(Received for publication, December 28, 1995)
From the
The D-type cyclins promote progression through the G phase of the cell cycle and may provide a link between growth
factors and the cell cycle machinery. We determined the nucleotide
sequence of the 5`-flanking region of the human cyclin D2 and cyclin D3
genes and identified the transcription start sites. Analysis of the
upstream sequences required for transcription of the cyclin D2 and
cyclin D3 genes in continuously dividing cells revealed marked
differences in their regulatory elements. In the cyclin D2 gene
positive elements were localized between positions -306 and
-114 relative to the ATG codon at +1. Additional positive
elements were localized between -444 and -345, whereas
sequences that reduced transcription were identified between
nucleotides -1624 and -892. In the cyclin D3 gene all of
the positive elements required for maximal transcription were localized
between nucleotides -366 and -167, and no negative elements
were found. The activities of a reporter gene linked to the upstream
regulatory sequences of the cyclin D2 gene but not the cyclin D3 gene
were induced when starved cells were serum stimulated. This suggests
that although the abundance of both the cyclin D2 and cyclin D3 mRNAs
is increased by serum stimulation, only the cyclin D2 gene is
up-regulated at the transcriptional level. Sequences between
nucleotides -306 and -1624 of the cyclin D2 gene were
necessary for serum inducibility.
Progression of eukaryotic cells through the cell cycle is
controlled by the activity of cyclin dependent kinases (cdks) ()(Morgan, 1995). Activation of cdks requires binding of a
cyclin and phosphorylation by the Cdk-activating kinase (Morgan, 1995).
In addition, the activity of the Cdk-cyclin complex is inhibited by
binding to cdk inhibitory proteins (Morgan, 1995; Nasmyth and Hunt,
1993). In mammalian cells the commitment to divide is made in the
G
phase of the cell cycle in response to various stimuli,
including growth factors. Passage through the G
phase is
regulated by the D- and E-type cyclins in combination with Cdk2, Cdk3,
Cdk4, and Cdk6 (Sherr, 1993). The primary target of the G
cyclin-Cdk complexes is thought to be the retinoblastoma protein
(pRB) and/or the related proteins p107 and p130 (Sherr, 1993). In turn,
the phosphorylation state of pRB regulates the activity of the E2F
family of transcription factors (La Thangue, 1994; Nevins, 1992). These
transcription factors have been shown to induce the expression of
several genes that are required for DNA replication in S phase (La
Thangue, 1994; Nevins, 1992).
The D-type cyclins are expressed
earlier in G than cyclin E, thereby implicating them in the
earliest events leading to cell division (Sherr, 1993). Unlike other
members of the cyclin family, cyclins D1, D2, and D3 have unique cell-
and tissue-specific patterns of expression (see Sherr(1993) and
references therein). Microinjection of cyclin D1 antibodies or
antisense plasmids prevents passage of cells through G
(Baldin et al., 1993; Tam et al., 1994),
whereas overexpression of cyclin D1 shortens G
(Resnitzky et al., 1994). These data suggest that cyclin D1 is required
for progression through G
. Similarly, overexpression of
cyclin D2 and cyclin D3 shortens the G
phase (Kato and
Sherr, 1993). The mRNA for the D-cyclins have been shown to be induced
by serum and growth factors (Ando et al., 1993; Matsushime et al., 1991; Won et al., 1992), and in the case of
the cyclin D3 and cyclin D1 genes this was shown to be due at least in
part to an increase in transcription rates (Won et al., 1992).
The increase in cyclin D mRNA correlated with increased protein levels
(Ando et al., 1993) and preceded entry into S phase (Baldin et al., 1993; Won et al., 1992). Thus it has been
suggested that the D-cyclins provide a link between growth factors and
the cell cycle. There is evidence that the three D-type cyclins are not
functionally redundant but play distinct roles in the cell cycle
(Sherr, 1993). Recently cyclin D1 (Motokura et al., 1991) and
also cyclin D2 (Hanna et al., 1993) have been implicated in
cancer.
