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Dual Function of the Extracellular Matrix: Stimulatory for Cell Cycle Progression of Naive T Cells and Antiapoptotic for Tissue-Derived Memory T Cells¹

Andreas Sturm^2,*,, Kimberley A. Krivacic^2,, Claudio Fiocchi^*, and Alan D. Levine^3,*,,

Departments of ^* Medicine, Pathology, and Pharmacology, University Hospitals of Cleveland, Case Western Reserve University School of Medicine, Cleveland, OH 44106; and Division of Gastroenterology and Hepatology, Department of Medicine, Charité-Campus Virchow Clinic, Berlin, Germany

	Abstract

Top Abstract Introduction Materials and Methods Results Discussion References

 
Tissue T cells encounter Ag in a distinct microenvironment, where they are embedded in the interstitial extracellular matrix (ECM). In contrast, while naive T cells are exposed to Ag in the lymph node, immediately after naive T cells are activated they must extravasate into the ECM to function effectively. Because integrin-mediated adhesion to the ECM modulates cell cycle progression and survival in adherent nonimmune cells, we hypothesize that blood and tissue-derived T cells have similarly adapted their behavior to their first or continued encounter with ECM. T cells from peripheral blood (PBT) and tissue (the intestinal lamina propria T cell (LPT)) were stimulated with anti-CD3-coated beads in the presence or absence of native ECM derived from intestinal fibroblasts, plate-immobilized fibronectin, or collagen type I. Native ECM and collagen, but not fibronectin, induced in anti-CD3 activated PBT a 4- to 5-fold increase in the entry, progression, and completion of the cell cycle over that triggered by anti-CD3 alone. Neutralizing ₁ integrin Abs abrogated this increase. None of these ECM proteins stimulated cell cycle progression in LPT. In contrast, anti-CD3 activation of LPT in the presence of native ECM and fibronectin reduced activation-induced cell death by 40%. These results demonstrate that naive and effector/memory T cells respond differently upon exposure to specific ECM components. When naive PBT encounter Ag in the context of ECM, their progression through the cell cycle is enhanced, favoring clonal expansion; while tissue T cell longevity may be mediated by interactions with the ECM.

	Introduction

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Progression through the cell cycle and clonal expansion are critical for T cells to establish an effective immune response. While ligation of the TCR selects for the appropriate cell with the required antigenic specificity, engagement of other cell surface receptors further regulates the T cell response (1, 2, 3). Classically, T cell activation requires at least two distinct signals. The first is Ag-specific stimulation through the TCR by cognate peptide/MHC complexes on APC. The second signal is generally considered to be provided by costimulatory and accessory molecules found on APC, which bind to counterreceptors on the T cell, including CD28, CD2, and the ₂ integrin, LFA-1 (2, 3). In addition, components of the interstitial extracellular matrix (ECM)⁴ also modulate the response of the T cell and regulate its differentiation (4, 5, 6, 7).

T cell adhesion to the ECM is mediated by integrins, a large family of heterodimeric receptors consisting of eight subunits and 18 -chains that control matrix ligand specificity (8, 9). Most somatic cells are anchorage dependent (10, 11, 12), requiring integrin-mediated adhesion to transit through the G₁-S restriction point, by activating complexes of cyclin-dependent protein kinases (Cdk) and cyclins (13, 14, 15). In turn, the retinoblastoma protein (Rb) is hyperphosphorylated, thus allowing cell cycle progression (16). In addition, somatic cells receive survival signals when adherent, but undergo apoptosis upon loss of integrin-mediated cell attachment (17, 18, 19). Because of their ability to spend extended periods of time in circulation, until recently T lymphocytes were considered to be the exception to the rule of anchorage dependence (20). However, effector/memory T cells encounter their cognate antigenic peptide in vivo within tissues in the presence of a distinct three-dimensional extracellular (ECM) microenvironment, which may act as a dynamic participant in the activation of these cells (21). Therefore, we hypothesize that T cells also possess an integrin-mediated adhesion dependency that modulates their proliferation, survival, and differentiation.

The contribution of ₁ integrin costimulation to T cell activation is not fully understood. Several studies have shown that fibronectin, but not collagen type I, coimmobilized with stimulatory anti-CD3 Abs induces freshly isolated T cell proliferation (4, 5, 6). In contrast, other studies have demonstrated that collagen type I, in a dose-dependent manner, stimulated anti-CD3 activated freshly isolated T cell proliferation (4, 22). Others suggested that resting peripheral blood T cells (PBT) are not representative of the populations of extravascular effector cells that engage ECM components and reported that T cell lines generated in vitro were stimulated by collagen type I when coimmobilized with anti-CD3 (7). More recent evidence also suggests that integrin engagement modulates T cell apoptosis. For example, ₄₁ and _L2 integrin ligation enhances activation-induced cell death (AICD) in Ag-specific T cell clones (23), whereas the ₂₁ integrin inhibits AICD in Jurkat T cells (24). We recently reported that exposure to collagen during PBT activation rapidly generates an integrin expression profile and adhesive phenotype for ECM that mirrors the characteristics of tissue-derived T cells (25).

In the present study, we addressed these contradictory findings, by developing an in vitro model system that allows us to evaluate the costimulatory capacity of three-dimensional ECM derived from human intestinal fibroblasts (native ECM), purified fibronectin, or collagen on freshly isolated circulating naive PBT, effector/memory PBT, and tissue-derived intestinal T cells, the prototypical extravascular effector/memory T cells. In addition, we recreated the three-dimensional geometry of Ag presentation in tissue by delivering the signals for TCR activation by anti-CD3-coated polystyrene beads. We performed a detailed analysis of the ability of ₁ and ₂ integrins to regulate the key events of cell cycle progression, and showed that naive and memory T cell populations respond differently to specific components of the ECM. Circulating PBT are costimulated by native ECM or collagen type I for entry into and progression of the cell cycle, while tissue T cells are protected against apoptosis by fibronectin. These results highlight the unique functional role exerted by the ECM in differentially modulating T cell responses, adjusting the immune outcome depending upon the origin and state of maturation of the responding T cell subset.

	Materials and Methods

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Reagents and Abs

For T cell isolation, the mucosal strips derived from surgically resected colons were digested in collagenase type 3 (206 U/mg) and deoxyribonuclease type 1 (3228 U/mg), both from Worthington Biochemical (Lakewood, NJ). Recombinant human IL-2 (Chiron, Emeryville, CA), and anti-CD3 mAb (OKT3; Ortho Diagnostics, Raritan, NJ) were used for T cell activation. Polystyrene, sulfate-coated beads (5.3 µm; Interfacial Dynamics, Portland, OR) were used for T cell stimulation. Fluorescein-conjugated cyclin B1 was purchased from BD Pharmingen (San Diego, CA), anti-phosphohistone H3 Ab was obtained from Upstate Biotechnology (Lake Placid, NY), and secondary Alexa 350-labeled goat-anti-rabbit IgG was obtained from Molecular Probes (Eugene, OR). Propidium iodide (PI) was purchased from Calbiochem (San Diego, CA). All protease and phosphatase inhibitors used for Western blotting were purchased from Sigma-Aldrich (St. Louis, MO). The Abs against human retinoblastoma protein, cyclin A, p21, p27, and p53 were obtained from BD Pharmingen. For immunofluorescence, cells were stained with a mAb against human ₁ (clone 1973Z; Chemicon International, Temecula, CA). Anti-human integrin ₂ mAb clone P1E6, anti-human integrin ₃ mAb clone P1B5, anti-human integrin ₄ clone P4C2, anti-human integrin ₅ clone P1D6, and anti-human integrin ₁ clone P4C10 were purchased from Invitrogen Life Technologies (Grand Island, NY). Neutralizing Ab to human integrin ₂ (clone P1388Z) was purchased from Chemicon International.

