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The Journal of Immunology, 2006, 177: 787-795.
Copyright © 2006 by The American Association of Immunologists

Disruption of Cyclin D3 Blocks Proliferation of Normal B-1a Cells, but Loss of Cyclin D3 Is Compensated by Cyclin D2 in Cyclin D3-Deficient Mice1

Jennifer M. Mataraza*, Joseph R. Tumang{dagger}, Maria R. Gumina*, Sean M. Gurdak{dagger}, Thomas L. Rothstein2,{dagger},{ddagger} and Thomas C. Chiles3,*

* Department of Biology, Boston College, Chestnut Hill, MA 02467; {dagger} Department of Medicine, Boston University School of Medicine, Boston, MA 02118 and Immunobiology Unit, Evans Memorial Department of Clinical Research, Boston University Medical Center, Boston, MA 02118; and {ddagger} Department of Microbiology, Boston University School of Medicine, Boston, MA 02118


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Peritoneal B-1a cells differ from splenic B-2 cells in the molecular mechanisms that control G0-S progression. In contrast to B-2 cells, cyclin D2 is up-regulated in a rapid and transient manner in phorbol ester (PMA)-stimulated B-1a cells, whereas cyclin D3 does not accumulate until late G1 phase. This nonoverlapping expression of cyclins D2 and D3 suggests distinct functions for these proteins in B-1a cells. To investigate the contribution of cyclin D3 in the proliferation of B-1a cells, we transduced p16INK4a peptidyl mimetics (TAT-p16) into B-1a cells before cyclin D3 induction to specifically block cyclin D3-cyclin-dependent kinase 4/6 assembly. TAT-p16 inhibited DNA synthesis in B-1a cells stimulated by PMA, CD40L, or LPS as well as endogenous pRb phosphorylation by cyclin D-cyclin-dependent kinase 4/6. Unexpectedly, however, cyclin D3-deficient B-1a cells proliferated in a manner similar to wild-type B-1a cells following PMA or LPS stimulation. This was due, at least in part, to the compensatory sustained accumulation of cyclin D2 throughout G0-S progression. Taken together, experiments in which cyclin D3 was inhibited in real time demonstrate the key role this cyclin plays in normal B-1a cell mitogenesis, whereas experiments with cyclin D3-deficient B-1a cells show that cyclin D2 can compensate for cyclin D3 loss in mutant mice.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
The D-type cyclins are regulatory subunits of cyclin-dependent kinase (cdk)4 4 and cdk6 (1, 2, 3, 4, 5, 6). Unlike other cyclins whose protein levels oscillate during the cell cycle, accumulation of D-type cyclins is dependent on mitogenic and oncogenic signals (1, 2). Cyclin D-cdk4/6 complexes initiate phosphorylation of the retinoblastoma tumor suppressor protein (pRb) and pRb-related "pocket proteins" p107 and p130; this promotes dissociation of E2F from pRb, which reverses its G1-S inhibitory function, in part, by induction of E2F target genes (4, 5, 6, 7, 8, 9). In addition, several lines of evidence point to a second noncatalytic function of D-type cyclin-cdk complexes in G0-S progression by sequestering members of the Cip/Kip family (9). By contrast, the p16INK4a tumor suppressor protein uniquely interacts with cdk4 and cdk6, blocking cdk4/6-mediated pRb phosphorylation and leading to G1-phase arrest (10, 11, 12). Recent analyses of mice lacking D-type cyclins, or their catalytic partners cdk4 and cdk6, reveal that these proteins are critically required for proliferation only in selected tissues of the developing embryo, suggesting the existence of cyclin D-independent mechanisms to initiate mitogen-induced proliferation (13, 14, 15).

The three D-type cyclins (cyclins D1, D2, and D3) are encoded by separate genes that exhibit a high degree of amino acid homology and are expressed in adult tissues in a partially overlapping fashion (16, 17). Mice homologous for genetic disruptions in individual D-type cyclin genes are viable and display narrow, tissue-specific phenotypes, suggesting redundant function contributing to viable animals (18). Splenic B-2 lymphocytes express cyclins D2 and D3, but not cyclin D1 following BCR cross-linking (19, 20). Evaluation of bone marrow and spleen from cyclin D2-deficient mice reveals normal numbers of B220+IgM+ B lymphocytes (21, 22); however, the peritoneal CD5+ B-1 cell population is severely diminished in cyclin D2-deficient mice (22). Cyclin D2-deficient B-2 cells also exhibit impaired proliferation in response to BCR cross-linking; that proliferation is not completely blocked may be due to redundancy with cyclin D3 in this tissue (23). Although little is known about the regulation and function of cyclin D3 in B cell subsets, B-2 cells deficient in c-Rel exhibit diminished cyclin D3 induction in response to BCR cross-linking (24). Interestingly, the cyclin D3 gene promoter contains multiple transcription factor recognition sites, including NF-{kappa}B-binding sites (25).

B-1 cells constitute a unique set of B lymphocytes, distinguished from B-2 cells by numerous phenotypic and functional characteristics (26, 27, 28, 29). B-1 cells are further subdivided into CD5+ (B-1a) or CD5– (B-1b) cells and in adult mice represent the principal lymphocyte population in the peritoneal cavity (26, 28, 30). B-1a cells are responsible for the majority of nonimmune serum IgM and contribute considerable amounts of "resting" IgA (30). B-1a cells are associated with several human disease states characterized by aberrant B cell growth. In mice, monoclonal expansions of B-1a cells that resemble human chronic lymphocytic leukemia develop as a function of age; further, New Zealand Black mice, which contain increased numbers of B-1a cells, develop several types of B-1a-derived lymphoid malignancies in early life (31, 32, 33).

In adult animals, peritoneal B-1a cells are self-replenishing, giving rise to their own progeny, following establishment early during ontogeny, whereas B-2 cells arise continually from surface Ig-negative stem cell precursors and fail to proliferate following maturation in the absence of exogenous stimulation (30, 34). Ex vivo B-1a cells fail to proliferate following BCR cross-linking, which drives B-2 cells into S phase (35, 36). Conversely, B-1a cells enter S phase in response to PMA and do so unusually rapidly, whereas B-2 cells require the combination of PMA and a calcium ionophore (35, 37). These results establish that B-1a cells differ from B-2 cells in the molecular mechanisms that control G0-S phase progression. In support of this, cyclins D2 and D3 are expressed in a nonoverlapping manner during G0-S progression in PMA-stimulated B-1a cells (38, 39). Notably, cyclin D2 is up-regulated in a rapid and transient fashion in B-1a cells, whereas cyclin D3 induction and assembly into active cdk4/6-containing complexes occurs after cyclin D2 degradation and parallels peak endogenous pRb phosphorylation in late G1 phase (39). In marked distinction, mitogenic stimulation of B-2 cells leads to coordinate induction of cyclins D2 and D3 (19, 20, 40). Thus, in B-1a cells, but not B-2 cells, stimulated expression of cyclin D2 and cyclin D3 is temporally distinct. Taken together, these observations suggest that cyclin D3 is uniquely positioned in B-1a cells to mediate transition through the G1-S boundary, which may, as well, reflect its role in conventional B-2 cells where cyclin D3 expression overlaps that of cyclin D2. To investigate the need for cyclin D3-cdk complexes for G0-S phase progression in normal B-1a cells, we used protein transduction technology to specifically block assembly and activation of cyclin D3 holoenzyme complexes in normal peritoneal B-1a cells, and we examined the proliferative responses of B-1a cells from cyclin D3-deficient mice.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Preparation of murine B-1a and B-2 lymphocytes

The generation of cyclin D3-deficient mice has been described (41). BALB/cByJ mice were purchased from The Jackson Laboratory and housed at Boston University Medical Center or Boston College. The studies described below were reviewed and approved by institutional animal care and use committees at both institutions. Mice were cared for and handled at all times in accordance with National Institutes of Health and University guidelines. Unseparated cells were obtained by peritoneal washout and splenic disruption, were stained with immunofluorescent Abs directed against B220 and CD5, and were subjected to FACS at 4°C using a Mo-Flo flow cytometer (DakoCytomation) to yield purified peritoneal B-1a (B220+CD5+) and splenic B-2 (B220+CD5–) cells, including the use of an anti-CD8 "dump" channel for B cell purification, as previously described (42). Sort-purified B cell populations were reanalyzed by immunofluorescent staining with Abs directed against CD3 and CD14. Sort-purified peritoneal B-1a cells and splenic B-2 cells were found to be >95% (SEM ± 0.98%) and >96% (SEM ± 0.84%) pure, respectively. In addition, both populations contained <1.8% and 1.2% CD3+ or CD14+ cells, respectively. B cells were cultured in RPMI 1640 medium supplemented with 10% heat-inactivated FCS (Atlanta Biologicals) as previously described (39).