The proliferation of vascular smooth muscle cells (VSMC)
plays a central role in several disease processes, including restenosis
and atherosclerosis (Hermans et al., 1991; Liu et
al., 1989; Ross, 1993). We were interested in studying the
earliest G events that lead to cell division in VSMC. Given
that the D-cyclins appear to be activated by growth factors at the
transcriptional level in several cell types, we set out to define the
regulatory elements that control transcription of the human cyclin D2
and cyclin D3 genes and to define the response of these genes to serum
and growth factors.
Figure 1: Northern blot showing time course of serum induction of cyclin D2 and cyclin D3 mRNAs. Panel A, Northern blot analysis for cyclin D2. RNA was prepared from rat VSMC that had been serum starved and then stimulated with serum for the length of time (h) indicated above each lane. 0, RNA from starved cells. Panel B, similar Northern blot analysis for cyclin D3.
Figure 2:
Nucleotide sequence of the upstream region
of the human cyclin D2 gene. The sequence is numbered relative to the
translation start site (+1). Locations of 5` deletion clones used
in this study are indicated above the sequence by a and the
nucleotide position of the 5` end of the deletion. Also marked above
the sequence are the locations of relevant restriction enzyme sites.
The boxed area indicates the location of a DNase I footprint.
Potential transcription factor binding sites are indicated below the
sequence. For a description of these factors refer to Faisst and Meyer
(1992) and references therein. The ATG translation initiation codon is
also boxed. Transcription start sites determined by
ribonuclease protection are indicated by the stars, and those
determined by primer extension are indicated by the # mark.
Figure 3:
Nucleotide sequence of the upstream region
of the human cyclin D3 gene. The symbols used are the same as
in Fig. 2, except that indicates the location of a DNase
I-hypersensitive site.
Figure 4: Mapping the 5` end of the human cyclin D2 and cyclin D3 mRNA by ribonuclease protection. Panel A, an antisense RNA probe spanning nt -1 to -306 of the cyclin D2 gene was hybridized to 40 µg of yeast RNA (lanes 1 and 4), 40 µg of human lung RNA (lanes 2 and 5), or 20 µg of MRC-5 RNA (lane 3) at 58 °C (lanes 1-3) or 68 °C (lanes 4 and 5). After digestion with ribonucleases the protected fragments were analyzed on a sequencing gel. A sequencing ladder run alongside the samples was used to calculate the size of the protected fragments. Protected fragments are indicated by the arrows, and the star indicates some undigested probe. Panel B, an antisense probe spanning nucleotides +40 to -279 from the cyclin D3 gene was hybridized to 20 µg of yeast RNA (lanes 1 and 3) or 20 µg of MRC-5 RNA (lanes 2 and 4) at 55 °C (lanes 1 and 2) or 62 °C (lanes 3 and 4). The arrows indicate specific protected fragments.
A probe spanning nucleotides -279 to +40 of the cyclin D3 gene was used in a ribonuclease protection assay with RNA from MRC-5 cells (Fig. 4, panel B). At least two major and four minor protected fragments were detected, and the locations of the start sites predicted by these bands are indicated by the stars above the sequence in Fig. 3. This indicates that the cyclin D3 gene also utilizes multiple transcription start sites. Attempts to map the 5` end of the cyclin D3 mRNA by primer extension analysis with several different primers gave inconsistent results that did not correspond to the ribonuclease protection data. This may have been caused by secondary structure in the mRNA due to its high G/C content in this region. The 5` end of the longest published cDNA sequence (Motokura et al., 1992) corresponds to position -162 on the sequence in Fig. 3which is 15 nt upstream of the 5`-most start site detected by ribonuclease protection. This discrepancy could be a result of the different sources of RNA used in the synthesis of the cDNA library and the ribonuclease protection assay.