Preparation of T lymphocytes from intestinal lamina propria and peripheral blood

Intestinal T cells were isolated from surgical specimen obtained from patients admitted for bowel resection, for malignant and nonmalignant conditions, including colon cancer, benign polyps, rectal prolapse, and diverticulosis. All specimens were histologically normal. Healthy volunteers served as the source of PBT. Lamina propria T cells (LPT) were isolated as previously described (26). Briefly, the dissected intestinal mucosa was freed of mucus and epithelial cells in sequential washing steps with DTT and EDTA, and digested overnight at 37°C with collagenase and deoxyribonuclease. Mononuclear cells were separated from the crude cell suspension by layering on a Ficoll-Hypaque density gradient. For LPT purification, macrophage-depleted lamina propria mononuclear cells were incubated for 30 min at 4°C with magnetically labeled anti-CD19, -CD14, and -CD16 Abs directed against B lymphocytes, monocytes, and neutrophils, respectively (MACS; Miltenyi Biotec, Auburn, CA). T cells were then collected by negative selection using MACS. PBT were isolated from heparinized venous blood using Ficoll-Hypaque density gradient and monocyte depleted. The same process of negative selection described for LPT was used to isolate PBT. As assessed by flow cytometry, the purified LPT and PBT population contained >90 and >95% CD3⁺ cells, respectively. For isolation of naive (CD45RA⁺) and memory (CD45RO⁺) PBT, PBMC were submitted to the same process of negative selection described for LPT in combination with CD45RO Abs to select the CD45RA⁺ population, or CD45RA Abs to select the CD45RO⁺ population, respectively (CD45 RA and RO Abs from Miltenyi Biotec). The purity of these naive and memory populations was >97%.

Isolation and culture of human intestinal fibroblasts (HIF)

HIF were isolated as described previously (27). Briefly, surgical specimens from patients undergoing bowel resection for colon cancer were washed in calcium- and magnesium-free HBSS (BioWhittaker, Walkersville, MD). Strips of normal mucosa, at least 10 cm from the margin, were dissected, cut in small fragments, and laid on the bottom of scored tissue culture dishes. Medium consisting of DMEM supplemented with 10% heat-inactivated FCS, 2.5% L-glutamine, 1% mixture of penicillin, streptomycin, and Fungizone (PSF), and 25 mM HEPES buffer (all from BioWhittaker) was added to the dishes, which were incubated at 37°C in humidified 5% CO₂. Fibroblast growth emanating from the fragments was observed within 3–5 days. At 3–4 wk, subconfluent fibroblasts were detached with a trypsin-versene mixture (BioWhittaker) and established in long-term cultures. Cells were fed twice weekly and split at confluence every 4–7 days.

Preparation of native ECM-coated wells

HIF, established from colonic mucosal explants obtained from control donors, were grown in flat-bottom 24-well tissue culture plates (Costar, Cambridge, MA). At confluence, monolayers were maintained in culture for an additional 12 days in the presence of 100 U/ml human recombinant TNF- (R&D Systems, Minneapolis, MN). Fibroblast monolayers were washed with HBSS and then lysed using 0.5% Triton X-100 (Sigma-Aldrich) in Dulbecco’s PBS (DPBS) with calcium and magnesium (BioWhittaker) for 10 min, followed by 0.25 N ammonium hydroxide (Fisher Scientific, Hanover Park, IL) for 2 min (28). The resultant native ECM was washed six times with 2 ml of DPBS.

Flow cytometry

Analysis of cell cycle phase distribution was performed after staining for cyclin B1, phosphohistone H3, and PI followed by flow cytometry as previously described (29). Briefly, cells were washed twice with PBS, adjusted to 1 x 10⁶ cells/sample and fixed in 90% methanol at –20°C. After fixation, cells were washed twice with PBS, incubated with a polyclonal rabbit anti-pH3 Ab for 45 min at 4°C, followed by an incubation with a goat-anti-rabbit AlexaFluor-350 Ab for 45 min at 4°C. Thereafter, cells were washed and incubated for 45 min at 4°C with cyclin B1-FITC-conjugated mAb. After the final wash, cells were resuspended in PBS and 5 µl of RNase (0.6 µg/ml, 30–60 Kunitz units; Sigma-Aldrich), incubated at 37°C for 15 min and then chilled on ice. One hundred twenty-five microliters of PI (200 µg/ml) were added before analysis by flow cytometry. Each analysis was performed on at least 25,000 events. Integrin surface receptor expression was determined after staining with Abs against human integrin _1–5 and ₁ for 45 min at 4°C, followed by FITC-goat anti-mouse Abs (BioSource International, Camarillo, CA). Cells were then washed twice in 1% BSA-PBS and fixed in 1% paraformaldehyde. Each analysis was performed on at least 10,000 events. The background level of immunofluorescence was determined by incubating cells with FITC- or PE-conjugated mouse IgG. Cell fluorescence was measured with the Elite ESP and Epics XL flow cytometers (Beckman Coulter, Miami, FL) using UV and/or 488-nm excitation and band pass filters optimized for individual fluorochromes. Flow cytometry data were analyzed with WinList (Verity Software House, Topsham, ME). As an alternate determination of apoptosis, cells were stained with a FITC-labeled mAb against annexin V (BD Pharmingen) to detect externalization of phosphatidylserine and PI to identify necrotic cells. Fluorescence on a minimum of 15,000 cells was analyzed by flow cytometry using the CellQuest software program (BD Pharmingen).

Western blotting

Cells were washed twice in cold PBS, and lysed in cell lysis buffer (1% Triton-X, 0.5% Nonidet P-40, 0.1% SDS, 0.5% sodium deoxycholate, 5 mM EDTA, a 50 mM phosphatase and 50 mM protease inhibitor mixture, 1 mM PMSF, 100 µg/ml trypsin-chymotrypsin inhibitor, 100 µg/ml chymostatin in PBS). The concentration of proteins in each lysate was measured using the Bio-Rad protein assay (Hercules, CA). Equivalent amounts of protein (10 µg) were fractionated on a 4–20% Tris-glycine gel and electrotransferred to a nitrocellulose membrane (NOVEX, San Diego, CA). Membranes were blocked overnight at 4°C with 5% milk in 0.1% Tween 20-PBS (Fisher Scientific), followed by incubation for 60 min at room temperature with the indicated primary Ab. The membranes were washed six times with 0.1% Tween 20-PBS and then incubated for 1 h with the appropriate HRP-conjugated secondary Ab (Santa Cruz Biotechnology), washed again, and incubated with a chemiluminescent substrate (Super Signal; Pierce, Rockford, IL) for 5 min. The membranes were then exposed to film (Amersham, Arlington Heights, IL).

