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p53 negatively regulates intestinal immunity by delaying mucosal T cell cycling
Andreas Sturm,1 Jugoh Itoh,1 James W. Jacobberger,2 and Claudio Fiocchi1
1Division of Gastroenterology, Department of Medicine, and2 Ireland Cancer Research Center, University Hospitals of Cleveland, Case Western Reserve University School of Medicine, Cleveland, Ohio, USA
Address correspondence to: Claudio Fiocchi, Division of Gastroenterology, University Hospitals of Cleveland, Case Western Reserve University School of Medicine (BRB 425), 10900 Euclid Avenue, Cleveland, Ohio 44106-4952, USA. Phone: (216) 368-1668; Fax: (216) 368-1674; E-mail: cxf18@po.cwru.edu.
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1Division of Gastroenterology, Department of Medicine, and2 Ireland Cancer Research Center, University Hospitals of Cleveland, Case Western Reserve University School of Medicine, Cleveland, Ohio, USA
Address correspondence to: Claudio Fiocchi, Division of Gastroenterology, University Hospitals of Cleveland, Case Western Reserve University School of Medicine (BRB 425), 10900 Euclid Avenue, Cleveland, Ohio 44106-4952, USA. Phone: (216) 368-1668; Fax: (216) 368-1674; E-mail: cxf18@po.cwru.edu.
Find articles by Itoh, J. in: JCI | PubMed | Google Scholar
1Division of Gastroenterology, Department of Medicine, and2 Ireland Cancer Research Center, University Hospitals of Cleveland, Case Western Reserve University School of Medicine, Cleveland, Ohio, USA
Address correspondence to: Claudio Fiocchi, Division of Gastroenterology, University Hospitals of Cleveland, Case Western Reserve University School of Medicine (BRB 425), 10900 Euclid Avenue, Cleveland, Ohio 44106-4952, USA. Phone: (216) 368-1668; Fax: (216) 368-1674; E-mail: cxf18@po.cwru.edu.
Find articles by Jacobberger, J. in: JCI | PubMed | Google Scholar
1Division of Gastroenterology, Department of Medicine, and2 Ireland Cancer Research Center, University Hospitals of Cleveland, Case Western Reserve University School of Medicine, Cleveland, Ohio, USA
Address correspondence to: Claudio Fiocchi, Division of Gastroenterology, University Hospitals of Cleveland, Case Western Reserve University School of Medicine (BRB 425), 10900 Euclid Avenue, Cleveland, Ohio 44106-4952, USA. Phone: (216) 368-1668; Fax: (216) 368-1674; E-mail: cxf18@po.cwru.edu.
Find articles by Fiocchi, C. in: JCI | PubMed | Google Scholar
Published June 1, 2002 - More info
J Clin Invest. 2002;109(11):1481–1492. https://doi.org/10.1172/JCI14967.
© 2002 The American Society for Clinical Investigation
Received: January 3, 2002; Accepted: May 1, 2002
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Methods
Reagents and antibodies. For T cell isolation, the mucosa strips were digested in collagenase type 3 (206 U/mg) and deoxyribonuclease type 1 (3228 U/mg), both from Worthington Biochemical Corp. (Lakewood, New Jersey, USA). Recombinant human IL-2 (Chiron Corp., Emeryville, California, USA) and mAb’s to CD3 (OKT3; Ortho Diagnostic Systems Inc., Raritan, New Jersey, USA), CD2 (T112 and T113; generously provided by Ellis Reinherz, Dana Farber Cancer Institute, Boston, Massachusetts, USA), and CD28 (ANC28.1/5D10; Ancell Corp., Bayport, Minnesota, USA) were used for T cell activation. FITC-conjugated cyclin B1 and bromodeoxyuridine (BrdU) were purchased from BD Pharmingen (San Diego, California, USA), anti–phospho–histone H3 antibody was obtained from Upstate Biotechnology Inc. (Lake Placid, New York, USA), and secondary Alexa Fluor 350–labeled goat anti-rabbit IgG was obtained from Molecular Probes Inc. (Eugene, Oregon, USA). The Vybrant CFDA SE Cell Tracer Kit was also obtained from Molecular Probes Inc. Propidium iodide (PI) was purchased from Calbiochem-Novabiochem Corp. (San Diego, California, USA). All protease and phosphatase inhibitors used for Western blotting and rhodamine123 were purchased from Sigma-Aldrich (St. Louis, Missouri, USA). The antibodies against CDK4 and cyclin D2 were obtained from Santa Cruz Biotechnology Inc. (Santa Cruz, California, USA), and those against human cyclin D3, retinoblastoma protein, p130 (Rb2), p107, cyclins A and B1, p21, and p27 were obtained from BD Pharmingen. The ribonuclease protection assay hcc-1 was also obtained from BD Pharmingen. Nocodazole was purchased from Biomol Research Laboratories (Plymouth Meeting, Pennsylvania, USA).