TAT-p16 peptides

Peptides were synthesized by the Biotechnology and Genomics Research Center (Utah State University, Logan, UT). The 32-mer peptides contain an NH2-terminal 11 residue TAT protein transduction domain (YGRKKRRQRRR) immediately followed by a glycine residue and either a 20-mer wild-type p16 sequence (DAAREGFLATLVVLHRAGAR) or a charge-match control sequence (ARGRALTAHVDRLGEFVAAL), as described (43, 44).

Cell cycle analysis and proliferation assay

B cells (105) were resuspended in 300 µl of PBS containing 0.1% (v/v) Triton X-100, 200 µg/ml DNase-free RNase A, and 20 µg/ml propidium iodide (42). B cell fluorescence was then acquired with a BD FACSCanto flow cytometer (BD Biosciences) and the data were analyzed by ModFit LT software (Verity Software House). To measure proliferation, B cells (1–2 x 104 in 0.2 ml) were cultured in quadruplicate and stimulated as indicated in the figure legends. DNA synthesis was measured by incubating B cells with 0.5 µCi [3H]thymidine (20 Ci/mmol; New England Nuclear) during the last 6 h of culture. Cells were then harvested onto glass fiber filters and [3H]thymidine incorporation into DNA was quantitated by liquid scintillation spectroscopy.

Western blotting

B cells were solubilized in Triton X-100 buffer (20 mM Tris (pH 7.4), 100 mM NaCl, 0.1% Triton X-100) containing 2.5 µg/ml leupeptin/aprotinin, 10 mM beta-glycerophosphate, 1 mM PMSF, 1 mM NaF, and 1 mM Na3VO4. Insoluble debris was removed by centrifugation at 15,000 x g for 15 min (4°C). Lysate protein was separated by electrophoresis through a 10% polyacrylamide SDS gel (SDS-PAGE) and transferred to an Immobilon-P membrane. The membrane was blocked in TBS-T (20 mM Tris (pH 7.6), 137 mM NaCl, and 0.05% Tween 20) containing 5% nonfat dry milk for 5 h and then incubated overnight (4°C) with primary Ab at 1 µg/ml in TBS-T. The membrane was washed several times in TBS-T, incubated with a 1/2500 dilution of anti-rabbit or anti-mouse IgG-coupled HRP Ab (60 min) and developed by ECL.

Fluorescence microscopy

For imaging the TAT-p16-FITC, slides were washed once with PBS and then mounted with Aqua Polymount. For imaging D-type cyclins, lymphocytes (2.5 x 105) were fixed with methanol for 10 min at –20°C and then permeabilized with 0.1% Triton X-100 at room temperature. Cells were blocked for 30 min with 2% BSA in PBS and then washed several times with PBS. Detection of cyclin D2 and cyclin D3 was performed by incubating cells with 1/50 anti-cyclin D2 or anti-cyclin D3 mAbs at 4°C, respectively. After several washes with PBS, cells were incubated with a 1/200 FITC-conjugated Affinipure goat anti-mouse Ab (Jackson ImmunoResearch Laboratories) for 2 h at room temperature. The slides were then dried, mounted with Aqua Polymount, and immunofluorescence images were captured with a Leica confocal microscope.

Gene expression

RNA was prepared from B cells using Ultraspec reagent (BiotecX) and was DNase treated. cDNA was prepared using an iScript cDNA Synthesis kit (Bio-Rad), and normalized by PCR for beta2-microglobulin expression as previously described (42). Gene expression was then assessed by real-time PCR (Stratagene) using the following primers (forward/reverse): beta2-microglobulin (CCCGCCTCACATTGAAATCC/GCGTATGTATCAGTCTCAGTGG); cyclin D2 (TGGGCTTCAGCAGGATGATG/ACGGAACTGCTGCAGGCTGT); cyclin E1 (TGGATTTGCTGGACAAAGCC/TGGATTTGCTGGACAAAGCC); cyclin E2 (TAGGAATTGTTGGCCACCTGT/AATCTGGCAGAGGTGAGGGAT); E2F-1 (GAGGCTGGATCTGGAGACTGA/CAAGAAGCGTTTGGTGGTCA); and VH11 (GCAATAAACTACGCACCATCCA/TGTCCTCCGATCGCACATT). Primers for GAPDH and a GAPDH TaqMan probe were obtained from Applied Biosystems.

ELISPOT

Spontaneous secretion of IgM was measured as described previously (45). Naive, FACS-sorted B-1 and B-2 cells, and FACS-sorted B-2 cells stimulated with LPS at 25 µg/ml for 48 h, were distributed at various concentrations onto Multiscreen-IP plates (Millipore) precoated with goat anti-mouse Ig (H+L; Southern Biotechnology Associates) and then incubated for 3 h at 37°C and 5% CO2. Plates were treated with alkaline phosphatase-conjugated goat anti-mouse IgM (Southern Biotechnology Associates) and developed with 5-bromo-4-chloro-3-indolyl phosphate/p-NBT chloride substrate (KPL). Spots reflecting Ig-secreting cells were enumerated using Phoretix Expression software (NonLinear Dynamics).

Reagents and Abs

Protein G agarose and anti-mouse IgG-coupled HRP, anti-mouse cdk4, anti-cyclin D2, and anti-cyclin D3 Abs (Abs) were purchased from Santa Cruz Biotechnology. The phospho-pRb (Ser807/811) and phospho-cdk2 (Thr172) Abs were obtained from Cell Signaling Technologies. Rabbit complement and Lympholyte M were purchased from Accurate Chemical and Scientific. All other chemicals were obtained from Sigma-Aldrich. Fluorescent-labeled Abs directed against B220, CD5, CD23, Mac-1, CD4, CD8, CD3, and CD14 for FACS and flow cytometric analysis were obtained from BD Pharmingen. ECL reagents were obtained from Kirkegaard & Perry Laboratories. PMA and LPS from Salmonella typhimurium (LPS) were obtained from Sigma-Aldrich and used at 300 ng/ml and 25 µg/ml, respectively. Soluble rCD40L was obtained from transfected J558L cells that secrete a chimeric CD40L/CD8{alpha} fusion protein and prepared as previously described (46, 47). Anti-CD8{alpha} Ab was obtained from the supernatant of 53-6-72 hybridoma cells and was used to cross-link rCD40L (47). CD40L was used at a 1/10 dilution of supernatant and anti-CD8{alpha} Ab was used at a 1/40 dilution of supernatant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
TAT-p16INK4a peptidyl mimetics are transduced into ex vivo peritoneal B-1a cells and disrupt endogenous D-type cyclin-cdk4/6 complexes

Our strategy for determining whether cyclin D3-cdk4/6 complexes are required for proliferation of B-1a cells was to precisely target cyclin D3-cdk4/6 holoenzymes by transducing p16INK4a peptidyl mimetics into ex vivo B-1a cells. Lane and coworkers (43) demonstrated that a 20-mer peptide derived from the third ankyrin-like repeat of the p16INK4a tumor suppressor protein selectively bound to cdk4 and cdk6 and inhibited cdk4/6-mediated pRb phosphorylation in vitro. To mediate efficient transduction into B-1a cells, we coupled the p16INK4a peptidyl mimetic to an 11-mer peptide consisting of the NH2-terminal HIV TAT protein domain (denoted herein as TAT-p16 wild type) (44). Flow cytometry revealed transduction of nearly 100% of B-1a cells that had been incubated with TAT-p16-FITC wild-type peptide for 120 min (Fig. 1A). Confocal microscopy of parallel B-1a cells confirmed intracellular transduction of TAT-p16-FITC wild-type peptide into B-1a cells (Fig. 1B).