T cell stimulation

Anti-CD3-coated beads were made by incubating 15 x 10⁶ polystyrene, sulfate-coated beads (5.3 µm) overnight at room temperature, gently rocking with the following concentrations of OKT3 mAb: 0, 10, 25, 50, 75, or 100 µg/ml in 0.2 M carbonate-bicarbonate buffer, pH 9.5. Beads were then washed three times with DPBS, and resuspended in RPMI 1640, and used immediately. Flat-bottom 96-well plates were precoated overnight at 4°C with 0, 5, 20, 100, or 500 µg/ml fibronectin in DPBS, or with collagen type I in 0.1 M acetic acid, and were then washed three times with DPBS. Freshly isolated T cells and anti-CD3-coated beads were then mixed at a 1:1 ratio in serum-free media and 1.3 x 10⁵ T cells were added to each well of a 96-well plate or 3 x 10⁶ cells in each well of a 24-well plate in the absence or presence of the indicated matrix, and incubated for 4 days at 37°C in a humidified incubator containing 5% CO₂. Cells were then harvested, fixed in 90% methanol, and stained for flow cytometry. To measure proliferation, cells were cultured in 96-well plates at 2 x 10⁵ cells/well at 37°C in a humidified incubator containing 5% CO₂ for 96 h. One-half of a microcurie of [³H]thymidine per well (DuPont/NEN, Boston, MA) was incorporated for the final 20 h of culture. Cells were harvested using a Harvester 96 (Tomtec, Orange, CT) onto glass filter mats and counted in a beta-scintillation counter (Wallac, Gaithersburg, MD). Suboptimal stimulation for PBT and LPT was observed for beads coated with 25 µg/ml OKT3. Maximal or optimal proliferation was induced with beads coated with 50 µg/ml, and a less than optimal response was noted with beads coated with 75 or 100 µg/ml OKT3.

Statistical analysis

Statistical analysis was performed using the paired Student t test (Microsoft Excel, Redmond, WA). Results are expressed as mean ± SEM, and significance was inferred with p values <0.05.

	Results

Top Abstract Introduction Materials and Methods Results Discussion References

 
Native ECM and collagen provide potent costimulation for cell cycle entry and progression by freshly isolated PBT

We have previously shown that activation of freshly isolated PBT in the presence of ECM components rapidly alters both the surface expression and functional status of integrins on PBT, thereby conditioning them to assume a more adhesive phenotype inherent to tissue-derived T cells (25). Therefore, we investigated the functional response of PBT when their TCR was cross-linked concurrently with their exposure to native ECM or two individual ECM components. PBT were activated by anti-CD3-coated polystyrene beads, used as surrogate APCs, and plate immobilized native ECM, fibronectin, collagen type I, or BSA as a control, and we assessed the role of integrin-dependent costimulation on cell cycle progression (Fig. 1). The aim of this in vitro model is to mimic the in vivo setting in which TCR and ₁ integrin substrates are delivered to the T cell on two opposing surfaces, with the goal of addressing the ongoing controversy of whether integrins transduce signals required for cell cycle progression or simply provide the adhesive contact needed by the TCR to engage its cognate ligand (4, 5, 6). Resting T cells reside in the G₀/G₁ phase of the cell cycle and to progress through the cell cycle, they must enter and complete the S and G₂/M phases. Progression through the S phase is controlled by cyclin A, and cyclin B1 drives the cell through late G₂ phase up to mitosis, ensuring the initiation of cell division (30). The level of cyclin B1 was examined by flow cytometry in conjunction with PI staining to exactly localize its change within the cell cycle. Furthermore, we measured the phosphorylation of histone H3, a protein whose modification is exclusively detectable during unfolding of DNA in mitosis (31). When PBT were stimulated in the presence of native ECM or immobilized collagen type I, but not fibronectin, a significant stimulation of cyclin B1 expression was induced (Fig. 1; p < 0.01), indicating a potent costimulatory signal for PBT to enter the G₂/M phase. PBT did not enter the cell cycle in the presence of ECM without engagement of the TCR/CD3 complex (data not shown). Confirmation of the entry of the cells into the S phase was obtained by pulsing the cells with [³H]thymidine for their final 16 h of culture. Both collagen and native ECM stimulated a 7- to 8-fold increase (20,000 cpm) in proliferation when compared with suboptimal CD3 cross-linking in the absence of matrix or CD3 cross-linking in the presence of fibronectin (2,500 cpm). Although the data in Fig. 1 illustrate the stimulation of cell cycle progression induced by collagen and native ECM at a fixed concentration of OKT3 on day 4, it is important to appreciate that cell cycle progression was monotonically increased over the 4-day period. In addition, the ability of collagen to costimulate T cell cycle progression was dose dependent and occurred at suboptimal, optimal, and superoptimal concentrations of TCR cross-linking in the presence and absence of serum (data not shown). We previously reported that collagen type I enhances the surface expression of its cognate integrin ₁₁ on freshly isolated PBT (25), an observation consistent with our current finding that collagen stimulates entry of PBT into cell cycle progression.

View larger version (23K): [in this window] [in a new window]   FIGURE 1. Native ECM and collagen, but not fibronectin stimulate TCR-activated peripheral T cell entry and passage through the cell cycle. Freshly isolated PBT (3 x 105) were stimulated with predetermined suboptimal concentrations of bead-bound anti-CD3 (25 µg/ml OKT3) in the absence or presence of native ECM, or graded concentrations of immobilized fibronectin or collagen type I. After 4 days, cells were stained for flow cytometric analysis of cyclin B1, phosphohistone H3, and DNA content (>2n, i.e. S/G2/M phases). A, Representative dot plots depict the increase in cell cycle entry (DNA content, x-axis) and progression (cyclin B1, y-axis) induced by matrix. The mean percentage of cells positive for cyclin B1 (), phosphohistone H3 () and in the S/G2/M phase () are shown for PBT activated in the presence of native ECM (B), fibronectin (C), and collagen (D).

Having established that individual ECM components differentially regulate PBT cell cycle progression when stimulated with anti-CD3 Abs, we investigated the levels of key regulatory molecules of the cell cycle in matrix-activated PBT. Immunoblot analysis showed that the phosphorylation of Rb, which is essential for the G₁/S phase transition and passage beyond the restriction point, increased when CD3-activated PBT were cultured with collagen or native ECM, but not fibronectin (Fig. 2). Similarly, strong up-regulation of cyclin A was induced in CD3-cross-linked PBT by collagen or native ECM, but not fibronectin (Fig. 2). In addition, expression of the key cell cycle regulator p21 increased after costimulation with collagen type I or native ECM, but not fibronectin in CD3-activated PBT, a finding consistent with the role that p21 plays in cell proliferation (32). In contrast, neither collagen, fibronectin, nor native ECM altered p27 protein expression. p53 expression and phosphorylation (pp53) increased only modestly in anti-CD3 activated PBT by collagen or native ECM, but not fibronectin (Fig. 2).

View larger version (98K): [in this window] [in a new window]   FIGURE 2. Native ECM and collagen differentially regulate the expression or phosphorylation of cell cycle promoters and inhibitors. Fresh PBT were stimulated with predetermined suboptimal concentrations of bead-bound anti-CD3 (25 µg/ml OKT3) in the presence of BSA-coated plastic culture plates, 5 or 500 µg/ml fibronectin or collagen type I, or native ECM for 3 days, after which expression of cell cycle regulators was assessed by immunoblotting. Each panel is representative of four to six different experiments. GAPDH was used as an internal standard.

