c-Rel regulation of the cell cycle in primary mouse B lymphocytes

Constance Y. Hsia1, Shuhua Cheng2, Alexander M. Owyang1, Steven F. Dowdy3 and Hsiou-Chi Liou2

1 Immunology Program and 2 Division of Immunology, Department of Medicine, Weill Medical College of Cornell University, 515 East 71st Street, New York, NY 10021, USA 3 Howard Hughes Medical Institute, Department of Cellular and Molecular Medicine, University of California at San Diego School of Medicine, La Jolla, CA 92093, USA

Correspondence to: H.-C. Liou; E-mail: hcliou{at}med.cornell.edu
Transmitting editor: L. H. Glimcher


    Abstract
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 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Surface-expressed BCR mediates the proliferation and expansion of antigen-specific B lymphocytes during a humoral immune response. Although several studies extensively characterize BCR proliferative signaling, the mechanisms linking these pathways to the cell cycle remain elusive. Using knockout mice, we show that c-Rel, a proto-oncogenic member of the NF-{kappa}B transcription factor family, is essential to BCR-mediated proliferation and cell cycle progression. Splenic B cells obtained from gene-targeted c-Rel knockout mice display a defective proliferation response to antigen receptor cross-linking, resulting in G1 arrest. At the molecular level, we see that BCR stimulation of resting c-Rel–/– B cells fails to induce proper cyclin D3 and cyclin E expression, thereby negatively impacting G1 phase cyclin-dependent kinase (CDK) activity. c-Rel-deficient B cells also exhibit incomplete phosphorylation of the Retinoblastoma protein (pRb) and poor expression of E2Fs, thus impeding the G1 to S phase transition. Down-regulation of the pRb-related p130 protein during the G0 to G1 transition and removal of the CDK inhibitor p27KIP1 in late G1 parallel that of wild-type cells, suggesting that Rel-deficient B cells can exit the G0 resting state and enter G1 phase normally. Finally, we demonstrate that restoration of proliferation can be achieved partially upon reintroduction of cyclin E using a protein transduction method to reconstitute primary B cells. Collectively, these studies emphasize the importance of c-Rel in lymphocyte proliferation and oncogenesis, and highlight a requirement for c-Rel in establishing an effective humoral immune response.

Keywords: antigen receptor, B cell, c-Rel knockout, cyclin, NF-{kappa}B


    Introduction
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
BCR-mediated proliferation is crucial to the initiation of a targeted immune response. During lymphocyte activation and clonal expansion, phase-regulated de novo synthesis of cell cycle proteins is rapid (1,2), indicating an important role for transcriptional pathways in eliciting antigen-induced B cell proliferation. Numerous studies detailing the membrane-proximal signaling events of the BCR are described in the literature (3), yet the identification of transcription factors which couple BCR signaling directly to the cell cycle machinery is essentially unknown. In vitro studies and knockout mice reveal a critical function for the NF-{kappa}B/Rel transcription factor family (46). Mice deficient for p50 or RelB, for example, exhibit reduced B cell proliferation in response to lipopolysaccharide (LPS) or CD40 signals (79), while deletion of the lymphoid-specific member c-Rel leads to multiple B and T lymphocyte defects (1013). Although the addition of exogenous IL-2 can restore proliferation to c-Rel-deficient T cells, c-Rel–/– B cell responses are not rescued by co-culture with wild-type B cells, suggesting that cell-autonomous defects underlie c-Rel-mediated BCR proliferative responses which cannot be complemented by soluble factors alone.

Recently a link between NF-{kappa}B/Rel and cell cycle control was described by Hinz et al. which identified a {kappa}B element in the mouse cyclin D1 promoter. In these studies, activation of a reporter gene fused downstream of the cyclin D1 promoter could be induced by co-transfection of COS-7 cells with p50 and p65 heterodimers (NF-{kappa}B1) (14). Cyclin D1 expression, however, has not been detected in mature peripheral mouse B lymphocytes stimulated through the BCR by treatment with anti-IgM, in contrast to other cyclins such as D2, D3, E and A, all of which are detectably induced upon BCR cross-linking [C. Hsia and H.-C. Liou, unpublished results and (2)]. It remains to be determined whether NF-{kappa}B/Rel regulates any of these latter genes and, if so, whether the effects are direct or indirect consequences of NF-{kappa}B/Rel transcription.

Other mammalian cell cycle proteins include cyclin-dependent kinases (CDK) and CDK inhibitors (CKI) which cooperate with rate-limiting cyclins to selectively modulate cell cycle progression in response to various stimuli (15). Proliferative signals transduced from mitogenic stimulation of cell surface receptors, for example, trigger the expression of cyclins and removal of CKI, leading to the formation of active CDK complexes as cells advance past the G1 restriction point into S phase, in essence driving the proliferation response. Target substrates of CDK include the Retinoblastoma protein (pRb) family of tumor suppressor proteins which upon hyper-phosphorylation result in release of free E2F transcription factors in the nucleus (1619). Several DNA replication genes have been demonstrated to be E2F-dependent targets (20,21), thus overexpression of E2F genes can promote cell cycle progression and DNA synthesis in G0 quiescent or G1-arrested cells. Furthermore, in vitro evidence suggests that E2F can also regulate the expression of cyclins E and A (20,2225), as well as autoregulate the expression of E2F genes themselves (26,27), thereby underscoring the necessity for strict transcriptional control in orchestrating G1 to S phase transitions.

