Journal of Cell Science 105, 113-122 (1993)
Printed in Great Britain © The Company of Biologists Limited 1993
113
Inhibition of transcription blocks cell cycle progression of NIH3T3
fibroblasts specifically in G1
Sabine Adolph, Sabine Brüsselbach and Rolf Müller*
Institut für Molekularbiologie und Tumorforschung (IMT), Philipps-Universität Marburg, Emil-Mannkopff-Strasse 2,
D-3550 Marburg, Germany
*Author for correspondence
SUMMARY
We have analysed the role of RNA polymerase II-dependent transcription in cell cycle progression. Time-lapse
video recording and cytogenetic analysis were used to
determine the sensitivity of NIH3T3 cells to the RNA
polymerase II inhibitor -amanitin at different stages of
the cell cycle. Our results show that -amanitin blocks
cells specifically in G1, irrespective of the concentration
within the range of 3 to 30 g/ml. This indicates that
transcription in G 1 is required to overcome a restriction
point located in this phase of the cell cycle. In agreement with this conclusion is the requirement for an
uninhibited protein synthesis during G1 progression. In
addition, the insensitivity of S-phase cells to RNA polymerase II inhibition suggests that the transcription of
genes thought to be normally induced during S/G2 is not
required for the completion of an ongoing cell cycle.
S/G2 progression was however clearly dependent on
protein synthesis. This suggests that cells exposed to amanitin are able to complete their cell cycle because
sufficiently high levels of mRNA are present in S/G2 due
to basal level transcription, or are left from preceding
cell cycles. It is therefore unlikely that transcriptional
regulation in S or G2 plays a crucial role in the control
of cell cycle progression in NIH3T3 cells.
INTRODUCTION
progression past the R point (Campisi et al., 1982; Pardee,
1989). Although candidate ‘R proteins’ have been identified (Croy and Pardee, 1983; Pardee, 1989), their function
in cell cycle progression remains unknown. It is tempting
to speculate, however, that the R point may be under the
control of the tumor suppressor gene product pRB, whose
negative regulatory function is abrogated by phosphorylation events occurring in late G1, and/or by the recently identified G1-specific cyclins and cdc2-related protein kinases
(for reviews see Hamel et al., 1992; Weinberg, 1991). The
R point in mammalian cells may thus be analogous to the
START point in S. cerevisiae not only because of its central role in cell cycle progression, but also with respect to
the underlying molecular control mechanisms (for reviews
see Herskowitz et al., 1991; Nasmyth et al., 1991).
Kinetic analyses performed by time-lapse cinematography of NIH3T3 cells have led to conclusions that can easily
be reconciled with the R point model described above
(Zetterberg and Larsson, 1985). In these experiments, the
position of cells at different points between mitosis and Sphase was determined and the sensitivity of these cells to
serum deprivation and the inhibition of protein synthesis
was analysed (by following the cells to the next mitosis).
These experiments clearly showed that cells younger than
When mitogens are withdrawn from cells or protein synthesis is partially blocked by an inhibitor of translation, they
will arrest in a state of quiescence (G0) or in the G1-phase
of the cell cycle, unless they have reached a restriction point
close to the beginning of S-phase, referred to as the R point
(Pardee, 1974; for reviews see Baserga, 1985; Norbury and
Nurse, 1992; Pardee, 1989; Soprano and Consenza, 1992).
This point has been localised in different systems anywhere
between 12 min and 3 h before S-phase entry, and its variability has been suggested to be largely responsible for the
variations in cell cycle length seen in individual cells
(Pardee, 1974; Rubin and Steiner, 1975; Yen and Pardee,
1978). Beyond the R point cells no longer require mitogens
and are less sensitive to protein synthesis inhibitors, suggesting a key role for this control point in mammalian cell
cycle progression. This conclusion is in agreement with the
observation that the R point is not, or only poorly, functional in transformed cells (Medrano and Pardee, 1980;
Pardee, 1974). The R point has been postulated to be controlled by (a) labile protein(s) with a half-life of 2-3 h,
called the ‘R protein’, which is synthesised earlier in G1
and whose expression at a critical level is a prerequisite for
Key words: cell cycle progression, α-amanitin, transcription, gene
expression, restriction point
114
S. Adolph, S. Brüsselbach and R. Müller
3.5 h were highly dependent on both the presence of mitogens and maximal protein synthesis, and that the length of
this period of G1 was more or less constant. The authors
termed this phase the post-mitotic or G1pm-phase. The cells
then entered the presynthetic or G1ps-phase, which was
independent of serum, but highly variable in length; some
cells entering S-phase almost immediately after leaving
G1pm, others remaining in G1ps for up to 10 h. It is very
likely that the R point discussed above defines the switch
from G1pm to G1ps. Very similar observations were also
made with human diploid fibroblasts (Larsson et al., 1989),
showing that the described behaviour is not a peculiarity of
NIH3T3 cells.