ECM-induced cell cycle progression in PBT is mediated by ₁ integrins

The ability of native ECM and collagen type I to costimulate cell cycle progression of CD3-activated PBT suggested an integrin-dependent mechanism. We have previously shown that PBT adhesion to native ECM and collagen type I is mediated by the ₁ integrin family and that adhesion to collagen type I is almost exclusively mediated by the ₁ subunit (25). Freshly isolated PBT were costimulated with anti-CD3-coated beads and either native ECM (Fig. 3A) or collagen type I (Fig. 3B) in the absence or presence of a mAb that blocks adhesion through the ₁ or ₁ integrin chain. PBT entry into the cell cycle, as determined by the expression of cyclin B1 or the percentage of cells in the S/G₂/M phases, was significantly inhibited by neutralizing the ₁ family of integrins (p < 0.01). The presence of ₁ integrin blocking Ab during PBT stimulation with bead-bound OKT3 alone did not inhibit cell cycling (data not shown), indicating the lack of a requirement for ₁ integrin engagement in the absence of matrix. Furthermore, costimulation by type I collagen was largely mediated by the ₁ subunit, in that >70% of PBT cyclin B1 expression and entry into the S/G₂/M phases was inhibited by a neutralizing anti-₁ Ab (Fig. 3B, p < 0.01). Consistent with the identification of collagen-induced cell cycle progression being mediated by the ₁₁ integrin, neutralizing Abs to ₄ or ₅ chains did not inhibit cyclin B1 expression and entry into the S/G₂/M phases after costimulation (Fig. 3B). In contrast, neutralizing anti-₁ Ab did not inhibit PBT cell cycle progression induced by the native ECM, underscoring the importance of investigating a complex matrix of natural origin.

View larger version (17K): [in this window] [in a new window]   FIGURE 3. Native ECM-induced cell cycle progression is mediated through 1 integrins, and purified collagen stimulation is primarily mediated through 11. Fresh PBT were stimulated with 25 µg/ml bead-bound OKT3 and either native ECM (A) or 20 µg/ml collagen (B). After 4 days, the cells were analyzed by flow cytometry for cyclin B1 expression () and DNA content () S/G2/M phase). At the initiation of culture, saturating concentrations of blocking Abs to 1, 4, 5, and 1 were added as indicated. Each panel is representative of three different experiments.

We routinely observed that high density T cell cultures (>1,200,000 cells/ml) required a lower concentration of bead-bound OKT3 to achieve the same percentage of cells progressing through the cell cycle, when compared with a low density culture (300,000–600,000 cells/ml; data not shown). Therefore, we assessed whether the presence of ECM components would collaborate with the cell-to-cell contacts likely to occur in a high density culture. Freshly isolated PBT were cultured on BSA-coated plastic, fibronectin, collagen type I, or native ECM in the presence of anti-CD3-coated beads for 4 days. In the presence of plate-immobilized collagen type I or native ECM, the size and number of PBT homotypic clusters, as observed by transmitted light microscopy, were decidedly increased compared with that of cells cultured in either BSA-coated plastic or fibronectin (Fig. 4). In fact, the number and size of PBT cell clusters in the presence of fibronectin were decreased when compared with those in the BSA-coated control wells (Fig. 4). PBT activated in the presence of fibronectin acquired a spread morphology and remained as isolated cells on the bottom of the wells, while PBT activated on native ECM or collagen assumed a spread morphology but formed large aggregates. Similarly, when PBT were activated in the presence of collagen or native ECM, but not fibronectin, cell size significantly increased, as determined by increased forward scatter on a flow cytometer (p < 0.05; data not shown).

View larger version (150K): [in this window] [in a new window]   FIGURE 4. Collagen stimulates T cell homotypic clustering. Fresh PBT were stimulated with 25 µg/ml bead-bound OKT3 in the presence of BSA-coated plastic culture plates, 20 µg/ml fibronectin, 20 µg/ml collagen type I, or native ECM and on day 4, representative phase contrast microscopic images (x100 magnification) were recorded. Each panel is representative of four to six different experiments.

Engagement of ₁ integrins by the ECM compensates for the loss of ₂-mediated cell-cell adhesion and supports PBT cell cycle progression

Because anti-CD3-induced cell cycle progression is dependent on T cell density, the interaction between the ₁ and ₂ integrin costimulatory pathways to promote S phase entry in quiescent T cells was investigated. Freshly purified PBT, stimulated by bead-bound OKT3 alone, rapidly formed homotypic clusters, which were entirely dependent on ₂ integrin-mediated cell-cell adhesion, as shown by the complete inhibition of cell clustering in the presence of blocking ₂ Ab (Fig. 5, A and B). Similarly, saturating concentrations of ₂ Abs inhibited cell cycle progression of activated PBT (Fig. 5C). The inhibition of PBT cell cycle progression by blocking ₂-mediated cell clustering could be reversed by activating T cells in the presence of various concentrations of collagen type I (Fig. 5C), but not fibronectin (data not shown). These findings demonstrate that under conditions where ₂ integrins are inhibited, ligation of the ₁ subset of integrins by collagen type I during TCR triggering provides the critical alternate second signal necessary for PBT cell cycle progression in vitro.

View larger version (44K): [in this window] [in a new window]   FIGURE 5. 1 integrin-mediated adhesion overcomes the requirement for cell-cell contact. Fresh PBT were stimulated with 25 µg/ml bead-bound OKT3 in the absence (A) or presence (B) of functional blocking Ab directed against the 2 chain. On day 4, representative phase contrast microscopic images from three donors (x100 magnification) were recorded (C). Parallel PBT cultures, activated with bead-bound OKT3 in the presence of a blocking Ab to 2 integrin, were incubated with increasing concentrations of collagen type I. Expression of cyclin B1 () and DNA content (, S/G2/M phase) on day 4 was determined by flow cytometry. Each panel is representative of three different experiments.

The ability of collagen to induce cell cycle progression in activated PBT is somewhat paradoxical in light of previous reports suggesting that collagen is sequestered from the T cells within the secondary lymph nodes by reticular cells (21, 33). Therefore, consistent with our previous demonstration of distinctive adherence properties of tissue T cells to the ECM (25), we hypothesized that PBT are first exposed to collagen after extravasating into their final tissue of residence. In support of this possibility, it was previously reported that mitogen-stimulated PBT lines derived in vitro, used as models of effector/memory T cells, are more potently simulated by collagen type I than fibronectin (7). We chose to examine the effects of native ECM, fibronectin, and collagen on freshly isolated LPT, a population of effector/memory cells (34) that maintain a CD4⁺ to CD8⁺ ratio similar to that of PBT (35), which reside in continuous contact within the interstitial matrix and express collagen-binding ₁ integrins (25). LPT were stimulated with anti-CD3- coated polystyrene beads in the absence and presence of native ECM, fibronectin, and collagen type I. Surprisingly, native ECM, fibronectin, or collagen type I had no effect on anti-CD3 activated LPT cell cycle progression (Fig. 6), even though a large percentage of LPT are positive for ₁₁, ₂₁, ₃₁, and ₄₁ integrins (see Fig. 8 below) (25). Similarly, native ECM, fibronectin, or collagen type I had no effect on LPT proliferation as measured by [³H]thymidine incorporation (data not shown).

