(Received for publication, November 21, 1996, and in revised form, January 23, 1997)
From the Department of Tumor Cell Biology, St. Jude Children's Research Hospital, Memphis, Tennessee 38105
DT40 lymphoma B-cells normally express cyclins D1
and D2 but not D3. When cyclin D1 expression was extinguished in these
cells by gene knockout, specific alterations in their ability to
transit the cell cycle were observed. These changes are exemplified by a delay of approximately 2 h in their progression through a normal 14-h cell cycle. This delay results in an increase in the number of
cells in the G2/M phase population, most likely due
to triggering of checkpoints in G2/M, inability to enter
G1 normally, and/or alterations of crucial event(s) in
early G1. The defect(s) in the cell cycle of these D1
"knockout" cells can be rescued by overexpression of any normal
mouse D-type cyclin but not by a mutant mouse cyclin D1 protein that
lacks the LXCXE motif at its amino terminus.
These data suggest that the cell cycle alterations observed in the
D1/
cells are a direct effect of the absence of the cyclin D1
protein and support the hypothesis that the D-type cyclins have
separate, but overlapping, functions. Elimination of cyclin D1 also
resulted in enhanced sensitivity to radiation, resulting in a
significant increase in apoptotic cells. Expression of any normal
murine D-type cyclin in the D1
/
cells reversed this phenotype.
Intriguingly, expression of the mutant cyclin D1 in the D1
/
cells
partially restored resistance to radiation-induced apoptosis. Thus,
there may be distinct differences in cyclin D1 complexes and/or its
target(s) in proliferating and apoptotic DT40 lymphoma B-cells.
Regulation of the vertebrate cell cycle requires the periodic formation, activation, and inactivation of unique protein kinase complexes that consist of cyclin (regulatory) and cyclin-dependent kinase (CDK1; catalytic) subunits. Cell cycle-dependent fluctuations in the levels of many of the cyclin proteins are thought to contribute, at least in part, to the activation of these enzymes (1-4). The cyclins are required to regulate many of the p34cdc2-related cyclin-dependent protein kinases (CDKs), which have diverse functions prominently linked to the control of cell division (5-16). For example, cyclin B participates in the regulation of the G2/M transition by its association with p34cdc2, whereas cyclin A appears to be essential for the completion of S phase and entry into G2 phase in complexes with both p34cdc2 and cdk2 (17-27). In contrast, complexes formed between the D-type cyclins and both cdk4 and cdk6 integrate growth factor signals and the cell cycle, allowing cells to progress normally through G1 phase (10, 12, 28-34). The genetic alterations that occur in this pathway during oncogenesis appear to involve many of its components, including the D-type cyclins, cyclin-dependent protein kinases (cdks), and cyclin-dependent kinase inhibitors (CKIs).
Unlike other G1 cyclins described thus far (e.g. cyclin C and cyclin E), the D-type cyclins are highly homologous to one another and their expression overlaps during G1 phase, suggesting that this group of cyclins may be somewhat redundant in their function. However, it has also been noted that not all D-type cyclins are expressed in each tissue, suggesting that their function may be linked to the specific tissues in which they are expressed. This particular cell cycle pathway is subject to a number of alterations during tumorigenesis, presumably due to its importance in response to mitogenic stimulation (31, 35-43).
The biologic role of multiple D-type cyclins has not yet been resolved.
It has been established that each of the D-type cyclins contain a
functional pRb-binding motif and that these cyclins are induced in
response to mitogens in a cell lineage-specific manner (3, 4). However,
cyclins D2 and D3 bind preferentially to pRb, whereas cyclin D1·pRb
complexes are less stable (44, 45). Microinjection of cyclin D1
antibodies and/or antisense oligonucleotides into normal human diploid
fibroblasts, NIH3T3, and Rat-2 cells revealed that a sub-group of these
cells could not enter S phase (46, 47). The ability of these
microinjected cells to pass the restriction point and begin DNA
replication was directly related to the time that had elapsed between
the readdition of serum and microinjection of cyclin D1 antibodies or
antisense oligonucleotides. Thus, by decreasing the level of functional
cyclin D1 protein prior to the restriction point in late G1
phase, the ability of these cells to progress through the cell cycle
was impaired. These agents have no apparent effect on cell cycle once
cells have passed the restriction point late in G1 phase of
the cell cycle. Finally, it has been shown that ectopic expression of
cyclin D1 inhibits MyoD and myogenin-mediated skeletal muscle
differentiation indirectly (48). To determine the role of cyclin D1 in
muscle differentiation, experiments were designed to determine whether
MyoD·myogenin and/or pRb were targets of cyclin D1·CDK
phosphorylation. Mutation of CDK phosphorylation sites in myogenin had
no effect on the ability of cyclin D1 to inhibit differentiation.
Ectopic MyoD expression in fibroblasts can induce muscle-specific gene
expression, as long as wild-type pRb is also expressed (48). In pRb
/
fibroblasts expressing MyoD and a mutated pRb (which cannot be
hyperphosphorylated), ectopic cyclin D1 expression continues to inhibit
muscle-specific gene expression, whereas ectopically expressed cyclins
A and E have no effect. These data suggest that the initiation of
muscle-specific gene expression can be blocked by two distinct
pathways, one of which is dependent on pRb hyperphosphorylation and
the other is not.
