Journal of General Virology (2003), 84, 51–60
DOI 10.1099/vir.0.18702-0
Human cytomegalovirus immediate-early
protein 2 (IE2)-mediated activation of cyclin
E is cell-cycle-independent and forces
S-phase entry in IE2-arrested cells
Lüder Wiebusch, Jasmin Asmar, Ralf Uecker and Christian Hagemeier
Department of Pediatrics, Laboratory for Molecular Biology, Charité, CCM-Ziegelstr. 5–9,
Humboldt-University, Berlin, Germany
Correspondence
Christian Hagemeier
[email protected]
Received 10 July 2002
Accepted 4 October 2002
In human cytomegalovirus (HCMV) infection, the isolated expression of the viral immediate-early
protein 2 (IE2) 86 kDa regulatory protein coincides with an up-regulation of cyclin E gene
expression, both in fibroblasts and U373 cells. Since IE2 also interferes with cell-cycle progression,
it is unclear whether IE2 is a genuine activator of cyclin E or whether IE2-arrested cells contain
elevated levels of cyclin E primarily as a consequence of them being arrested at the beginning of S
phase. It is important to distinguish between these possibilities in order to define and analyse at a
mechanistic level the proliferative and anti-proliferative capacities of IE2. Here we have shown that
IE2 can activate cyclin E independent of the cell-cycle state and can therefore function as a genuine
activator of cyclin E gene expression. A mutant of IE2 that failed to activate cyclin E also failed to
promote G1/S transition. Instead, cells became arrested in G1. S-phase entry could be rescued in
these cells by co-expression of cyclin E, but these cells still arrested in early S phase, as is the case
with wild-type IE2. Our data demonstrate that IE2 can promote two independent cell-cycle
functions at the same time: (i) the induction of G1/S transition via up-regulation of cyclin E, and (ii) a
block in cell-cycle progression in early S phase. In G1, the proliferative activity of IE2 appears to be
dominant over the anti-proliferative force, whereas after G1/S transition, this situation is reversed.
INTRODUCTION
The lytic infection cycle of human cytomegalovirus
(HCMV) is accompanied by a characteristic deregulation
of the host cell cycle (Fortunato et al., 2000; Kalejta & Shenk,
2002). Cellular DNA synthesis is disturbed by the virus,
resulting in cell-cycle arrest at or near the G1/S transition
(Bresnahan et al., 1996; Dittmer & Mocarski, 1997; Lu &
Shenk, 1996). At the same time, many S-phase-promoting
activities are induced, even under conditions of growth
factor deprivation, contact inhibition and differentiation
(Bresnahan et al., 1996; Jault et al., 1995; Sinclair et al.,
2000). In particular, cyclin E–cdk2 kinase activity, which
drives G1/S transition in the normal cell cycle (Sauer &
Lehner, 1995), is constitutively up-regulated during HCMV
infection (Jault et al., 1995). Consistent with this, the cyclin
E–cdk2 substrates pRb and p130 have been found to be
hyperphosphorylated (Jault et al., 1995; McElroy et al., 2000;
Sinclair et al., 2000) and Rb/E2F-controlled genes, such
as dihydrofolate reductase and thymidylate synthase, are
transcriptionally activated in infected cells (Gribaudo et al.,
2000; Wade et al., 1992). All in all, this leads to an S-phase
metabolism with greatly increased nucleotide pools in
HCMV-infected cells (Biron et al., 1986). However, whereas
cyclin E–cdk2 activity and biosynthesis of nucleotides are
0001-8702 G 2003 SGM
both essential host-cell functions for HCMV replication
(Bresnahan et al., 1997a; Gribaudo et al., 2000), ongoing
cellular replication appears to counteract progress of the
infectious cycle (Salvant et al., 1998).
The HCMV 86 kDa immediate-early protein 2 (IE2) is not
only an essential transcriptional activator of early viral gene
expression (Gebert et al., 1997; Heider et al., 2002; Marchini
et al., 2001) but also a cell-cycle regulator that can reproduce
some important features of the deregulated cell-cycle
phenotype in HCMV infection. It blocks cell-cycle progression at the beginning of S phase in various cell types
(Murphy et al., 2000; Wiebusch & Hagemeier, 1999, 2001),
including primary fibroblasts (Kronschnabl et al., 2002).
This arrest resembles the specific inhibition of DNA
replication by chemical inhibitors such as hydroxyurea
(HU) or aphidicoline since it is dominant over cyclin–cdk
activities and leaves the Rb–E2F pathway untouched
(Wiebusch & Hagemeier, 2001). Moreover, cyclin E expression and activity are up-regulated in IE2-arrested cells
(Wiebusch & Hagemeier, 2001) and the mRNA levels
of numerous genes involved in nucleotide synthesis
and DNA replication are elevated (Song & Stinski, 2002).