View larger version (23K): [in this window] [in a new window]   FIGURE 6. Tissue-derived T cells are not stimulated by native ECM, fibronectin, or collagen. Freshly isolated LPT were stimulated with predetermined suboptimal concentrations of bead-bound anti-CD3 (25 µg/ml OKT3) in the absence or presence of native ECM, or graded concentrations of immobilized fibronectin or collagen type I. After 4 days, cells were stained for flow cytometric analysis of cyclin B1, phosphohistone H3, and DNA content. A, Representative dot plots demonstrating no change in cell cycle entry (S and G2/M phase, x-axis) and progression (cyclin B1, y-axis) induced by matrix are shown. The mean percentage of cells positive for cyclin B1 (), phosphohistone H3 () and in the S/G2/M phase () are shown for LPT activated in the presence of native ECM (B), fibronectin (C), and collagen (D). Each panel is representative of four to six different experiments.

View larger version (34K): [in this window] [in a new window]   FIGURE 8. Integrin expression is distinct in naive PBT, effector/memory PBT, and LPT. Flow cytometric analysis of 1 integrin surface expression (1 through 5) on freshly isolated naive PBT, effector/memory PBT, and LPT was performed. Negative control cells were gated to contain <3% positive cells. Both PBT and LPT populations were >90% CD3 positive. Representative histograms, the number of percent-positive cells, and the mean fluorescence intensity (MFI) of four experiments are shown.

The insensitivity of LPT to ECM-induced T cell proliferation may be due to either their residency in the interstitial matrix of the lamina propria or their effector/memory phenotype. To address this point, PBT were fractionated by negative selection into naive CD45RA⁺ and effector/memory CD45RO⁺ populations, and stimulated with anti-CD3-coated beads in the absence or presence of native ECM or increasing concentrations of fibronectin and collagen type I. The assignment of CD45RA⁺ T cells as naive was confirmed by demonstrating that this isolated cell population was also L-selectin (92 ± 6.2%) and CCR7 (88 ± 7.8%) positive. Native ECM and collagen costimulated an anti-CD3 response in naive CD45RA⁺ but not CD45R0⁺ effector/memory PBT, as shown by increased expression of cyclin B1 and phosphorylation of histone H3 (Fig. 7). This costimulation of naive PBT entry and completion of the cell cycle by collagen was dose dependent. No combination of anti-CD3 beads and fibronectin affected naive PBT. Similarly, no combination of anti-CD3-coated beads and matrix stimulated CD45RO⁺ PBT, suggesting this resistance to proliferation may be due to the effector/memory phenotype of the T cell, and is independent of their tissue origin. We conclude that when naive CD45RA⁺ PBT are activated by their cognate antigenic peptide/MHC complex in the presence of collagen, engagement of ₁ integrins provides costimulatory signals that enhance the entry and progression of the cell cycle.

View larger version (32K): [in this window] [in a new window]   FIGURE 7. Naive, but not effector/memory, populations of PBT enter the cell cycle in response to TCR cross-linking in the presence of native ECM and collagen. PBT were fractionated into naive (CD45RA+) and effector/memory (CD45RO+) populations by negative magnetic selection. Each T cell subset was analyzed for cell cycle progression in the absence and presence of ECM as described in the legend for Fig. 1. A, Representative flow cytometric analysis of cyclin B1 (y-axis) vs DNA content (x-axis) for native ECM, fibronectin, and collagen type I. The mean percentage of cells positive for cyclin B1 (), phosphohistone H3 () and in the S/G2/M phase () are shown for naive (B) and effector/memory (C) PBT activated in the presence of native ECM and collagen.

Distinctive ₁ integrin expression profiles among naive PBT, effector/memory PBT, and LPT

Our findings that native ECM and collagen type I promote cell cycle progression of naive, but not effector/memory PBT or LPT, suggest that the surface expression of ₁ integrins on these three T cell populations may differ. These differences may account for their divergent response to the ECM. Therefore, we compared the expression of the integrin subunits ₁-₅ and ₁ on freshly isolated CD45RA⁺ PBT, CD45RO⁺ PBT, and LPT, which are almost exclusively CD45RO⁺ (34). Both effector/memory PBT and LPT were composed of a homogeneous population of cells expressing the ₁ integrin subunit with distinct levels of expression (Fig. 8). In contrast, naive CD45RA⁺ PBT showed a bimodal expression of the ₁ chain, consisting of both a bright and dim population of cells (36), which differs from an earlier report attributing the bimodal distribution to CD45RO⁺ cells (37). However, the overall percentage of ₁ integrin expression and the mean fluorescence intensity is similar among all T cell populations examined. Expression of individual integrin chains showed that a larger percentage of LPT express ₁ and ₃ on their surfaces, but were almost devoid of ₅ surface expression when compared with that of both CD45RA⁺ and CD45RO⁺ PBT. There are also modest differences in integrin expression between CD45RA⁺ and CD45RO⁺ PBT, in that a higher percentage of naive PBT express ₃ and a higher percentage of effector/memory PBT express ₁ when compared with one another. However, the overall level of expression among these T cell subsets was similar, as noted by minimal differences in the mean fluorescence intensity of the integrin-positive cells. In summary, the expression of the ECM-specific ₁ integrin family differs between naive and effector/memory circulating T cells and clearly differs between effector/memory circulating PBT and tissue LPT. However, differences in integrin surface expression do not account for the distinct costimulatory responses in these various T cell populations, as collagen costimulation promoted cell cycle progression of both the ₁ bright and dim population of PBT. These results indicate that not only the phenotype of the T cell (effector/memory vs naive), but also the origin of the cell, including its prior exposure to the surrounding interstitium, determine its integrin surface expression.

Native ECM and specifically fibronectin inhibit AICD in tissue-derived memory T cells

We have previously reported that freshly isolated unstimulated LPT undergo a higher spontaneous rate of apoptosis in culture than do CD45RA⁺ and CD45RO⁺ PBT (38). In addition, the proportion of apoptotic LPT increases dramatically following anti-CD3 stimulation, whereas there was no significant increase in apoptosis in stimulated PBT (38). It is well established that most mesenchymal cells receive survival signals when adhered to an ECM and undergo apoptosis on the loss of integrin-mediated cell attachment (18). Because LPT are not costimulated to enter the cell cycle by native ECM or its components, we hypothesized that a major function of the interstitial matrix is to provide survival signals for those effector/memory T cells that are embedded within the ECM. Therefore, we assessed whether the presence of ECM during LPT activation would reduce apoptosis. Fresh PBT and LPT were stimulated by OKT3 Ab in the presence of native ECM, fibronectin, or collagen type I, and the degree of apoptosis quantified by PI staining, as previously described (39) (Fig. 9). One half of LPT underwent apoptosis following CD3 stimulation in the absence of ECM (BSA control). In contrast, activating LPT in the presence of native ECM or fibronectin afforded a significant protection against apoptosis (40%). No protection from AICD was provided by collagen. Not surprisingly, very few PBT underwent AICD (<7%), and this low level of apoptosis was unaltered in the presence of ECM. To confirm and extend these observations, we evaluated the protection afforded by fibronectin from AICD-induced apoptosis to LPT by measuring the expression of phosphatidylserine (annexin V staining) on the outer leaflet of the dying T cell. Fibronectin, but not collagen, protected LPT from apoptosis (by 45–50%) even in the presence of neutralizing Abs against the ₁ and ₁ through ₅ integrin subunits. These findings demonstrate that upon activation, tissue T cells, which are in continuous contact with components of the ECM, depend on their interstitial microenvironment for survival, possibly to preserve their locally needed immune effector function.