Isolation of T lymphocytes from intestinal lamina propria and peripheral blood. Intestinal T cells were isolated from surgical specimens obtained from patients admitted for bowel resection for malignant or nonmalignant conditions including colon cancer, benign polyps, rectal prolapse, and diverticulosis. All specimens were histologically normal. LPTs were isolated as previously described (24). 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. If necessary, further purification was archived by subsequent Percoll density gradients. For LPT purification, macrophage-depleted lamina propria mononuclear cells were incubated for 30 minutes at 4°C with magnetically labeled anti-CD19, -CD14, and -CD16 antibodies directed against B lymphocytes, monocytes, and neutrophils, respectively (Miltenyi Biotec Inc., Auburn, California, USA). T cells were then collected by negative selection using a magnetic cell sorting system (MACS; Miltenyi Biotec Inc.).
PBTs were isolated from heparinized venous blood of healthy volunteers using a Ficoll-Hypaque density gradient and were monocyte-depleted. For isolation of naive (CD45RA+) and memory (CD45RO+) PBTs, PBMCs were submitted to the same process of negative selection described for LPTs, in combination with CD45RO antibodies to select the CD45RA+ population, or CD45RA antibodies to select the CD45RO+ population. CD45RA and CD45RO antibodies were from Miltenyi Biotec Inc.
As assessed by flow cytometry, the purified LPT and PBT populations contained >92% and >95% CD3+ cells, respectively. LPTs were 91% CD45RO+, and PBTs were 54% naive T cells (CD45RA+). The CD45RO-depleted PBT population was >98% CD45RA+ and less than 2% CD45RO+, while the CD45RA-depleted PBT population was >98% CD45RO+ and less than 2% CD45RA+.
Cells were cultured in complete medium (RPMI 1640, 10% FCS, 1.5% HEPES buffer, 2.5% penicillin-streptomycin; all from BioWhittaker Inc., Walkersville, Maryland, USA) for 3 days in a humidified incubator containing 5% CO2, alone or in the presence of cross-linked plate-bound anti-CD3 mAb (OKT3; 10 μg/ml) or soluble anti-CD2 mAb pairs (T112 and T113; 1:1000). PI content, telomerase activity, and clonal expansion of PBTs submitted to the protocol used for LPTs were comparable to those of freshly isolated PBTs.
Flow cytometry. Cell fluorescence was measured with the Elite ESP and Epics XL flow cytometers (Beckman Coulter Inc., Miami, Florida, USA) using ultraviolet and/or 488-nm excitation and band pass filters optimized for individual fluorochromes. Flow cytometry data were analyzed with WinList (Verity Software House Inc., Topsham, Maine, USA).
Analysis of cell cycle phase distribution. Flow cytometry was performed after staining for DNA content, cyclin B1, and mitotic cells essentially as previously described (25). Briefly, cells were washed twice with PBS, adjusted to 1 × 106 cells per sample, and fixed in 90% methanol at –20°C. After fixation, cells were washed twice with PBS and incubated with a polyclonal rabbit anti–phospho–histone H3 antibody for 45 minutes at 4°C, followed by an incubation with a goat anti-rabbit Alexa Fluor 350 antibody for 45 minutes at 4°C. Thereafter cells were washed and incubated for 45 minutes at 4°C with a 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 minutes, and then chilled on ice. One hundred twenty-five microliters of PI (200 μg/ml) were added prior to analysis by flow cytometry. Each analysis was performed on at least 25,000 events.
Cell surface staining for phenotypic analysis. Cell phenotype was analyzed using mAb’s conjugated to CD45RA–R-phycoerythrin (CD45A-RPE), CD45RO-FITC, and CD3-RPE (all from DAKO Corp., Carpinteria, California, USA). The background level of immunofluorescence was determined by incubating cells with FITC- or RPE-conjugated mouse IgG. After 30 minutes’ incubation on ice, cells were washed twice in 1% BSA-PBS and fixed in 1% paraformaldehyde. Each analysis was performed on at least 10,000 events.