Several models of D-type cyclin function have been proposed based on the available data, but they all envision these cyclins as growth sensors (3, 4, 45, 46, 49, 50). The first model suggests that the D-type cyclins are tissue-specific regulatory subunits that are merely redundant in their function. A second model suggests that the functions of D-type cyclins are related to their sequential expression pattern during the cell cycle. More recent studies with cyclin D1 gene "knockouts" in mice have supported the former possibility (49, 50). Specifically, proper development of the retina and breast ductal epithelium was perturbed, suggesting that their normal maturation depends upon the presence of cyclin D1. This mouse model has provided significant insight into the requirements of cyclin D1 for normal development, but a specific requirement for this cyclin during cell cycle progression per se, has not been established.
We now report the successful utilization of the chicken lymphoma B-cell
line, DT40, to eliminate the expression of specific G1
cyclins. This particular cell line undergoes gene-specific recombination at frequencies that are ~100-fold higher than those in
mouse embryonic stem cells (51). DT40 cells provide a unique reagent to
easily and quickly examine the necessity for each of these cyclins
individually and collectively, as well as interactions between the
various cyclins and other cell cycle proteins. These lymphoma B-cells
normally express cyclin D1 and D2 but not cyclin D3. Elimination of
cyclin D1 in DT40 cells is not lethal; however, ablation of cyclin D1
gene expression results in marked alterations in the ability of these
cells to transit the cell cycle which, in turn, affects their normal
growth rate. Furthermore, loss of cyclin D1 did not result in a
compensatory increase in the levels of cyclin D2 in these cells.
Overexpression of normal mouse cyclin D1, D2, or D3 in the D1 /
cells restored normal growth and cell cycle progression. However, a
mutant murine cyclin D1 lacking the LXCXE motif
(the so-called retinoblastoma protein (pRb)-binding motif that enhances
D-type cyclin interaction with pRb) did not rescue the D1
/
phenotype (44, 45).
Here we show that elimination of cyclin D1 leads to a specific cell
cycle phenotype and that these cells can be rescued by expression of
any one of the three murine D-type cyclins, but not by a murine cyclin
D1 mutant missing the LXCXE motif. The rescue of
the D1 /
DT40 cells by murine cyclin D3 is somewhat intriguing,
since avians do not express a cyclin D3 homologue in any cell line or
tissue examined thus far (52, 53, and this study). Elimination of
cyclin D1 in DT40 cells also resulted in a significant increase in
apoptotic death following either UV or
irradiation. Finally,
expression of either murine cyclin D1, D2, or D3 in the DT40 D1
/
cells resulted in decreased sensitivity to radiation-induced apoptosis,
indistinguishable to what was observed in the parental DT40 cell line.
Expression of the mutant cyclin D1 partially rescued cells from
radiation-induced apoptosis, suggesting that the mechanism(s) involving
cyclin D1 in this pathway is distinct from its role in normal cell
cycle progression. Thus, in avian B-lymphoma cells cyclin D1 appears to
fully suppress apoptosis, whereas a mutant cyclin D1 partially rescued
these cells, suggesting that the targets of cyclin D1·CDK activity in apoptotic cells may be distinct from those in proliferating cells.
Agarose was purchased from FMC Bioproducts
(Rockland, ME). Restriction and modification enzymes were obtained from
New England Biolabs (Beverly, MA) and Boehringer Mannheim. Urea and
cell culture reagents, including chicken serum, were purchased from
Life Technologies, Inc. Sequenase and DNA sequencing reagents were
purchased from U. S. Biochemical Corp. All radioisotopes were purchased
from DuPont NEN. All chemicals were purchased from either Sigma or Fisher. The chicken cosmid genomic library was obtained from Clonetech (Palo Alto, CA). The PrimeIt labeling kit and Duralose membranes were
purchased from Stratagene (La Jolla, CA). Selection drugs were
purchased from Life Technologies, Inc., Sigma, or Boehringer Mannheim.
Fluorescein-conjugated goat anti-mouse and goat anti-rabbit antibodies
were obtained from Boehringer Mannheim or Southern Biotechnology.
Rabbit polyclonal anti-cyclin D1 antibody was kindly provided by Dr.
Charles Sherr (see Ref. 2), and anti-BudR antibodies were obtained from
Boehringer Mannheim. The DNA-specific dye 4,6-diamidino-2-phenylindole was obtained from Boehringer Mannheim. Oligonucleotides for DNA sequencing were synthesized on an Applied Biosystems 394 DNA/RNA Synthesizer by the Molecular Resources Facility of St. Jude Children's Research Hospital.
Cyclin D1 cDNAs were isolated from a UG9 T-cell cDNA
library (55) by low stringency hybridization with a human cyclin D1 cDNA, kindly provided by Dr. Steve Reed (56). The nucleotide sequence of these cDNAs was determined as described previously (57). Oligonucleotides were spaced approximately 80-100 base pairs
apart spanning the entire cDNA. A full-length cyclin D1 cDNA
was transcribed and translated in vitro, producing a 36-kDa protein that could be immunoprecipitated with the mouse polyclonal cyclin D1 antibody (see Fig. 4, panel A). The corresponding
cyclin D1 (Gallus gallus ccnd1) gene was isolated by
screening a Cornish White Rock chicken cosmid library (Stratagene) with
the full-length cyclin D1 cDNA as described previously (58). A
13-kb BamHI restriction fragment containing exons 1-3 of
the cyclin D1 gene was subcloned into the pKS plasmid and either this
DNA, or cosmid DNA, was used for double-strand DNA sequence analysis of
the gene. The same oligonucleotides used for establishing the cDNA
sequence were used to establish the molecular organization (intron/exon
boundaries, exons) of the gene by sequencing the DNA strands in both
directions. Genomic DNA was isolated as described previously (59).