View larger version (12K): [in this window] [in a new window]   FIGURE 9. Native ECM and fibronectin, but not collagen, inhibit apoptosis in TCR-activated tissue-derived T cells. Freshly isolated LPT were stimulated with predetermined suboptimal concentrations of bead-bound anti-CD3 (25 µg/ml OKT3) in the absence or presence of native ECM, 20 µg/ml fibronectin, or 20 µg/ml collagen type I. After 4 days, cells were stained for flow cytometric analysis of DNA content. The percentage of cells containing subdiploid DNA content is indicated (n = 3–4).

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

Clonal expansion is an essential component of a T cell response. Previous reports have investigated the role of both collagen and fibronectin engagement of surface integrins as potential accessory signals during T cell activation (4, 5, 6, 22). These studies were limited by the lack of availability of critical cell cycle reagents, and [³H]thymidine incorporation, which is believed to evaluate cell proliferation, was used as the sole indicator of cell growth. Because entry into and completion of the cell cycle is tightly regulated (43, 44, 45), we focused on individual checkpoints that control cell cycle progression (46). Resting T cells reside in the G₀/G₁ phase and must enter and complete the S and G₂/M phases. Entry and progression through the S phase is controlled by Rb phosphorylation and cyclin A (47), while cyclin B1 regulates the late G₂ phase up to mitosis, ensuring the initiation of cell division (30). Completion of cell division is marked by the phosphorylation of histone H3 (31, 48). In this report, we demonstrate that native ECM or collagen costimulation of PBT increases Rb phosphorylation and therefore stimulates cell cycle progression beyond the restriction point (47). Together, our results establish that engagement of the TCR in the presence of ECM enhances clonal expansion of an Ag-inexperienced PBT.

Previous reports by different investigators disagree on the individual abilities of collagen or fibronectin to modulate PBT proliferation (4, 5, 6, 22). Two investigations demonstrated that collagen displayed no costimulatory activity, while fibronectin did (5, 6, 7), which contrasts with our and other studies. In the majority of these reports, both anti-CD3 and the matrix protein were coimmobilized onto the surface of a plastic culture dish (4, 5, 6, 22). It is likely that adhesion to fibronectin itself via ₅₁, not costimulation by fibronectin, mediated the enhanced T cell proliferation, as suggested by the authors (5, 6, 7). In other reports, the investigators were unable to detect T cell proliferation with anti-CD3 alone (4, 5), an unlikely scenario. To address this controversy, we chose an in vitro model that separates events initiated by adhesion from those attributable to costimulation. Under physiological conditions, opposing surfaces of the migrating T cell encounter ECM and APC, therefore, we respected the geometry of the adherent PBT by engaging their TCR with beads coated with anti-CD3 Ab. At every concentration of beads tested, both native ECM and collagen type I exhibited a concentration-dependent costimulation of PBT cell cycle progression. Reviewing earlier reports revealed that when those investigators used anti-CD3-coated beads, instead of coimmobilization of Ab and matrix, the T cell proliferative response to fibronectin was lost (5, 6), supporting our findings. Furthermore, other investigators, coimmobilizing anti-CD3 and matrix, did observe that collagen costimulated T cell proliferation (4, 6, 22), similar to our results. In light of our recent report that ₅₁ is solely responsible for adhesion of PBT to fibronectin (25), an alternate interpretation of the earlier data, suggested and supported by Geginat et al. (20), is that increased adhesion to the plate via fibronectin expands the surface area of the T cell in contact with anti-CD3 and thus enhances the effective concentration of CD3 cross-linking in the culture.

Integrin-dependent cell cycle progression in response to growth factor stimulation is observed in anchorage-dependent cell types (49), and in anti-CD3 activated PBT that require _L2 (LFA-1) integrin driven homotypic cell clustering. When T cells are activated, ₂ integrin engagement is a requisite for the induction of G₁ cyclins D2 and D3, activation of Cdk6, and phosphorylation of Rb (20). In agreement, we find that higher PBT cell density requires less activation to enter the cell cycle due to increased engagement of _L2. However, costimulation by ₁ integrins (i.e., collagen type I engagement) is not observed at high cell density, due to the increased cell-cell adhesion mediated by ₂ integrins. In this light, earlier reports attributing a costimulatory activity to fibronectin after coimmobilization with anti-CD3 may be reinterpreted as showing that fibronectin enhanced the capacity for ₂-mediated cell-cell contact by increasing the local T cell density at the site of adhesion.

Loss of PBT cell cycle progression due to the inhibition of ₂ engagement is partially reversed by increased ₁ engagement by collagen or native ECM. This suggests three possible mechanisms by which collagen costimulates cell cycle progression: 1) engagement of ₁ integrins provides the necessary costimulatory signals needed; 2) ₁ integrin ligation increases the affinity/avidity of the ₂ integrin family, and the resulting increase in homotypic clustering drives costimulation; 3) a combination of ₁ integrin engagement by collagen and the increase in ₂ integrin engagement by its ligand, ICAM, together deliver the necessary signals for cell cycle progression. These results illustrate three major effects of integrin engagement: 1) activation through the TCR/CD3 complex converts integrins from the inactive to the active conformation (25); 2) integrin cross-talk modulates T cell responses and they compensate for one another (50); and 3) collagen engagement of ₁ integrins or cross-linking ₂ integrins by cell-cell contact is essential for progression through the G₁ phase of the cell cycle beyond the restriction point (20).

Only the naive population of PBT responds to native ECM and collagen costimulation, yet these cells bear a similar integrin surface expression profile as nonresponding effector/memory PBT. Therefore, integrin expression alone cannot modulate T cell entry into the cell cycle. Instead, the combination of T cell phenotype coupled to ECM exposure during T cell activation determines a functional response. We previously reported that freshly isolated PBT, preconditioned with collagen, increase their surface expression of ₁ and ₃ (25) and acquire the ability to adhere strongly to collagen, which then costimulates cell cycle progression. In fact, blood T cells may never completely interact with all of the components of the interstitial ECM when migrating through a lymph node (LN) (33, 51), as it was proposed that collagen, in particular, is sequestered from the blood T cell by a complex reticular network of dendritic cells that shield LN T cells from this protein (33, 51). This proposed structure of the LN suggests that in vivo the first contact of a CD45RA⁺ T cell with collagen may be directly within the inflamed tissue. It was reported that naive T cells reside exclusively in secondary lymphoid tissues, such as the spleen and lymph nodes, in mice that were not exposed to Ag (40). Upon Ag exposure, freshly activated naive T cells proliferate in the LN and then migrate to the lungs, liver, gut, and salivary glands (40), where they will be exposed to collagen embedded within the interstitial matrix. Based on our results, we hypothesize that the entry of these CD45RA⁺ T cells into only those tissues in which the cognate Ag is present will stimulate an additional round of proliferation during which costimulation by interstitial collagen further enhances cell cycle progression.