Figure 1
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  		FIGURE 1. Transduction of TAT-p16 wild-type peptide into B cells. A, B-1a cells were incubated for 120 min in the presence of 10 µM TAT-p16-FITC peptide, washed once in PBS, and analyzed by flow cytometry (TAT-p16). Untreated cells were analyzed for comparison (Control). B, Parallel B cells were visualized by microscopy as described in Materials and Methods. Both bright field (left panel) and fluorescence (right panel) images were collected at x100 magnification. The data are representative for three independent experiments.

 
To demonstrate in vivo efficacy, 10 µM TAT-p16 wild-type peptides were transduced into PMA-stimulated B-1a cells, which express cyclin D3-cdk4 complexes as evidenced by immunoreactive cyclin D3 in nondenatured anti-cdk4 immunoprecipitates (Fig. 2A, Control) (39). Transduction of TAT-p16 wild-type peptide into B-1a cells before cyclin D3 induction prevented cyclin D3-cdk4 complex assembly (Fig. 2A, p16WT); a control charge-matched TAT-p16 (TAT-p16 mutant) peptide, that is unable to bind to cdk4 or cdk6, did not block cyclin D3-cdk4 complex assembly (Fig. 2A, p16Mut). Similar results were obtained when evaluating the efficacy of TAT-p16 peptides with respect to endogenous cyclin D3-cdk6 complexes (data not shown). Asynchronously growing murine Bal17 B cells were analyzed in parallel, as they constitutively express assembled cyclin D2-cdk4 complexes (Fig. 2B, Control). Transduction of TAT-p16 wild-type peptide disrupted endogenous cyclin D2-cdk4 complexes, whereas the TAT-p16 mutant peptide had minimal effect (Fig. 2B). As a control for specificity, we measured cdk-activating kinase-mediated Thr172 phosphorylation of cdk2; this modification occurs subsequent to the nuclear translocation of cyclin E-cdk2 and, thus, serves as an indicator for the presence of cyclin E-cdk2 complexes. Transduction of TAT-p16 wild-type peptide into PMA-stimulated B-1a cells before cyclin E-cdk2 assembly did not measurably affect PMA-induced Thr172 phosphorylation of cdk2 (Fig. 2C, PMA).


Figure 2
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  		FIGURE 2. TAT-p16 wild-type peptide blocks formation of cyclin D3-cdk4 complexes in B-1a cells. A, Peritoneal B-1a cells were stimulated with 300 ng/ml PMA to induce the expression of cyclin D3 and assembly of cyclin D3-cdk4 complexes (Control). In parallel sets of B-1a cells, 10 µM TAT-p16 wild-type (p16 WT) or TAT-p16 mutant (p16 Mut) peptides were added at 14 h after PMA, before cyclin D3 induction. At 20 h, B-1a cells were collected and immunoprecipitated (IP) with 1.5 µg of anti-cdk4 Ab as described (39 ). The immune complexes were separated by SDS-PAGE and Western blotted with anti-cyclin D3 Ab. The blot was also probed with anti-cdk4 Ab to ensure that equal amounts of cdk4 were immunoprecipitated. B, Asynchronously growing Bal17 B cells were similarly treated with the TAT-p16 peptides for 4 h; B cells were then collected, immunoprecipitated (IP) with 1.5 µg of anti-cyclin D2 Ab, and the immune complexes were analyzed by Western blot with anti-cdk4 Ab. The blot was also probed with anti-cyclin D2 Ab to ensure that equal amounts of cyclin D2 were immunoprecipitated. C, B-1a cells were cultured in medium alone (M) or stimulated with 300 ng/ml PMA for 24 h to induce cyclin E-cdk2 assembly. Where indicated, 10 µM TAT-p16 wild-type (p16 WT) or TAT-p16 mutant (p16 Mut) peptides were added during the last 4 h of PMA treatment. Phosphorylation of cdk2 on Thr172 was detected by Western blotting with an anti-phospho(Thr172)-cdk2 Ab. The blot was stripped and reprobed with an anti-beta-actin Ab. The data are representative of two independent experiments.

 
Transduction of TAT-p16 wild-type peptide into normal peritoneal B-1a cells inhibits proliferation

To directly evaluate the contribution of cyclin D3-cdk4/6 complexes in B-1a cell proliferation, we took advantage of the nonoverlapping expression of cyclins D2 and D3 in PMA-stimulated B-1a cells (38, 39). Blocking the assembly of temporally expressed cyclin D3-cdk4/6 complexes was achieved by transducing the TAT-p16 wild-type peptide into B-1a cells at 14 h after stimulation with PMA. This time corresponds to a point in the cell cycle wherein cyclin D2 protein is not detectable and cyclin D3 protein induction has not yet begun (initially detectable at 17 h). B-1a cells transduced with TAT-p16 wild-type peptide exhibited a >70% reduction in PMA-stimulated tritiated thymidine incorporation in comparison to control B-1a cells or B-1a cells transduced with TAT-p16 mutant peptide (Fig. 3A). Similar results were obtained with B-1a cells stimulated with LPS or CD40L (Fig. 3A). Of note, incubation of nonstimulated B-1a cells with TAT-p16 peptides did not alter the basal level of tritiated thymidine incorporation (Fig. 3A, Medium). To corroborate these findings, we analyzed B-1a cells for cell cycle position by propidium iodide staining and flow cytometry. Transduction of TAT-p16 mutant peptide into PMA-stimulated B-1a cells had a minimal effect on the percentage of B-1a cells in S/G2+M phases of the cell cycle in comparison to control B-1a cells (Fig. 3B, PMA). By contrast, transduction of TAT-p16 wild-type peptide reduced the percentage of S/G2+M B-1a cells by 70% in comparison to B-1a cells transduced with TAT-p16 mutant peptide. Similar results were obtained with LPS- or CD40L-stimulated B-1a cells (Fig. 3B).


Figure 3
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  		FIGURE 3. TAT-p16 wild-type peptide inhibits DNA synthesis in B-1a cells in response to PMA, LPS, or CD40L. B-1a cells were cultured in medium alone, 300 ng/ml PMA, 25 µg/ml LPS, or CD40L for 24 h. Where indicated, 10 µM TAT-p16 wild-type (WT) or TAT-p16 mutant (Mut) peptides were added at 14-h postmitogen addition. Control (C) denotes B-1a cells cultured in the absence of added peptides. A, DNA synthesis was monitored by tritiated thymidine incorporation. Mean results are shown, along with lines indicating SEs of the means (n = 4). B, B-1a cells were stained with propidium iodide and analyzed by flow cytometry as described in Materials and Methods. The data are represented as the percentage of B-1a cells in the S + G2-M phase of the cell cycle. The data are representative of three independent experiments. C, B-1a cells were cultured in medium alone (M) or stimulated with 300 ng/ml PMA in the absence or presence of 10 µM TAT-p16 wild-type (p16WT) or TAT-p16 mutant (p16Mut) peptides as described in A. Whole cell extracts were prepared, and Western blotting was performed with an anti-phospho-pRbSer807/811 Ab. The blot was stripped and reprobed with anti-pRb Ab.