Gene Constructs
The constructs containing the inserted drug
selection cassettes were made by digestion of the 13-kb
BamHI subclone that contained exons 1-3 of the
Ggccnd1 gene, with the enzyme BstI that opened this fragment at a single site in exon 2 (Fig. 2, panel A).
This exon encodes a major portion of the cyclin box region.
Approximately 5.5 kb of genomic DNA were left intact on the 5 side of
this insertion, and approximately 7.5 kb of genomic DNA remained on the
3
side. Restriction fragments containing either the neo or hyg gene under the control of a
-actin gene promoter
(kindly provided by Dr. C. Thompson) were cloned into this
BstI site by blunt-end ligation using Klenow enzyme. Further
details concerning the selectable marker genes can be found elsewhere
(60, 61). The integrity of the constructs was confirmed by DNA
sequencing, demonstrating that exon 2 was disrupted and that the
remainder of the gene was intact. Homologous recombination of these
constructs into the Gccnd1 locus of the DT40 cells was
demonstrated by the increase in size of a 15-kb XbaI
fragment that contains exons 1 and 2 of the gene but that also contains
a unique 5
region (5
-flanking probe) not found in the 13-kb
BamHI subclone used for generating the disruption constructs
(Fig. 2, panel A). This probe could then be used to analyze
genomic DNA from single cell clones after XbaI digestion.
Insertion of the neo cassette (2 kb in size) generated a
17-kb XbaI hybridizing fragment (Fig. 2, panels C
and D), and insertion of the hyg gene cassette (3 kb in size) generated an 18-kb XbaI hybridizing fragment
(Fig. 2, panels C and D).
Targeted Disruption and Analysis of Cyclin D1 Expression
DT40 lymphoma B cells (kindly provided by Dr. C-L.
Chen) were grown in suspension cell culture in Dulbecco's modified
Eagle's medium supplemented with 10% fetal calf serum, 1% chicken
serum, penicillin, streptomycin, and glutamine. pCycD1-neo
and pCycD1-hyg were linearized and transfected into DT40
cells by electroporation (550 V, 25 microfarads). Twenty-four hours
after DNA transfection, the appropriate concentration of selection drug
(2 mg/ml G418 or 1.5 mg/ml hygromycin) was added to the culture medium,
and the cells were selected for ~14 days. Single cells, isolated by flow sorting, were expanded into individual clones. Genomic DNA was
isolated from multiple single cell-derived clones, digested with the
appropriate restriction enzymes and hybridized with the cyclin D1
5-flanking genomic probe (Fig. 2, panel A) to screen for
homologous recombinants. Disruption of one, heterozygote (+/
), or
both, homozygote (
/
), cyclin D1 (Ggccnd1) alleles was
observed. For all experiments, multiple single cell clones of
/+ and
/
cyclin D1 disruptions were examined by both Southern and Northern blotting. RNA was isolated for Northern blotting, and the blots were
hybridized to the cyclin D1 cDNA probe as described previously (57,
62). Equal loading of RNA samples was verified by reprobing these blots
with a human
-actin cDNA probe. The probes were prepared using a
random labeling kit (Stratagene) as specified by the manufacturer. The
blots were visualized by using a Molecular Dynamics 400A
PhosphorImager. Exposure times were as follows: 12 h for the
cyclin D1 cDNA, 2 h for the cyclin B2 cDNA, and 20 min for
the
-actin cDNA. Cyclin D1 protein expression was examined by
immunoprecipitation of [35S]methionine-labeled cell
lysates using either mouse polyclonal cyclin D1, D2, or D3 antiserum as
described previously by others (28). As controls for these experiments,
the mouse preimmune sera were used in all experiments, and an
appropriate unlabeled cyclin D-GST fusion protein was used as a
competitor to demonstrate specificity of the various D-type cyclin
immunoprecipitations in DT40 cells (Fig. 4, panel A).
Cells were selected in 2 mg/ml G418 or
1.5 mg/ml hygromycin and then maintained in cell culture under
continuous drug selection at the same concentrations. For the growth
curves, 2 × 105 cells were plated in triplicate and
grown in the presence of 10% serum. Each point represents the average
of three determinations ± the standard error of the mean. Growth
curves were obtained for each of the cell lines for 7 days at 1-day
intervals. DT40 cells were synchronized by first using centrifugal
elutriation to enrich the G1 phase population of cells.
These cells were then blocked at the G1/S boundary using 25 mg/ml aphidicolin for 10 h. These synchronized cells were removed
from the drug and then placed into culture media containing BudR,
harvested at 2-h intervals, and replicating (BudR +) cells stained and
counted. Cytospin slides were prepared and stained with an anti-BudR
antibody and a fluorescein labeled secondary antibody according to the
protocol from the manufacturer (Boehringer Mannheim). The cells were
also stained with 4,6-diamidino-2-phenylindole, and the number of
metaphase cells was determined as well. For the BudR incorporation and
mitotic index experiments, two separate clonal D1
/
cell lines were examined (clones 1 and 3). For cell cycle analysis, asynchronously growing cells were stained with propidium iodide, and their DNA content
was determined using the fluorescence activated cell sorter. This
analysis was performed as described previously (62) using asynchronously growing cultures by analyzing approximately 50,000 cells
from polyclonal, or single cell, populations for each sample. The
percentage of cells in each phase of the cell cycle was determined from
the histograms using the MODFIT cell cycle analysis program.
Rescue of the cyclin D1 /
DT40 cells was
accomplished by transfecting a clonal D1
/
cell line (clone 1) with
either a RSV-mouse CycD1 cDNA, RSV-mouse CycD2, RSV-mouse CycD3, or
the RSV-mutant mouse CycD1 cDNA, which have been reported
previously (28, 30, 47). These cDNAs were co-transfected with a
puromycin (pur) selectable drug resistance marker to allow
selection of stable integrants. Since all clonal D1
/
DT40 cell
lines had similar, if not identical, cell cycle alterations, only one
was selected for transfection of the two mouse cDNAs (clone 1).