Upon mitogen stimulation of quiescent fibroblasts a large
number of genes are transcriptionally activated (for reviews
see Bravo, 1990; Hershman, 1991). The diversity of the
genomic response to mitogen stimulation is underscored by
the fact that genes associated with many different cellular
processes are activated, including secreted factors, proteins
involved in building or degrading the extracellular matrix,
metabolic enzymes and other proteins associated with
energy metabolism and transport processes, enzymes
involved in nucleotide and DNA synthesis, proteins
involved in signal transduction, and transcription factors. In
addition, a number of cell cycle genes, most notably of the
cdk and cyclin families, have been identified in recent years
(Lew and Reed, 1992; Meyerson et al., 1992). Many of
these genes and their products are activated at specific
stages of the cell cycle after mitogen stimulation of quiescent cells or after the release from metabolic blocks. These
genes include those encoding the S- and M-phase-specific
cyclin A and the mitotic B-type cyclins (Pines and Hunter,
1989, 1990), the cdc2 protein kinase (Lee and Nurse, 1987)
and the cdc25 protein phosphatase (Sadhu et al., 1990),
whose function is indispensable for S and/or G2 progression (Riabowol et al., 1989; Walker and Maller, 1991;
Girard et al., 1991; Hoffmann et al., 1993; for a review on
the role of these genes in S. pombe see Forsburg and Nurse,
1991). Very little is known, however, about the transcription of these genes and their role during cell cycle progression in normally cycling, non-synchronised mammalian
cells. It thus remains unclear which stages of the cell cycle
are dependent on the expression of specific sets of genes.
This is an important issue since it is likely that such genes
encode the proteins required to overcome the restriction
points in the cell cycle. As a first step in addressing these
open questions we have determined, by time-lapse video
recording and cytogenetic analysis of bromodeoxyuridinesubstituted chromosomes, the sensitivity of NIH3T3 cells
at different stages of the cell cycle to the RNA polymerase
II inhibitor α-amanitin (Wieland and Faulstich, 1991). We
find that only the G1-phase is sensitive to inhibition by αamanitin, which is in agreement with the observation that
G1 is also the most sensitive phase with respect to the inhibition of protein biosynthesis by cycloheximide. Our results
also suggest that an unimpaired transcription of many cell
cycle genes normally induced during S/G2 is not required
for the completion of an ongoing cell cycle. These findings
place transcriptionally controlled restriction points predominantly or even exclusively in G1.
MATERIALS AND METHODS
Materials
Cycloheximide and α-amanitin were obtained from Sigma. The
antibody for BrdU detection was purchased from Partec and used
at a 1:500 dilution. The antibody to detect c-Fos protein has been
described elsewhere (Verrier et al., 1986) and was used at a 1:30
dilution. The second antibodies for indirect immunohistochemical
staining were either a Cy3- or a HRP-coupled rabbit anti-mouse
IgG (Dianova).
Time-lapse video recording (TLV)
NIH3T3 cells were cultured under standard condition (5% CO2,
37°C) in DMEM with 10% FCS. For video recording, about 105
cells were seeded into a 25 cm2 plastic dish, cultured for one day
in a CO 2 incubator and then transferred to an inverted microscope
(Zeiss, Axiovert 35) equipped with a temperature- and CO2-controlled stage. Cell proliferation was recorded by a CCD camera
(CF 15 MC, Kappa) coupled to a video recorder (AG 7350, Panasonic) for 40 to 60 h. After observing the cells for 18 to 24 h, the
inhibitors (α-amanitin or cycloheximide) were applied and cells
were recorded for another 24 to 30 h. At least 100 individual cells
per film were followed from mitosis to mitosis and the intermitotic times (tc) were determined.