 
To determine the status of endogenous pRb phosphorylation in B-1a cells treated with TAT-p16 wild-type peptide, whole cell extracts were prepared and immunoblotted with anti-phospho(Ser807/811)pRb Ab that specifically detects cyclin D-cdk4/6-mediated phosphorylation of pRb. Unstimulated B-1a cells did not express measurable pRb phosphorylation, whereas PMA stimulated B-1a cells exhibited abundant phosphorylation of endogenous pRb on Ser807/811 (Fig. 3C). Phosphorylation of endogenous pRb on Ser807/811 was markedly inhibited following transduction of TAT-p16 wild-type, but not TAT-p16 mutant, peptides into B-1a cells (Fig. 3C). Total pRb protein levels were not altered by transduction of the TAT-p16 peptides. These results indicate that TAT-p16 wild-type peptide blocks endogenous pRb phosphorylation by cyclin D-cdk4/6 complexes in PMA stimulated B-1a cells.

Cyclin D3-deficient mice have normal peripheral B-1 and B-2 lymphocyte compartments

Because the results above indicate that cyclin D3-cdk4/6 complexes are required for mitogenic stimulation of normal B-1a cells, we were interested in determining whether B-1a cell development and proliferation were affected by loss of cyclin D3 (41). To pursue this, we initially evaluated the splenic and peritoneal lymphoid compartments of cyclin D3-deficient mice. To confirm the absence of cyclin D3 in D3-deficient animals, total splenic lymphocytes were stimulated with a combination of mitogens known to induce cyclins D2 and D3 in both T and B lymphocyte populations (19, 20, 24, 40); whereas up-regulation of both cyclins D2 and D3 was apparent in spleen cells from wild-type mice, only cyclin D2 was induced in cyclin D3-deficient splenic lymphocytes and no cyclin D3 protein was observed (Fig. 4A). Cyclin D3-deficient spleens contained a decreased number of total lymphocytes as compared with spleens obtained from wild-type littermate animals, which immunofluorescent staining showed was largely attributable to decreased numbers of B-2 cells (Fig. 4, B and C). However, there was no difference between cyclin D3-deficient mice and wild-type littermates in the total number of peritoneal cells, and, more specifically, the numbers of B-1a cells (CD5+B220lowMac-1+), B-1b cells (CD5–B220lowMac-1+), B-2 cells, T cells and macrophages, recovered by peritoneal washout (Fig. 4, B and D).


Figure 4
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  		FIGURE 4. Characteristics of B-1a cell lymphoid compartments in wild-type and cyclin D3-deficient mice. A, Total splenic lymphocytes were isolated from wild-type (+/+) and cyclin D3-deficient mice (–/–) and were either left untreated (M) or were stimulated (S) with a mitogenic combination consisting of 300 ng/ml PMA plus 400 ng/ml ionomycin and 25 µg/ml LPS. At 24 h, lymphocytes were collected, detergent extracts were prepared, and then Western blotting was performed with anti-cyclin D2 or anti-cyclin D3 Abs. The blot was stripped and reprobed with an anti-beta-actin Ab to verify equal loading of each lane. B, Peritoneal washouts (peritoneum) and spleen cell suspensions (spleen) were obtained from cyclin D3-deficient mice (–/–, {square}) and wild-type littermate control animals (+/+, {cjs2108}), and total cell numbers were determined. Mean results are shown, along with lines indicating SEs of the means (n = 6). C, Spleen cell suspensions were obtained from cyclin D3-deficient (–/–) and littermate control mice (+/+). The distribution of splenic T cells (T-S; B220–CD5+), B-2 cells (B-2S; B220+CD5–), and MZ cells (MZ-S; CD21highCD23low) was determined by immunofluorescent staining and flow cytometric analysis and converted to cell number based on initial cell counts. Mean numbers of cells in each lymphoid population are shown, along with lines indicating SEs of the means (n = 6 except for MZ-S where n = 2 and the line depicts the range of values). D, Peritoneal washout cells were obtained from cyclin D3-deficient (–/–) and littermate control mice (+/+). The distribution of peritoneal B-1a cells (B-1aP; B220lowCD5+Mac-1+), B-1b cells (B-1bP; B220lowCD5–Mac-1+), B-2 cells (B-2P; B220+CD23+), T cells (T-P; B220–CD5+Mac-1–), and macrophages (M0-P); forward and side scatter high, Mac-1+) was determined by immunofluorescent staining and flow cytometric analysis and converted to cell number based on initial cell counts. Mean numbers of cells in each lymphoid population are shown, along with lines indicating SEs of the means (n = 6).

 
Furthermore, as determined by real-time PCR, there was no difference between cyclin D3-deficient and wild-type animals in the typically high level of peritoneal B-1a cell VH11 expression as compared with the relatively low level expression found in splenic B-2 cells (Fig. 5A). In addition, as shown by ELISPOT (Fig. 5B), there was no difference between cyclin D3-deficient and control animals in the typically high level of spontaneous B-1a cell IgM secretion as compared with the relatively low level characteristic of naive B-2 cells which, as expected, was increased by LPS stimulation for 2 days. Thus, by several measures, the development and function of cyclin D3-deficient peritoneal B-1a cells is normal.


Figure 5
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  		FIGURE 5. Functional aspects of B-1a cells in cyclin D3-deficient mice. A, Splenic B-2 (B-2S) and peritoneal B-1 (B-1P) cells were isolated by cell sorting from cyclin D3-deficient (–/–) and littermate control mice (+/+). VH11 expression was determined by real-time PCR and normalized to expression of GAPDH using primer sets described in Materials and Methods. Mean normalized values for three independent experiments are shown along with lines indicating SEs of the means. B, Splenic B-2 (B-2S) and peritoneal B-1 (B-1P) cells were isolated by cell sorting from cyclin D3-deficient (–/–) and littermate control mice (+/+). Naive B cells ({cjs2108}), and B-2S cells stimulated with 25 µg/ml LPS for 48 h ({blacksquare}), were applied to precoated plates that were incubated for 3 h, after which IgM secretion was assessed by ELISPOT assay, as described in Materials and Methods. Results are shown for two mice per group, with lines indicating the range of values. C, B-1a cells from wild-type (WT) or cyclin D3-deficient (KO) were cultured in medium alone (inset) or stimulated with 300 ng/ml PMA or 25 µg/ml LPS for the times indicated. Incorporation of tritiated thymidine was assessed for the final 6 h of culture as described in Materials and Methods. Results represent mean values of triplicate cultures with lines indicating SEs of the means. The data are representative of three independent experiments.

 
We next evaluated the role of cyclin D3 in peritoneal B-1a cell proliferation induced by PMA. In contrast to the results obtained when cyclin D3-cdk4/6 complex assembly was prevented by transduction of TAT-p16 wild-type peptide into normal B-1a cells, we found that cyclin D3-deficient B-1a cells responded comparably to wild-type B cells stimulated by PMA (Fig. 5C). In addition, the proliferation of cyclin D3-deficient B-1a cells in response to LPS was similar to wild-type B-1a cells (Fig. 5C). In data not shown, similar results were obtained with B-1a cell populations stimulated with CD40L.