Putative transcription factor
binding sites were identified in the cyclin D2 and cyclin D3 upstream
sequences using the TFD data base (Ghosh, 1993) and the Macpattern
Program (Fuchs, 1994) and are indicated below the sequence in Fig. 2and Fig. 3. All of the transcription start sites
identified in the cyclin D2 gene, with the exception of the two 3`-most
ones, lie within, or a few nt away from a sequence matching the CAP
site from other genes. The CAP site is the loosely conserved sequence
within which transcription initiates. One of the transcription start
sites (band 5 in Fig. 4, panel A) also lies
within a consensus CAAT box (binding site for NFY/CTF) while another
pair of start sites that correspond to the strongest protected
fragments (bands 3 and 4 in Fig. 4, panel
A) lie within a potential AP2 binding site. However, no TATA box
was evident in the vicinity of the transcription start sites. Putative
binding sites for Sp1, C/EBP, PEA3, PuF, NFB, SIF, E2F, GCF, and
AP1 were also identified in the cyclin D2 upstream sequence. In
addition, a region with sequence identity at 9 of 12 positions to the
binding site of a serum-inducible protein from the cyclin D1 promoter
(Herber et al., 1994) was identified between nt -587 and
-575. Inspection of the cyclin D3 upstream sequence showed no
canonical TATA box close to the transcription start sites, but there
were potential binding sites for Sp1 and AP2 about 10 nt downstream of
the two most 5` start sites. In addition one of the two major start
sites (labeled 3 in Fig. 4, panel B) lies
within a potential AP2 site. Putative binding sites for CTF/NFY, C/EPB,
PEA3, PuF, AP1, p53, and Myb were also identified in the cyclin D3
upstream sequence.
Figure 5: Deletion analysis of cyclin D2 gene regulatory sequences. The left side of the figure shows diagrammatic representations of the luciferase reporter gene constructs, and the right side shows the activity of those constructs after transient transfection into rat VSMC. The constructs are named according to the length of cyclin D2 sequence upstream of the start site. D2 Rev contains the cyclin D2 sequences inserted in the reverse orientation. The boxed area below the arrow indicates the region containing the transcription start sites. The shaded box represents a negative regulatory sequence, and the stippled boxes are positive regulatory sequences. The cross-hatched box represents the luciferase gene. Luciferase activity (corrected for differences in transfection efficiency) is presented as a percentage of the activity of the D2 -1624 construct, whose activity was set at 100% and is the mean of four independent transfections with standard deviations as shown. The D2 Rev construct gave a negative number because its activity was less than that of the promoterless luciferase vector, the activity of which was subtracted from all data.
Sequences between -3300 and -112 from
the cyclin D3 gene (D3 -3300) also directed efficient luciferase
activity in rat VSMC (Fig. 6). The activity of the D3
-3300 construct was on average 3-4 10
relative light units/9.6-cm
well, which was 10-fold
higher than that of the D2 -1624 construct in the same cells.
When the sequences between -3300 and -112 from the cyclin
D3 gene were inserted in the reverse orientation (D3 Rev), activity was
only 8% of that seen with the correct orientation. Deletion of
sequences -1017 to -112 which included the transcription
start sites (D3
-1017/-112) completely abolished
promoter activity. Deletion to the SmaI site at approximately
-1090 resulted in a small increase in activity which was not
statistically significant. Stepwise deletions down to position
-366 had only a minimal effect (approximately 20% reduction) on
promoter activity. However, further deletion to nt -264 reduced
promoter activity to 22% of the full-length plasmid, indicating that
strong positive elements are located between -366 and -264.
Further deletion to nt -167 eliminated all remaining promoter
activity. In conclusion, sequences between -366 and -167
contain all of the regulatory elements required for maximal expression
of the cyclin D3 gene in dividing cells.
Figure 6: Deletion analysis of cyclin D3 gene regulatory sequences. The left side of the figure is a representation of cyclin D3 luciferase constructs that, when transiently transfected into rat VSMC, gave rise to the luciferase activity shown on the right of the figure. The symbols used are the same as those in Fig. 5. The arrow indicates the approximate location of the six transcription start sites that span a 75-nt region.
A subset of the same
deletion constructs were assayed for promoter activity by transfection
in to the normal human lung fibroblast cell line MRC-5 (Table 1).
Deletion of the sequence between -1624 and -444 from the
cyclin D2 promoter resulted in a nearly 2-fold increase in promoter
activity. Although consistent with the finding of a negative element in
this region in VSMC (Fig. 6, panel A) the magnitude of
the effect was less. Deletion of sequences between -444 and
-345 resulted in a 3.4-fold drop in promoter activity confirming
the presence of the positive element in this region which had been seen
in VSMC. Deletion to -306 did not alter promoter activity,
whereas the further removal of sequences to -114 eliminated most
of the activity, consistent with what had been seen in VSMC. The
results with the cyclin D3 promoter in MRC-5 cells were also similar to
those observed in VSMC. Deletion of sequences between -3300 and
-366 had little effect on promoter activity, although a slight
but inconsistent increase was observed. Further deletion to position
-264 resulted in a 2-fold reduction in activity down to 68% of
the full-length construct. This differs quantitatively from the results
in VSMC where D3 -264 had only 22% of the activity of the
full-length construct. Deletion down to -167 eliminated most of
the remaining promoter activity, consistent with what had been observed
in VSMC.