Our current studies investigate the relationship of c-Rel to control the mature B cell cycle. c-Rel–/– B cells arrest in G1 and fail to enter S phase, potentially resulting from inadequate induction of positive cell cycle regulators or insufficient removal of negative regulators, and possibly a combination of the two. While it is hypothesized that c-Rel deficiency may cause direct disturbances in cell cycle gene transcription, evidence to support this claim has not been formally established. Nevertheless, our data indicate that loss of G1 cyclin expression may contribute to insufficient levels of CDK activity and dysregulation of the pRb/E2F pathway in anti-IgM-treated c-Rel–/– B cells, thus perpetuating a hypoproliferative phenotype. Reintroduction of G1 cyclins into c-Rel–/– primary B cells using a protein transduction approach can partially restore proliferative responses. Together, these studies are the first to demonstrate that the c-Rel proto-oncogene is involved in BCR-mediated cell cycle progression and proliferation via regulation of positive-acting elements of the cell cycle.


    Methods
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 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Mice
Homozygous c-Rel null mice were generated through gene-targeted deletion as previously described (10), and have been backcrossed onto a C57BL/6 background for >6 generations. All experiments were conducted using <12-week-old age-matched wild-type and c-Rel knockout mice maintained under specific pathogen-free conditions.

Cell culture and preparation of cell extracts
Splenic resting B cells were purified and analyzed for purity as previously described (11). Greater than 95% pure B cells were cultured in RPMI 1640 containing 1% L-glutamine (Cellgro, Herndon, VA) supplemented with 10% heat-inactivated low endotoxin FCS (Cellgro), 1% penicillin (Life Sciences/BRL/Invitrogen, Carlsbad, CA), 1% streptomycin (Life Sciences/BRL) and 50 µM ß-mercaptoethanol (Sigma, St Louis, MO). Stimulations were carried out with control media or media containing 10 µg/ml F(ab')2 goat anti-mouse µ chain (anti-IgM) (Jackson Immunotech, West Grove, PA). The CH31 mature B cell line was cultured in RPMI 1640 containing 1% L-glutamine (Cellgro) supplemented with 10% heat-inactivated FCS (Cellgro), 1 mM sodium pyruvate, 1 x non-essential amino acids, 1% penicillin, 1% streptomycin (all Life Sciences/BRL/Invitrogen) and 50 µM ß-mercaptoethanol. Stimulations of CH31 cells were carried out in the presence or absence of 10 µg/ml anti-IgM or anti-CD40 [hybridoma 1C10; kindly provided by Drs Edward Clark, Maureen Howard and A. Heath (28)]. Whole-cell lysates were generated by lysing cells in 1 x RIPA buffer for Western blots (29) or in pRb lysis buffer for immunoprecipitation and kinase assays, and sonicating 6 x 8 pulses with a Branson 250 microtip sonicator (VWR Scientific, Bridgeport, NJ) at 20% duty cycle. Supernatants were separated from pellets and stored at –80°C. pRb lysis buffer contains 50 mM HEPES (pH 7.5), 150 mM NaCl, 1 mM EDTA, 2.5 mM EGTA, 1 mM dithiothreitol, 0.1% Tween 20, 10% Glycerol, 10 mM ß-glycerophosphate, 1 mM sodium fluoride, 0.1 mM Na3VO4, 0.2 mM PMSF, 5 µg/ml aprotinin, 5 µg/ml leupeptin and 2.5 mM sodium pyrophosphate.

Immunoblotting
Protein concentrations were determined by Bradford assay. Between 30 and 40 µg of whole-cell lysate were loaded onto SDS–polyacrylamide gels and transferred to a PVDF membrane (Millipore, Bedford, MA) by a semidry method. Blots were probed with the following commercial antibodies diluted into 1% non-fat milk, TBS containing 0.05% Tween 20 (TBS-T): cyclin D2 (M-20; sc-593), cyclin D3 (C-16; sc-182), cyclin E (M-20; sc-481), cyclin A (H-432; sc-751), E2F1 (C-20; sc-193), E2F2 (C-20; sc-633), E2F3 (N-20); sc-879), E2F4 (C-20; sc-866), CDK2 (M2; sc-163), CDK4 (C-22; sc-260), CDK6 (C-21; sc-177), p27 (C-19; sc-528), p130 (C-20; sc-317x) and p107 (C-18; sc-317x) all from Santa Cruz Biotechnology (Santa Cruz, CA); pRb (G3-245; 14001A) from PharMingen (San Diego, CA); and anti-hemagglutinin (HA) (12CA5) a gift from Dr Martin Scott. Horseradish peroxidase conjugated anti-rabbit secondary antibody (NA934) and anti-mouse secondary antibody (NA931) were purchased from Amersham/Pharmacia (Piscataway, NJ). The ECL Plus chemiluminescence detection system was used to visualize Western blots (RPN 2132 from Amersham). In all experiments, equal protein loading was controlled for by either stripping blots in 63 mM Tris–HCl (pH 6.8), 2% SDS (w/v) and 100 mM ß-mercaptoethanol, at 50°C for 30 min, followed by 3 washes with TBS-T, and reprobing blots with anti-CDK2 antibody (Santa Cruz Biotechnology). All Western blot experiments have been confirmed in multiple experiments using separate sets of cell lysates. The data presented in the figures are representative of several experiments with similar results.

Immunoprecipitation and kinase assays
Kinase assays were performed essentially as described in (29). Briefly, the 86-kDa GST–Rb substrate containing the E2F pocket binding residues was prepared from pGEX-Rb (aminoacids 379–928) Escherichia coli (30), and the purified protein incubated with immunoprecipitated lysates in the presence of 50 mM HEPES (pH 7.5), 10 mM MgCl2, 1 mM dithiothreitol, 2.5 mM EGTA, 10 mM ß-glycerophosphate, 0.1 mM Na3VO4, 1 mM sodium fluoride, 2.5 mM sodium pyrophosphate, 24 mM ATP and 10 µCi [{gamma}-32P]ATP (3000 Ci/mmol; NEN/Perkin Elmer, Boston, MA) at 37°C for 30 min. Reactions were then run on 10% SDS–polyacrylamide gels, dried onto filter paper (Whatman, Kent, UK) and exposed on Kodak Bio-Max film. Histone H1 kinase assays were performed by incubating immunoprecipitated lysates in 20 mM Tris, pH 7.4, 7.5 mM MgCl2, 1 mM dithiothreitol, containing 0.05 µg histone H1 (Boehringer Mannheim/Roche, Indianapolis, IN), 33 µM ATP, pH 7.0 and 10 µCi [{gamma}-32P]ATP (3000 Ci/mmol; NEN DuPont) at 37°C for 30 min. Reactions were then run on 12% SDS–polyacrylamide gels, dried and exposed to film as mentioned above.