Data analysis and representation
To analyse the distribution of cell cycle lengths of untreated cells
we chose the method of Shields and Smith (Shields and Smith,
1977). These authors defined the proportion of cells which have
an intermitotic time greater or equal to t as α(t) and plotted logα
against the intermitotic time (tc). Since there is a minimum intermitotic time (TB), no cells will divide in a period less then TB,
leading to the corresponding value of α=100%. According to the
transition probability model of cell cycle progression, a theoretical plot of logα against time would have two components, a lag
phase equal to TB, and an exponential phase which will be linear
in the logα plot. This presentation of the data proved especially
useful to compare the tc values of cells hit by the drug during an
ongoing cell cycle. In addition, the cells’ age at the time of drug
application (tx) as well as the time they needed to finish a proper
mitosis after drug application (ty) were listed and plotted against
each other.
Cytogenetic techniques
Cells were grown directly on microscope slides in Quadriperm
dishes (Heraeus) and metaphase chromosomes were prepared in
situ by the standard technique. To determine the duration of the
G2/M-phase, tG∑, bromodeoxyuridine (BrdU; 20 µg/ml) was
applied for increasing times to the cell culture (from 1 to 6 h)
before chromosome preparation. The incorporated BrdU was
detected by the anti-BrdU antibody technique as described (Vogel
et al., 1986). The frequency of unlabelled metaphase plates gives
the frequency of cells which were in G 2 at the time of BrdU application. To determine the duration of G2/M and S, tS+G∑, BrdU was
applied for increasing times to the cell cultures from 7 to 15 h
before chromosome preparation. The incorporated BrdU was visualised by fluorescence microscopy either directly by acridine
orange staining (Dutrillaux et al., 1973) or by Hoechst staining
(Latt, 1973). Cells which incorporated BrdU over their total length
of S will have pale, weakly stained chromosomes due to quenching of fluorescence emission by the incorporated BrdU. Those
cells which started to replicate their DNA before the application
of BrdU will show brilliant, fluorescing bands on their chromosomes. The frequency of metaphase plates with completely pale
chromosomes corresponds to the frequency of G1 cells at the time
of BrdU application. The proliferation of cells in the presence of
Restriction of cell cycle progression
the inhibitor was also analysed by replication analysis of
metaphase chromosomes. Cells were seeded onto slides in
Quadriperm dishes and labelled for the first S-phase with BrdU
(25 h). Thereafter, medium was changed and supplemented with
thymidine (10 µM) and the inhibitors. About 2 days later, chromosomes were harvested in situ as described above and the incorporated BrdU was detected by the antibody technique. The segregation of the original BrdU-substituted chromatids in the
thymidine makes it possible to calculate the number of cycles (Sphases) the cells completed during drug application.
Measurement of protein biosynthesis
Cells were pulse-labelled for 2 h in medium containing
[3H]isoleucine (NEN, 2 µCi/ml), 10% of the normal concentration of unlabelled isoleucine and different concentrations of cycloheximide. The incorporated [3H]isoleucine was determined in the
acid-insoluble material by placing it in scintillation fluid and
counting in a Beckman scintillation counter. Radioactivity of
untreated cells was taken as 100%.
115
Table 1. Cell cycle parameters of untreated NIH-3T3
cells
tc (h)
Frequency
(%)*
tG∑/M
(h)†
t S+G∑/M
(h)†
tS
(h)‡
tG œ
(h)‡
≥10
≥11
≥12
≥13
≥14
≥15
≥16
≥18
≥20
≥30
97
90
80
62
45
28
20
10
5
2
1.7-2.4
2.0-2.7
2.4-3.4
2.7-3.5
3.2-4.0
3.6-5.2
3.8-5.8
4.2-7.0
4.9-8.0
5.7-8.7
7.0-8.2
7.2-9.0
7.8-9.2
8.1-9.6
8.8-10.0
9.4-10.8
10.0-11.2
11.8-12.2
12.2-13.0
12.8-13.5
5.3-5.8
5.2-6.3
5.4-5.8
5.4-6.1
5.6-6.0
5.8-5.6
6.2-5.4
7.6-5.2
7.3-5.0
7.1-4.8
1.8-3.0
2.0-3.8
2.8-4.2
3.4-4.9
4.0-5.2
4.2-5.6
4.8-6.0
5.8-6.2
7.0-7.8
≥7.0
*Determined by time-lapse analysis for t c and by cytogenetic analysis
for tG∑/M or tS+G ∑/M.