Analysis of cell division. Analysis of cell division by dye dilution was performed using the Vybrant CFDA SE Cell Tracer Kit. Cells were washed twice with cold PBS, resuspended in PBS with 5 μM carboxyfluorescein diacetate succinimidyl ester (CFDA SE) per 1 × 106 cells, and incubated for 15 minutes at 4°C in the dark. The staining was quenched by adding 5× cell culture medium containing 10% FCS. After staining, cells were cultured alone (unstimulated), or with soluble anti-CD2 mAb pairs (T112 and T113; 1:1000) or cross-linked plate-bound anti-CD3 mAb (OKT3; 10 μg/ml), each with CD28 (5 μg/ml) and IL-2 (20 U/ml). After 4 days, cells were harvested, washed twice in cold PBS, fixed with 1% paraformaldehyde, and analyzed by flow cytometry.
Measurement of DNA synthesis time, potential doubling times, G2/M enter fractions, and tritiated thymidine uptake. Analysis of cell kinetics was performed using two-color flow cytometric techniques relying on S phase labeling with BrdU or stathmokinetic experiments relying on the accumulation of cells in the M phase by nocodazole. For the determination of S phase duration and potential doubling times, cells were grown for 3 days with or without the respective stimuli and then incubated for 60 minutes with 20 μM BrdU, which was then replaced by thymidine. At designated time points, cells were harvested and fixed with 90% methanol. The BrdU-labeled nuclei were then stained with an FITC-conjugated mAb against BrdU (BD Pharmingen). The nuclei were also stained with PI following the above-mentioned protocols. The denaturation of DNA allowing antibody binding to the incorporated BrdU was achieved by an acid treatment using a previously described protocol (26). Mathematical analysis was performed according to the methods of Begg et al. (27) and White et al. (28). The movement of BrdU-labeled cells across S phase relative to the position of G1 and G2+M was calculated by
Equation 11
where RM = relative movement, FG1 = unlabeled G1 mean red fluorescence, FG2+M = unlabeled G2+M mean red fluorescence, and FS = mean red fluorescence of the BrdU-labeled cells at time t. S phase duration (TS) was calculated as the time for one unit relative movement. The potential doubling time was computed by
Equation 22
with v defined as ln[1 + flu(t)/1 – fld(t)/2], where flu(t) = fraction of labeled, undivided cells at time t, and fld(t) = fraction of labeled, divided cells at time t.
Cells were incubated with or without the respective stimulus for 3 days. Mitosis was then inhibited by adding 100 ng/ml nocodazole. At timed intervals, cells were harvested and fixed for flow cytometric analysis. The G2/M enter rates were calculated as the percentage of cells that left or entered the respective cell cycles at the designated time points. These experiments were complemented by measurement of tritiated thymidine uptake as previously described (14).
Assessment of energy consumption by determining mitochondrial polarization status. Rhodamine123 is a fluorescent cationic dye that accumulates in the mitochondrial matrix because of its charge and solubility in both the inner mitochondrial membrane and matrix space (29). Based on the observation that accumulation of this lipophilic dye is in proportion to ΔΨ, oxygen consumption, and ATP production (30) and that de-energizing of the mitochondria decreases rhodamine123 fluorescence (31), we measured the mitochondrial membrane potential of PBTs and LPTs. Cells were stimulated with CD2 or CD3 for 4 days, harvested, washed, and resuspended in 1 ml of rhodamine123 (10 μg/ml) for 30 minutes at 37°C in the dark. The samples were washed twice in cold PBS, and fluorescence analysis by flow cytometry using an argon ion laser with an emission filter at 530 nm was performed immediately without fixation. PBTs and LPTs without stimulation and unstained samples were used as controls.
Western blotting. Treated cells were washed twice in cold PBS and lysed in cell lysis buffer (1% Triton X, 0.5% NP-40, 0.1% SDS, 0.5% sodium deoxycholate, 5 mM EDTA, 50 mM phosphatase and 50 mM protease inhibitor cocktail, 1 mM PMSF, 100 μg/ml trypsin-chymotrypsin inhibitor, and 100 μg/ml chymostatin in PBS). The concentration of proteins in each lysate was measured using the Bio-Rad protein assay (Bio-Rad Laboratories Inc., Hercules, California, USA). Equivalent amounts of protein (10 μg) were fractionated on a 4–20% Tris-glycine gel and electrotransferred to a Nitrocellulose membrane (Novex, San Diego, California, USA). Membranes were blocked overnight at 4°C with 5% milk in 0.1% Tween-20–PBS (Fisher Scientific Co., Hanover Park, Illinois, USA), followed by incubation for 60 minutes at room temperature with the indicated primary antibody. The membranes were washed six times with 0.1% Tween-20–PBS and then incubated for 1 hour with the appropriate horseradish peroxidase–conjugated secondary antibody (Santa Cruz Biotechnology Inc.), washed again, and incubated with chemiluminescent substrate (Super Signal; Pierce Chemical Co., Rockford, Illinois, USA) for 5 minutes. The membranes were then exposed to film (Amersham Life Sciences Inc., Arlington Heights, Illinois, USA). The level of individual proteins in each band was measured by densitometry and expressed as arbitrary densitometry units normalized to their respective GAPDH. This allows comparison within one protein, but not between proteins.