Other findings also point towards IE2 as a factor with an
ambivalent role in regulating cell proliferation: IE2 relieves
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51
L. Wiebusch and others
the flat-cell phenotype of pRb-transfected Saos cells
(Fortunato et al., 1997) and rescues the defect of ts13
cells in cyclin transcription (Lukac et al., 1997) – in both
cases the cell cycle is still blocked. IE2 even forces quiescent REF-52 cells to re-enter the cell cycle and synthesize
at least limited amounts of DNA (Castillo et al., 2000).
Thus, there is considerable evidence that IE2 triggers both
proliferative and anti-proliferative cellular activities but it
is unknown how these functions integrate mechanistically
on IE2.
Activation of cyclin E–cdk2 in HCMV-infected cells is
mainly achieved by a strong increase in cyclin E transcription (Bresnahan et al., 1998; Salvant et al., 1998). Other
HCMV-induced alterations that are known to contribute to
the cyclin E–cdk2 activation are the translocation of cdk2
from the cytoplasm into the nucleus (Bresnahan et al.,
1997b) and the enhanced degradation of the cdk inhibitor
p21WAF1,CIP1 (Chen et al., 2001). How IE2 could contribute
towards cyclin E–cdk2 regulation is not fully understood.
One report investigating the requirements for cyclin E
activation in infected cells questioned the importance of IE2.
It demonstrated that cyclin E induction depended on early
viral gene expression and was mainly associated with a
change in cyclin E promoter occupancy concerning the
composition of E2F–pocket protein complexes (McElroy
et al., 2000). In contrast, in vitro experiments showing binding of IE2 to the cyclin E promoter (Bresnahan et al., 1998)
and a functional IE2–p21WAF1,CIP1 interaction (Sinclair
et al., 2000) have suggested a direct involvement of IE2 in
cyclin E–cdk2 activation on the transcriptional as well as the
post-translational level. However, even the fact that IE2
transfection leads to an up-regulation of cyclin E promoter
activity (Bresnahan et al., 1998), cyclin E mRNA level
(Song & Stinski, 2002; Wiebusch & Hagemeier, 2001), cyclin
E protein level and cyclin E-associated kinase activity
(Wiebusch & Hagemeier, 2001) cannot unequivocally
qualify cyclin E as a primary IE2 target. Since the beginning
of S phase where IE2-expressing cells accumulate coincides
with maximum cyclin E expression and maximum cyclin
E–cdk2 activity (Dulic et al., 1992; Koff et al., 1992), it is also
conceivable that cyclin E only serves as a cell-cycle marker
for IE2-arrested cells.
Therefore, the work presented here is aimed at examining
whether IE2 can serve as a genuine activator of cyclin E
gene expression and looking at the possible functional
implications of such an activation in IE2-expressing cells.
METHODS
Cells. U373-MG cells and primary human embryonic lung fibro-
blasts were grown in Dulbecco’s modified Eagle’s medium supplemented with 5 % newborn calf serum, 5 % foetal calf serum,
100 U penicillin ml21 and 100 mg streptomycin ml21. Every 3 days,
cells were split in a 1 : 3 ratio to maintain a subconfluent state.
Plasmids. pSG5 (Green et al., 1988) and the expression plamids
for CD20, IE2(195-579) and cyclin E [pSG5-CD20, pSG5-3HAIE2(195-579) and pCMX-CycE] have been previously described
52
(Wiebusch & Hagemeier, 2001). For IE2 expression, the vector
pSG5-IE2-His was used containing full-length IE2 cDNA of AD169
origin C-terminally fused to a tag of six histidine codons.
DNA transfections. Cells were transfected using the calcium phos-
phate co-precipitation method, as previously described (Wiebusch &
Hagemeier, 1999). The DNA precipitates were left on the cells for
14–16 h. The transfection medium was replaced with fresh culture
medium supplemented with 1 mM mimosine, 1 mM HU or 10 mM
5-bromo-29-deoxyuridine (BrdU) as indicated. Nocodazole was
added at a final concentration of 50 ng ml21 24 h after removal of
the DNA precipitates. The time-points of cell harvest are specified in
the figure legends.