†Determined by cytogenetic analysis of metaphase chromosomes after
BrdU exposure.
Values from several experiments were combined.
‡Calculated as: tS=tS+G∑/M−tG∑ /M and tGœ=tc −tS+G ∑/M.
Measurement of mRNA synthesis
The inhibition of RNA polymerase II transcription by α-amanitin
was analysed by c-Fos induction in newly stimulated 3T3 cells
after serum starvation. At various times before stimulation, serumstarved cells were treated with α-amanitin (10 µg/ml), and 1 h
after serum stimulation the level of Fos protein in individual cells
was determined immunohistochemically (Verrier et al., 1986) and
quantitated as described (Quantimet, Leica; Lucibello et al., 1993).
RESULTS
Cell cycle parameters of untreated cells
We first determined the intermitotic times (tc) of more than
400 untreated cells from several experiments by time-lapse
video recording (TLV) to establish the normal distribution
of the cell cycle lengths in the population of NIH3T3 cells
used in the present study. The shortest tc determined was
8.5 h (in 0.5% of the cells). Half of the population needed
13 to 14 h to complete a cell cycle, and 90% of the cells
showed tc values of 10 to 18 h. Only 5% had a cell cycle
longer than 21 h, the longest found being 34 h.
After culturing the cells for different times in BrdU, cytogenetic analysis of the metaphase plates enabled us to determine the frequency of BrdU-unlabelled metaphase plates.
In addition, we established the fraction of cells that replicated their DNA in BrdU over the total length of S-phase.
These two frequencies correspond to the duration of G2/M
(tG∑/M) and S+G2/M (tS+G∑/M), respectively. In several
experiments, about 100 metaphase plates were analysed for
each BrdU exposure. These results are presented in Table
1. The shortest tG∑/M was 1.7 h. After 3.5 h exposure, 50%
of the metaphase plates were BrdU-labelled, and after 8 h
the corresponding value was 95%. The shortest tS+G∑/M was
7 h, and no cell, out of 200 metaphases, needed longer than
14 h to complete S+G2/M. Nine to ten hours were necessary for 50% of the cells to pass through S+G2/M. Therefore, the average value for tG∑/M is 3.5 h and for tS+G∑/M
9.5 h.
Both methods, TLV and the cytogenetic analysis, are
used to study individual cells. The former technique yields
exact tc values without any information about the duration
of the different cell cycle phases; on the other hand, the
cytogenetic analysis gives the tG∑/M and tS+G∑/M values without reference to the corresponding tc values. The duration
of G1 (tGœ) can only be estimated as tc−tS+G∑/M. The two
data sets were therefore combined assuming that tc values
and the lengths of cell cycle phases correlate, i.e. that the
cells with the shortest tc also have the shortest tG∑ and tS+G∑
values, and conversely, that long tc, tG∑ and tS+G∑ values
are also linked. Table 1 summarises the tc, tG∑ and tS+G∑
values measured as described above and the corresponding
calculated tS and tGœ values. This way the average tS and
tGœ values were found to be 6 h and 4.5 h, respectively. The
minimal time for G1 can be calculated to be 2 h.
One goal of our investigation was to inhibit RNA polymerase II transcription by α-amanitin and to identify phases
in the cell cycle that differ in their sensitivity to the drug.