RNA preparation and RNase protection assay. Total RNA was extracted using the guanidinium thiocyanate method (32). The RNase protection assay for quantification of cell cycle regulator (CDKs 1–4, p27, p21, PISSLRE, and p16) mRNA was performed according to the manufacturer’s instructions (BD Pharmingen). Briefly, a human cell cycle regulator multiprobe template set (hcc-1) was labeled with α-32P uridine triphosphate (NEN Life Science Products Inc., Boston, Massachusetts, USA). Five micrograms of total RNA were hybridized with the α-32P labeled probe. The remaining RNA was digested, and the extracted product was resolved on a 0.4-mm 4% urea-polyacrylamide gel. The signal was analyzed using a phosphoimaging system for quantification (Bio-Rad Laboratories Inc.). GAPDH and L32 were used as internal control to normalize for loading differences.
Measurement of telomerase activity. Telomerase activity was measured using a photometric enzyme immunoassay for quantitative determination of telomerase activity (Telo TAGG Telomerase PCR ELISA; Roche Diagnostics GmbH, Mannheim, Germany) based on the telomeric repeat amplification protocol (TRAP) method described by Kim and Wu (33). 2 × 105 cells were lysed, incubated on ice for 20 minutes, and then centrifuged at 16,000 g for 20 minutes at 4°C. The TRAP was performed according to the manufacturer’s instructions. Briefly, telomeric repeats were added to a biotin-labeled primer during the first reaction. The elongation products were amplified by PCR. An aliquot of PCR product was denatured, hybridized to a digoxigenin-labeled, telomeric repeat–specific probe, and bound to a streptavidin-coated 96-well plate. An antibody to digoxigenin, conjugated to peroxidase, was subsequently added, and binding to digoxigenin was visualized by virtue of the ability of the enzyme to metabolize tetramethyl benzidine to produce a colored reaction product. The absorbance of the samples was measured at 450 nm (reference wavelength 690 nm) using a microplate reader (Bio-Tek Instruments Inc., Winooski, Vermont, USA) within 30 minutes after addition of the stop reagent. Mean OD values were recorded as relative telomerase activity. The immortalized human kidney cell line 293 was used as positive control. Inactivation of telomerase protein at 85°C for 10 minutes was used as negative control. For the generation of the telomeric-mediated six-nucleotide ladder, cell lysates were prepared and amplified as described above. One-third of the amplified products (1000 cell equivalents) were then separated on a 12% nondenaturing polyacrylamide gel and stained with SYBR green I nucleic acid stain (Molecular Probes Inc.).
p53 oligodeoxynucleotides and transfection protocol. Phosphorothioate p53 antisense oligonucleotide (5′-CCCTGCTCCCCCCTGGCTCC-3′) and scrambled control (5′-CGGTGATCTCCAGAGTATGC-3′) oligonucleotides as negative control were obtained from Calbiochem-Novabiochem Corp. (34, 35). Transfection was performed using cationic lipopolyamines, which have been shown to ensure an efficient transport of oligonucleotides in eukaryotic cells (36). Briefly, in every experiment 20 × 106 PBTs or LPTs were washed and then resuspended in 10 ml Opti-MEM medium (Invitrogen Corp., Carlsbad, California, USA) containing 1 μl/ml Oligofectamine (Invitrogen Corp.) premixed with the p53 or scrambled control antisense at a final concentration of 4 μg DNA/ml. After a 4-hour incubation at 37°C and 5% CO2, 5 ml of growth medium containing 30% FCS was added and cells were stimulated for 3 days as described. Samples incubated with Oligofectamine or antisense alone and nontransfected cells were used as controls.
Statistical analysis. Statistical analysis was performed using the Student’s t test. Results are expressed as mean ± SEM, and significance was inferred with P values less than 0.05.