Cell sorting. Transfected cells were labelled with a CD20 antibody
(clone 2H7; Pharmingen) and separated from untransfected cells
by magnetic affinity cell sorting (MACS), as previously described
(Wiebusch & Hagemeier, 2001). To control the sorting efficiency,
aliquots of sorted and unsorted cells were stained with an FITCconjugated secondary antibody (F0313; DAKO) and analysed
by flow cytometry for CD20 expression. To control for IE2 coexpression, cells were permeabilized by overnight incubation in
70 % ethanol in PBS, blocked in 0?5 % BSA in PBS and labelled
with a primary IE1/2-specific antibody (clone E13; Argene) and a
secondary FITC-conjugated mouse IgG1-specific antibody (clone
A85-1; Pharmingen). Both antibody-binding reactions were carried
out for 15 min at room temperature using 5 mg antibodies ml21
in 0?5 % BSA in PBS, followed by a washing step with 0?5 % BSA
in PBS. Finally, cells were suspended in PBS and analysed by flow
cytometry.
Cell-cycle analysis. The overall DNA content of transfected cells
was determined by propidium iodide staining and flow cytometry,
as previously described (Wiebusch & Hagemeier, 1999). The BrdU
incorporation assay for detection of newly synthesized DNA was
carried out as described by Wiebusch & Hagemeier (2001). Briefly,
following CD20-directed cell sorting, BrdU-treated cells were permeabilized in 70 % ethanol and stained by indirect immunofluorescence using an anti-BrdU antibody (clone 3D4; Pharmingen) and
an isotype-specific secondary antibody (clone A85-1; Pharmingen)
to avoid cross-reaction with the CD20 antibody used for cell sorting.
Cells were then co-stained with propidium iodide and analysed by
flow cytometry.
Immunoblot analysis and kinase assay. After sorting, cell
extracts of CD20-positive cells were prepared as previously described
(Wiebusch & Hagemeier, 2001). These were used for immunoblot
analysis and cyclin E kinase assays. Both assays were performed
exactly as previously described (Wiebusch & Hagemeier, 2001)
employing the cyclin E monoclonal antibodies HE12 (Santa Cruz)
for immunoblotting and HE111 (Santa Cruz) for immunoprecipitation of cyclin E kinase activity. Blots were routinely controlled
for equal loading and protein transfer by Ponceau S staining. To
control the expression level of IE2 and its mutant IE2(195-579),
blots were probed with a rabbit antiserum recognizing the Cterminal region of IE2 (a kind gift from Thomas Stamminger,
Erlangen, Germany).
Ribonuclease protection assay (RPA). Total RNAs from CD20positive cells were isolated using Trizol reagent (Gibco) according
to the manufacturer’s instructions. Two mg of each preparation
was taken as input in a multiprobe RPA (Riboquant; Pharmingen).
A defined pool of radioactively labelled probes was generated by
in vitro transcription from the template set hCyc-2 (Pharmingen)
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Journal of General Virology 84
HCMV IE2 activation of cyclin E
and hybridized with the input RNA. The samples were subsequently digested with an RNase A/T1 mixture, purified by ethanol
precipitation, separated by denaturing PAGE and subjected to autoradiography.
RESULTS
We set up an experimental system to determine whether
IE2 was a genuine activator of cyclin E or whether cyclin E
up-regulation was a mere consequence of the early S-phase
block in IE2-expressing cells. This system allowed us to study
the effect of IE2 on cyclin E gene expression prior to the
physiological and cell-cycle-dependent induction of cyclin E.
This distinction is important because it would allow us to
assign a proliferative capacity to IE2 (and suggest a mechanistic explanation of it) in addition to the previously shown
cell-cycle arrest function of this essential virus regulator.
Thus, we applied an experimental system that we have
successfully used before (Wiebusch & Hagemeier, 2001),
which relies on the co-transfection of cDNAs for IE2 and the
cell-surface marker CD20 into U373 cells, a cell line in which
HCMV can replicate. This enabled us: (i) to determine the
cell-cycle stage of IE2-expressing cells, and (ii) to physically
separate IE2-expressing cells via MACS for biochemical
analysis (see Methods; Wiebusch & Hagemeier, 2001). In
addition, we made use of two drugs that are known to arrest
cell-cycle progression (Fig. 1A): mimosine, which can block
cells in a dose-dependent manner in G1, and hydroxyurea
(HU), which inhibits ongoing DNA synthesis in early S
phase (Krude, 1999, Wiebusch & Hagemeier, 2001). Since
cells take up transfected DNA primarily during mitosis
at times when the nuclear membrane breaks down, these
cells become functionally synchronized by the transfection
procedure itself (Adams et al., 1997; Rodriguez et al., 1999;
unpublished observations). Therefore, in the presence of
mimosine or HU, control-transfected, i.e. CD20-positive,
cells synchronously ran into a cell-cycle block that appeared
to be characterized by a G1-like DNA content (Fig. 1B,
top panel). However, HU-treated cells had elevated levels
of cyclin E protein (Fig. 1C, lane 2) reflecting the fact that
these cells were arrested right at the beginning of S phase,
which could be directly demonstrated by these cells having
incorporated a small amount of BrdU (data not shown;
Wiebusch & Hagemeier, 2001). In contrast, mimosinearrested cells were truly blocked in G1 (for BrdU incorporation analysis, see below) and importantly, these cells
had non-elevated levels of cyclin E protein (Fig. 1C, lane 3).