TAT fusion protein transduction and flow cytometry
TAT–cyclin E and TAT–cyclin D3 fusion proteins were generated by cloning mouse cyclin E and mouse cyclin D3 genes into a (His)6 HA-tagged TAT vector obtained from Dr Steven Dowdy (32). Both genes were cloned using gene-specific PCR primers 5'-GAGAATTCTCATTCTGTCTCCTGCTCGCTG and 5'-TTCATCTCGAGATGAAGGAGGACGGCGGCG (cyclin E) and 5'-GTACCGGTGAGCTGCTGTGTTGCGAG and 5'-AGGAATTCCTACAGGTGAATGGCTGTG (cyclin D3) to amplify sequences from mouse cDNA, and the PCR products then digested and cloned into XhoI and EcoRI sites (TAT–cyclin E) and AgeI and EcoRI sites (TAT–cyclin D3) of the TAT vector. TAT–green fluorescence protein (GFP) was generated by subcloning a GFP-containing fragment from the SFG-NTP-ires-EGFPsp vector (kindly provided by Dr Michel Sadelain) into the EcoRI and NcoI sites of the TAT vector. Fusion proteins were grown in BL21 E. coli, purified over a nickel agarose column (Qiagen, Valencia, CA), denatured in 8 M guanidine hydrochloride (Life Sciences/BRL/Invitrogen) and desalted by a PD10 column (Amersham/Pharmacia). Typically 1 mg/ml solutions were achieved and protein degradation monitored by Coomassie blue staining of SDS–PAGE gels. Purified protein preparations were then added to primary B cell cultures at various concentrations for the indicated times. To avoid the contaminating effects of bacterial LPS, we measured the amount of LPS in various TAT fusion protein solutions and determined the maximum tolerable level of LPS carryover to be <25 ng/ml. All solutions tested had levels below this amount or were discarded entirely. Polymyxin B sulfate (20 µg/ml; Sigma) was included in all assays, however, as a precaution to prevent the activation of cells by any residual LPS. TAT–GFP was used as a negative control in thymidine incorporation assays.

[3H]Thymidine incorporation
Cells were plated in triplicate in a 96-well plate at a density of 2 x 105 cells/well in 200 µl of control media or media containing 10 µg/ml anti-IgM. Following a 42 h incubation period, each well was pulsed with 0.5 µCi [3H]thymidine (Amersham) and incubated for an additional 6 h, then the entire plate frozen at –20°C. To quantitate [3H]thymidine incorporation, plates were thawed, cells harvested and measured by scintillation counter (Wallac, Turku, Finland).

Propidium iodide staining
Splenic resting B cells were cultured at 5 x 105 cells/well in 96-well plates for 2 days (48 h) in the presence of 10 µg/ml anti-IgM. Cells were then collected, washed, and stained with 50 µg/ml propidium iodide, 20 µg/ml RNase A, 0.1% Triton X-100 and 0.1% sodium citrate, and analyzed by flow cytometry using CellQuest software (BD PharMingen, San Diego, CA). Cells with 2N DNA content were considered as G0 or G1 phase; cells with >2N but <4N DNA content were considered as S phase; cells with 4N DNA content were considered G2 or M phase.

RT-PCR
Splenic resting B cells were cultured at 5 x 106 cells/ml in six-well plates for 2 days (48 h) in the presence or absence of 10 µg/ml anti-IgM. Cells were then collected, washed and RNA extracted by the STAT-60 method (Tel-Test, Friendswood, TX) following the manufacturer’s protocol. Purified RNA (2 µg) was then reverse transcribed into cDNA by Superscript II (Life Technologies) at 42°C for 1 h using 0.5 µM oligo(dT) to prime each reaction. PCR reactions were carried out using primers specific for each gene: 5'-AGCGCTGCGAGGAGGATGTCTT-3' and 5'-GCCAGGAAGTCGTGCGCAATC-3' for cyclin D3; 5'-GAGCCTCCCCACTTCCCGTCTT-3' and 5'-CCGGAGCAAG CGCCATCTGT-3' for cyclin E; 5'-TACCTGGACAAAGCACC TCCAC-3' and 5'-TCCACGGGTGTTGCGTATGTG-3' for CDK4; and 5'- AGTGGAGATTGTTGCCATCAA-3' and 5'-AG CAGTTGGTGGTGCAGGA-3' for GAPDH. Limiting dilutions of each cDNA sample were assayed for GAPDH to verify linearity of PCR results.


    Results
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 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
c-Rel–/– B cells fail to enter S phase and arrest in G1
Studies with c-Rel knockout mice have revealed a gross failure to generate lymphocyte proliferation responses upon BCR cross-linking, LPS treatment and CD40 ligation. BCR cross-linking by the F(ab')2 fragment of the IgM heavy chain (anti-IgM) normally induces DNA synthesis of resting splenic B cells over a period of 72 h. However, c-Rel knockout B cells respond very weakly to this signal, indicating poor mobilization of DNA replication machinery in the absence of c-Rel (11,12). Propidium iodide staining shows that c-Rel–/– B cells have higher sensitivity to anti-IgM-induced apoptosis (13,32), thus explaining the reduced numbers of proliferating cells. However, within the viable population of cells, there also appears to be an increased number of cells in G0/G1 relative to S and G2/M phases.