However, multiple single cell clones or pooled clone samples were
subsequently isolated from each of these transfections and selections
(clones 1/56 and 1/57 for normal mouse cyclin D1, a pool of positive
clones for normal mouse cyclin D2, normal mouse cyclin D3, and for the mutant mouse cyclin D1) for further analysis. Two of the original cyclin D1
/
cell clones, as well as two clones from the same D1
/
clone into which the murine D1 cDNA expression construct was
introduced, were analyzed for cyclin D1 protein expression (Fig. 4,
panel C). Similarly, clonal cell lines of the same D1
/
cells containing either the murine cyclin D2 or D3 expression construct
were analyzed for their expression of these respective proteins (Fig.
4, panel D). Two of the original DT40 D1
/
clonal cell
lines, as well as one expressing either normal murine cyclin D1, D2, or
D3 protein, were analyzed further by flow cytometric analysis. The
mutant mouse cyclin D1 protein migrates as a smaller molecular weight
protein on SDS-polyacrylamide gel electrophoresis due to the deletion
of its amino terminus, which includes the pRb-binding motif
LXCXE (Fig. 4, panel C (30)).
Normal and D1 /
cells
were exposed to varying levels of UV or
irradiation to assess
whether there are differences in the level of programmed cell death
between these different cell lines. Others have previously shown (63,
64) that DT40 cells are a radiation-sensitive lymphoma B cell line that
requires PLC-
2 activation, via the combined activities of the
Bruton's tyrosine kinase and the SYK protein kinase. Apoptosis was
induced by exposing the DT40, D1
/
, D1
/
[+ murine D1],
D1
/
[+ murine D2], or D1
/
[+ murine D3] cell lines to
increasing amounts of either UV or
irradiation (63-65). Three
different single cell clones were used for these analyses. For the UV
radiation studies, DT40 cells were removed from culture (growing at a
density of ~106/ml), plated on polylysine-coated
coverslips (25 µg/ml), and allowed to attach for 30 min to 1 h,
and washed twice with phosphate-buffered saline. These cells were then
irradiated with varying amounts of either UV (UVC; 500, 1000, 1500, and
2000 J/m2). For
irradiation (4-Gy or 8-Gy
-rays),
the cells were treated in culture at a density of 5 × 105/ml, as described by others (65, 66). Both the UV and
irradiated cells were returned to the incubator following addition
of fresh media. Cells were then harvested 4-8 h following exposure to
radiation, and the percentage of apoptotic cells was determined by
TUNEL analysis and/or Hoescht staining of nuclei.
The cyclin D1 cDNA was isolated by screening a UG-9
T-cell cDNA library using the human cyclin D1 cDNA as a probe
(56). This particular T-cell line expresses cyclin D1 and cyclin D2 but
not cyclin D3.2 Similarly, we found that
the DT40 B-cell line expressed cyclins D1 and D2 but not D3 (data not
shown; Fig. 3). Mullner and colleagues (53) have suggested that
chickens may lack the cyclin D3 gene and the absence of an avian cyclin
D3 homologue may indicate an evolutionary divergence in the tissue
specificity of D-type cyclin expression. Our results appear to support
this hypothesis. Ten positive cyclin D1 cDNA clones were isolated
and purified. Six of these cDNAs had inserts ~1.3 kb and were
used for subsequent determination of the nucleotide sequence and
predicted open reading frame (Fig. 1). Comparison of the
avian, human, and murine cyclin D1 open reading frames demonstrates a
high level of sequence conservation (86% identity with human and
mouse) between the proteins, particularly in the conserved "cyclin
box" region (1). The corresponding gene was isolated from a chicken
cosmid library, and its structure was determined in its entirety by
restriction endonuclease and DNA sequence analysis (Fig.
2, panel A). The size, ~10 kb, and organization of the avian cyclin D1 gene (Ggccnd1) is
identical to the human (CCND1) and mouse (ccnd1)
homologues (67, 68).
A major portion of the Ggccnd1 gene was subcloned for
further analysis and construction of insertion mutants to eliminate expression of this gene in DT40 cells. A 13-kb BamHI
fragment containing exons 1-3 of the Ggccnd1 gene was used
for generating disruption constructs, and a single copy ~3-kb
BamHI fragment located 5 of the larger BamHI
fragment, but outside the region used to disrupt this allele by
homologous recombination, was used for genomic DNA analysis (Fig. 2,
panel A). We took advantage of a unique BstI
restriction site found only in exon 2 of the cyclin D1 gene to generate
insertion mutants, containing either neomycin (neo) or
hygromycin (hyg) selectable marker genes that inactivate the
normal gene (Fig. 2, panel B). Insertion of either of these
selectable marker gene cassettes into the BstI restriction site would alter the normal mobility of a 15-kb XbaI genomic
fragment that is detected by the 5
-flanking probe (Fig. 2, panel
A). Introduction of the neo gene cassette by homologous
recombination into this region of the cyclin D1 gene generates a 17-kb
XbaI fragment when hybridized with the 5
-flanking probe,
while the insertion of the hyg gene cassette generates an
18-kb XbaI fragment using the same probe (Fig. 2,
panel C). As shown in Fig. 2, panel D, the wild-type DT40 cells (+/+) contain two copies of the normal 15-kb XbaI hybridizing fragment, whereas cells containing one
normal allele and one allele disrupted by the introduction of the
neo gene (+/
, heterozygote DT40 cells) into the cyclin D1
gene contain 15- and 17-kb XbaI hybridizing fragments, as
expected. In DT40 cells containing both the neo and
hyg gene cassettes homologously recombined into the cyclin
D1 locus (
/
, homozygotes), the wild-type 15-kb XbaI
hybridizing fragment is completely absent, and it has been replaced by
the 17- and 18-kb XbaI fragments, respectively (Fig. 2,
panel D). Additional proof of cyclin D1 elimination is provided by virtue of Northern blotting (Fig. 3) and
protein analysis (Fig. 4).