Although mimosine has also been reported to block cells
in early S phase (Hughes & Cook, 1996), this was clearly not
the case in the transfected, CD20-positive population, since
after transfection these cells synchronously leave mitosis and
pass into G1 before becoming arrested in this phase.
The validity of our experimental approach was further supported by analysing CD20-negative, i.e. non-transfected,
cells, which reflected the non-synchronized population.
Here, HU treatment resulted in arrested cells not only at the
beginning of S phase but also, as expected, during later stages
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Fig. 1. Mimosine arrests cells before cyclin E expression peaks
at the G1/S transition. Proliferating U373 cells were transiently
transfected with pSG5 and a CD20 expression plasmid
(pSG5-CD20). After transfection, mimosine (Mimo) or hydroxyurea (HU) were applied to the cells. After 32 h, cells were
harvested and sorted by MACS technology. (A) Schematic diagram of the temporal order of transfection events and drug
effects in the cell cycle. (B) Cell-cycle profiles of the different
CD20-positive and CD20-negative cell fractions obtained by
propidium iodide staining and flow cytometry analysis. (C)
Extracts of the same cell fractions examined for cyclin E
expression by immunoblot analysis.
of S (Fig. 1B, lower panel). Equally, the mimosine-treated
culture not only contained cells in G1 but also cells in early S,
as one would expect from the mode of action of this drug
on a proliferating cell population. Importantly, the analysis
of cyclin E protein levels was consistent with this notion. As
expected, CD20-negative HU-treated cells had high levels of
cyclin E, whereas the mimosine-arrested population had
intermediate levels of cyclin E, reflecting both the majority
of G1 phase cells and cells that were hit by this drug in early S
phase (Fig. 1C, lanes 4–6).
Together these data showed that in our experimental system
of functionally synchronized cells, mimosine not only
arrested transfected cells in G1 but, furthermore, the time
point of arrest within G1 was prior to the physiological boost
of cyclin E induction late in G1. This assay system should
therefore allow us to measure any genuine effect of IE2 on
the expression level of cyclin E.
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L. Wiebusch and others
Cyclin E activation by IE2 is independent of the
cell-cycle stage
We next transfected cells with either an IE2 expression
plasmid or its parental empty plasmid, both in the presence and absence of mimosine, to test for IE2-mediated
cell-cycle-independent activation of cyclin E. First, we asked
whether IE2 expression would interfere with the mimosinedependent cell-cycle block in G1. Whereas in the absence of
mimosine control-transfected cells were evenly distributed
over S phase, IE2-expressing cells were blocked at the beginning of S (Fig. 2A, plots 1 and 2; B) as shown by BrdU
incorporation assays. These results are consistent with
previous data (Wiebusch & Hagemeier, 2001). In contrast,
mimosine treatment, as expected from the data presented
in Fig. 1, led to a clear-cut block of cell-cycle progression in
G1, irrespective of the presence or absence of IE2 (Fig. 2A,
plots 3 and 4; B). Thus, IE2 expression did not overcome
the mimosine-induced G1 arrest.
We then used aliquots of these cells to look for cyclin E gene
expression. Consistent with previously published results
(Wiebusch & Hagemeier, 2001), IE2 up-regulates cyclin E
in the absence of mimosine, which can be observed by
increased steady-state levels of mRNA (Fig. 3A, lanes 2 and
3), protein (Fig. 3B, lanes 1 and 2) and kinase activity
(Fig. 3C, lanes 1 and 2), the latter measured as cyclin
E-dependent phosphorylation of histone H1. A virtually
identical picture of an IE2-mediated cyclin E induction
was observed when the experiment was performed in
the presence of mimosine (Fig. 3A, lane 5; B, C, lane 4),
although mimosine itself clearly arrested cells in G1 (Fig. 2A,
plot 3) prior to cyclin E induction (Fig. 3A, lane 4; B, C,
lane 3). This demonstrates that IE2 can actively (and
independently from G1/S transition) up-regulate cyclin
E protein levels, which results in the untimely induction
of cyclin E-associated kinase activity under the experimental conditions. The activation appears to occur
primarily at the level of transcription, although additional
post-transcriptional mechanisms of activation cannot be
excluded.
Cyclin E up-regulation by IE2 also occurs in
primary fibroblasts
Fig. 2. IE2-expressing cells are arrested in G1 by mimosine.
