incomplete down-modulation of the TCR. As reported for the internalization of the TCR following the recognition of its cognate ligand (13, 16), this partial, Ab-induced receptor down-regulation was significantly enhanced by the coengagement of integrins. Conversely, the presence of soluble function-blocking anti-LFA-1 mAbs in the culture did not significantly alter the extent or the kinetics of internalization of the TCR at any anti-CD3 concentration tested (1–10 µg; data not shown). Since our model of costimulation is based on the use of saturating amounts of anti-CD3, these findings suggest that enhanced TCR triggering followed by receptor complex internalization are not responsible for the observed integrin-dependent S phase entry.

View larger version (21K): [in this window] [in a new window]   FIGURE 3. The rate of TCR internalization is not altered by integrin coengagement at saturating anti-CD3 concentration. Cells were either plated on poly-L-lysine (squares) or stimulated with immobilized, biotinylated anti-CD3 mAb at saturating (10 µg/ml; circles) or suboptimal (5 µg/ml; diamonds) concentrations with (filled symbols) or without (open symbols) coimmobilized ICAM-1 for the indicated time periods. Cells were then harvested, incubated for 30 min with biotinylated anti-CD3 mAb on ice to saturate surface CD3 complexes, and stained with FITC-conjugated streptavidin followed by flow cytometry. Results are expressed as the percent reduction of mean fluorescence levels compared with those in untreated cells and represent the mean ± SD of four separate experiments.

In most cell types, mitogenic signals are relayed into the nucleus through the activation and nuclear translocation of MAP kinases (28). Previous studies in nonlymphoid cells have shown that aggregation of integrins, alone or in combination with growth factors, contributes to the activation of both ERK and JNK (5, 29, 30, 31), two well-characterized members of the family. Fig. 4A shows that at the time of maximal stimulation both ERK and JNK activities were potentiated when the TCR and leukocyte integrins were coengaged in quiescent T cells. The integrin-dependent, but not the TCR-dependent, component of MAP kinase activation appeared to require an intact actin cytoskeleton and not just receptor aggregation, as judged by its complete inhibition in CCD-treated cells (Fig. 4A) and by the failure of soluble anti-integrin Ab to induce enzyme activation (not shown). Since the selective inhibition of ERK by the MEK inhibitor PD98059 completely prevented S phase entry in our model (see below), we concluded that ERK activation is required for T cell proliferation induced by TCR and integrin coengagement, and asked whether the synergistic and/or prolonged activation of these kinase pathways in costimulated cells was sufficient per se to promote the onset of DNA replication that occurs exclusively under this condition. To this aim, we compared the activation levels and kinetics of ERK and JNK in T cells that were stimulated as shown in Fig. 1, that is by coengagement of the TCR and integrins or by immobilized anti-CD3 Ab alone with or without soluble anti-integrin Ab to prevent homotypic clustering. Fig. 4, B and C, shows that although the simultaneous engagement of the two receptors indeed resulted in a more pronounced activation of both kinase pathways, the extent and kinetics of ERK and JNK activity were virtually superimposable in the other two conditions despite the fact that S phase entry was completely prevented by blocking integrin-dependent homotypic clustering. These findings suggest that integrin-dependent costimulation in TCR-stimulated T cells cannot be uniquely explained by a synergistic effect on MAP kinase activation. The activation profiles of p38 kinase, a recently described MAP kinase family member that has been reported to be important for T cell proliferation (32, 33), followed closely those of ERK under the conditions used in our study (not shown), suggesting that none of the major eukaryotic MAP kinase pathways is primarily mediating the costimulatory effect of integrins on cell cycle progression of normal T cells.

View larger version (23K): [in this window] [in a new window]   FIGURE 4. ERK-2 and JNK activation by TCR- and integrin-dependent signals. A, Purified resting T cells were stimulated for 20 min as indicated with (lanes 3, 5, and 7) or without (lanes 1,2, 4, and 6) pretreatment (30 min at 37°C) with 10 µM cytochalasin D. ERK activation was measured by an electrophoretic mobility shift assay. JNK activity was assessed with an in vitro kinase assay as described in Materials and Methods. B and C, Time course of JNK (B) and ERK-2 (C) activation. Cells were left untreated (open squares) or were stimulated for the indicated times with immobilized anti-CD3 mAb (filled circles), coimmobilized anti-CD3 mAb and ICAM-1 (filled squares), or immobilized anti-CD3 mAb in the presence of blocking soluble anti-LFA-1 mAb (open circles). ERK and JNK activation was determined as described in A. Results represent the mean ± SD of four separate experiments.