Deregulation of cyclin D2 expression in cyclin D3-deficient B-1a cells

To understand the molecular basis of the apparently normal proliferation of cyclin D3-deficient B-1a cells in response to PMA, we compared the gene expression pattern of cyclin D2 and cyclin E in wild-type and cyclin D3-deficient B-1a cells. These analyses revealed little difference in the expression of mRNAs encoding cyclins E1 and E2 in the PMA-stimulated B-1a cell populations (Fig. 6A). In B-2 cells, E2F-1 induction by BCR cross-linking corresponds to late G1 phase of the cell cycle and coincides with pRb hyperphosphorylation (23). The expression of E2F-1 mRNA in PMA-stimulated B-1a cells from cyclin D3-deficient mice was comparable to wild-type B-1a cells (Fig. 6A). In agreement with our previous results in normal B-1a cells stimulated with PMA (38), cyclin D2 mRNA was elevated at 2 h and reached a maximal level within 4 h, which corresponded to a 1.3- and 2-fold increase above nonstimulated B-1a cells, respectively (Fig. 6A). However, in PMA-stimulated cyclin D3-deficient B-1a cells, we detected a 4.5- and 8.3-fold increase in cyclin D2 mRNA above the levels observed in nonstimulated B-1a cells at 2 and 4 h, respectively. In keeping with this, in PMA-stimulated cyclin D3-deficient B-1a cells, the level of endogenous cyclin D2 protein measured at 4 h was greater than that of parallel PMA-stimulated B-1a cells from control wild-type mice (~4.5-fold, based on scanning densitometry of the ECL exposed film obtained after Western blot of B-1a cell lysates) (Fig. 6B).


Figure 6
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  		FIGURE 6. Expression of cyclin D2 is dysregulated in PMA-stimulated cyclin D3-deficient B-1a cells. A, Expression levels of cyclins D2, E1, and E2 as well as E2F-1 in PMA-stimulated wild-type (+/+) (•) and cyclin D3-deficient (–/–) ({circ}) B-1a cells relative to beta2-microglobulin were determined by real-time PCR as described in Materials and Methods. Results are representative of two independent experiments. B, B-1a cells were isolated from wild-type (+/+) and cyclin D3-deficient mice (–/–) and cultured in medium alone (M) or stimulated with 300 ng/ml PMA (P) for 4 h. Whole cell extracts were prepared, and Western blotting was performed with anti-cyclin D2 Ab. The blot was stripped and reprobed with anti-beta-actin Ab. Parallel B-1a cells were cultured in medium alone (M) or stimulated with 300 ng/ml PMA (P) for 21 h and then Western blotted with anti-phospho-pRbSer807/811 Ab. C, B-1a cells from wild-type (cyclin D3+/+) and cyclin D3-deficient (cyclin D3–/–) mice were cultured in medium alone (M) or stimulated with 300 ng/ml PMA for 4 and 21 h. B-1a cells were collected and prepared for indirect immunofluorescence staining of cyclin D2 and cyclin D3 as described in Materials and Methods. The data are representative of two independent experiments. D, Cyclin D3-deficient B-1a cells were cultured in medium alone or stimulated with 300 ng/ml PMA for 24 h. Where indicated, 10 µM TAT-p16 wild-type (WT) or TAT-p16 mutant (Mut) peptides were added together with PMA. C denotes control B-1a cells cultured in the absence of added peptide. DNA synthesis was monitored by tritiated thymidine incorporation. Results represent mean values of triplicate cultures with lines indicating SEs of the means. The data are representative of two independent experiments.

 
To obtain further evidence that this elevated induction of cyclin D2 may act to compensate for the loss of cyclin D3, we determined the expression of cyclin D2 protein in wild-type and mutant B-1a cells by indirect immunofluorescence microscopy. As expected, cyclins D2 and D3 were not detected in nonstimulated B-1a cells (Fig. 6C, lane M); the weak fluorescence signal detected in nonstimulated B-1a cells was similar in intensity to that of parallel B-1a cells stained with a control isotype mAb (data not shown). Stimulation of wild-type B-1a cells with PMA resulted in the nonoverlapping expression of cyclin D2 and cyclin D3 at 4 and 21 h, respectively (Fig. 6C). In contrast, immunofluorescent staining of cyclin D2 in cyclin D3-deficient B-1a cells revealed that cyclin D2 was expressed at both 4 and 21 h post-PMA stimulation. It should be mentioned that similar results were obtained using flow cytometry to evaluate intracellular D-type cyclin expression in wild-type and cyclin D3-deficient B-1a cells (data not shown). Taken together, these results suggest that in contrast to wild-type B-1a cells, wherein cyclin D2 protein is induced in a rapid and transient manner, expression of cyclin D2 remains elevated throughout the G0-S interval in cyclin D3-deficient B-1a cells stimulated with PMA.

Evidence that cyclin D2 protein was functional in cyclin D3-deficient B-1a cells was provided by the measurable level of endogenous pRb phosphorylation on cyclin D-cdk4/6-targeted residues, detected by Western blotting of B-1a cell lysates at 21 h following PMA stimulation (Fig. 6B, lower panel); this level of pRb phosphorylation was comparable to wild-type B-1a cells stimulated with PMA. It is important to note that in addition to cyclin D3, cyclin D1 was not expressed in PMA stimulated cyclin D3-deficient B-1a cells (data not shown). Further evidence that PMA-induced proliferation of B-1a cells in the absence of cyclin D3 may result from compensation by cyclin D2 was obtained by transduction of TAT-p16 wild-type peptide into cyclin D3-deficient B-1a cells, wherein only cyclin D2 expression is detectable, that resulted in an ~80% reduction of PMA-stimulated tritiated thymidine incorporation (in comparison to control B-1a cells or B-1a cells transduced with TAT-p16 mutant peptide) (Fig. 6D).


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
To summarize the findings herein, the temporal disjunction between cyclin D2 and cyclin D3 expression in B-1a cells provided the means to dissect the role of late G1-phase cyclin D3 in mediating mitogenic responses. Inhibition of cyclin D3-cdk4/6 complexes with TAT-p16 wild-type peptide blocked normal B-1a cell proliferation in response to PMA, LPS, and CD40L, indicating that unfettered early and transient up-regulation of cyclin D2 is insufficient to drive normal B-1a cells into S phase of the cell cycle. However, complete loss of cyclin D3 did not adversely affect B-1a cell proliferation, because of a countervailing compensatory increase in cyclin D2 mRNA and sustained accumulation of functioning cyclin D2 protein in cyclin D3-null B-1a cells after stimulation with PMA. Thus, dysregulated cyclin D2 can compensate for the loss of cyclin D3 in fostering B-1a cell cycle progression in knock out mice. Our results also strike a cautionary note by demonstrating that compensation by cyclin D2 at the molecular or cellular level in cyclin D3-deficient mice can mask the normal function that cyclin D3 provides in the mitogenesis of B-1a cells.