U373 cells were transfected with an IE2 expression plasmid
(IE2) or its parental plasmid pSG5 (control). pSG5-CD20 was
co-transfected as a selection marker. After transfection, BrdU
was added to the cells, with or without mimosine. After an
incubation period of 32 h, cells were harvested and doublestained with propidium iodide and an FITC-labelled anti-BrdU
antibody to enable FACS analysis of DNA content and BrdU
incorporation. (A) Density dot plots of CD20-positive cells. (B)
Quantification of BrdU-positive cells shown as a bar chart.
Data from a single representative experiment are shown.
54
To exclude the possibility that IE2 up-regulation of cyclin E
is cell-type-specific or a consequence of the transformed
phenotype of U373 cells, we extended our analysis to
primary lung fibroblasts. We co-transfected the CD20
surface marker together with IE2 and prepared extracts
for immunoblot and ribonuclease protection analysis from
transfected and MACS-sorted fibroblasts. Since the transfection efficiency of primary fibroblasts was very low (3 % in
Fig. 4A, top right-hand panel), we first wanted to ensure
that transfected cells were expressing both the CD20 marker
and IE2. Fig. 4(A) shows that MACS allowed a very
significant enrichment of transfected cells (compare top
and bottom right-hand panels) and that most CD20-sorted
cells indeed also expressed IE2 (89 %, bottom right-hand
panel). In these cells, IE2 expression led to a marked increase
in cyclin E expression at both the mRNA (Fig. 4B, lane 3)
and protein levels (Fig. 4C, lane 2) when compared with
sorted control-transfected primary fibroblasts (Fig. 4B, lane
2; C, lane 1). In contrast, a transactivation-deficient mutant
of IE2 (Wiebusch & Hagemeier, 1999) lacking the first 194
amino acids, IE2(195-579), was not able to up-regulate
cyclin E (Fig. 4B, lane 4; C, lane 3), although it was expressed
to the same extent as full-length IE2 (see Fig. 5C; Wiebusch
& Hagemeier, 1999). This mutant also failed to activate
cyclin E in U373 cells (Wiebusch & Hagemeier, 2001), but
was still competent in inducing a cell-cycle block (Wiebusch
& Hagemeier, 1999; see below). These data support the view
of transcriptional activation of cyclin E gene expression by
IE2, which is consistent with the result shown in Fig. 3 and
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HCMV IE2 activation of cyclin E
Fig. 3. Cyclin E is activated by IE2 in mimosine-treated cells.
U373 cells were transfected as described in Fig. 2. Following
transfection, cells were treated with mimosine (Mimo) for 32 h
or left untreated. After cell harvest, CD20-positive cells were
separated and analysed for expression and activity of cyclin E.
(A) Cyclin E mRNA levels measured by multiprobe ribonuclease protection assay. (B) Cyclin E protein levels determined
by immunoblot analysis. (C) Cyclin E-associated kinase activity
assayed in vitro using immunoprecipitated (IP) cyclin E protein
complexes and the substrate histone H1.
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Fig. 4. IE2 up-regulates cyclin E in primary fibroblasts. Human
embryonic primary lung fibroblasts were transfected with
pSG5-CD20, in combination with pSG5 (control) or plasmids
expressing IE2 or the IE2 mutant IE2(195-579). CD20-positive
cells were isolated and analysed for cyclin E expression. (A)
Cells transfected with CD20 and the indicated expression plasmids and analysed for co-expression of CD20 and IE2 by flow
cytometry. (B) Cyclin E protein levels determined by immunoblot analysis of total cell extracts. (C) Cyclin E mRNA levels
analysed by ribonuclease protection assay.
in agreement with other reports (Bresnahan et al., 1996;
Song & Stinski, 2002). Moreover, and in contrast to a
previous study (McElroy et al., 2000), it has been shown
here for the first time that this activation also resulted in
elevated cyclin E protein levels in primary fibroblasts. Thus,
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L. Wiebusch and others
Fig. 5. Cyclin E activation is responsible for S-phase entry in IE2-expressing cells. U373 cells were transiently transfected
with pSG5-CD20 together with plasmids expressing IE2, IE2(195-579), or cyclin E, alone or in the indicated combinations.
As a control, pSG5-CD20 was co-transfected with pSG5. In all transfections, BrdU was included in the culture medium after
removal of DNA precipitates. Cells were harvested 32 h later and sorted for CD20 expression. The cell-cycle distribution of
CD20-positive cells was analysed by anti-BrdU–FITC and propidium iodide staining. (A) Plots of BrdU signal against the
overall DNA content. (B) Quantification of the cell-cycle phases and comparison of the ratios between the G0/G1 and early S
phase. (C) Analysis of the expression of IE2, IE2(195-579) and exogenous cyclin E by immunoblotting of CD20-positive cells.