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Results
Distinct cell cycle kinetics of PBTs and LPTs. Differences in PBT and LPT proliferation in response to stimulation of the CD3 and CD2 pathways have been largely defined based on results of tritiated thymidine uptake (10, 13, 14, 37). Using this method, we confirmed this differential reactivity by showing that LPTs displayed a significantly reduced proliferation in response to a 3-day anti-CD3 stimulation (P < 0.05) and enhanced proliferation in response to a 3-day anti-CD2 stimulation (P < 0.005) compared with equally stimulated PBTs (Figure 1). Although simple and reproducible, this method only measures DNA synthesis during the S phase and provides no information on the fraction of cells going through the other phases, including resting (G0/G1), mitotic (G2/M), dividing (M), and apoptotic cells (3). To determine whether PBTs and LPTs differ in regard to distribution in each cell cycle phase, and whether their different composition of naive (CD45RA+) and memory (CD45RO+) cells affects the results, we performed a complete analysis of their cycling profiles in response to CD3 and CD2 activation. Immediately after isolation, more than 98% of PBTs and LPTs were in G0/G1, regardless of naive or memory status. After a 3-day activation with CD3 mAb, the percentage of CD45RA+ PBTs in the G0/G1 phase dropped to 70% ± 3.1%, while those in the S and G2/M phase increased to 30% ± 3.5% (Figure 2). After the same period of time, 77% ± 4.1% of the CD45RO+ PBTs were in the G0/G1 and 23% ± 2.9% were in the S and G2/M phase. In contrast, 80% ± 4.3% of LPTs remained resting in the G0/G1 phase and 20% ± 2.4% cells advanced to the S and G2/M phase (Figure 2). When activation was induced with anti-CD2 mAb, only 5% ± 1.7% of CD45RA+ PBTs and 15% ± 2% of the CD45RO+ PBTs cycled, whereas LPTs behaved like CD3-activated LPTs (Figure 2). Interestingly, the number of LPTs cycling in response to both CD3 and CD2 stimulation remained high at later time points (4–6 days), while the number of CD3-activated CD45RA+ and CD45RO+ PBTs continued to drop (Figure 2).
Figure 1Differential proliferation of PBTs and LPTs in response to anti-CD3 and anti-CD2 stimulation. PBTs and LPTs were cultured in the absence and presence of immobilized anti-CD3 mAb or soluble anti-CD2 (T112 and T113) mAb for 72 hours, and pulsed with 3H thymidine for the last 18 hours, prior to harvesting and counting in a beta counter. Data are expressed as mean ± SE of eight separate experiments. *P < 0.05, **P < 0.005.
Figure 2Distinct cell cycle profiles of CD45RA+ PBTs, CD45RO+ PBTs, and LPTs. The number of CD3-activated LPTs in the S and G2/M phase peaks at 4–6 days, in contrast to CD45RA+ and CD45RO+ PBTs, which peak at 2–3 days. CD2-activated LPTs in the S and G2/M phase peak at 4–6 days, in contrast to CD45RA+ PBTs, which mostly remain in the G0/G1 phase, and CD45RO+ PBTs, which peak at 3 days and then decline. Freshly isolated PBTs and LPTs were cultured with anti-CD3 or -CD2 mAb, and each phase of the cell cycle was assessed by measuring DNA content by PI staining followed by flow cytometry. The graphs are representative of eight different experiments.
Next, we performed stathmokinetic analysis to determine the time taken by PBTs and LPTs to traverse each cycle phase. This was accomplished using nocodazole as a metaphase inhibitor and BrdU incorporation to model and measure cell cycle kinetics (27, 38, 39). The G0/G1, S, and G2/M phase traverse times of PBTs activated by CD3 signaling were substantially faster than when cells were activated through CD2 (Table 1). In LPTs, the times the cells spent in each phase were comparable in response to CD3 and CD2 activation, but longer than that of CD3-activated PBTs (Table 1). Calculation of potential doubling times (Tpot) showed that it takes 1 day (24 hours) for CD3-activated PBTs to double, but 4 days (104 hours) days for CD2-activated PBTs. In contrast, it takes nearly 2 days for LPTs to double, regardless of CD3 (47 hours) or CD2 (39 hours) stimulation (Table 1).