To determine whether anti-IgM-treated c-Rel–/– B cells actively undergo G1 phase arrest disproportionately compared to wild-type cells, we re-analyzed the viable cell population alone (>=2N DNA content) and found that these cells fail to enter S phase, instead accumulating in G0/G1 (Fig. 1). While 49% of wild-type cells have entered S–G2/M phase by day 2, only 15% of c-Rel–/– B cells have entered S–G2/M and the remaining 85% of cells are stuck in G0/G1. This cell cycle arrest persists into day 3 and day 4 when most of the wild-type B cells have already begun to divide and reappear as G1 phase cells. Thus, it is evident from these studies that BCR activation induces c-Rel-dependent proliferation pathways that are clearly distinct from c-Rel-mediated survival pathways. We therefore speculate that G0, G1 and G1/S phase cell cycle genes are potential targets of c-Rel regulation during BCR-induced proliferation.



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Fig. 1. c-Rel–/– B cells arrest in G1. Purified splenic B cells were cultured in 10 µg/ml anti-IgM over a duration of 4 days (90 h) and harvested at the indicated times. Cells were washed and stained with propidium iodide, and then analyzed for cell cycle by flow cytometry. Viable cells with >=2N DNA content were evaluated as follows: cells containing 2N DNA content were considered G0/G1 phase, while cells having >2N DNA content were considered S–G2/M phase. The calculated percentages of viable cells in G0/G1 or S–G2/M phases are shown and are representative of multiple experiments. Compared to wild-type B cells, c-Rel–/– cells fail to enter S–G2/M phases and instead accumulate in G0/G1.

 
G1 CDK activity is impaired in c-Rel-deficient B cells
Because CDK activity is essential to G1 progression, we wanted to determine whether CDK activity is affected by the absence of c-Rel. CDK4 and CDK6 kinase activity can be measured by radioactive assay using an 86-kDa fusion protein substrate GST–Rb which contains amino acid residues corresponding to CDK4 and CDK6 phosphorylation sites in the E2F binding pocket of pRb (30,33). Whole-cell extracts generated from both wild-type and c-Rel-deficient mouse splenic B lymphocytes cultured in the presence or absence of anti-IgM over 48 h (see Methods) were immunoprecipitated for CDK4 and CDK6 protein. Kinase activity was then assayed on GST–Rb in the presence of [{gamma}-32P]ATP. As seen in Fig. 2, we observed partial loss of CDK4 kinase activity in knockout lysates at 12 h (cf. lanes 2 and 6, Fig. 2A), while CDK6 activity was significantly reduced at 36 h compared to maximal kinase activity seen in wild-type lysates (cf. 12 and 16, Fig. 2A).



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Fig. 2. G1 and G1/S CDK kinase activity is impaired in c-Rel–/– B cells. Whole-cell extracts were prepared from splenic B cells stimulated with 10 µg/ml anti-IgM for the designated time. Lysates were immunoprecipitated (IP) using antibodies against CDK4, CDK6 or CDK2, and kinase assays performed using either GST–Rb (A) or histone H1 (B) as substrates. Polyclonal rabbit Ig anti-sera served as a negative (–) control, while the presence of non-specific bands indicates even loading (load). Western blot analysis was performed using antibodies specific for (C) CDK4 and CDK6 or (D) CDK2. Non-specific bands (load) indicate equal loading. Anti-CD40-treated CH31 cells (C + {alpha}-CD40) and anti-IgM-treated CH31 cells (C + {alpha}-IgM) not as control lysates to demonstrate that CDK protein expression is not affected by B cell mitogenic stimuli.

 
Downstream CDK activity corresponding to G1/S and S phases was measured by CDK2 immunoprecipitation and kinase assay using histone H1 as a substrate. Wild-type B cells responded with robust CDK2 activity at 30 and 60 h, but c-Rel–/– cells failed to give any measurable response above background (Fig. 2B). Together these results support the finding that absence of c-Rel impedes proliferative signaling by failing to induce CDK kinase activity necessary for G1 to S phase progression.

To verify that loss of CDK activity was not a consequence of diminished CDK protein levels, we measured CDK2, CDK4 and CDK6 protein expression by immunoblotting. No discrepancies were found in the expression patterns of CDK proteins over a time course of 48 h anti-IgM treatment (Fig. 2C and D) including controls from the B cell line CH31. These results are consistent with previous reports of constitutive CDK expression in primary B cells (2,34). This data suggests that observed deficiencies in CDK kinase activity might result from defects in cyclin or CKI regulation. To test this possibility, we next examined expression levels of G1 cyclins and p27KIP1.

Expression of G1 cyclins is impaired in c-Rel–/– B cells, but p27KIP1 turnover is normal
CDK kinase activity is positively controlled by association with cyclins and negatively regulated by formation of CDK–CKI inhibitor complexes. Inducible expression of cyclins is therefore a crucial rate-limiting step to G1 progression, while removal of inhibitors is necessary for full CDK activation. Because c-Rel-deficient B cells fail to enter S phase, we chose to monitor cell cycle regulators activated during or prior to S phase. Several cyclin genes, listed here in the order they are expressed upon BCR cross-linking by anti-IgM, can be detected at the protein level during G1 and S phases: cyclin D2 > D3 > E > A (2,34,35).

Whole-cell extracts were generated from purified resting B lymphocytes cultured in the presence or absence of anti-IgM over a 48 h duration (see Methods). Comparing wild-type with knockout cells, we observed a slight delay in cyclin D2 induction at 12 h (lanes 2 and 7, Fig. 3A), a significant reduction of cyclin D3 protein beginning at 24 h (lanes 3–5 and 8–10 Fig. 3B), diminished expression of cyclin E from 24 to 45 h (lanes 3–5 and 8–10, Fig. 3C), and loss of cyclin A from 36 to 45 h (lanes 4–5 and 9–10, Fig. 3D) in c-Rel–/– knockout B cells. These results point to multiple defects occurring after entry into G1 (as evidenced by normal cyclin D2 up-regulation) but prior to induction of S phase due to loss of mid-late G1 and S phase cyclins. Loss of cyclin D2 expression at 48 h is presumed to be a consequence of poor cell viability at this time point.