Effects of integrin-dependent costimulation on c-fos and c-jun expression

As a later readout for MAP kinase activation in TCR- and integrin-stimulated T cells, we analyzed the expression of the c-jun and c-fos immediate early gene products, since enhanced transcription of both genes is believed to be an important downstream event following the activation of JNK and ERK, respectively (28, 34). Fig. 5 demonstrates that stimulation of quiescent T cells with immobilized anti-CD3 Ab led to a rapid up-regulation of both proto-oncogene products, reaching a plateau at 2–8 h poststimulation and gradually decreasing at later time points. As expected, the abrogation of integrin-dependent homotypic clustering, which prevents T cells from entering S phase, did not significantly affect the kinetics or the absolute expression levels of either proto-oncogene product in TCR-stimulated cells. Interestingly, the expression levels of both proto-oncogene products were only slightly enhanced by coengaging leukocyte integrins, particularly at late time points after stimulation.

View larger version (27K): [in this window] [in a new window]   FIGURE 5. Integrin-dependent costimulation does not involve the modulation of c-Fos and c-Jun expression in TCR-stimulated T cells. Cells were stimulated with immobilized anti-CD3 mAb in the absence or the presence of either soluble, function-blocking anti-LFA-1 mAb or coimmobilized ICAM-1 for the indicated times. The c-Jun and c-Fos protein levels were assessed by Western blotting. Similar results were obtained in four separate experiments.

The experiments reported above suggest that although integrin-dependent cell spreading is a requisite for optimal MAP kinase activation, a critical component of integrin-mediated signaling is acting at a later step in the sequence of events leading to the onset of DNA replication in TCR-stimulated cells. To more precisely map the integrin-dependent requirement within G₁, we added CCD (Fig. 6) or excess soluble function-blocking anti-integrin Ab (not shown) at various times following the mitogenic stimulus. The MEK inhibitor PD98059 was used as a probe for ERK function within the same time period. The results of this experiment (Fig. 6) clearly indicate that an intact actin-based cytoskeleton is required for up to 8 h postmitogenic stimulation in normal T cells. Such anchorage dependence coincides with the time required for TCR-stimulated cells to transit through G₁ and complete the necessary biochemical modifications of cell cycle-related proteins that allow the initiation of DNA replication (see below). Notably, the selective inhibition of ERK critically affects cell cycle progression for the initial 2–4 h poststimulation, following which cells appear to progress through the cell cycle independently of ERK activity.

View larger version (16K): [in this window] [in a new window]   FIGURE 6. A sustained integrin-dependent spreading is required throughout G1 to initiate DNA replication in TCR-stimulated T cells. The MEK inhibitor PD98059 (50 µM) or CCD (10 µM) was added at the indicated time points to T cells stimulated with coimmobilized anti-CD3 mAb plus ICAM-1. Progression into the cell cycle was assessed by propidium iodide staining 72 h poststimulation under all conditions, and the results are expressed as the percentage of S/G2/M cycling cells compared with that in untreated controls. Values represent the mean ± SD of four separate experiments.

Accumulating evidence indicates that in eukaryotes the main molecular switch allowing mitogen-stimulated cells to enter S phase is the inactivating phosphorylation of pRb, which occurs in mid to late G₁ (19). This process is effected by G₁ phase Cdks, whose catalytic activities are tightly regulated by threshold levels of complexed G₁ cyclins and Cdk inhibitors (Cki). Fig. 7 shows that primary T cells undergoing proliferation in response to immobilized anti-CD3 Ab displayed significant hyperphosphorylation of pRb (as detected by the appearance of slower migrating bands), clearly detectable at 16 h poststimulation. Concomitantly with pRb hyperphosphorylation, we observed a marked, progressive increase in the levels of G₁ cyclins (D2 and D3) and Cdk6, and a moderate increase in the levels of Cdk4, which were paralleled by a sharp reduction (averaging two- to threefold) of the levels of the Cki p27^kip1. Conversely, the levels of the other G₁ phase Cdk, Cdk2, did not significantly increase compared with that of the reference protein ERK-2 throughout the observation period (not shown). Notably, under conditions in which S phase entry was prevented by blocking integrin-dependent homotypic clustering (Fig. 7), pRb remained in a hypophosphorylated state despite moderate to high levels of cyclin D3 and D2, respectively. In addition, p27^kip1 levels remained high and comparable to those observed in unstimulated cells. Coengaging leukocyte integrins strongly enhanced pRb hyperphosphorylation, which was accompanied by enhanced cyclin D3 and strongly reduced p27^kip1 levels. Conversely, the expression levels of cyclin D2 and the G₁ Cdks were not significantly enhanced by coimmobilized ICAM-1. Consistent with these findings, the catalytic activities of the two major pRb kinases, Cdk4 and Cdk6, closely correlated with pRb hyperphosphorylation in T cells traversing G₁ under the various experimental conditions (Fig. 8).