In contrast, LPT are completely embedded in the ECM (52), causing the cell to polarize and forcing the T cell to encounter its cognate peptide on one pole of the cell (53). Once T cells are removed from the interstitial matrix, they rapidly undergo AICD (38), which is reduced by the presence of native ECM or fibronectin. A role for integrin cross-linking as an inhibitor of apoptosis has long been appreciated with adherent cells (54, 55), and only recently has the effect of ECM been recognized for activated CD4⁺ murine T cells (56). Collagen engaging the ₂₁ integrin inhibits anti-CD3-induced, Fas-mediated cell death in Jurkat cells (24). We extended these studies by demonstrating that in freshly isolated PBT, which are resistant to AICD (38), collagen has no effect on apoptosis but instead stimulates cell cycle progression. In contrast, purified fibronectin is antiapoptotic for LPT, similar to the report that ₄₁ engagement by fibronectin rescues thymocytes from glucocorticoid-induced cell death (57). Similarly, activation of ₁ integrins with a cross-linking Ab has been reported to inhibit anti-CD3-induced apoptosis and increase proliferation of intestinal CD4⁺ T cells by 62% (58). It is tempting to speculate that fibronectin is the active component of native ECM that mediates its antiapoptotic effect on LPT. However, we approach this possibility with caution because blocking Abs to ₁ integrins do not inhibit the antiapoptotic activity of purified fibronectin. This latter finding is somewhat surprising and suggests that the antiapoptotic effect of ECM is extremely complex. After secondary antigenic activation, the apoptotic death of effector and memory T cells seems counterintuitive, as their survival in injured or infected tissue is essential. Therefore, increased survival of tissue T cells due to their interaction with the interstitial ECM is likely to contribute to and preserve normal immunity. However, if T cell activation and survival is inappropriately extended in the intestinal mucosa by integrin interactions with fibronectin, chronic inflammation may result (59). Our findings stress the importance of recognizing that both the topology and the composition of the ECM display immunoregulatory functions, directing naive and memory T cells toward distinct responses.

	Acknowledgments

 
We thank Gail West, R. Michael Sramkoski, and Megan Gottlieb for technical assistance. We also thank the Departments of Surgery and Pathology, University Hospitals of Cleveland, the Colorectal Surgery Department of the Cleveland Clinic Foundation, and the Departments of Surgery and Pathology, Charite, Campus Virchow Clinic (Berlin, Germany). The Cooperative Human Tissue Procurement Facility of the University Hospitals of Cleveland provided some tissue samples.

	Footnotes

 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

¹ This work was supported by grants from the National Institutes of Health (DK30399 and DK50984 (to C.F.) and DK-54213 (to A.D.L.)), the Deutsche Forschungsgemeinschaft and the Charité Berlin, Germany (STU247/2-1 and 247/3-1 (to A.S.)), and National Institutes of Health National Research Service Award CA-73515 (to K.A.K.).

² A.S. and K.A.K. contributed equally to this work.

³ Address correspondence and reprint requests to Dr. Alan D. Levine, Department of Medicine, Case Western Reserve University School of Medicine, 10900 Euclid Avenue, Cleveland, OH 44106-4952. E-mail address: alan.levine{at}case.edu

⁴ Abbreviations used in this paper: ECM, extracellular matrix; Cdk, cyclin-dependent kinase; Rb, retinoblastoma protein; PBT, peripheral blood T cell; AICD, activation-induced cell death; PI, propidium iodide; LPT, lamina propria T cell; HIF, human intestinal fibroblast; DPBS, Dulbecco’s PBS; LN, lymph node.

Received for publication November 10, 2003. Accepted for publication July 13, 2004.

	References

Top Abstract Introduction Materials and Methods Results Discussion References

 