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Fig. 3. c-Rel–/– deficiency leads to loss or delay of G1 and S phase cyclin induction in B cells, but p27KIP1 down-regulation is unaffected. Wild-type and c-Rel–/– resting splenic B cells were cultured with 10 µg/ml anti-IgM for the indicated times and whole-cell extracts prepared as described in Methods. Western blots were performed using (A) anti-cyclin D2, (B) anti-cyclin D3, (C) anti-cyclin E, (D) anti-cyclin A and (E) anti-p27KIP1 rabbit polyclonal antibodies. Protein loading was controlled by stripping and reprobing with (F) anti-CDK2 polyclonal antibody. (G) Cyclin E transcription is defective in anti-IgM-treated c-Rel–/– B cells. Splenic B cells were cultured at 5 x 106/ml in the presence of anti-IgM for 48 h and harvested at the indicated timepoints for RNA extraction. mRNA was reverse transcribed into cDNA, and cyclin D3, cyclin E and CDK4 genes were amplified using specific primers. GAPDH was included as a control. Cyclin E transcripts are reduced in the absence of c-Rel, while cyclin D3 and CDK4 transcripts are unaffected.

 
Removal of CKI inhibition is also necessary to drive cells into proliferation, since cyclin induction alone is not sufficient to activate CDK in the presence of CKI-mediated inhibition. Inhibition is enforced by the INK4 and p21CIP1/p27KIP1 families throughout G1 (15). In resting B cells, however, p27KIP1 serves as the predominant inhibitor species, whereas other inhibitors are expressed at low or undetectable levels [(2,36,37) and data not shown). Our results showed no difference between wild-type and c-Rel–/– B cells in p27KIP1 turnover (Fig. 3E), suggesting that loss of CDK2 activity in c-Rel-deficient B cells is not due to elevated or sustained p27KIP1 levels. Instead, we suspect that decreased CDK activity is a result of low cyclin induction. These results clearly uncouple BCR-induced p27KIP1 degradation from c-Rel-mediated cyclin up-regulation, and demonstrate that c-Rel-dependent and -independent mechanisms of BCR proliferative signaling can be distinguished molecularly through the cell cycle pathway.

Cyclin E transcription is reduced in c-Rel–/– B cells
Because c-Rel knockout B cells fail to enter S phase, we wished to examine potential transcriptional defects of the G1 cyclins, in particular cyclins D3 and E, since both these genes exhibited deficient protein expression in the absence of c-Rel. To determine whether reduction of cyclin D3 and E levels in anti-IgM-treated c-Rel–/– B cells was a direct result of decreased gene transcription, we used RT-PCR to measure mRNA levels over a period of 48 h. As shown in Fig. 3(G) we detected comparable amounts of cyclin D3 transcription in wild-type and c-Rel–/– B cells upon anti-IgM stimulation; however, cyclin E transcription was considerably reduced in the absence of c-Rel compared to wild-type. The CDK4 gene, whose protein expression is constant in B cells (see Fig. 2C), also showed no difference in RNA transcript levels between wild-type and c-Rel knockout cells. Hence, the defects in cyclin D3 expression appear to be post-transcriptionally dependent upon c-Rel, whereas the defects in cyclin E expression are transcriptionally affected.

pRb phosphorylation is delayed in activated c-Rel–/– B cells
To further confirm the observed defects in CDK activity, pRb phosphorylation was monitored. pRb suppresses cell cycle progression during G1 by blocking E2F transcriptional activity at target gene promoters. Inactivation of pRb is mediated largely through hyper-phosphorylation of critical serine/threonine residues in the E2F binding pocket by CDK kinases (15,16,18). Hence, we reasoned that loss of CDK activity in c-Rel–/– B cells (see Fig. 2A and B) would result in suboptimal pRb phosphorylation.

As shown in Fig. 4, wild-type lysates display complete hyper-phosphorylation of pRb at 36 h (lane 6, Fig. 4A), and begin to undergo dephosphorylation of pRb by 48 h (lane 7, Fig. 4A), thus resetting the pRb–E2F cycle for the next phase. c-Rel–/– cells, in contrast, display incomplete hyper-phosphorylation at 36 h (cf. lanes 6 and 12, Fig. 4A) and, interestingly, also exhibit reduced overall expression of pRb protein. To determine if this resulted from heterogeneous protein loading, we stripped the blot and reprobed for STAT6, which is expressed constitutively in all mature B cells including the CH31 control cell line (Fig. 4D). Despite a slight underloading of lanes 12 and 13, our results indicate that reduced pRb expression in knockout cells is not due to artifact, but might instead be related to loss of E2F expression since pRb gene regulation is also reported to be E2F dependent (see below) (22,38).



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Fig. 4. pRb family proteins show poor responsiveness to anti-IgM-induced hyper-phosphorylation in c-Rel-deficient B cells. Western blot analysis of pRb family proteins reveals incomplete phosphorylation and induction for pRb and p107, but not p130. (A) At 36 h anti-IgM stimulation, c-Rel–/– B cells display incomplete pRb hyper-phosphorylation compared to wild-type. Hyper-phosphorylated pRb migrates at a higher position on the blot. The blot was then stripped and reprobed with (D) anti-STAT6 mAb as a loading control. (B) p107 induction is impaired at 24–48 h (lanes 5–7 and 11–13), while (C) p130 down-regulation is comparable between wild-type and c-Rel–/– cells. Loading was verified by the presence of non-specific bands (data not shown). Anti-CD40-treated CH31 (C + {alpha}-CD40) and anti-IgM-treated CH31 (C + {alpha}-IgM) control lysates are included to represent even blotting of the membrane.