View larger version (73K): [in this window] [in a new window]   FIGURE 7. Cell cycle-related protein expression and pRb inactivation as a function of integrin engagement in TCR-stimulated T cells. Resting T cells were stimulated with immobilized anti-CD3 mAb in the absence or the presence of function-blocking, soluble anti-LFA-1 mAb or coimmobilized ICAM-1 for the indicated time periods, followed by lysis and immunoblot analysis with mAbs specific for the indicated proteins. Results are representative of six separate experiments.

View larger version (39K): [in this window] [in a new window]   FIGURE 8. The catalytic activities of the pRb kinases Cdk4 and Cdk6 depend on integrin coengagement in TCR-stimulated T cells. Resting, primary T cells were stimulated for 24 h as indicated. Following cell lysis, Cdk4 and Cdk6 were immunoprecipitated, and in vitro kinase assays were performed using recombinant GST-pRb as an exogenous substrate in the presence of 32P-labeled ATP. The activation state of the endogenous pRb on total cell lysates was evaluated under the same experimental conditions with an electrophoretic mobility shift assay (see Fig. 7).

Several independent reports have recently shown that p27^kip1 degradation in primary T lymphocytes is mediated by IL-2 (35, 36) and represents the main biochemical event underlying S phase entry in IL-2-responsive T cells exposed to this mitogenic cytokine (35). We therefore asked whether the marked integrin-dependent reduction in p27^kip1 levels observed in our model was an indirect consequence of a more efficient production of IL-2 in cells costimulated via integrins and the TCR. We used several approaches to address this question, and the results are collectively shown in Table I. The coengagement of the TCR and integrins in primary T cells indeed resulted in marked increases in IL-2R -chain expression and IL-2 production; IL-2 production was first detectable at 4–8 h poststimulation and reached a plateau at 8–12 h poststimulation. Conversely, the inhibition of integrin-dependent homotypic clustering by soluble anti-integrin Abs completely abrogated the production of IL-2 observed in anti-CD3-stimulated cells. To further confirm that IL-2 production was a limiting step in the integrin-dependent G₁ to S transition of TCR-stimulated T cells, we performed complementation experiments in which low doses (20 U/ml) of exogenous IL-2 were added to the cultures under the various conditions. Table I clearly demonstrates that the addition of exogenous IL-2 completely rescues S phase entry of T cells in which integrin-dependent homotypic clustering has been blocked by soluble anti-integrin Abs. Interestingly, exogenous IL-2 did not induce cell cycle progression in T cells stimulated with soluble anti-CD3 Abs or via integrin alone (not shown), indicating that TCR triggering or aggregation of integrins per se does not confer IL-2 sensitivity. Final evidence that endogenously produced IL-2 in costimulated T cells is indeed responsible for their entry into S phase was obtained using a neutralizing anti-IL-2 antiserum that efficiently (>60%) blocked DNA replication in costimulated T cells and completely prevented the effects of exogenous IL-2 in the complementation experiments described above (Table I). These results clearly demonstrate that integrin-dependent IL-2 production is both necessary and sufficient to drive IL-2-sensitive cells into S phase.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

To address these issues in detail we explored the contribution of ß₂ integrin engagement throughout the G₀ and G₁ phases of the cell cycle in TCR-stimulated primary T lymphocytes. Our results suggest that leukocyte integrins complement TCR-driven mitogenic signals not as a result of their direct clustering, but as an indirect response to integrin-dependent organization of the actin cytoskeleton. The integrity of the actin cytoskeleton and a sustained spread cell shape appear to be required both at an early phase (0–2 h) following TCR triggering, to allow efficient MAP kinase activation and the acquisition of IL-2 sensitivity, and at a later stage in the process (4–8 h), to coordinately effect the expression of genes (such as IL-2) required for G₁ to S transition in primary T lymphocytes.