Complete elimination of the 15-kb
XbaI restriction fragment containing the normal
Ggccnd1 gene was easily obtained in a number of single cell
DT40 /
clones (Fig. 2, panel D, lanes 4-9). When expression of the cognate cyclin D1 mRNA (~1.3 kb in size) was examined by Northern blotting using these same single cell clones, as
well as a number of additional clones (15 total), expression of the
mRNA was undetectable in
/
homozygotes and barely detectable in
+/
heterozygotes (Fig. 3). However, a cross-hybridizing cyclin D2
mRNA (~6 kb) was readily detected in all cell lines (54, 58).
Consistent with the results of others, we were not able to detect an
avian cyclin D3 mRNA species, or a corresponding gene, in DT40
cells (data not shown) (52, 53). As with human and mouse, there is a
relatively high degree of sequence homology between avian cyclin D1 and
cyclin D2 (Fig. 1) (58). Concomitantly, expression of the normal cyclin
D1 protein was also extinguished when examined by immunoprecipitation
(IP) of metabolically labeled cell lysates (Fig. 4). In a control
experiment, the mouse polyclonal cyclin D1 antibody was shown to IP the
chicken in vitro transcribed and translated (IVTT) protein,
as well as the [35S]methionine-labeled cyclin D1 protein
from the DT40 cells (Fig. 4, panel A). This ability to IP
the cyclin D1 protein from DT40 cells was directly competed by
preincubation with cold competitor mouse cyclin D1 protein (Fig. 4,
panel A), demonstrating specificity. A comparison of the
preimmune serum control with the mouse polyclonal D1 antisera using the
same [35S]methioninie-labeled +/+ wild-type DT40 cell
lysates is also shown (Fig. 4, panel B, DT40). Analysis of
two of the
/
cyclin D1 single cell clones shown above (clones 1 and
2) by metabolic labeling followed by cyclin D1 IP, demonstrated that
neither of these cell lines expressed this protein (Fig. 4, panel
B). However, expression of either the wild-type or mutant murine
cyclin D1 cDNAs in these same
/
cells (the mutant protein is
truncated and no longer contains the pRb-binding motif
LXCXE (44, 45)) can easily be detected using this
strategy (Fig. 4, panel C, normal murine cyclin D1, clones
1/56 and 1/57; mutant murine cyclin D1, clones 1/2 and 1/17).
Additional experiments involving the expression of either the murine
cyclin D2 or D3 cDNA, and its corresponding protein, in the same D1
/
DT40 cells yielded similar results (Fig. 4, panel D).
The murine polyclonal antisera for cyclin D2 was capable of
immunoprecipitating both the IVTT-labeled avian cyclin D2 and the
metabolically labeled murine cyclin D2 overexpressed in the D1
/
cells (Fig. 4, panel D). When excess cold murine cyclin D2-GST fusion protein was added to the cyclin D2 antibody, the immunoprecipitation of the [35S]methionine-labeled IVTT
protein was effectively competed (data not shown). Since an avian
cyclin D3 homologue has not been isolated, a cyclin D3 IVTT control was
not available for the corresponding immunoprecipitation of the
metabolically labeled murine cyclin D3 protein expressed in the D1
/
cells (Fig. 4, panel D). Steady-state levels of the
various murine cyclins (D1, D2, and D3) were comparable in single cell
clones, as judged by [35S]methionine labeling (Fig. 4 and
data not shown). These metabolic labeling and immunoprecipitation
experiments demonstrate that the cyclin D1 protein is absent in the D1
/
DT40 cells, as well as confirm the expression of the normal
murine cyclin D1, D2, D3, and mutant cyclin D1 in the same D1
/
DT40 cells.
Cyclin D1 elimination
is not lethal, presumably because cyclin D2 continues to be expressed
in these cells. This is consistent with results reported in homozygous
cyclin D1 /
mice, which are also viable (49, 50). However,
alterations in the cellular growth characteristics (Fig.
5) and normal progression through the cell cycle (Table
I) were noted by flow cytometric analysis of
asynchronously growing cells. The cell cycle phenotype of these cyclin
D1
/
cells, as might be expected, is distinct from that reported
for modest overexpression of cyclin D1 in fibroblasts (47, 69, 70).