 
Other members of the pRb pocket protein family include p130 and p107, which can associate with specific E2F members to form additional repressor complexes distinct from pRb complexes. p130 is predominantly expressed in quiescent or growth-arrested G0 cells, while p107 is found in actively dividing G1-progressing cells. It has been proposed that p130 complexes containing E2F4 and E2F5 mediate repression of genes affecting G1 progression, while p107–E2F4 and pRb–E2F1/2/3 complexes restrict G1-progressing cells from entering S phase prematurely (1,39). Mitogenic signals effectively remove these forms of repression by first triggering phosphorylation and degradation of p130 in quiescent G0 cells, and then later promoting the expression and subsequent phosphorylation of pRb and p107 in late G1.

Our results show that anti-IgM stimulation of c-Rel–/– B cells promotes normal phosphorylation and removal of p130 as cells exit the G0 resting state to enter G1 (Fig. 4B). c-Rel–/– B cells, however, fail to induce proper expression of p107 in late G1, evident by 24, 36, and 48 h treatment with anti-IgM (cf. lanes 5–7 and 11–13, Fig. 4C). The observed loss of p107 induction, similar to the reduced expression levels of pRb, may in fact arise from poor E2F expression as well (see below) since E2F activity is thought to play a role in pRb expression. Together our experiments indicate that turnover of p130 and exit from a G0 resting state is unaffected by the absence of c-Rel; however, c-Rel is required for proper expression and phosphorylation of pRb and p107 during G1 progression.

E2F expression levels are reduced or delayed in activated c-Rel–/– B cells
The impaired CDK kinase activity and pRb/p107 phosphorylation defects very likely affect E2F activity since pRb family pocket proteins control E2F transcriptional activation. Because E2Fs can also conceivably autoregulate the expression of other E2F family members (22,26,27), as well as control expression of late G1 and S phase cyclins E and A (23,40) (whose expression is reduced in c-Rel–/– B cells), we suspected that E2F-dependent events would be impaired in c-Rel-deficient B cells. These findings, coupled with the important role E2Fs play in controlling S phase entry, prompted us to look at E2F activity and protein expression.

As depicted in Fig. 5, E2F1, E2F2, E2F3, E2F4 and E2F5 protein levels are all significantly reduced in the absence of c-Rel (Fig. 5). These findings agree with loss of CDK kinase activity (see Fig. 2A and B), and observed deficiencies cyclin E and cyclin A expression (see Fig. 3C and D). In addition, these results also correlate with reductions in pRb and p107 protein levels (see Fig. 4A and C) whose expression shows E2F dependence as well (26,27,38). Inadequate E2F activation can promote G1 arrest even in the absence of pRb/p107, thereby providing a plausible explanation of the G1 block in c-Rel-deficient B cells despite reduced pRb and p107 levels. Together our results suggest that the multiple proliferation defects observed in late G1 of c-Rel–/– B cells may result from insufficient E2F activity and expression, and underscore the relevance of E2F mediated effects in c-Rel-dependent B cell proliferation.



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Fig. 5. E2F1, E2F2, E2F3 and E2F4 expression levels are reduced in stimulated c-Rel–/– B cells. Western blots show reduced expression of E2F1, E2F2, E2F3, E2F4 and E2F5 levels. Blots were performed using E2F-specific antibodies as indicated. Blots were stripped and reprobed with anti-CDK2 antibody to verify equal loading.

 
TAT–cyclin E protein transduction into c-Rel–/– B cells partially restores proliferation
Since cyclin expression and CDK activity is reduced in c-Rel–/– B cells, we hypothesized that reintroduction of cyclins should restore proliferative responses to knockout B cells by restoring CDK kinase activity. This would in turn effect the activation of downstream E2Fs which are directly linked to the transcription of DNA replication genes and S phase cyclins. Difficulties associated with DNA transfection of primary mouse B cells, however, have made attempts to reconstitute c-Rel knockout cells remarkably unsuccessful. In an effort to circumvent this problem, we undertook a protein transduction method that utilizes a peptide derived from the HIV TAT protein to transduce fusion molecules across the cell membrane (31,41).

Bacterially expressed TAT–GFP, TAT–cyclin D3 and TAT–cyclin E proteins were purified to homogeneity, and then assayed for activity in primary B cells. As shown in Fig. 6, each protein can efficiently transduce both wild-type and c-Rel–/– primary mouse B lymphocytes (>85% cells are GFP+ within 7 h) (Fig. 6A and data not shown). Moreover, the intrinsic fluorescence properties of GFP suggest that proper protein refolding has occurred in TAT-transduced GFP+ cells. TAT fusion proteins transduced into primary B cells can also be detected by immunoblotting with an anti-HA antibody (Fig. 6B), demonstrating the relative stability of these proteins since degradation products are not detected in these blots. Furthermore, TAT–cyclin D3 and TAT–cyclin E can remain stable in cultured B cells for >48 h (data not shown).