Since TCR internalization has recently been reported to correlate strictly with the efficiency of T cell activation (16, 26, 27), we first analyzed whether coengagement of integrins alters the extent and kinetics of Ab-induced TCR internalization. We showed that despite a remarkable effect of costimulation in facilitating G₁ to S transition in TCR-stimulated T cells, the rate of TCR internalization was not affected by simultaneous engagement of integrins with the specific ligand ICAM-1. These data are in agreement with recent reports indicating that the requirements for inducing TCR down-regulation and T cell activation show little or no correlation (13, 41). However, as anti-CD3 Ab was used as a surrogate Ag in our study, we cannot exclude that coengagement of integrin may affect the internalization of TCRs recognizing a physiologic ligand. Indeed, a recent report shows that LFA-1-deficient T cells display a reduced down-regulation of the TCR when exposed to limiting doses of Ag (16).

Activation of the MAP kinase cascade following engagement of the TCR is a prerequisite for efficient IL-2 transcription in response to antigenic stimulation in the Jurkat T cell line (42). Our findings indicate that in primary T cells, TCR-induced activation of the three main mammalian MAP kinase members (ERK, JNK, and p38) is strongly dependent on cell spreading and actin cytoskeleton reorganization, as previously reported for growth factor-stimulated nonlymphoid cells in the case of ERK (5). Further, our results demonstrate that ligand-engaged ß₂ integrins, similar to ß₁ and ß₄ integrins, do effect the activation of both ERK (30, 43, 44) and JNK (45) and synergize with the TCR in promoting efficient MAP kinase activation. However, our results also suggest that differences in the extent, kinetics, or subcellular localization of activated MAP kinases do not entirely account for the costimulatory effect of integrins on TCR-induced G₁ to S transition in primary T lymphocytes. Consistent with this observation, we demonstrate that integrin-dependent costimulation and cytoskeletal integrity are also required at a relatively late stage (i.e., 8 h) during cell cycle progression, well beyond the exhaustion of TCR-induced activation of MAP kinases, as shown by CCD and soluble anti-integrin Ab blocking experiments. These results confirm and extend previous findings obtained using mitogen-stimulated human fibroblasts (46) and suggest that T lymphocytes, like most somatic cells, can be considered anchorage dependent for proliferation in response to mitogenic stimuli. However, in our model of T cell stimulation, and most likely under physiological conditions in which T cells form conjugates with APCs, anchorage appears to be mainly provided by integrin-dependent intercellular adhesion as opposed to cell-extracellular matrix adhesion.

As in our model integrin-dependent adhesion was clearly essential for progression through the G₁ phase of the cell cycle, which ends with hyperphosphorylation of pRb, we hypothesized that integrin stimulation is directly or indirectly required to complete the necessary biochemical modifications of cell cycle-related proteins that ultimately lead to pRb hyperphosphorylation and transit through the restriction point R. These include increased expression of D-type cyclins, phosphorylation followed by ubiquitin-dependent degradation of Cdk inhibitors, and selected phosphorylations and dephosphorylations of Cdks that culminate in their increased catalytic activity (19, 47). Interestingly, among the components of G₁ phase-related Cdk complexes whose levels were analyzed in this study, the only sizable variations observed that could be ascribed directly to integrin-dependent stimulation were a moderate increase in D3 cyclin expression and a sharp decrease in the level of the Cdk inhibitor p27^kip. These events were paralleled by increased catalytic activity of Cdk4 and Cdk6 kinases. Our findings extend previous results obtained in nonlymphoid cells, showing that anchorage is involved in regulating the transcriptional and translational levels of D-type cyclins and in reducing the steady state levels of p27^kip (48). Notably, recent reports have shown that p27^kip governs Cdk activity during G₁ to S transition in T lymphocytes (35), and that p27^kip degradation (36) as well as D-type cyclin expression, are IL-2-dependent processes that underlie the catalytic activation of cyclin/Cdk complexes at the G₁/S boundary (24, 49, 50). This prompted us to analyze whether integrin-dependent costimulation was critically affecting the levels of IL-2 produced in response to TCR triggering. Several independent observations confirm that production of limiting amounts of IL-2 is the most relevant contribution of integrin-dependent adhesion in TCR-stimulated cells. First, abrogation of integrin-dependent homotypic clustering completely prevents IL-2 production in our model. Second, the defect in G₁ to S transition observed by blocking integrin-dependent adhesion can be complemented by addition of low doses of exogenous IL-2, suggesting that in the absence of integrin costimulation, TCR-stimulated T cells acquire sensitivity to the growth factor. Third, a neutralizing anti-IL-2 antiserum markedly inhibits T cell proliferation induced by integrin-dependent costimulation. Finally, increased steady state levels of IL-2 mRNA, mostly due to post-transcriptional regulation of its half-life, can be observed in TCR-stimulated T cells upon coengagement of integrins (G. J. Wang, J. Geginat, R. Pardi, and J. R. Bender, manuscript in preparation). Interestingly, our results show that, unlike D3 cyclin, D2 cyclin expression is relatively independent of IL-2 production and integrin stimulation, thus confirming previous reports showing that these G₁-specific cyclins are independently regulated in mitogen-stimulated T lymphocytes (24, 49).