Specifically, the total length of the cell cycle in these DT40 D1
/
cells is increased by 2-3 h in two separate clones (1 and 3),
increasing the cell doubling time from 14 to 16-17 h (Table
II and Fig. 5). Additionally, the percentage of
asynchronously growing cells in G2/M phase is increased,
and the percentage of cells in S phase is concomitantly decreased (Table I). In one of the mouse cyclin D1 knockout studies, fibroblasts derived from the D1
/
mice, but not normal controls, were also delayed in G2/M phase when quiescent D1
/
cells were
restimulated by the addition of serum growth factors (50). Even though
the effects of cyclin D1 elimination can be easily seen in
asynchronously growing cell populations, they are more dramatic using
synchronized cyclin D1
/
DT40 cells (Table II; cells were
synchronized at the G1/S boundary as described under
"Experimental Procedures"). Once again, two separate cyclin D1
/
clones (1 and 3) were used for this analysis. Progression through
the cell cycle in these various cell populations was analyzed by both
incorporation of BudR into cellular DNA and determination of their
mitotic index (MI). BudR labeling experiments demonstrated that both of
the
/
cell lines did not incorporate the labeled nucleotide into their DNA as quickly as the wild-type DT40 cells (e.g. 42.6 or 44.2% incorporation at 6 h post-BudR in the D1
/
clones as
compared with 76.3% incorporation at 6 h in wild-type DT40 cells;
Table II). Similarly, when the MI for each cell line was monitored, the
wild-type DT40 cells reached a peak in their MI at 10 h
post-G1/S release, whereas both
/
clones had a much
broader peak in their MI (spanning ~6 h), which peaked at 12 h
post-G1/S release (Table II). These results provide two
distinct parameters of cell cycle progression demonstrating that the
growth of the D1
/
DT40 cells was delayed by ~2-3 h.
|
|
Having generated viable clonal cell lines
lacking normal cyclin D1, but which continue to express normal levels
of cyclin D2 (confirmed by IP of endogenous cyclin D2 from
metabolically labeled wild-type and D1 /
DT40 cells; data not
shown), we reasoned that the observed changes in cell cycle progression
could be reversed by reinstating cyclin D1 expression in these cells.
We also reasoned that if D-type cyclin function were redundant,
overexpression of all three D-type cyclins might rescue the observed
phenotype of the D1
/
DT40 cells. Therefore, we introduced either
normal murine cyclin D1, D2, and D3 cDNAs or a D1 mutant cDNA
(lacking the LXCXE motif (44, 45)) into the D1
/
DT40 cells (clone 1). Expression of these cDNAs was driven by
an RSV promoter, as described previously (25). These investigators
demonstrated that this particular promoter generated the highest level
of "tolerable" cyclin D1 expression in murine fibroblasts and that
exogenously expressed cyclin D1 could shorten the duration of
G1 phase specifically. Expression of murine cyclin D1, D2,
D3, or the mutant D1 was confirmed by analysis of protein expression in
single cell clones (Fig. 4, panels C and D).
Normal cell cycle synchrony was restored in the D1
/
cells
(i.e. no apparent block in G2/M) by all three D-type cyclins but not by expression of the mutant mouse cyclin D1
cDNA (in which the amino-terminal LXCXE
pRb-binding sequence has been deleted (45) (Table I)). This mutant
cyclin D1 cDNA construct also failed to alter normal cell cycle
progression when it was overexpressed in rodent fibroblasts (47). Of
particular significance is the fact that expression of any one of the
murine D-type cyclin proteins was capable of rescuing cells from the apparent G2/M delay and not merely shortening the time
spent in G1 phase (Table I). Expression of the three murine
D-type cyclin proteins was comparable in all of the single cell clones
isolated and examined (Fig. 4, panels C and D).
This suggests that the characteristics of the cyclin D1
/
phenotype
are due to absence of cyclin D1 specifically but that this phenotype
can be rescued by expression of any of the normal murine D-type cyclin
proteins. It should be noted that cyclins D2 and D3 interact with pRb
more efficiently than cyclin D1 (10, 44, 45), suggesting that the
LXCXE motif in cyclin D1 may mediate the binding
of pRb-related proteins (e.g. p107, p130) or some unknown
protein. Apparently normal pRb was detected in DT40 cells; Western
blotting of DT40 lysates with a human pRb antibody demonstrated a broad
band at ~104 kDa, the reported molecular mass for avian pRb (data not shown) (71, 72). The inability of the mutant murine cyclin D1, but not
the normal murine cyclins D1, D2, and D3, to rescue these D1
/
cells suggests that whatever interacts with the
LXCXE motif of cyclin D1, its loss results in an
altered cell cycle. These results are also consistent with the studies
of Skapek et al. (48) demonstrating that cyclins A and E
block muscle gene expression via phosphorylation of pRb but cyclin D1
does not; one interpretation of these data is that a pRb-independent
pathway of skeletal muscle differentiation is regulated by cyclin D1. These results are consistent with what we have observed in the DT40 D1
/
cells. Further biochemical studies of various cell cycle
components (i.e. cyclins, CDKs, and CKIs) will require
reagents, such as antibodies, that react with the avian proteins, since these proteins do not routinely cross-react with the available human
and mouse antisera (data not shown) (52).
Considerable controversy exists regarding the possible
role(s) of the D-type cyclins, particularly cyclin D1, in programmed cell death (73-78). Evidence from several different cell lines in
which cyclin D1 levels are manipulated, either by overexpression or
antisense (including rodent neuronal, mouse mammary epithelial, NIH 3T3
mouse fibroblasts, p53 /
mouse embryo fibroblasts, rat fibroblasts,
and human HCE7 esophageal carcinoma), suggests that cyclin D1
expression is required for apoptosis (73, 75-78). Conversely, others
(32) have shown that the D-type cyclins function as growth factor
sensors that can partially suppress apoptosis in myeloid, interleukin-3-dependent blood cells. Selective tissue
atrophy and loss of proliferative capacity have also been observed in the retina and mammary epithelium of cyclin D1
/
mice, consistent with the hypothesis that cyclin D1 acts as a tissue-specific growth factor sensor (49, 50). In addition, overexpression of cyclin D1 in
rodent fibroblasts, or even concomitant expression of both cyclins D1
and E, has not been associated with increased levels of programmed cell
death (47, 68, 70). In a separate study, Sofer-Levi and Resnitzky (78)
used a tetracycline-responsive promoter to direct expression of cyclin
D1 in serum-starved rat fibroblasts. Induction of cyclin D1 protein
expression in these serum-starved fibroblasts resulted in significant
apoptosis. However, in the presence of normal serum, induction of
cyclin D1 did not result in a significant increase in apoptotic cells.