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Fig. 6. TAT fusion proteins can efficiently transduce primary mouse B cells; TAT–cyclin E partially restores c-Rel–/– proliferation. (A) Flow cytometry shows that >85% of primary wild-type B cells are transduced with 300 nM TAT–GFP by 7 h. (B) Primary wild-type B cells were transduced with either 300 nM TAT–cyclin D3 (TAT–cycD3) or 300 nM TAT–cyclin E (TAT–cycE) in the presence of 10 µg/ml anti-IgM for 24 h. Cells were then harvested, washed and lysed, and Western blots performed using anti-HA antibody to detect HA-tagged TAT fusion proteins which had entered cells. (C and D) Purified splenic B cells (2 x 105/well) were plated in triplicate into a 96-well plate and cultured at 37°C for 48 h in the presence or absence of 10 µg/ml anti-IgM, 20 µg/ml Polymyxin B (LPS chelator) and either PBS, 300 nM TAT–GFP, 100 nM TAT–cyclin E, 300 nM TAT–cyclin E, 50 nM TAT–cyclin D3, 150 nM TAT–cyclin D3 or 300 nM TAT–cyclin D3. At 48 h, cells were pulsed with 0.5 µCi [3H]thymidine and then cultured for an additional 6 h before determining the amount of [3H]thymidine incorporation. TAT–cyclin E increases the proliferation of both wild-type and c-Rel–/– B cells in a dose-dependent manner, while TAT–cyclin D3 has negligible effect on c-Rel–/– B cells. TAT–GFP acts as a negative control. (E) Cells were cultured as described in (C and D), and harvested on days 1, 2 and 3 for cell cycle analysis using propidium iodide staining and analysis by flow cytometry. Only day 2 data is shown. Cells were subdivided into viable (>=2N DNA) and apoptotic populations (<2N DNA), and the viable population analyzed for cell cycle distribution. Both TAT–cyclin E (TAT–cycE) and TAT–cyclin D3 (TAT–cycD3) increase the percentage of cells entering S–G2/M phases (>2N DNA).

 
In primary B cell cultures, however, only TAT–cyclin E (Fig. 6C), and not TAT–cyclin D3 (Fig. 6D) or TAT–GFP negative control protein, is capable of partially restoring proliferation responses to c-Rel-deficient B cells. Although TAT–cyclin E can also moderately improve wild-type B cell proliferation, c-Rel–/– B cell responses are significantly enhanced with addition of TAT–cyclin E. At 100 and 300 nM, TAT–cyclin E increases [3H]thymidine incorporation 3.8- and 5.4-fold respectively over PBS controls in anti-IgM-treated c-Rel–/– B cells, corresponding to 29 and 34% restoration of wild-type proliferation responses. In contrast, TAT–cyclin D3 has a negligible effect on both wild-type and c-Rel–/– B cells (error bars indicate no significant difference). The combination of TAT–cyclin D3 and TAT–cyclin E proteins did not further enhance the proliferative response over TAT–cyclin E alone (data not shown).

Since anti-IgM induces a high degree of apoptosis in c-Rel–/– B cells, however, the effects of TAT–cyclin D3 are potentially masked by disproportionately low numbers of viable cells able to respond to the TAT–cyclin D3 protein. To investigate this possibility, we analyzed the effects of TAT–cyclin D3, TAT–cyclin E and the control TAT–GFP protein using propidium iodide to discriminate viable cells from dead cells, as well as identify various cell cycle populations based on the DNA content of individual cells. The results in Fig. 6(E) show that TAT–cyclin D3 has a small but significant contribution to the proliferation of both wild-type (14% increase in S–G2/M) and c-Rel–/– B cells (7% increase in S–G2/M) over TAT–GFP controls, which is observable in the viable cell population (>=2N) after apoptotic cells have been excluded (Fig. 6E). The effects of TAT–cyclin E are even greater (19% increase for wild-type in S–G2/M and 21% increase for c-Rel knockout in S–G2/M), while TAT–GFP has little effect on proliferation. These results indicate a major role for cyclin–CDK activity in propagating BCR proliferative signals through c-Rel, but also suggest that other c-Rel-dependent events are required for full engagement of cell cycle progression.


    Discussion
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
We have demonstrated that c-Rel-deficient B cells exhibit diminished expression of several cell cycle regulators, including cyclin D3, cyclin E, cyclin A, pRb, p107, as well as a number of E2F proteins. Furthermore, CDK2, CDK4 and CDK6 kinase activities are significantly impaired, correlating with the reduced phosphorylation of pRb family pocket proteins and diminished E2F activity. The combination of these defects may explain the observed G1 arrest of c-Rel-deficient B lymphocytes and implicates a crucial role for c-Rel in BCR-mediated proliferation. To bypass these proliferative impairments we demonstrate that a downstream cell cycle regulator, cyclin E, can be efficiently delivered by TAT transduction into primary c-Rel-deficient B cells to partially restore anti-IgM-induced proliferation. Interestingly, other NF-{kappa}B/Rel members do not compensate for c-Rel–/– B cell defects (data not shown) (10,11).

It is possible that c-Rel may have direct involvement in the transcriptional regulation of cyclins, perhaps in cooperation with other transcription factors, such as the STAT and E2F families. Promoter analysis of the cyclin D3 gene reveals a putative {kappa}B consensus site GGGGCTTCTC at –21 bp (42), making it a potential target gene of c-Rel. Meanwhile, STAT 5 deficiency in T lymphocytes produces a similar phenotype as c-Rel knockout B cells, including profound defects in G1 cyclin expression (43). Our attempts to show cyclin D3 activation by reporter assays and RNase protection experiments, however, have not been able to substantiate a clear role for c-Rel involvement (data not shown). Furthermore, RT-PCR experiments indicate comparable levels of cyclin D3 mRNA are expressed in anti-IgM stimulated wild-type and c-Rel–/– B cells, suggesting that c-Rel-dependent post-transcriptional mechanisms control cyclin D3 protein levels in these cells. One such possibility is the ubiquitin-mediated proteolytic pathway by which several cell cycle genes are regulated, including the D-type cyclins (44,45).

Kinetically, the expression of cyclin E and cyclin A follow the induction of c-Rel substantially later, and may be subject to regulation by c-Rel-dependent secondary transcription factors such as E2Fs and STATs rather than directly by c-Rel itself. Certainly the fact that cyclins D2, D3, E and A show only partial loss in c-Rel-deficient B cells implies that additional factors participate in cyclin induction. Sub-maximal cyclin induction, however, may not be capable of sustaining threshold CDK activity necessary to drive cells past the G1 restriction point. These observations suggest that other modulators of CDK activity, such as CDK-activating kinase and CDC25A phosphatase, might also be dysregulated in the absence of c-Rel, and should be examined for activity or expression in further study of c-Rel knockout B cells.