Our previous findings raised the possibility that cyclin D3 is uniquely positioned to mediate G1-to-S phase progression in B-1a cells (38, 39). Notably, PMA stimulation of B-1a cells resulted in an early and transient induction of cyclin D2; the assembled cyclin D2-cdk4/6 complexes were transiently active, but only accounted for a relatively minor amount of the total endogenous pRb phosphorylation (38). Evidence for a second D-type cyclin functioning in G1-S progression was supported by our findings that cdk4/6-mediated phosphorylation of endogenous pRb increased dramatically in late G1 phase (during a time period wherein cyclin D2 protein and its associated pRb kinase activity was not detectable); this coincided with the induction of cyclin D3 and its assembly with cdk4/6 into active pRb phosphorylating complexes (39). In the present work, we were able to rapidly and efficiently transduce TAT-p16 peptides into primary B-1a cells. This methodology allows for biochemical manipulation of the entire ex vivo B-1a cell population at precisely timed intervals and avoids many of the problems associated with transduction of expression plasmids, such as artifacts induced by the unregulated expression of recombinant proteins (44). p16 is known to bind to cdk4 and cdk6 and, consistent with that, TAT-p16 peptide, but not a charge-matched mutant TAT-p16 peptide, blocked cyclin D3-cdk4/6 assembly and resulted in a loss of PMA-induced phosphorylation of endogenous pRb specifically on D-type cyclin-cdk4/6-targeted residues. Transduction of TAT-p16 peptide into normal B-1a cells results in inhibition of proliferation, as evidenced by reduced PMA-stimulated tritiated thymidine incorporation and reduced percentage of B-1a cells in S/G2+M phases of the cell cycle. Similar results were obtained with B-1a cells stimulated via LPS or CD40L.

Our results also indicate that in normal B-1a cells where cyclin D3-cdk4/6 complex assembly has been selectively blocked by TAT-p16 peptide, the early and transient induction of cyclin D2 is not sufficient to mediate proliferation induced by PMA, LPS, or CD40L. This might reflect the relatively low pRb kinase activity associated with cyclin D2-cdk4/6 complexes and/or the transient nature of cyclin D2 holoenzyme activation in stimulated B-1a cells (38). In the absence of additional pRb phosphorylation (mediated via cyclin D3-cdk4/6), the activity and duration of cyclin D2/cdk-mediated pRb phosphorylation alone may not be of sufficient strength to drive progression through the G1-S transition. On a related point, it is highly unlikely that TAT-p16 peptide blocked B-1a cell proliferation by interfering with levels of cyclin D2 that may be below the level of detection, because the pRb kinase activity of cyclin D2 is relatively low as compared with cyclin D3 and no measurable cyclin D2 kinase activity has been detected at 17 h after stimulation. Alternatively, cyclin D2 function may not be directly involved in proliferation, but rather may be limited to promoting B-1a cell growth (i.e., accumulation of cell mass), occurring in early G1 phase of the cell cycle (48). It should be mentioned that while the analysis of cyclin D2-deficient mice has revealed significantly decreased numbers of peritoneal B-1a cells, the molecular mechanism(s) underlying this loss remain to be established (22).

It is recognized that although D-type cyclins show high amino acid homology within the cdk-binding region (75–78%), the extent of homology outside of this region is 39–47% (16, 17). Thus, we cannot rule out the possibility that separate from mediating pRb phosphorylation and G1-S phase progression, cyclin D3 may carry out additional functions in B-1a cells. For example, D-type cyclins exhibit cdk-independent functions that act as either negative or positive regulators of transcription factors, such as STAT3 and cyclin D-interacting myb-like protein-1, in addition to having a role in cell cycle regulation (reviewed in Ref. 2). Cyclin D3 was identified as a negative regulator of the hemopoietic transcription factor acute myeloid leukemia 1 (AML1), presumably by a mechanism that involves displacement of core-binding factor beta from AML1, thereby inhibiting AML1’s DNA binding to target gene promoters (49). Recent reports by Gu and coworkers (50, 51) have served to extend the list of cyclin D3 partner proteins beyond transcriptional regulators to include the signal transduction protein kinase ERK3, and the translational regulator eIF3k. Notwithstanding, our results provide the first direct evidence that cyclin D3 in the context of assembled cyclin D3-cdk complexes is required for the G1-S phase progression in stimulated, normal B-1a cells.

We further analyzed the function of cyclin D3 by isolating and then stimulating B-1a cells from cyclin D3-deficient mice. Mice homozygous for a mutant allele containing a targeted deletion of the first two coding exons of cyclin D3 are viable, but suffer from defects in thymocyte development characterized by reduced CD4+CD8+ double-positive T cells (41). Cyclin D3-null thymocytes fail to undergo the proliferative burst associated with the CD4–CD8– double-negative 3 to double-negative 4 transition. Our analyses herein of cyclin D3-deficient splenocytes revealed a decrease over wild-type littermates in the total cell number produced, in large part, by a decrease in the number of splenic B-2 (and not marginal zone) B cells that remains unexplained. Importantly, we found that cyclin D3 deficiency did not impact the peritoneal B-1a cell compartment, as the numbers of peritoneal B-1a and B-1b cells in cyclin D3-deficient mice were comparable to wild-type littermates. Furthermore, increased VH usage and spontaneous IgM secretion were similar for peritoneal B-1a cells in cyclin D3-deficient mice as compared with wild-type littermates. Consistent with this, we found that the serum levels of IgA and IgM were not significantly altered in cyclin D3-null mice in comparison to wild-type mice (data not shown). These results suggest that cyclin D3 is dispensable for B-1a cell development, self-renewal, and function. As noted above, this contrasts with the situation in cyclin D2-deficient mice, wherein a dramatic decrease in the number of peritoneal CD5+ B cells has been reported (22). Thus, cyclin D2, but not cyclin D3 provides a nonredundant function in the development and/or self-renewal of peritoneal B-1a cells in the animal.

Our analysis of the potential for ex vivo peritoneal B-1a cells to proliferate in response to PMA indicated that cyclin D3-deficient B-1a cells proliferate at a normal tempo in comparison to wild-type B-1a cells. Although our results with the use of TAT-p16 peptides in normal B-1a cells demonstrate that cyclin D3 is important for mediating proliferative signals from several mitogens (e.g., PMA, LPS, and CD40L), the findings in cyclin D3-deficient mice suggest that cyclin D3 is dispensable for B-1a cell proliferation in ex vivo primary cultures stimulated with PMA or LPS and under certain conditions. Such conditions include cyclin D2 gene expression increased 4-fold above the level found in wild-type B-1a cells stimulated by PMA. Furthermore, the elevated induction of cyclin D2 mRNA in cyclin D3-deficient B-1a cells is associated with early and sustained expression of cyclin D2 protein throughout G0-S progression. This sustained expression of cyclin D2 contrasts with the early and short-lived induction of cyclin D2 in wild-type B-1a cells stimulated with PMA.

The possibility that cyclin D2 may functionally replace cyclin D3 is strengthened by the observation that the level of endogenous pRb phosphorylation on cdk4/6-targeted residues in cyclin D3-deficient B-1a cells is comparable to that of wild-type B-1a cells similarly stimulated with PMA for 21 h. Additional support for the notion that cyclin D2 can compensate for cyclin D3 loss in driving B-1a cell proliferation by PMA was obtained in that DNA synthesis was blocked by transduction of TAT-16 wild-type, but not TAT-p16 mutant, peptides into cyclin D3-deficient B-1a cells. Although we cannot entirely rule out the possibility of a compensatory role by a cyclin D-independent pathway in PMA-induced B-1a cell proliferation, it is noteworthy that no measurable changes in the timing and level of gene expression for the E-type cyclins were observed between wild-type and cyclin D3-deficient B-1a cells stimulated with PMA. Taken together, we interpret these findings as indicating that, although cyclin D3 normally fulfills a critical role in late G1 phase, in the absence of cyclin D3, PMA can drive B-1a cell proliferation via sustained expression of cyclin D2.

Currently, the molecular mechanism underlying the sustained expression of cyclin D2 in cyclin D3-deficient mice is unknown. This compensatory accumulation of cyclin D2 raises the possibility of a negative feedback loop, in which cyclin D3 limits the duration of cyclin D2 accumulation in normal B-1a cells. Genetic ablation of cyclin D3 would then be envisaged to relieve this restriction, thereby allowing for sustained accumulation of cyclin D2 throughout the G0-S interval. It remains unclear whether the apparent compensatory mechanism for cyclin D2 up-regulation identified here is present (but not activated) in wild-type B-1a cells, or only comes into play when cyclin D3 is completely absent. Additional experiments are underway to understand the molecular basis for the sustained accumulation of cyclin D2 in cyclin D3-deficient B-1a cells.