Therefore, our interest focused on those cells that were hit
during an ongoing cell cycle. The age of a given cell at the
time of drug application (tx) as well as the time that cell
needed after drug application to complete the cell cycle (ty)
were determined by TLV. In addition, we sought to establish a technique for assigning a given tx value to a cell cycle
phase. For this purpose, tx and ty values were plotted against
each other. Fig. 1 shows such a typical tx/ty-plot for mocktreated cells. Each line represents cells showing the same
cell cycle length tc=tx+ty, hit at a cell age of x hours and
completing their cell cycle y hours after inhibitor application. The shaded area covers the expected tc values from
10 to 18 h, corresponding to 90% of untreated cells. Since
ty also corresponds to the time of BrdU application in the
cytogenetic analysis it is possible to include the distribution of G2, S, and G1 in this plot. We incorporated tG∑/M
and tS+G∑/M from Table 1 as ty values into this plot and calculated the corresponding tx values as tc−ty. This enabled
us to estimate the position of individual cells in the cell
cycle at the time of drug application. Any cell dividing in
the left-most field (left of the upper dotted line) was likely
to be in G1 at the time of the treatment. Any cell dividing
in the field between the dotted lines was likely to be in S,
and any cell dividing in the right-most field (right of the
lower dotted line) should have been in G2 at the time of
116
S. Adolph, S. Brüsselbach and R. Müller
Fig. 1. Typical tx/ty-plot of mock-treated cells. tx values were
determined by TLV and are equivalent to the cell age at treatment.
The ty values represent the time cells needed to complete the cell
cycle after drug application as determined by TLV. In addition, ty
values correspond to the time of BrdU exposure prior to
cytogenetic analysis. The total cell cycle length of a given cell is
equal to tc=tx+ty. Cells with the same cell cycle length (tc) are
represented by lines. To determine the borders of G1/S and S/G2
the values of tS+G∑/M (j) and tG∑/M (+) from Table 1 were
included as ty values. The corresponding tx values were calculated
by correlating the frequency of tc values with the frequencies of
tS+G∑/M and tG∑/M. The broad dotted lines represent the
approximate border of G1/S and S/G2. The shaded field marks the
area of tc values from 10 to 18 h, corresponding to 90% untreated
cells.
drug application. The validity of this interpretation was confirmed by experiments where the same cells observed by
TLV were analysed for incorporation of BrdU (data not
shown; also see Fig. 7). A cell with the same age at treatment but a different ty value can either have a normal but
exceptionally high tc, or might have a prolonged ty value
caused by the drug. Since it is impossible to decide this by
analysing single cells, we used the following definition to
identify a sensitive cell cycle phase: none, or only a few
cells of the same age at treatment, divide in the expected
field of the tx/ty-plot, and the fraction of cells that reach the
end of recording without division is significantly increased.
Influence of -amanitin and cycloheximide on
proliferation, transcription and protein
biosynthesis
To determine the kinetics of α-amanitin on RNA polymerase II-dependent transcription in NIH3T3 cells, we
measured the level of c-Fos in individual cells one hour
after stimulation with serum following different times of αamanitin pretreatment (Fig. 2a). The c-fos gene was chosen
because its transcription is rapidly induced by mitogens and
its mRNA is also rapidly translated: maximum protein
levels are reached 1 hour after serum stimulation of
Fig. 2. Inhibition of c-Fos induction by α-amanitin and total
protein synthesis by cycloheximide. (a) Serum-deprived NIH3T3
cells were stimulated with 10% FCS after different times of αamanitin pretreatment (10 µg/ml). Cells were fixed (‘harvest’) 1 h
after stimulation and stained for Fos expression by indirect
immunofluorescence. The fluorescence intensity in 300 individual
cells was measured by digital image analysis (Quantimet, Leica)
and average fluorescence levels were plotted against time.
(b) NIH3T3 cells growing in DMEM plus 10% FCS were
metabolically labelled with [3H]isoleucine in the presence of
different concentrations of cycloheximide. The radioactivity in
acid-precipitated material was determined and compared to the
incorporation of [3H]isoleucine into cells in the absence of
cycloheximide (100%).
NIH3T3 cells. Addition of 10.0 µg/ml α-amanitin 3 h prior
to serum stimulation was sufficient to block fos expression
by ≥90% and half maximum inhibition was seen after about
80 min. Evaluation of the immunofluorescence analysis also
showed that α-amanitin affected c-fos expression in all cells
of the population to a similar extent (data not shown). This
suggests that cells dividing after ≥2 h of exposure to the
drug should also be affected in their RNA polymerase II
transcription.