The availability of avian D1
/
B-cells allowed us to examine this
question directly.
UV and radiation-induced apoptosis occurred in a
dose-dependent manner in both DT40 control and DT40 D1
/
single cell clones (Table III). The percentage of
cells undergoing programmed cell death resulting from 500 to 1,500 J/m2 of UVC or
radiation was substantially increased in
DT40 D1
/
cells (10-15% increase after UVC exposure and a
15-25% increase after
radiation; Table III). To determine whether
this increased sensitivity to radiation, reflected as increased
programmed cell death, was linked to the expression of D-type cyclins,
we examined the effect(s) of
radiation on D1
/
cells in which
murine D-type cyclins were expressed. These experiments involved the
same D1
/
+D-type cyclin(s) cell lines described above, in which
cell cycle abnormalities resulting from ablation of cyclin D1 were
rescued by expression of any murine D-type cyclin. Expression of mouse cyclin D1 in DT40
/
cells (Fig. 4, panel C, D1
/
clone 1/57) eliminated the increased sensitivity to
irradiation
exhibited by the D1
/
cells (Table III). In fact, the percentage of
apoptotic cells in the D1
/
+ murine D1 cells was nearly identical
to the parental cells. Similarly, when D1
/
+ murine D2 or D1
/
+ murine D3 cells were used (Fig. 4, panel D, D1
/
clone
1 + D2/D3), enhanced sensitivity to
irradiation, as reflected by the number of apoptotic cells, was apparently eliminated (Table III).
Intriguingly, expression of the mouse mutant cyclin D1 (missing the
LXCXE motif) partially, but not completely,
restored cellular resistance to radiation-induced apoptosis (Table
III). Since the increased sensitivity of the D1
/
cells to
radiation-induced apoptosis can be effectively reversed by expression
of any normal D-type cyclin, it is likely that these proteins do, in
fact, suppress apoptosis in DT40 cells. Furthermore, the ability of the
mutant cyclin D1 protein to partially suppress radiation-induced
apoptosis but not cell cycle abnormalities (Tables I and III) suggests
that proliferating and apoptotic cells may contain distinct cyclin D1
complexes. Whether these complexes are composed of different polypeptides and/or distinct post-translational modifications is not
known at this time. Further studies with the D1
/
DT40 cell lines
will help to elucidate whether the observed defect in G1
checkpoint control following radiation-induced DNA damage is due
entirely to the disruption of normal p21waf1/cip1cyclin D1
interactions or inappropriate interactions between p21waf1/cip1
and other cell cycle machinery in these cells. However, this response
is consistent with data from p21waf1/cip1
/
mice,
demonstrating that these mice are defective in G1
checkpoint control (79), as well as the developmental abnormalities
observed in D1
/
mice (49, 50) and the inactivation of cdks
following UV irradiation (80).
|
The specific biologic functions of the three D-type cyclins remain
somewhat of an enigma. Mice lacking cyclin D1 by gene targeting in
embryonic stem cells have recently been generated, but the results of
this study reflect a more generalized requirement of this particular
cyclin during development than a specific requirement during the cell
cycle (49, 50). Furthermore, interpretation of the effect(s) of gene
"knockouts" in animal models can be complicated by the expression
of functionally redundant proteins that may mask the phenotype in
certain tissues (81). We have now generated avian B-cell lines that
lack cyclin D1 but retain normal cyclin D2 expression, which exhibit
defects in normal cell cycle progression. The growth of the D1 /
cells is slightly retarded (~20% slower than wild-type DT40 cells),
presumably due to a delay in G2/M phase. A similar delay in
G2/M phase was observed when mouse embryo fibroblasts
derived from the D1
/
mice were analyzed (50). The cell cycle
"defects" observed in the DT40 D1
/
cells can be specifically
rescued by the expression of normal murine cyclin D1, D2, or D3 but not
by a mutant cyclin D1 protein that lacks the
LXCXE pRb-binding motif (44). The fact that
overexpression of the murine cyclin D3 protein, for which there is no
apparent avian homologue, rescues the D1
/
phenotype in DT40 cells
strongly supports the hypothesis that the D-type cyclins have
overlapping functions (3, 4, 82). Our results are also consistent with
a specific requirement for cyclin D1 for normal transit through the
cell cycle and that this requirement cannot be compensated for by the
endogenous cyclin D2 protein. A normal cell cycle can only be acquired
by exogenous expression of any one of the D-type cyclins. The molecular
basis of this rescue is unknown at this time. One can speculate that
the observed effect(s) on the cell cycle are due to inappropriate
level(s) and/or association(s) between D-type cyclin interactors and
other components of the cell cycle machinery (e.g.
inappropriate levels of p21waf1/cip1,
p27Kip1, INK4 or cdk4/6 proteins may affect the
activity of other cyclins, such as cyclin E). Further biochemical
analyses of these cells will require the production of appropriate
antibody reagents for avian cyclins, cdks, and ckis.