Lanes correspond to plots 1–6 in (A).
IE2-mediated up-regulation of the cyclin E protein is neither
cell-type-specific nor dependent on cells with a transformed
phenotype.
Cyclin E activation is responsible for S-phase
entry of IE2-expressing cells
The fact that IE2 and IE2(195-579) can both induce a
cell-cycle arrest but only full-length IE2 activates cyclin
E opens up the question of the biological significance of
the cyclin E activation by IE2. In order to address this
question, we further characterized cells arrested by IE2
and IE2(195-579) using BrdU incorporation assays (Fig. 5A,
B). Control-transfected cells were evenly distributed over
S phase, whereas IE2-arrested cells were found to be
located right at the beginning of S phase (Fig. 5A, plots 1
and 2), as previously shown. Surprisingly, lack of BrdU
incorporation in cells expressing the mutant form of IE2
suggested that this mutant blocks cell-cycle progression
in G1 rather than early S phase (Fig. 5A, plot 3). The
IE2(195-579)-expressing population contained only half as
many cells in early S phase compared with cells expressing
IE2, which was primarily due to a respective increase of cells
in G1 (Fig. 5B).
56
Given that the IE2 mutant protein failed to activate cyclin E
(see above), this suggested to us that IE2-mediated cyclin E
induction is a prerequisite for S-phase entry. Conversely,
lack of cyclin E expression in IE2(195-579)-transfected
cells appeared to go hand in hand with a lack of S-phase
entry. To test this hypothesis directly, we co-expressed cyclin
E and IE2(195-579) and asked whether, under these conditions, cells would now enter S phase as measured by
BrdU incorporation. As predicted, cyclin E was found
to completely rescue the S-phase entry of IE2(195-579)expressing cells (Fig. 5A, plot 4). At the same time, fulllength IE2 was also able to rescue S-phase entry when
co-expressed with the IE2 mutant form (Fig. 5A, plot 6),
also implying that IE2(195-579) does not function as a
dominant negative form of IE2. Quantification of the
rescue experiments showed that co-expression of cyclin E
induced a more pronounced S-phase entry in IE2(195-579)transfected cells than co-expression of IE2 (49 % versus
38 %, Fig. 5B). This was most likely due to the significantly
higher levels of cyclin E protein in these cells (data not
shown). In accordance with this notion, a significantly
higher proportion of S phase cells was also seen after the sole
transfection of cyclin E (59 %) versus IE2 (40 %). These
findings further support the view that IE2-mediated cyclin E
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HCMV IE2 activation of cyclin E
induction drives IE2(195-579)-arrested cells across the G1/S
border.
Although S-phase entry in IE2(195-579)-expressing cells
could be rescued by raising cyclin E protein levels as
demonstrated, these cells did not progress through S phase
and into G2 (Fig. 5A, B). Instead, they remained in an early
S phase compartment, like cells arrested by IE2 [Fig. 5B,
compare the G0/G1 : early S ratios between cells expressing IE2 (1 : 1?2), IE2(195-579) plus cyclin E (1 : 2?3)
and IE2(195-579) plus IE2 (1 : 1?1)]. Again, the direct
overexpression of cyclin E appeared to empty the G1 compartment more rapidly than IE2. As demonstrated by
immunoblot analysis, we could exclude cross-regulation
of the IE2 mutant by cyclin E (Fig. 5C, left-hand panel,
lanes 3 and 4) and of cyclin E by the IE2 mutant (Fig. 5C,
right-hand panel, lanes 4 and 5) to account for the differences in BrdU incorporation.
Together these data suggest that IE2, by transcriptionally
up-regulating cyclin E, promotes G1/S transition before
cells finally arrest in early S phase. Thus, in addition to the
demonstrated cell-cycle-arrest function of IE2, we can also
assign an independent and genuine proliferative capacity
to IE2. This view is consistent with the observation of
the untimely induction of cyclin E in mimosine-arrested
IE2-expressing cells shown in Fig. 3.
IE2(195-579) retains the ability to arrest cells in
early S phase
The rescue experiments shown in Fig. 5 suggested that cells
finally arrest in early S phase after traversing the G1/S border.