1. Semnani, R. T., T. B. Nutman, P. Hochman, S. Shaw, G. A. van Seventer. 1994. Costimulation by purified intercellular adhesion molecule 1 and lymphocyte function-associated antigen 3 induces distinct proliferation, cytokine and cell surface antigen profiles in human "naive" and "memory" CD4⁺ T cells. J. Exp. Med. 180:2125.[Abstract/Free Full Text]
2. Croft, M., C. Dubey. 1997. Accessory molecule and costimulation requirements for CD4 T cell response. Crit. Rev. Immunol. 17:89.[Medline]
3. Damle, N. K., K. Klussman, P. S. Linsley, A. Aruffo. 1992. Differential costimulatory effects of adhesion molecules B7, ICAM-1, LFA-3, and VCAM-1 on resting and antigen-primed CD4⁺ T lymphocytes. J. Immunol. 148:1985.[Abstract]
4. Matsuyama, T., A. Yamada, J. Kay, K. M. Yamada, S. K. Akiyama, S. F. Schlossman, C. Morimoto. 1989. Activation of CD4 cells by fibronectin and anti-CD3 antibody: a synergistic effect mediated by the VLA-5 fibronectin receptor complex. J. Exp. Med. 170:1133.[Abstract/Free Full Text]
5. Davis, L. S., N. Oppenheimer-Marks, J. L. Bednarczyk, B. W. McIntyre, P. E. Lipsky. 1990. Fibronectin promotes proliferation of naive and memory T cells by signaling through both the VLA-4 and VLA-5 integrin molecules. J. Immunol. 145:785.[Abstract]
6. Shimizu, Y., G. A. van Seventer, K. J. Horgan, S. Shaw. 1990. Costimulation of proliferative responses of resting CD4⁺ T cells by the interaction of VLA-4 and VLA-5 with fibronectin or VLA-6 with laminin. J. Immunol. 145:59.[Abstract]
7. Rao, W. H., J. M. Hales, R. D. Camp. 2000. Potent costimulation of effector T lymphocytes by human collagen type I. J. Immunol. 165:4935.[Abstract/Free Full Text]
8. Hemler, M. E.. 1990. VLA proteins in the integrin family: structures, functions, and their role on leukocytes. Annu. Rev. Immunol. 8:365.[Medline]
9. Shimizu, Y., G. A. Van Seventer, K. J. Horgan, S. Shaw. 1990. Regulated expression and binding of three VLA (₁) integrin receptors on T cells. Nature 345:250.[Medline]
10. Assoian, R. K.. 1997. Anchorage-dependent cell cycle progression. J. Cell Biol. 136:1.[Free Full Text]
11. Schwartz, M. A., R. K. Assoian. 2001. Integrins and cell proliferation: regulation of cyclin-dependent kinases via cytoplasmic signaling pathways. J. Cell Sci. 114:2553.
12. Zhu, X., M. Ohtsubo, R. M. Bohmer, J. M. Roberts, R. K. Assoian. 1996. Adhesion-dependent cell cycle progression linked to the expression of cyclin D1, activation of cyclin E-cdk2, and phosphorylation of the retinoblastoma protein. J. Cell Biol. 133:391.[Abstract/Free Full Text]
13. Lees, E.. 1995. Cyclin dependent kinase regulation. Curr. Opin. Cell Biol. 7:773.[Medline]
14. Morgan, D. O.. 1995. Principles of CDK regulation. Nature 374:131.[Medline]
15. Grana, X., E. P. Reddy. 1995. Cell cycle control in mammalian cells: role of cyclins, cyclin dependent kinases (CDKs), growth suppressor genes and cyclin-dependent kinase inhibitors (CKIs). Oncogene 11:211.[Medline]
16. Lipinski, M. M., T. Jacks. 1999. The retinoblastoma gene family in differentiation and development. Oncogene 18:7873.[Medline]
17. Frisch, S. M., E. Ruoslahti. 1997. Integrins and anoikis. Curr. Opin. Cell Biol. 9:701.[Medline]
18. Frisch, S. M., R. A. Screaton. 2001. Anoikis mechanisms. Curr. Opin. Cell Biol. 13:555.[Medline]
19. Grossmann, J.. 2002. Molecular mechanisms of "detachment-induced apoptosis–Anoikis". Apoptosis 7:247.[Medline]
20. Geginat, J., G. Bossi, J. R. Bender, R. Pardi. 1999. Anchorage dependence of mitogen-induced G₁ to S transition in primary T lymphocytes. J. Immunol. 162:5085.[Abstract/Free Full Text]
21. Gretz, J. E., E. P. Kaldjian, A. O. Anderson, S. Shaw. 1996. Sophisticated strategies for information encounter in the lymph node: the reticular network as a conduit of soluble information and a highway for cell traffic. J. Immunol. 157:495.[Abstract]
22. Dang, N. H., Y. Torimoto, S. F. Schlossman, C. Morimoto. 1990. Human CD4 helper T cell activation: functional involvement of two distinct collagen receptors, 1F7 and VLA integrin family. J. Exp. Med. 172:649.[Abstract/Free Full Text]
23. Damle, N. K., K. Klussman, G. Leytze, A. Aruffo, P. S. Linsley, J. A. Ledbetter. 1993. Costimulation with integrin ligands intercellular adhesion molecule-1 or vascular cell adhesion molecule-1 augments activation-induced death of antigen-specific CD4⁺ T lymphocytes. J. Immunol. 151:2368.[Abstract]
24. Aoudjit, F., K. Vuori. 2000. Engagement of the ₂₁ integrin inhibits Fas ligand expression and activation-induced cell death in T cells in a focal adhesion kinase-dependent manner. Blood 95:2044.[Abstract/Free Full Text]
25. Krivacic, K. A., A. D. Levine. 2003. Extracellular matrix conditions T cells for adhesion to tissue interstitium. J. Immunol. 170:5034.[Abstract/Free Full Text]
26. West, G. A., T. Matsuura, A. D. Levine, J. S. Klein, C. Fiocchi. 1996. Interleukin 4 in inflammatory bowel disease and mucosal immune reactivity. Gastroenterology 110:1683.[Medline]
27. Strong, S. A., T. T. Pizarro, J. S. Klein, F. Cominelli, C. Fiocchi. 1998. Proinflammatory cytokines differentially modulate their own expression in human intestinal mucosal mesenchymal cells. Gastroenterology 114:1244.[Medline]
28. Fridman, R., Y. Alon, F. Doljanski, Z. Fuks, I. Vlodavsky. 1985. Cell interaction with the extracellular matrices produced by endothelial cells and fibroblasts. Exp. Cell Res. 158:461.[Medline]
29. Jacobberger, J. W., R. M. Sramkoski, S. B. Wormsley, W. E. Bolton. 1999. Estimation of kinetic cell-cycle-related gene expression in G₁ and G₂ phases from immunofluorescence flow cytometry data. Cytometry 35:284.[Medline]
30. Sherr, C. J.. 1994. G₁ phase progression: cycling on cue. Cell 79:551.[Medline]
31. Wei, Y., L. Yu, J. Bowen, M. A. Gorovsky, C. D. Allis. 1999. Phosphorylation of histone H3 is required for proper chromosome condensation and segregation. Cell 97:99.[Medline]
32. Besson, A., V. W. Yong. 2000. Involvement of p21^Waf1/Cip1 in protein kinase C -induced cell cycle progression. Mol. Cell. Biol. 20:4580.[Abstract/Free Full Text]
33. Ebnet, K., E. P. Kaldjian, A. O. Anderson, S. Shaw. 1996. Orchestrated information transfer underlying leukocyte endothelial interactions. Annu. Rev. Immunol. 14:155.[Medline]
34. Schieferdecker, H. L., R. Ullrich, H. Hirseland, M. Zeitz. 1992. T cell differentiation antigens on lymphocytes in the human intestinal lamina propria. J. Immunol. 149:2816.[Abstract]
35. Selby, W. S., G. Janossy, M. Bofill, D. P. Jewell. 1984. Intestinal lymphocyte subpopulations in inflammatory bowel disease: an analysis by immunohistological and cell isolation technique. Gut 25:32.[Abstract/Free Full Text]
36. Raju, B., C. F. Tung, D. Cheng, N. Yousefzadeh, R. Condos, W. N. Rom, D. B. Tse. 2001. In situ activation of helper T cells in the lung. Infect. Immun. 69:4790.[Abstract/Free Full Text]
37. Horgan, K. J., G. E. Luce, Y. Tanaka, T. Schweighoffer, Y. Shimizu, S. O. Sharrow, S. Shaw. 1992. Differential expression of VLA-4 and VLA-1 discriminates multiple subsets of CD4⁺CD45R0⁺"memory" T cells. J. Immunol. 149:4082.[Abstract]
38. Sturm, A., J. Itoh, J. W. Jacobberger, C. Fiocchi. 2002. p53 negatively regulates intestinal immunity by delaying mucosal T cell cycling. J. Clin. Invest. 109:1481.[Medline]
39. Sturm, A., S. Mohr, C. Fiocchi. 2002. Critical role of caspases in the regulation of apoptosis and proliferation of mucosal T cells. Gastroenterology 122:1334.[Medline]
40. Reinhardt, R. L., A. Khoruts, R. Merica, T. Zell, M. K. Jenkins. 2001. Visualizing the generation of memory CD4 T cells in the whole body. Nature 410:101.[Medline]
41. Masopust, D., V. Vezys, A. L. Marzo, L. Lefrancois. 2001. Preferential localization of effector memory cells in nonlymphoid tissue. Science 291:2413.[Abstract/Free Full Text]
42. Rossetti, G., M. Collinge, J. R. Bender, R. Molteni, R. Pardi. 2002. Integrin-dependent regulation of gene expression in leukocytes. Immunol. Rev. 186:189.[Medline]
43. Stillman, B.. 1996. Cell cycle control of DNA replication. Science 274:1659.[Abstract/Free Full Text]
44. Bird, J. J., D. R. Brown, A. C. Mullen, N. H. Moskowitz, M. A. Mahowald, J. R. Sider, T. F. Gajewski, C. R. Wang, S. L. Reiner. 1998. Helper T cell differentiation is controlled by the cell cycle. Immunity 9:229.[Medline]
45. Elledge, S. J.. 1996. Cell cycle checkpoints: preventing an identity crisis. Science 274:1664.[Abstract/Free Full Text]
46. Balomenos, D., A. C. Martinez. 2000. Cell-cycle re