We would suggest that the cell cycle perturbations observed in the
cyclin D1 /
cells might be due to 1) an insufficient "threshold" level of cyclin D1 required for entry into
G1 phase from mitosis or 2) defects in replication
resulting from the absence of cyclin D1 that might trigger
G2 phase checkpoints and delay cells at this point until
such damage is repaired. These two possible mechanisms are suggested
since the most obvious outcome of cyclin D1 elimination, lengthening of
G1 phase, was not observed. The second possible explanation
is supported by data from this study in which D1
/
cells, blocked
at the G1/S boundary by centrifugal elutriation and
treatment with aphidicolin, are subject to a 2-3-h delay in
progression through the cell cycle when released from this cycle block
(Table II and Fig. 6). A delay in G2 phase
due to the activation of checkpoints before mitosis is triggered, possibly involving inappropriately associated
cyclin-dependent protein kinase inhibitory (CKI) subunits,
could potentially explain these observations. Whether the observed cell
cycle phenotype and enhanced programmed cell death associated with
these D1
/
B-lymphoma cells will be found in other cell types, or
is specific for this cell lineage, is not known at this time. Further
study of these cell lines, as well as the murine cyclin D1 knockout models (49, 50), should provide answers to these questions. The ability
of any of the D-type cyclins to "rescue" D1
/
DT40 cells
suggests that these cyclins are somewhat redundant in function. However, we cannot completely rule out the possibility that individual D-type cyclins have specific function(s). Additional analysis of the
cyclin D1
/
cell lines may help to resolve the exact relationship(s) between the various D-type cyclins and the proteins they are associated with in vivo (e.g. the CDKs, CDKIs, pRb/p107/p130
and c-Myc (29, 83, 84)).
A separate, but perhaps equally important issue, has arisen regarding
the possible role of the D-type cyclins in apoptosis. The availability
of D1 /
cells has allowed us to examine the possible relationship
between cyclin D1 expression and the induction of apoptosis. Several
studies have linked induction of cyclin D1 gene expression in
post-mitotic neurons and senescent fibroblasts to programmed cell death
(73, 75). In addition, cyclin D1 overexpression in specific tumor cell
lines results in enhanced apoptosis by selective agents (77). Finally,
two different studies have been published concerning cyclin D1-induced
apoptosis in cultured cells (75, 78). In cultured neuronal cells,
cyclin D1 overexpression, in the presence of normal growth factors
(10% serum), was capable of inducing apoptosis (75). However, in cultured fibroblasts, cyclin D1 overexpression induced apoptosis only
in the absence of growth factors (0.1% serum) (78). It should be noted
that the human cyclin D1 gene, CCND1, was originally identified by one group as the PRAD1 oncogene residing at
the breakpoint of a chromosome 11 inversion in parathyroid adenomas (85). As a consequence, these tumor cells express very high levels of
apparently normal cyclin D1 mRNA and protein. Thus, it is somewhat
paradoxical that elevated expression of cyclin D1 can be so well
tolerated in these tumor cells, whereas its overexpression in some
transformed cell lines enhances apoptosis (73, 75, 85). In fact, these
results parallel what has been observed with c-myc, an
oncogene that also induces apoptosis when overexpressed in
serum-deprived cells but not when normal growth factor levels are
maintained (85, 86).
Our results strongly support the role of cyclin D1 as a positive growth
regulator that suppresses apoptosis in proliferating cells but whose
inappropriate expression in factor-deprived cells may trigger cell
death by default. Independent studies of the effects of DNA damage
(e.g. UV and/or irradiation) on the cell cycle machinery
support the notion that D-type cyclins normally suppress apoptosis
(80). In response to UV irradiation, the activity of various CDKs was
inhibited in a p53-dependent manner by enhanced expression
of CDK inhibitors (e.g. p21Waf1/Cip1),
as well as phosphorylation of Thr-14 and Tyr-15 of
p34cdc2. Inhibition of CDK activities in response to radiation
apparently allows cells to either repair their DNA damage, by
suspending progress through the cell cycle until the damage is
repaired, or, alternatively, leads to their elimination by apoptosis if the damage is too extensive. The enhanced level of programmed cell
death observed in the DT40 D1
/
cell lines in response to
radiation-induced DNA damage, as well as the ability of exogenously expressed murine D-type cyclins to desensitize these cells, suggests that cyclin D1 may suppress apoptosis. Once again, we do not know whether this is a cell type-specific effect. Further studies of the
DT40 D1
/
cells could provide valuable insight into how cyclin D1
functions as a mediator of apoptosis and help determine whether this
reflects changes in cyclin D1 complexes with other proteins, such as
CDKs, CDKIs, and pRb/p107/p130, as cells transit the cell cycle. In
addition, it will now be of interest to determine whether elimination
of cyclin D2, on its own or concomitantly with cyclin D1, has a similar
effect(s) on the cell cycle and/or programmed cell death.
Finally, these studies demonstrate, persuasively, that the DT40 cell line is amenable to easy and rapid manipulation of specific genes that regulate the cell cycle and/or programmed cell death and that the resulting cell lines can be valuable reagents for the analysis of the function of these genes.
We especially thank Drs. Charles Sherr and Martine Roussel for providing the mouse cyclin D1, D2, and D3 antisera and murine GEX-cyclin fusion proteins, as well as the normal murine RSV-cyclin D1, D2, D3, and mutant D1 expression constructs and helpful discussions. We also thank Dr. Steve Reed for providing the human cyclin D1 cDNA; Drs. Chen-lo Chen and Max Cooper for providing the DT40 cells; Dr. Craig Thompson for providing the drug selection cassettes; Dr. Martin Berchtold for information regarding cell cycle synchronization of DT40 cells; and Drs. E. Mullner, H. Dolznig, and H. Beug for communicating their results prior to publication. We acknowledge the technical assistance of Gail Richmond, and the advice and comments of our colleagues in the Kidd and Lahti laboratories, as well as the Tumor Cell Biology Department.