Figure 7:
Footprint analysis of the cyclin D3
promoter. Panel A, DNase I footprinting of a PstI-HindIII fragment (see Fig. 3for location
of the PstI site) from the cyclin D3 gene promoter spanning nt
-664 to -112 and labeled at the HindIII end (3`
end of fragment). The four left lanes (GATC) are a sequencing
ladder generated with a primer complementary to the labeled end of the
probe. The fragment was incubated with 20 µg of HeLa nuclear
extract (lanes 1 and 2) or no nuclear extract (lanes 3 and 4) and digested with 50 ng (lane
1), 150 ng (lane 2), 7.5 ng (lane 3), or 15 ng (lane 4) of DNase I. Panel B, a XmaI-HindIII fragment prepared from D3
-496 and spanning nucleotides -496 to -112 from the
cyclin D3 gene was end labeled at the XmaI site (5` end) and
subjected to DNase I footprint analysis. The probe was incubated with
10 µg (lanes 1 and 2) or 30 µg (lanes 3 and 4) of HeLa nuclear extract or no nuclear extract (lanes 5-7) and digested with 25 ng (lane 1),
50 ng (lanes 2 and 3), 150 ng (lane 4), 5 ng (lane 5), 7.5 ng (lane 6), or 15 ng (lane 7)
of DNase I. The arrow and the star indicate the
location of a site that is made hypersensitive to cutting by DNase I in
the presence of nuclear extract.
Figure 8:
Effects of the cyclin D2 negative element
on heterologous promoters. The line drawing on the left side of the figure shows the restriction map of the 730-bp SacI-PstI fragment from the cyclin D2 gene. Open
rectangles below this represent DNA fragments from this region
which were cloned in either the forward (F) or reverse (R) orientation upstream of either the cyclin D3 promoter in
the construct D3 -664 or the SV40 promoter. The fragments
were cloned into two locations: immediately upstream of the promoter (P) or 2.9 kb upstream of the promoter (D).
Constructs were transiently transfected into rat VSMC and assayed for
luciferase activity. Results are presented as a percentage of the
activity of the promoter alone. The data are the mean of four
independent transfections with standard deviations as
shown.
Interestingly, two smaller fragments from within the SacI-PstI fragment reduced transcription from the cyclin D3 promoter to 14 and 32% of control (Fig. 8). Thus the negative regulatory element appears to contain at least two redundant elements.
Figure 9: Serum inducibility of cyclin D2 and cyclin D3 gene constructs. Each bar represents the mean fold induction by serum of the luciferase constructs shown below the graph. The data are the means of four independent experiments with standard deviations indicated by the error bars. Luciferase activities were corrected for differences in transfection efficiency before calculation of the fold induction. The statistical significance of differences between the fold induction of different constructs was determined using an unpaired Student's t test.
In contrast to cyclin D2, none of the cyclin D3 constructs exhibited serum induction that was significantly above that of the SV40 promoter (Fig. 9).