We then investigated the influence of α-amanitin on the
proliferation of NIH3T3 cells to be able to choose appro-
Restriction of cell cycle progression
Table 2. Proliferation of cells after treatment with amanitin and cycloheximide for 2 days
Fraction of cells that passed through
n number of S-phases after drug application
Untreated
α-Amanitin
1.0
3.0
9.0
15.0
Cycloheximide
0.01
0.05
0.1
0.5
1.0
Number of
metaphases
n≥4
n=3
n=2
n=1
n=0
48
36
16
0
0
185
2
0
0
0
21
7
0
0
64
36
0
0
4
57
0
0
0
0
100
0
55
14
<5
none
13
0
0
0
0
47
23
0
0
0
37
47
0
0
0
10
30
100
0
0
0
0
0
100
0
136
91
14
<5
none
= Cumulative fraction of cells (%)
Treatment
(concn in µg/ml)
Freshly seeded cells were pulse-labelled for the first S-phase (25 h) with
BrdU, then the medium was changed and supplemented with thymidine
plus either α-amanitin or cycloheximide at the concentrations indicated
in the Table. Two days later (51 h for α-amanitin and 43 h for
cycloheximide, respectively) metaphase chromosomes were prepared and
the incorporated BrdU was detected immunohistochemically. Segregation
of the BrdU-labelled chromatids allowed us to calculate the number of
replication cycles in thymidine.
117
100
50
20
CHX1.0
10
5
2
C
CHX0.02
1
0
5
10
15
20
25
30
35
ama30
40
45
50
tc (h)
priate concentrations of the drugs for subsequent experiments. The proliferation of cells in the presence of α-amanitin as analysed by the cytogenetic approach clearly
depended on the concentration used (Table 2). Almost half
of the untreated cells replicated at least 4 times in 51 h,
and 13% progressed through only 2 consecutive S-phases.
At a concentration of 1.0 µg/ml α-amanitin most of the
cells progressed through only 2 consecutive S-phases and
9.0 µg/ml were sufficient to prevent completion of a subsequent cell cycle. The mitotic index was strongly
decreased for concentrations ≥3.0 µg/ml. No metaphase
spreads at all could be found at 15.0 µg/ml, and none of
the nuclei (≥2000) showed incorporation of a visible
amount of thymidine during the subsequent S-phase. This
led to the conclusion that 15 µg/ml α-amanitin were sufficient to block entry into the subsequent S-phase.
Since one aspect of the present study was to compare the
role of protein and RNA synthesis in cell cycle progression, the protein biosynthesis inhibitor cycloheximide was
included in all subsequent experiments. The proliferation of
cells in the presence of cycloheximide was analysed in the
same way as described above for α-amanitin. A concentration of 0.01 µg/ml cycloheximide (corresponding to
about 40% reduction of protein biosynthesis; Fig. 2b) had
only a marginal effect on cell proliferation (Table 2). Application of 0.05 µg/ml cycloheximide (giving about 60%
reduction of protein biosynthesis) for 2 days reduced the
number of consecutive S-phases to 2 and 0.5 µg/ml (reduction of 85% in protein synthesis) were sufficient to prevent
the completion of any further S-phase during the 2 days of
the analysis. The mitotic index decreased clearly after application of 0.1 µg/ml or more. By these criteria 0.5 and 1.0
µg/ml cycloheximide have a comparable effect on cell proliferation to 9.0 and 15.0 µg/ml α-amanitin, respectively.
Fig. 3. Distribution of tc values of cells exposed to cycloheximide
or α-amanitin. Only those cells were included that started their
cell cycle before the start of treatment. The fraction of cells (α)
with a tc value equal to or bigger than a given value (on the
ordinate) is plotted as logα against tc. Since there is a minimum
intermitotic time (TB), no cells will divide in a period less then TB
(marked by vertical lines at the top),giving a value of α=100%.
The small dots represent cells that reached the end of recording
without division. C, untreated control cells; CHX 0.02 and CHX
1.0, cells treated with 0.02 and 1.0 µg/ml cycloheximide,
respectively; ama30, cells treated with 30 µg/ml α-amanitin.