Different levels of cell cycle promoters in PBTs and LPTs. Having established that PBTs and LPTs display important differences in cell cycle kinetics, we next investigated how levels of key regulatory molecules responsible for initiating and advancing each phase of the cell cycle varied in the two cell populations (40, 41). Western blot analysis showed that CDK4 and cyclin D3, the main regulators of the G1 phase, were upregulated only in CD3-activated PBTs, not in PBTs activated by CD2 or in LPTs activated by CD3 or CD2 mAb (Figure 3a). Similar results were obtained for cyclin D2 (data not shown). The analysis of Rb, which is essential for the G1/S phase transition, showed that increase in phosphorylation was marked in CD3-activated, but small in CD2-activated, PBTs (Figure 3a). This regulatory profile was entirely different in LPTs, since engagement of either the CD2 or the CD3 pathway led to a comparable increase of Rb phosphorylation. The levels of p107 and p130, the other two members of the Rb protein family, did not significantly change in either cell type, regardless of the stimulation pathway used (data not shown).
Figure 3(a) Different levels of cell cycle promoters in PBTs and LPTs. Immunoblot analysis shows that upregulation of CDK4 and cyclin D3 is only detected in CD3-activated PBTs, but not in PBTs or LPTs activated through the CD2 and CD3 pathways. Rb phosphorylation (pRb) was minimal in CD2-activated PBTs but markedly increased after CD3 activation, whereas, in LPTs, engagement of either the CD2 or CD3 pathway led to strong Rb phosphorylation. Strong upregulation of cyclin A in PBTs was induced by CD3 activation, whereas CD2 activation was required for an equally strong upregulation in LPTs. (b) Different levels of cell cycle inhibitors in PBTs and LPTs. The level of p21 varies little in unstimulated and stimulated PBTs and LPTs, except for a moderate increase in CD3-activated PBTs. Levels of the inhibitor p27 are downregulated in CD3-stimulated PBTs but not CD3-stimulated LPTs. Activation through the CD2 pathway induces a striking upregulation of p53 in LPTs but only a moderate increase in PBTs, whereas CD3 stimulation induces a lower and comparable p53 upregulation in PBTs and LPTs. Freshly isolated PBTs and LPTs were cultured in the absence and presence of CD3 or CD2 mAb for 3 days, after which expression of cell cycle inhibitors was assessed by Western blotting. Each bar represents the densitometry units of individual protein bands normalized to their own GAPDH. Each panel is representative of five to seven different experiments.
We also analyzed cyclins A and B1, responsible for leading the cells to mitosis after they passed the restriction point at the G1/S interphase. Distinct activation pathways appeared to be responsible for progression of PBTs and LPTs through the S phase. In fact, strong upregulation of cyclin A in PBTs was only induced by CD3 activation, whereas CD2 activation was required for an equally strong upregulation of cyclin A in LPTs (Figure 3a). Cyclin B1 was examined by flow cytometry in conjunction with PI staining to exactly localize its increase within the cell cycle. The expression of cyclin B1 in the G2/M phase of CD3-activated PBTs and CD2-activated LPTs paralleled the expression of cyclin A during the previous phase of the cell cycle (Figure 3). Furthermore, cells positive for phosphorylated histone H3, a mitosis marker exclusively detectable during unfolding of DNA in mitosis (42), were present in CD3-activated but essentially absent in CD2-activated PBTs, while equal numbers were found in CD2- and CD3-activated LPTs (Figure 4 and Table 1). The variations observed in protein levels of cell cycle promoters were confirmed at the mRNA level by ribonuclease protection assay (data not shown).
Figure 4Different levels of cyclin B1 and phospho–histone H3 in PBTs and LPTs. Flow cytometric analysis shows increase of cyclin B1 expression in the G2/M phase of anti-CD3–activated PBTs and anti-CD3– and anti-CD2–activated LPTs. The right upper inserts represent the phospho–histone H3–positive cells (y axis), and the arrows indicate the mitotic cell population. Freshly isolated PBTs and LPTs were cultured in the absence and presence of CD3 or CD2 mAb for 3 days, after which cyclin B1 and H3 expression and DNA content were examined by flow cytometry. The figure is representative of seven different experiments.
Different levels of cell cycle inhibitors in PBTs and LPTs. To complement the study of cell cycle promoters, we next determined expression levels of the key cell cycle inhibitors p21, p27, and p53. p21 levels did not significantly differ in unstimulated and stimulated PBTs and LPTs, except for an increase in CD3-activated PBTs, a finding in agreement with the role that p21 also plays in cell proliferation (Figure 3b) (43). Levels of the inhibitor p27 were downregulated in CD3-stimulated PBTs but not CD3-stimulated LPTs. In contrast with these inhibitors, major differences in the levels of p53 were observed that further confirmed the existence of distinct cell cycle regulatory patterns between PBTs and LPTs. Activation through the CD2 pathway induced a striking upregulation of p53 in LPTs but only a modest increase in PBTs, whereas CD3 stimulation induced a lower and similar upregulation in PBTs and LPTs (Figure 3b). Differences observed in protein levels of cell cycle promoters were confirmed at the mRNA level by ribonuclease protection assay (data not shown).