   Acknowledgments
 
We thank Dr. Peter Sicinski (Department of Cancer Biology, Dana-Farber Cancer Institute and Department of Pathology, Harvard Medical School, Boston, MA) for providing the cyclin D3-deficient mice. We also thank Blair Bleiman for studies of D-type cyclin expression in B-1a cells by flow cytometry.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   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 U.S. Public Health Service Grants AI-49994 (to T.C.C.) and AI-60896 (to T.L.R.). Back

2 Current address: Center for Oncology and Cell Biology, The Feinstein Institute for Medical Research, 350 Community Drive, Manhasset, NY 11030. Back

3 Address correspondence and reprint requests to Dr. Thomas C. Chiles, Department of Biology, Boston College, 414 Higgins Hall, Chestnut Hill, MA 02467. E-mail address: Chilest{at}bc.edu Back

4 Abbreviations used in this paper: cdk, cyclin-dependent kinase; MZ, marginal zone; AML1, acute myeloid leukemia 1. Back

Received for publication December 20, 2005. Accepted for publication April 19, 2006.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

  1. Matsushime, H., M. F. Roussel, R. A. Ashmun, C. J. Sherr. 1991. Colony-stimulating factor 1 regulates novel cyclins during the G1 phase of the cell cycle. Cell. 65: 701-713. [Medline]
  2. Kozar, K., P. Sicinski. 2005. Cell cycle progression without cyclin D-CDK4 and cyclin D-CDK6 complexes. Cell Cycle 4: 388-391. [Medline]
  3. Bates, S., L. Bonetta, D. Macallan, D. Parry, A. Holder, C. Dickson, G. Peters. 1994. CDK6 (PLSTIRE) and CDK4 (PSK-J3) are a distinct subset of the cyclin-dependent kinases that associate with cyclin D1. Oncogene 9: 71-79. [Medline]
  4. Kato, J. Y., M. Matsuoka, D. K. Strom, C. J. Sherr. 1994. Regulation of cyclin D-dependent kinase 4 (cdk4) by cdk4-activating kinase. Mol. Cell. Biol. 14: 2713-2721. [Abstract/Free Full Text]
  5. Meyerson, M., E. Harlow. 1994. Identification of G1 kinase activity for cdk6, a novel cyclin D partner. Mol. Cell. Biol. 14: 2077-2086. [Abstract/Free Full Text]
  6. Matsushime, H., M. E. Ewen, D. K. Strom, J.-Y. Kato, S. K. Hanks, M. F. Roussel, C. J. Sherr. 1992. Identification and properties of an atypical catalytic subunit (p34PSK-J3/Cdk4) for mammalian D-type G1 cyclins. Cell 71: 323-334. [Medline]
  7. Lundberg, A. S., R. A. Weinberg. 1998. Functional inactivation of the retinoblastoma protein requires sequential modification by at least two distinct cyclin-cdk complexes. Mol. Cell. Biol. 18: 753-761. [Abstract/Free Full Text]
  8. Harbour, J. W., R. Luo, A. Dei Santi, A. Postigo, D. Dean. 1999. Cdk phosphorylation triggers sequential intramolecular interactions that progressively block Rb functions as cells move through G1. Cell 98: 859-869. [Medline]
  9. Sherr, C. J., J. M. Roberts. 1999. CDK inhibitors: positive and negative regulators of G1-phase progression. Genes Dev. 13: 1501-1512. [Free Full Text]
  10. Ortega, S., M. Malumbres, M. Barbacid. 2002. Cyclin D-dependent kinases, INK4 inhibitors and cancer. Biochim. Biophys. Acta 1602: 73-87. [Medline]
  11. Serrano, M., G. J. Hannon, D. Beach. 1993. A new regulatory motif in cell cycle control causing specific inhibition of cyclin D/CDK4. Nature 366: 704-707. [Medline]
  12. Bruce, J. L., R. K. Hurford, Jr, M. Classon, J. Koh, N. Dyson. 2000. Requirements for cell cycle arrest by p16INK4a. Mol. Cell 6: 737-742. [Medline]
  13. Sherr, C. J., J. M. Roberts. 2004. Living with or without cyclins and cyclin-dependent kinases. Genes Dev. 18: 2699-2711. [Abstract/Free Full Text]
  14. Malumbres, M., R. Sotillo, D. Santamaria, J. Galan, A. Cerezo, S. Ortega, P. Dubus, M. Barbacid. 2004. Mammalian cells cycle without the D-type cyclin-dependent kinases Cdk4 and Cdk6. Cell 118: 493-504. [Medline]
  15. Kozar, K., M. A. Ciemerych, V. I. Rebel, H. Shigematsu, A. Zagozdzon, E. Sicinska, Y. Geng, Q. Yu, S. Bhattacharya, R. T. Bronson, et al 2004. Mouse development and cell proliferation in the absence of D-cyclins. Cell 118: 477-491. [Medline]
  16. Inaba, T., H. Matsushime, M. Valentine, M. F. Roussel, C. J. Sherr, A. T. Look. 1992. Genomic organization, chromosomal localization, and independent expression of human cyclin D genes. Genomics 13: 565-574. [Medline]
  17. Xiong, Y., J. Menninger, D. Beach, D. C. Ward. 1992. Molecular cloning and chromosomal mapping of CCND genes encoding human D-type cyclins. Genomics 13: 575-584. [Medline]
  18. Sicinski, P., J. L. Donaher, Y. Geng, S. B. Parker, H. Gardner, M. Y. Park, R. L. Robker, J. S. Richards, L. K. McGinnis, J. D. Biggers, et al 1996. Cyclin D2 is an FSH-responsive gene involved in gonadal cell proliferation and oncogenesis. Nature 384: 470-474.24. [Medline]
  19. Solvason, N., W. W. Wu, N. Kabra, X. Wu, E. Lees, M. C. Howard. 1996. Induction of cell cycle regulatory proteins in anti-immunoglobulin-stimulated mature B lymphocytes. J. Exp. Med. 184: 407-417. [Abstract/Free Full Text]
  20. Reid, S., E. C. Snow. 1996. The regulated expression of cell cycle-related proteins as B- lymphocytes enter and progress through the G1 cell cycle stage following delivery of complete versus partial activation stimuli. Mol. Immunol. 33: 1139-1151. [Medline]
  21. Mohamedali, A., I. Soeiro, N. C. Lea, J. Glassford, L. Banerji, G. J. Mufti, E. W.-F. Lam, S. B. Thomas. 2003. Cyclin D2 controls B cell progenitor numbers. J. Leukocyte Biol. 74: 1141-1145.
  22. Solvason, N., W. W. Wu, D. Parry, D. Mahony, E. W. Lam, J. Glassford, G. G. Klaus, P. Sicinski, R. Weinberg, Y. J. Liu, et al 2000. Cyclin D2 is essential for BCR-mediated proliferation and CD5 B cell development. Int. Immunol. 12: 631-638. [Abstract/Free Full Text]
  23. Lam, E. W., J. Glassford, L. Banerji, N. S. Thomas, P. Sicinski, G. G. Klaus. 2000. Cyclin D3 compensates for loss of cyclin D2 in mouse B-lymphocytes activated via the antigen receptor and CD40. J. Biol. Chem. 275: 3479-3484. [Abstract/Free Full Text]