In this study we have sequenced the 5` upstream regions of
the human cyclin D2 and cyclin D3 genes and mapped the start sites of
transcription. More than 10 transcription start sites spread over a
130-nt region were identified for the cyclin D2 gene; in the cyclin D3
gene 6 start sites covering 70 nt were detected. Both the cyclin D2 and
cyclin D3 genes are devoid of a TATA box, and this may explain the
existence of multiple transcription start sites since this motif is
thought to fix the site of transcription initiation. The existence of
more than one start site has also been reported for the cyclin A
(Henglein et al., 1994), cyclin D1 (Herber et al.,
1994) and cdk2()genes, all of which are also
devoid of a TATA box. The sequence at the site of transcription
initiation, or CAP site, exhibits loose conservation, having the
consensus sequence YYCAYYYYY, where Y is any pyrimidine (Azizkhan et al., 1993). Sequences matching the CAP sites from other
genes and this consensus sequence were found to lie close to or overlap
most of the transcription start sites identified in the cyclin D2 gene
(see Fig. 2). In the absence of a TATA box, binding sites for
transcription factors other than TATA binding protein are used to
initiate transcription (Azizkhan et al., 1993). These include
the factor YY1, as well as E2F and Sp1 (Azizkhan et al.,
1993). In the cyclin D2 gene the sequence surrounding the two most 5`
start sites, and a second cluster of start sites centered at nt
-165, match the consensus binding site for YY1 at eight of nine
positions (see Fig. 2), suggesting that this protein may be
directing initiation at these sites. Two more start sites lie within a
potential AP2 binding site, and yet another lies within a potential
CTF/NFY binding site. Thus, multiple proteins could be involved in
determining the start site of transcription. In the cyclin D3 gene one
of the major start sites lies within a potential binding site for AP2.
The presence of two potential Sp1 sites close to the other major start
site, one of which was bound by nuclear proteins from HeLa cells ( Fig. 3and Fig. 7), suggests that Sp1 may play a role in
transcription initiation from the cyclin D3 promoter.
By deletion
analysis we have mapped the regulatory elements responsible for
transcription of both the cyclin D2 and cyclin D3 genes. In the cyclin
D2 gene deletion analysis identified a negative regulatory element
localized between positions -1624 and -892. This element
also inhibited the activity of two heterologous promoters in a
position- and orientation-independent manner. The stimulatory effect of
the 5` deletions upon D2 promoter activity is unlikely to be caused by
less efficient transfection of larger plasmids since a similar
stimulation was not seen when even larger deletions of the cyclin D3
promoter were tested (Fig. 6). Furthermore, the fact that
deletion of cyclin D3 upstream sequences did not result in a
significant increase in expression makes it unlikely that the
stimulatory effect seen in the cyclin D2 deletions was caused by
bringing cryptic transcriptional enhancers within the cloning vector
closer to the promoter since these would be expected to affect the
cyclin D3 promoter in a similar manner. This explanation is also ruled
out by the fact that the negative element was functional when
transferred into the context of two other promoters and placed in two
different locations (Fig. 8). In these constructs cryptic
transcriptional enhancers within the vector would not have been brought
closer to the promoter. Hypothetically, negative regulation of
transcription, which has been documented for many genes, is a mechanism
by which the rate of transcription can be exquisitely controlled.
Specific repression of transcription can occur by several mechanisms.
In the simplest situation the binding of a negative transcription
factor prevents binding of a positive factor via steric hindrance.
Alternatively, negative factors bound to a negative element may affect
events occurring at a distance, either by interfering with
transcriptional activators or by inhibiting the basal transcription
machinery. Negative elements that interfere with specific activators
bound to the promoter will act on some but not all promoters. In
contrast, negative elements that interfere with the basal transcription
machinery will affect all promoters. Our results show that the negative
element from the cyclin D2 gene can inhibit the activity of both the
cyclin D3 promoter and the SV40 promoter. Thus it is more likely that
the cyclin D2 negative element functions by interfering with the basal
transcription complex. However, we cannot rule out the possibility that
the cyclin D2 negative element functions by inhibiting the activity of
a transcriptional activator common to the cyclin D2, cyclin D3, and
SV40 promoters. A negative element was also identified in the upstream
region of the human cyclin D1 gene (Herber et al., 1994),
although it reduced promoter activity by only 3-fold. Sequence
comparison identified a 12-nt region from within the cyclin D2 negative
element which matched a region of the cyclin D1 negative region at 10
positions, but the significance of this finding is unclear. The cyclin
D2 negative element may contribute to the serum inducibility of this
promoter since deletion of this element resulted in a reduction in
serum inducibility from 6.8- to 4.3-fold. This reduction is the result
of a greater increase in promoter activity after removal of the
negative element under starved conditions than under conditions of
serum stimulation. This suggests that one function of the negative
element may be to keep transcription repressed in the absence of
proliferative signals. Thus the negative element could potentially be
activated by extracellular stimuli that inhibit cell growth and/or
intracellular events in G or late cell cycle which
down-regulate G
events.