Importantly, p27KIP1 can inhibit both cyclin E–CDK2 and cyclin A–CDK2 complexes during G1 and S phases to prevent initiation of DNA replication. Irreversible degradation of p27KIP1 is therefore necessary to elicit progression past the G1 restriction point and must reliably coincide with the timing of S phase cyclin induction (4648). Our data imply that these two pathways are independently controlled. Since p27KIP1 down-regulation appears normal in c-Rel-deficient B lymphocytes, failure to proliferate is more likely to be attributed to a defect in cyclin up-regulation rather than an overabundance of CKI. This finding is intriguing given that other studies have demonstrated a requirement for CDK2 kinase activity in mediating p27KIP1 turnover (46,47,49). Because we observe normal p27KIP1 down-regulation in the absence of any detectable CDK2 activity, our data suggest that other CDK2-independent pathways can efficiently initiate p27KIP1 turnover.

Some CKI can even be induced by BCR signaling during late G1 (p19INK4D and p21CIP1) in what is now thought to be a mechanism to assemble stable cyclin–CDK2 complexes for S phase entry (2,15). We were unable to detect these proteins in resting B cells stimulated with anti-IgM, however, and do not believe that they play a significant role in mediating G1-arrest of c-Rel–/– B cells.

Because primary B cells are highly resistant to DNA transfection, various methods to reintroduce missing components into c-Rel knockout B cells have been considered and tested, but with little success. These methods either require the full proliferation of cells (i.e. retroviral infection), something not achievable in c-Rel–/– B cells, or else are prohibitively expensive and time consuming (i.e. generation and breeding of multiple transgenic mouse strains). The TAT fusion protein transduction method circumvents many of these problems and provides direct replacement of missing molecules to primary B cells. Our results with TAT–cyclin E show a clear improvement in the proliferative capacity of anti-IgM-treated c-Rel–/– B cells. In contrast, TAT–cyclin D3 has minimal effect on c-Rel–/– B cell proliferation, a result consistent with reports contending that cyclin D–CDK activity is insufficient to inactivate pRb at the G1 restriction point without adequate cyclin E–CDK2 activity (16). Nevertheless, our data also suggest that while cyclin E–CDK2 activity partially accounts for propagation of BCR proliferative signals through c-Rel, it appears that additional c-Rel-dependent events are required to achieve full cell cycle progression.

Finally, we conclude that in the absence of c-Rel, other probable defects impede the onset of cell cycle progression, such as the production of cytokines, co-stimulatory molecules and growth factor receptors. Mounting evidence suggests that cytokine signaling can promote cell cycle progression via STAT activation of critical regulators (43,50,51). Recent data suggest that cytokines not only synergize the effects of E2F-mediated transcription (52,53), but can also contribute to the E2F production pool itself (54). Furthermore, cytokines have been demonstrated to play a key role in c-Rel-dependent B cell responses following BCR activation (11,13). More importantly, co-culture experiments combining anti-IgM activated wild-type and knockout B cells have demonstrated a need for cell-autonomous signaling to restore c-Rel-dependent survival and proliferation defects (11,13). This last finding may reflect a requirement for a c-Rel-dependent B cell cytokine receptor and/or its downstream signaling pathway during BCR-mediated expansion.

Together, our data validate the hypothesis that the G1 arrest in BCR-activated c-Rel-deficient B cells results from multiple defects in positive regulators such as poor accumulation of cyclins and diminished activation of E2F transcription factors rather than insufficient degradation of negative regulators like p27KIP1. We note, however, that several other molecules have not been systematically tested or excluded yet. Nevertheless, our current model for BCR-induced proliferation is proposed as thus (Fig. 7). Ligation of the BCR induces both c-Rel-dependent and -independent signaling of cell cycle molecules. Activation of c-Rel leads to the transcription of immediate/early genes which may include direct effects on cyclin gene expression or act indirectly through secondary mediators. The accumulation of cyclins D2 (c-Rel independent) and D3 (c-Rel dependent) during G1 results in threshold CDK4 and CDK6 activity, and pRb/p130 phosphorylation. This leads to E2F de-repression and autoregulation (c-Rel dependent), boosting the induction of G1/S and S phase cyclins E and A respectively, as necessary for subsequent CDK2 activation. Meanwhile, BCR-induced p27KIP1 down-regulation during late G1 (c-Rel independent) further promotes CDK2 activation, thereby enhancing E2F activation via pRb/p107 hyper-phosphorylation, and ultimately resulting in transition across the G1 restriction point into S phase for DNA synthesis and lymphocyte proliferation.



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Fig. 7. Model: c-Rel induction of cell cycle regulators. See text.

 

    Acknowledgements
 
We would like to thank Drs Andrew Koff, Charles Sherr and Yue Xiong for sharing p27KIP1, p16INK4A, p18INK4C, p19INK4D and p21CIP1 antibodies with us; Drs Joan Massague and Stacey Blain for providing the GST–Rb bacterial clone; Dr Natalie Lissy for assistance in purification of TAT fusion proteins; Dr Selina Chen-Kiang for reading the manuscript, and Dr Joseph Tumang for constructing TAT–GFP. The work presented here was supported by NIH grants CA 68155, CA 90405 and 1 T32 A1 07621, the American Cancer Society Junior Investigator Award, the March of Dimes Basil O’Connor Scholarship, and the Leukemia and Lymphoma Society Scholarship.


    Abbreviations
 
CDK—cyclin-dependent kinase

CKI—cyclin-dependent kinase inhibitor

GFP—green fluorescence protein

GST—glutathione-S-transferase

HA—hemagglutinin

LPS—lipopolysaccharide

pRb—Retinoblastoma protein

TBS-T—Tris-buffered saline Tween


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 Methods
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