Differential clonal expansion and telomerase activity of PBTs and LPTs. We next measured clonal expansion to establish the actual capacity of PBTs and LPTs to divide in response to receptor activation. Using the CFDA lipophilic dye, we determined the number of cell divisions by measuring the dilution of the dye in each successive daughter population. Because of the distinct cell cycle kinetics observed in CD45RA+ PBTs, CD45RO+ PBTs, and LPTs, clonal expansion was measured in each of these populations. After 4-day stimulation with anti-CD3 mAb, three cell divisions comprising 70–90% of the original population were noted in CD45RA+ or CD45RO+ PBTs (Figure 5). During the same period of time, only 30% of equally stimulated LPTs divided, even though the number of cell divisions was the same. When clonal expansion in response to CD2 was investigated, an opposite pattern was observed. At least three divisions comprising more than 70% of the cells were induced in LPTs, whereas only 5–10% of CD45RA+ or CD45RO+ PBTs divided in response to CD2 stimulation (Figure 5).
Figure 5Clonal expansion of CD3- and CD2-activated CD45RA+ PBTs, CD45RO+ PBTs, and LPTs. After 4 days’ stimulation, CD3 activation induces a greater clonal expansion in both CD45RA+ and CD45RO+ PBTs than in LPTs, whereas in CD2-activated cells clonal expansion is strikingly greater in LPTs than in both CD45RA+ and CD45RO+ PBTs. The first peak to the right represents the undivided cell population. The numbers indicate the percentage of divided cells distributed in the subsequent peaks. Freshly isolated CD45RA+ and CD45RO+ PBTs and LPTs were incubated with 5 μM CFDA and cultured in the absence and presence of CD3 or CD2 mAb for 4 days, and cell divisions were determined by flow cytometry. The figure is representative of six to nine different experiments.
A step indispensable for cell division to occur is preservation of chromosome telomere length, a process strictly dependent on the activity of the enzyme telomerase (21). We therefore measured this enzyme activity in CD3- and CD2-stimulated PBTs and LPTs. Telomerase activity was essentially undetectable in unstimulated PBTs and LPTs and in CD2-stimulated PBTs but increased substantially in CD3-activated PBTs (Figure 6, a and b). In LPTs, telomerase activity was induced by both CD2 and CD3 activation, reaching only half of the level seen in CD3-activated PBTs.
Figure 6Telomerase activity of CD3- and CD2-activated PBTs and LPTs. (a) In PBTs, telomerase activity is induced only by CD3 activation, whereas both anti-CD3 and anti-CD2 activation induces enzymatic activity in LPTs. The level of activity is significantly greater in anti-CD3–stimulated PBTs than in both anti-CD3– and anti-CD2–activated LPTs (*P < 0.05). (b) Display of the telomeric-mediated six-nucleotide ladder. The intensity of expression closely correlates with the level of enzymatic activity seen in a. Freshly isolated PBTs and LPTs were cultured in the absence and presence of CD3 or CD2 mAb for 3 days, after which cells were lysed, amplified, and analyzed either by telomerase activity or Southern blot using a nondenaturing bis-acrylamide gel. 1, lysis buffer control; 2, heat-treated control (anti-CD3–activated PBTs). Each bar represents mean ± SEM of five to seven experiments.
Mitochondrial membrane potential in PBTs and LPTs. The marked differences observed in the capacity of PBTs and LPTs to cycle and divide in response to CD3 and CD2 receptor engagement suggest that the capacity to generate the energy required for these crucial processes may vary depending on the cell type or pathway of stimulation. To investigate this possibility we used the redistribution of the dye rhodamine123, a lipophilic cation that accumulates in the mitochondrial membrane (31). When we used the same cells used in the 4-day stimulation clonal expansion protocol, CD3-activated PBTs showed a clear increase in mitochondrial polarization potential compared with unstimulated cells (Figure 7). Intriguingly, a comparable increase was observed in both CD3- and CD2-activated LPTs, despite their marked differences in clonal expansion. In sharp contrast, CD2-activated PBTs showed no increase in mitochondrial polarization potential (Figure 7).