Two distinct positive regulatory elements were identified in the cyclin D2 upstream sequence. The first, which we consider to be the basal promoter, spans nucleotides -306 to -114 and has about 50% of the activity of the largest cyclin D2 construct. This region contains all of the transcription start sites as well as potential binding sites for CTF/NFY, AP2, and C/EBP. Sequences between -444 and -345 contain strong positive elements that increased the activity of the basal promoter by 10-fold in VSMC and by 3.4-fold in MRC-5 cells and thus seem to act like an enhancer. Putative binding sites for AP2 and PEA3 are present within this region. Interestingly, the addition of the cyclin D2 negative element almost neutralizes the effect of this enhancer upon promoter activity. Further experiments are needed to determine if the cyclin D2 negative element does in fact interfere with the function of the enhancer.
A single DNase I footprint was identified in the cyclin D2 promoter using HeLa cell nuclear extract. This footprint lies upstream of all of the positive elements identified by the deletion analysis, suggesting that it is not important for promoter activity in continuously dividing cells. However, we cannot rule out the possibility that it may be important for serum induction. The footprint contains sequences with similarity to the binding sites for PuF (a factor that binds to the c-myc promoter), AP2, and Sp1. Since HeLa nuclear extract contains Sp1 (Briggs et al., 1986) this may explain why this footprint was detected. Further work will be needed to identify the proteins interacting with the positive elements identified in the cyclin D2 promoter. In conclusion, cyclin D2 gene transcription is controlled by a combination of positive and negative elements and could potentially be modulated by changes in the activity of any of these elements.
The structure of the cyclin D3
promoter is very different from that of the cyclin D2 promoter; no
negative element was identified, and all of the sequences required for
full promoter activity were localized to a region between -366
and -167. Two DNase I footprints were detected in the cyclin D3
promoter. Both footprint I (-229 to -215) and footprint II
(-171 to -161) lie within the positive regulatory element
identified by deletion analysis, and both contain a sequence with
similarity to the binding site for Sp1. The role of these potential Sp1
sites in the transcription of the cyclin D3 gene remains to be
elucidated. Potential binding sites for Sp1 were also identified in the
promoters of the cyclin A (Henglein et al., 1994) and cyclin
D1 (Herber et al., 1994) genes as well as the cdk2 gene. The discovery of binding sites for Sp1, which is
a ubiquitously expressed factor, is consistent with the requirement for
expression of the cyclin genes in most cell types. Interestingly, a
DNase I-hypersensitive site was detected at position -133 of the
cyclin D3 gene. This is indicative of the binding of a nuclear protein
which alters the conformation of the DNA such that it becomes more
accessible to DNase I. This site lies within a region that, when
deleted, reduced promoter activity from 78 to 22% in VSMC and from 144
to 68% in MRC-5 cells, suggesting that it may be important for
transcription.
A similar analysis of the human cyclin D1 promoter in NIH3T3 cells identified positive elements between -848 and -742 and additional positive elements that were spread throughout the region between -742 and -29, rather than being localized to one particular region (Herber et al., 1994). Together these results show that there are marked differences in the arrangement of transcriptional regulatory elements between the three D-type cyclins. This suggests that transcription of these genes is independently regulated, supporting the idea that the three D-cyclins are not redundant but perform distinct functions. This was first suggested by the finding that the coding sequences of the human D-cyclins are more closely related to their mouse homologs than to each other (Inaba et al., 1992; Xiong et al., 1992). In addition, cyclins D2 and D3 form more stable complexes with the retinablastoma protein (pRB) than does cyclin D1 (Ewen et al., 1993; Kato et al., 1993). Furthermore, cyclins D2 and D3 efficiently activated Cdk2 in insect Sf9 cells, whereas cyclin D1 could not (Ewen et al., 1993); and unlike cyclins D2 and D3, cyclin D1 was unable to induce retinablastoma protein (pRB) phosphorylation in human diploid fibroblasts (Dowdy et al., 1993).