In order to test further whether the cell-cycle arrest imposed
by the mutant form of IE2 is dominant over cdk activity, as
for IE2, we used nocodazole treatment in the experimental
set-up described in Fig. 5. Nocodazole is a spindle poison
and blocks cells at the beginning of mitosis. Under this
treatment, cycling control-transfected cells primarily accumulated in G2/M as expected (Fig. 6, plot 1). In contrast,
a large proportion of IE2- and IE2(195-579)-expressing cells
remained arrested in S and G1, respectively (Fig. 6, plots
2 and 3). Importantly, the same was true for cells overexpressing cyclin E in addition to the mutant form of IE2
(Fig. 6, plot 4). This demonstrated that, although cyclin E
expression drives cells arrested by IE2(195-579) in G1 into S
phase (see Fig. 5A, plot 4), the majority of these cells do not
progress further through S phase and do not enter G2. Cyclin
E overexpression alone first accelerated G1/S transition and
then slowed down S-phase progression (Fig. 6, plot 5). This
resulted in a high proportion of a pan-S cell population that
slowly emptied into G2, which is consistent with previously
published observations (Resnitzky et al., 1994; Spruck et al.,
1999). As expected, cells expressing both IE2 and the IE2
mutant remained to a great extent in the G1/S compartment, demonstrating the cell-cycle arrest suggested from the
findings shown in Fig. 5.
In conclusion, our data suggest that IE2 has the capacity
to block cells in G1 but this cell-cycle block is counteracted
by the intrinsic activity of IE2 to up-regulate cyclin E.
Consequently, cells enter S phase where they finally arrest
due to an as yet unknown mechanism, despite high cyclin
E-associated kinase activity.
DISCUSSION
We have presented experimental evidence to demonstrate
for the first time that the IE2-mediated up-regulation of
endogenous cyclin E is cell-cycle independent, i.e. not a
consequence of the cell-cycle state of IE2-expressing cells
but rather actively promoted by the viral regulator. This
activation of cyclin E is necessary to push IE2-expressing
cells into S phase where they finally arrest since, rather
surprisingly, an IE2 mutant defective in cyclin E activation
was shown to block cell-cycle progression in G1. Therefore,
IE2 appears to integrate two opposing cell-cycle regulatory
activities: a proliferating activity promoting G1/S progression via cyclin E activation and an anti-proliferating activity
leading to an early S-phase arrest, as previously shown
(Murphy et al., 2000; Wiebusch & Hagemeier, 2001).
The initial question of this study, whether IE2 is a genuine
activator of cyclin E or not, was answered by experiments
employing the plant amino acid mimosine to arrest
IE2-expressing cells in G1 before they entered into the IE2
block and the time window of periodic cyclin E induction
around the G1/S transition phase (Figs 1–3). Mimosine is
known to arrest cells 2–4 h before the onset of S phase by as
yet undefined mechanisms (Krude, 1999). Since cyclin E
does not start to accumulate until 2 h before S-phase entry
(Ekholm et al., 2001), our finding of low cyclin E expression
Fig. 6. IE2(195-579) retains the ability to arrest cells in early S. U373 cells were transfected exactly as described in Fig. 5. At
24 h after removal of the DNA precipitates, nocodazole (Noco) was added to the cells for 72 h. Cells were then harvested,
sorted, stained with propidium iodide and analysed by flow cytometry. Shown are DNA histograms of CD20-positive cells.
Minor fractions of endomitotic cells with a chromosome number higher than 4n were excluded from the analysis.
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57
L. Wiebusch and others
in mimosine-arrested cells is fully consistent with what
one would expect from published evidence. Therefore, the
fact that IE2 promotes up-regulation of cyclin E in these
cells without having any obvious effect on the mimosine
arrest itself demonstrates that IE2 can act as a cell-cycleindependent activator of cyclin E.
How does IE2 activate cyclin E? An earlier study (Bresnahan
et al., 1998) suggested a model based on transient transfection assays employing cyclin E promoter constructs
whereby IE2 directly transactivates cyclin E transcription.
Two of our experimental results supported such a view.
Firstly, IE2 expression led to an increase in endogenous
cyclin E mRNA abundance (Fig. 3A, Fig. 4A), and secondly,
the IE2 mutant IE2(195-579), which lacks the transcriptional activator function of IE2, was deficient in
cyclin E activation (Fig. 4; Wiebusch & Hagemeier, 2001).
Since the same mutant still contains the major binding
regions for pRb and p21WAF1,CIP1 (Fortunato et al., 1997,
2000; Sinclair et al., 2000), it seems unlikely that an IE2
interaction with one of these proteins causes the untimely
up-regulation of cyclin E that we observed in U373 cells.