Figure 7Mitochondrial membrane potential in PBTs and LPTs. After 4 days’ stimulation through the CD2 or CD3 pathway, LPTs display a clear increase in the mitochondrial membrane potential, comparable to that observed in CD3-activated PBTs. CD2-activated PBTs show no significant increase. Freshly isolated PBTs and LPTs were cultured in the absence and presence of CD3 or CD2 mAb for 4 days, after which mitochondrial membrane potential was assessed by rhodamine123 staining followed by flow cytometry. Each graph is representative of five to eight experiments. Numbers indicate mean fluorescence intensity.
Inhibitory role of p53 in cell cycle progression and apoptosis of LPTs. The clear differences in cycle kinetics between PBTs and LPTs, combined with the striking activation-induced elevation of p53 in LPTs, suggested that p53 was responsible for the slow cycle progression of LPTs and their known increased susceptibility to apoptosis (16). To investigate this possibility, blockade of p53 was carried out using specific antisense oligonucleotides, and its impact on cell cycle progression as well as apoptosis was assessed.
Under the experimental condition previously used, treatment of PBTs and LPTs with p53 antisense oligonucleotides inhibited p53 expression in both cell types regardless of CD3 or CD2 activation, but the remaining phosphorylated p53 levels were clearly higher in LPTs than in PBTs (Figure 8). Scrambled control oligonucleotides had no effect on p53 expression. Since p53 and Rb are both critically involved in controlling the restriction point at the G1/S interphase (44), we also assessed the effect of p53 antisense oligonucleotides on Rb expression. In agreement with the role of p53 in cell cycle regulation, a substantial increase in Rb expression was noted in PBTs and LPTs, corresponding to the p53 blockade–induced downregulation (Figure 8).
Figure 8Effect of antisense treatment on p53 and Rb levels in PBTs and LPTs. Treatment with p53 antisense oligonucleotides decreases p53 expression but substantially increases Rb phosphorylation in CD3- or CD2-activated PBTs and LPTs. Freshly isolated PBTs and LPTs were cultured for 3 days in the presence or absence of p53 antisense oligonucleotides, scrambled controls, anti-CD3 mAb, or anti-CD2 mAb, after which p53 and Rb protein levels were assessed by Western blotting. Each panel is representative of five different experiments.
To define the functional significance of the different levels of p53 between activated PBTs and LPTs and confirm that p53 was actually responsible for the slow cycle progression of LPTs, we used BrdU incorporation to measure the relative movement through the S phase of cells treated with p53 antisense oligonucleotides. Regardless of the pathway of activation, blockade of p53 by antisense oligonucleotides was able to overcome the slow cycling of LPTs and enhanced their relative movement to the same high level as that of CD3-activated PBTs (Figure 9, a and b). Blockade of p53 in LPTs resulted in a 65% and 60% shortening of DNA synthesis time of CD3- and CD2-activated LPTs, respectively, while it decreased DNA synthesis time by 34% in CD3-activated, and 45% in CD2-activated, PBTs (Table 2).
Figure 9Effect of p53 antisense treatment on relative movement of PBTs and LPTs. (a) Treatment with p53 antisense (AS) oligonucleotides increases the relative movement of CD3-activated PBTs by 34% and of CD3-activated LPTs by 65%. (b) Treatment increases the relative movement of CD2-activated PBTs by 45% and of CD2-activated LPTs by 60%. Freshly isolated PBTs and LPTs were cultured in the presence or absence of p53 antisense oligonucleotide, scrambled controls, anti-CD3 mAb, or anti-CD2 mAb and, 3 days later, incubated with 20 μM BrdU and restimulated with the same mAb. Relative movement was calculated as described in Methods. Each line represents the mean ± SEM of five experiments. The relative movement after antisense treatment was significantly higher in both anti-CD3– and anti-CD2–activated PBTs and LPTs than in their respective nontransfected and scrambled controls (P < 0.05).
Because of the role of p53 in apoptosis (44, 45), we also investigated the effects of its blockade on apoptosis of CD3- and CD2-activated PBTs and LPTs. As expected, apoptosis was substantially greater in unstimulated LPTs than in unstimulated PBTs (Table 2) (16). Activation through the CD3 or CD2 pathway failed to significantly alter the level of PBT cell death, and no additional effect was observed with p53 antisense oligonucleotide treatment. In sharp contrast, activation of either pathway enhanced LPT apoptosis, and this response was markedly inhibited by p53 blockade (Table 2).
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