  24. Hsia, C. Y., S. Cheng, A. M. Owyang, S. F. Dowdy, H. C. Liou. 2002. c-Rel regulation of the cell cycle in primary mouse B lymphocytes. Int. Immunol. 14: 905-916. [Abstract/Free Full Text]
  25. Wang, Z., P. Sicinski, R. A. Weinberg, Y. Zhang, K. Ravid. 1996. Characterization of the mouse cyclin D3 gene: exon/intron organization and promoter activity. Genomics 35: 156-163. [Medline]
  26. Berland, R., H. H. Wortis. 2002. Origins and functions of B-1 cells with notes on the role of CD5. Annu. Rev. Immunol. 20: 253-300. [Medline]
  27. Herzenberg, L. A.. 2000. B-1 cells: the lineage question revisited. Immunol. Rev. 175: 9-22. [Medline]
  28. Rothstein, T. L.. 2002. Cutting edge commentary: two B-1 or not to be one. J. Immunol. 168: 4257-4261. [Abstract/Free Full Text]
  29. Hardy, R. R., K. Hayakawa. 2001. B cell development pathways. Annu. Rev. Immunol. 19: 595-621. [Medline]
  30. Herzenberg, L. A., A. M. Stall, P. A. Lalor, C. Sidman, W. A. Moore, D. R. Parks, L. A. Herzenberg. 1986. The Ly-1 B cell lineage. Immunol. Rev. 93: 81-102. [Medline]
  31. Caligaris-Cappio, F., M. Gobbi, M. Bofill, G. Janossy. 1982. Infrequent normal B lymphocytes express features of B-chronic lymphocytic leukemia. J. Exp. Med. 155: 623-628. [Abstract/Free Full Text]
  32. Stall, A. M., M. C. Farinas, D. M. Tarlinton, P. A. Lalor, L. A. Herzenberg, S. Strober. 1988. Ly-1 B-cell clones similar to human chronic lymphocytic leukemias routinely develop in older normal mice and young autoimmune (New Zealand Black-related) animals. Proc. Natl. Acad. Sci. USA 85: 7312-7316. [Abstract/Free Full Text]
  33. Marti, G. E., R. A. Metcalf, E. Raveche. 1995. The natural history of a lymphoproliferative disorder in aged NZB mice. Curr. Top. Microbiol. Immunol. 194: 117-126. [Medline]
  34. Hayakawa, K., R. R. Hardy, A. M. Stall, L. A. Herzenberg. 1986. Immunoglobulin bearing B cells reconstitute and maintain the murine Ly- 1 B cell lineage. Eur. J. Immunol. 16: 1313-1316. [Medline]
  35. Rothstein, T. L., D. L. Kolber. 1988. Anti-Ig antibody inhibits the phorbol ester-induced stimulation of peritoneal B cells. J. Immunol. 141: 4089-4093. [Abstract]
  36. Morris, D. L., T. L. Rothstein. 1993. Abnormal transcription factor induction through the surface immunoglobulin M receptor of B-1 lymphocytes. J. Exp. Med. 177: 857-861. [Abstract/Free Full Text]
  37. Rothstein, T. L., D. L. Kolber. 1988. Peritoneal B cells respond to phorbol esters in the absence of a co-mitogen. J. Immunol. 140: 2880-2885. [Abstract]
  38. Tanguay, D. A., T. P. Colarusso, S. Pavlovic, M. Irigoyen, R. G. Howard, J. Bartek, T. C. Chiles, T. L. Rothstein. 1999. Early induction of cyclin D2 expression in phorbol ester-responsive B-1a lymphocytes. J. Exp. Med. 189: 1685-1690. [Abstract/Free Full Text]
  39. Tanguay, D. A., T. P. Colarusso, C. Doughty, S. Pavlovic, T. L. Rothstein, T. C. Chiles. 2001. Differences in the signaling requirements for activation of assembled cyclin D3-cdk4 complexes in B-1a and B-2 lymphocytes. J. Immunol. 166: 4273-4277. [Abstract/Free Full Text]
  40. Tanguay, D. A., T. C. Chiles. 1996. Regulation of the catalytic subunit (p34PSK-J3/cdk4) for the major D- type cyclin in mature B lymphocytes. J. Immunol. 156: 539-548. [Abstract]
  41. Sicinska, E., I. Aifantis, L. Le Cam, W. Swat, C. Borowski, Q. Yu, A. A. Ferrando, S. D. Levin, Y. Geng, H. von Boehmer, P. Sicinki. 2003. Requirement for cyclin D3 in lymphocyte development and T cell leukemias. Cancer Cell 4: 451-461. [Medline]
  42. Tumang, J. R., W. D. Hastings, C. Bai, T. L. Rothstein. 2004. Peritoneal and splenic B-1a cells are separable by phenotypic, functional, and transcriptomic characteristics. Eur. J. Immunol. 34: 2158-2167. [Medline]
  43. Fahraeus, R., J. M. Paramio, K. L. Ball, S. Lain, D. P. Lane. 1996. Inhibition of pRb phosphorylation and cell cycle progression by a 20-resdiue peptide derived from p16CDKN2/INK4A. Curr. Biol. 6: 84-91. [Medline]
  44. Gius, D. R., S. A. Ezhevsky, M. Becker-Hapak, H. Nagahara, M. C. Wei, S. F. Dowdy. 1999. Transduced p16INK4a peptides inhibit hypophosphorylation of the retinoblastoma protein and cell cycle progression prior to activation of Cdk2 complexes in late G1. Cancer Res. 59: 2577-2580. [Abstract/Free Full Text]
  45. Tumang, J. R., R. Frances, S. G. Yeo, T. L. Rothstein. 2005. Cutting edge: spontaneously Ig-secreting B-1 cells violate the accepted paradigm for expression of differentiation-associated transcription factors. J. Immunol. 174: 3173-3177. [Abstract/Free Full Text]
  46. Lane, P., T. Brocker, S. Hubele, E. Padovan, A. Lanzavecchia, F. McConnell. 1993. Soluble CD40 ligand can replace the normal T cell-derived CD40 ligand signal to B cells in T cell-dependent activation. J. Exp. Med. 177: 1209-1213. [Abstract/Free Full Text]
  47. Francis, D. A., J. G. Karras, X. Y. Ke, R. Sen, T. L. Rothstein. 1995. Induction of the transcription factors NF-{kappa}B, AP-1 and NF-AT during B cell stimulation through the CD40 receptor. Int. Immunol. 7: 151-161. [Abstract/Free Full Text]
  48. Kushnuer, J. A., M. A. Ciemerych, E. Sicinska, L. M. Wartschow, M. Teta, S. Y. Long, P. Sicinski, M. F. White. 2005. Cyclins D2 and D1 are essential for postnatal pancreatic beta-cell growth. Mol. Cell. Biol. 25: 3752-3762. [Abstract/Free Full Text]
  49. Peterson, L. F., A. Boyapati, V. Ranganathan, A. Iwama, D. G. Tenen, S. Tsai, D.-E. Zhang. 2005. The hematopoietic transcription factor AML1 (RUNX1) is negatively regulated by the cell cycle protein cyclin D3. Mol. Cell. Biol. 25: 10205-10219. [Abstract/Free Full Text]
  50. Sun, M., Y. Wei, L. Yao, J. Xie, X. Chen, H. Wang, J. Jang, J. Gu. 2006. Identification of extracellular signal-regulated kinase 3 as a new interaction partner of cyclin D3. Biochem. Biophys. Res. Commun. 340: 209-214. [Medline]
  51. Shen, X., Y. Yang, W. Liu, M. Sun, J. Jaing, H. Zong, J. Gu. 2004. Identification of the p28 subunit of eukaryotic initiation factor 3(eIF3k) as a new interaction partner of cyclin D3. FEBS Lett. 573: 139-146. [Medline]



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