Furthermore, p21WAF1,CIP1 expression levels are very low
in this p53-negative cell line, arguing against the functional relevance of IE2 targeting p21WAF1,CIP1 in these
cells. However, considering the fact that cyclin E–cdk2
activity is a convergence point for multiple negative control
mechanisms in G1/S (Bartek & Lukas, 2001), it cannot
be excluded that IE2 generally exerts additional functions
at a post-transcriptional level contributing to the constitutive activation of cyclin E. For instance, IE2 activates
cyclin E–cdk2, even in p16INK4a-arrested cells (unpublished observation), where cyclin E–cdk2 is normally held
inactive by p21WAF1,CIP1 that has been set free from cyclinD–cdk4 complexes (Jiang et al., 1998). This supports the
view that under conditions of high p21WAF1,CIP1 availability,
as in differentiated cells, an IE2-mediated inactivation of
this cdk inhibitor may become important for cyclin E
activation (Sinclair et al., 2000). Similarly, it should be
interesting to investigate whether IE2 has an influence
on SCFCDC4-mediated proteolysis of cyclin E, which is
normally initiated shortly after S-phase entry (Strohmaier
et al., 2001).
A number of reports have shown that premature activation of cyclin E–cdk2 by constitutive overexpression of
cyclin E induces S phase, even in growth-arrested cells
(Leone et al., 1999; Lukas et al., 1997). Accordingly, the
ability of IE2 to up-regulate cyclin E expression independent
of the cell-cycle state may be one explanation of how IE2
expression leads to S-phase entry in quiescent cells (Castillo
et al., 2000). Since cyclin E–cdk2 is a key regulator of the
pocket protein/E2F pathway, which controls the transcription of numerous genes coupled to proliferation and
DNA synthesis (DeGregori et al., 1995), it is feasible that
the activation of E2F-responsive gene expression observed
in IE2-transfected fibroblasts (Song & Stinski, 2002) is, in
addition to the direct transactivation of certain genes by IE2,
58
the consequence of more general IE2 functions, such as
cyclin E–cdk2 activation or pRb inactivation (Hagemeier
et al., 1994).
There is considerable evidence that cyclin E overexpression
contributes to the development of many types of human
cancer (Donnellan & Chetty, 1999). Therefore, the finding
that IE2 activates cyclin E constitutively also suggests a latent
oncogenic activity for this viral regulator. Given the
functional relationship between IE2 and viral oncogenes
such as E1A and SV40 large T antigen [namely the interactions with pRb (Hagemeier et al., 1994) and p53 (Speir
et al., 1994)] and regarding the mutagenic (Shen et al., 1997)
and anti-apoptotic (Yu & Alwine, 2002; Zhu et al., 1995)
capabilities of IE2, one could predict that a – still
undiscovered – IE2 species specifically lacking the cellcycle-arrest function could have considerable oncogenic
potential.
Intriguingly, the lack of cyclin E activating function in
IE2(195-579) revealed a true G1 arrest activity that is
inherent to IE2. This activity normally appears to be overcome by the full-length IE2-mediated induction of cyclin E,
which pushes these cells into early S phase where they finally
arrest (Fig. 7). This underlines the importance of the
IE2-mediated cyclin E up-regulation.
Although moderate cyclin E overexpression has been
reported to be associated with a delay in S-phase progression
(Resnitzky et al., 1994), the constitutive cyclin E activity
Fig. 7. Opposing cell-cycle activities of IE2: cyclin E activation
and cell-cycle arrest. A transactivation defective mutant of IE2
(IE2mut) reveals the inherent ability of IE2 to arrest cell-cycle
progression in G1 (A). This G1 arrest activity is normally counteracted in the IE2 wt protein by its ability to up-regulate cyclin E.
As a consequence, cells cross the G1/S border into S phase
where they finally arrest (B). The dotted line indicates that
IE2mut retains the ability to arrest cells in early S phase when
G1/S transition in these cells is forced by overexpression of
exogenous cyclin E (see Fig. 6).
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Journal of General Virology 84
HCMV IE2 activation of cyclin E
in IE2-expressing cells cannot explain the long-term
cell-cycle arrest in those cells. Even when we overexpressed
cyclin E by transient transfection in U373 cells leading
to cyclin E protein levels nearly two orders of magnitudes
higher than in normal cells (Fig. 5C), we could easily
distinguish S-phase prolongation in these cells from S-phase
arrest in IE2-transfected cells (Fig. 6). This clearly demonstrates that IE2 has an activity capable of arresting cells in
S phase that is independent of cyclin E expression. Is the
same activity of IE2 responsible for the IE2(195-579)mediated G1 arrest and the full-length IE2-induced block in
S phase? In principle, two alternative models could explain
this finding: either IE2 possesses two independent arrest
functions or the same arrest function is active in G1 and
S but sensitive to high cyclin E–cdk2 activity in G1.
Elucidation of the molecular mechanism(s) underlying
the IE2 arrest phenotype should decide which of these
models is valid.
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ACKNOWLEDGEMENTS
We thank Thomas Stamminger for the IE2-specific antibody. The work
was supported by grants from the Deutsche Forschungsgemeinschaft
to C. H. (SFB421 and GA167/6-2).
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