The existing evidence concerning in vivo and in vitro T cell responses in leukocyte integrin-deficient (LFA-1^-/-) mice (16, 17, 18) essentially indicate that 1) LFA-1^-/^- animals have normal numbers of mature peripheral T cells, suggesting that thymic maturation is unaltered in the absence of leukocyte integrin-dependent signaling; 2) in vivo responses to some antigenic challenges (delayed-type hypersensitivity, syngeneic and allogeneic tumors), but not others (lymphocytic choriomeningitis virus and vesicular stomatitis virus infections), are reduced; 3) in vitro proliferative responses to TCR triggering are generally reduced, but can be restored by increasing the number and/or the affinity of TCR molecules engaged in Ag recognition or by inducing persistent T cell spreading using solid phase bound anti-TCR Abs in presence of APCs. How can these findings be reconciled with our results? While alternative adhesion/costimulatory molecules could be engaged at supraphysiologic doses of Ag, thus complementing the integrin-dependent step in T cell proliferation, the absence of integrin-mediated signaling during Ag priming in the thymus could allow T cell proliferation and survival in response to high affinity TCR-Ag interactions, thereby resulting in faulty negative selection. If this was the case, the TCR repertoire in LFA-1^-/- animals could be qualitatively altered and possibly skewed toward autoreactivity. Alternatively, it could be postulated that IL-2 production, which is critically affected by engagement of integrins in our model, is a relatively integrin-independent process during thymic maturation of T cell precursors.

In conclusion, our findings suggest that the existing models explaining how integrins costimulate cell cycle progression in primary T cells are not mutually exclusive. On the one hand, integrin-dependent spreading appears to be required for efficient activation of the MAP kinase cascades by the TCR. On the other hand, a sustained spreading mediated by ligand-engaged integrin is important throughout G₁ phase progression, when early signals delivered by the TCR are exhausted and critical transcriptional events, such as IL-2 production, are taking place. These results suggest that T lymphocytes represent a peculiar case of anchorage dependence, in which integrin-mediated adhesion is involved in a feedback mechanism controlling a sustained production of the essential growth factor, rather than in simple cooperation with the mitogenic cytokine to effect cell cycle progression and initiation of DNA replication.

	Acknowledgments

 
We thank I. de Curtis for useful comments and suggestions, P. Panina Bordignon for performing ELISAs, and S. Putignano for invaluable technical assistance.

	Footnotes

 
¹ This work was supported by a European Community predoctoral fellowship (ERB40021GT957566; to J.G.), in part by grants from Deutscher Akademischer Austauschdienst (to J.G.) and Telethon, Associazione Italiana per la Ricerca sul Cancro, and Ministero dell’Universitá e della Ricerca Scientifica e Tecnologica (to R.P.), and by National Institutes of Health Grant R01HL43331 (to J.R.B.).

² Address correspondence and reprint requests to Dr. Ruggero Pardi, Human Immunology Unit, DIBIT-Scientific Institute San Raffaele, via Olgettina 58, I-20132 Milan, Italy. E-mail address:

³ Abbreviations used in this paper: MAP, mitogen-activated protein; ERK, extracellular signal-regulated kinase; JNK, Jun N-terminal kinase; Cdk, cyclin-dependent kinase; CCD, cytochalasin D; GST, glutathione S-transferase; BrdU, 5-bromo-2'-deoxyuridine; pRb, retinoblastoma protein; Cki, cyclin-dependent kinase inhibitor; MEK, MAP/ERK kinase.

Received for publication November 9, 1998. Accepted for publication February 3, 1999.

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