We examined
serum inducibility of cyclin D2 and cyclin D3 reporter constructs in a
transient transfection assay. The conditions used in this assay are
similar to those in which we observed serum induction of the cyclin D2
and cyclin D3 mRNAs. However, two conditions that were necessary in
these transient transfection assays would be expected to lead to an
underestimate of the true level of serum inducibility of these
promoters. First, to minimize the loss of the reporter gene plasmid
that begins 4 days post-transfection the starvation period had to be
limited to 30 h. Second, when starved rat VSMC were restimulated with
serum less than 50% of the cells were found to have reentered the cell
cycle (data not shown). Despite these limitations we observed that
sequences between nt -1624 and -1 from the cyclin D2 gene
conferred 6.8-fold induction by serum (Fig. 9), which is
comparable to the induction of the endogenous rat cyclin D2 mRNA (Fig. 1, panel A). This demonstrates that increased
transcription from the cyclin D2 promoter is at least partly
responsible for the increase in cyclin D2 mRNA observed in response to
serum. The sequences required for serum induction lie between
-306 and -1624 but did not map to a single region within
this sequence. Sequences between -1624 and -444, which
contains the negative element, appear to contribute to serum induction
as do sequences between -444 and -306. A more comprehensive
analysis is needed to localize further the DNA sequences required for
serum induction. In the case of the cyclin D1 promoter (Herber et
al., 1994), all of the deletions tested were inducible by serum,
but a significant part of the serum inducibility could be attributed to
a 100-bp region around position -900 (Herber et al.,
1994). Within this region lie binding sites for a novel serum-inducible
protein (Herber et al., 1994) and the AP1 family of
transcription factors (Albanese et al., 1995; Herber et
al., 1994). This AP1 site was shown to mediate induction of cyclin
D1 transcription by transforming mutants of p21 (Albanese et al., 1995). A second signal transduction pathway appears to
function via a binding site for the ETS transcription factor within the
proximal 22 bp of the cyclin D1 promoter, which is itself regulated by
the activity of mitogen-activated protein kinase (Albanese et
al., 1995). This minimal cyclin D1 promoter with just 22 bp of
upstream sequence could be induced 3.5-fold by epidermal growth factor,
and this was mediated by mitogen-activated protein kinase (Albanese et al., 1995). This contrasts with our results, which show
that a cyclin D2 reporter construct that retains significant promoter
activity (D2
-306) is not inducible by serum supplemented
with basic fibroblast growth factor and epidermal growth factor. A
sequence with similarity to the binding site of the serum-inducible
protein from the cyclin D1 promoter (Herber et al., 1994) was
identified in the upstream region of the cyclin D2 gene between nt
-587 and -575. In this same region is the sequence TGAGTgA
(-582 to -576), which matches the AP1 site from the cyclin
D1 gene at six out of seven positions. These sequences lie within the
region of the cyclin D2 promoter which we have shown is required for
serum induction. Further experiments will be needed to determine the
functional significance of these regions of homology.
None of the cyclin D3 promoter constructs tested exhibited induction by serum and growth factors which was above that of the SV40 promoter. One interpretation of these data is that the induction of the cyclin D3 mRNA observed by Northern blot (Fig. 1) is due primarily to post-transcriptional events such as stabilization of the mRNA. However, we cannot formally rule out the possibility that sequences outside of the largest construct tested are required for transcriptional induction of the cyclin D3 gene.
Induction of the cyclin D2 and D3 mRNA occurs after the induction of the immediate-early genes, suggesting that products of the immediate-early genes may be required for this induction. Therefore, products of the c-fos, c-jun, c-myc, and c-myb genes are good candidates for transcription factors that might be involved in the activation of cyclin D2 gene transcription in response to serum. Indeed, we found putative binding sites for AP1 (fos, jun) and myb in the upstream sequence of the cyclin D2 gene. Further experiments are needed to determine which if any of these transcription factors are involved.
Our findings are consistent with the concept that, like the cyclin D1 gene, the transcription of the cyclin D2 gene is a downstream target of mitogen-activated signal transduction pathways, and they suggest fine control of cell cycle progression by transcriptional as well as post-translational regulation. It seems likely that the D-type cyclins may play different functional roles in various cell types by responding to different signal transduction pathways. This work provides the basis for understanding the exact molecular mechanisms that link mitogens to the cell cycle.
The nucleotide sequence(s) reported in this paper has been submitted to the GenBank(TM)/EMBL Data Bank with accession number(s) 447284 (cyclin D2) and 447285 (cyclin D3).