Influence of -amanitin and cycloheximide on tc
To compare the effect of the drugs on tc of an ongoing cell
cycle we plotted the distribution of tc as α-curves (Fig. 3)
as described in the Materials and Methods. Even the lowest
concentration of cycloheximide (0.02 µg/ml, reducing protein synthesis by 50%) resulted in a clear increase in tc.
This was even more pronounced at 1.0 µg/ml, which led to
a reduction in protein synthesis to 10%. In contrast, all concentrations of α-amanitin (3.0, 6.0, 30.0 µg/ml) gave the
same picture; about 50% showed a tc similar to cells treated
with 0.02 µg/ml cycloheximide, whereas the other 50%
gave rise to clearly increased tc values. The small dots in
Fig. 3, which represent cells that reached the end of recording without division, indicate that a large proportion of cells
treated with 30 µg/ml α-amanitin had cell cycle times that
might even be much longer.
Stage-specific inhibition of cell cycle progression
by -amanitin and cycloheximide
The influence of the cell age at treatment, tx, on the
inhibitory effect of the drug on an ongoing cell cycle was
118
S. Adolph, S. Brüsselbach and R. Müller
40
40
a
a
control
30
-amanitin, 3 µg/ml
30
≥24 h
20
20
G1
10
10
0
0
0
b
10
20
40
cycloheximide, 0.02 µg/ml
≥29 h
30
0
20
ty = Time until next mitosis (h)
ty = Time until next mitosis (h)
40
10
20
10
0
b
-amanitin, 6 µg/ml
30
≥24 h
20
G1
10
0
0
50
10
c
20
0
40
cycloheximide, 1 µg/ml
10
c
20
-amanitin, 30 µg/ml
≥42 h
40
≥24 h
30
30
20
20
G1
10
G1+S
10
0
0
10
20
tx = Cell age at treatment (h)
Fig. 4. Relationship between cell age (tx) and the time (ty)
required to complete the cell cycle after cycloheximide
application. (a) Untreated control cells; (b) cells treated with 0.02
µg cycloheximide/ml; (c) cells treated with 1 µg
cycloheximide/ml. Each solid square represents a dividing cell
that was hit at the age of x hours and needed y hours to complete
its cell cycle. The open squares represent cells that completed
their cell cycle before treatment. The shaded field marks the area
of tc values from 10 to 20 h, i.e. where ≥90% of the untreated cells
are found. The boxed area indicates cells that reached the end of
recording without division. The area expected for cells that were
in G1 or S at the time of exposure to the drug is marked in (c).
0
0
10
20
tx = Cell age at treatment (h)
Fig. 5. Relationship between cell age (tx) and the time (ty)
required to complete the cell cycle after α-amanitin application.
(a) 3 µg α-amanitin/ml; (b) 6 µg α-amanitin/ml; (c) 30 µg αamanitin/ml. For details see legend to Fig. 4.
determined by TLV. The results are presented in Figs 4 and
5 as tx/ty plots, as described above. Untreated control cells
divided in the expected area (compare Fig. 1) with only a
few cells showing tc values >20 h (Fig. 4a). The behaviour
of cells treated with a concentration of cycloheximide that
reduced protein synthesis to about 50% (0.02 µg/ml) is
shown in Fig. 4b. Many of the cells divided in the area
expected for untreated cells, but a number of cells showed
an increased ty, and thus tc. Increasing the cycloheximide
Restriction of cell cycle progression
Fig. 6. Fraction of non-dividing cells in different age groups. The
percentage of α-amanitin treated cells that reached the end of
recording without dividing is plotted against the cell age at
treatment. Each group of cells comprises a 2-hour interval. The
three experiments shown in Fig. 5 were combined for the plot
shown above.
to a concentration where 90% of the protein biosynthesis
was inhibited caused a complete depletion of cells in the
G1 and S fields; all these cells reached the end of the video
without dividing (Fig. 4c). Some of the cells with a cell age
at treatment of 9 to 14 h showed a significantly prolonged
total cell cycle time. Other cells of the same age class either
did not divide at all or completed their cell cycle with a
proper mitosis in the expected time. Such cells were probably in G2 at the time of treatment.
The three experiments with α-amanitin (3.0, 6.0, 30.0
µg/ml) gave very different tx/ty plots when compared to
cycloheximide. In addition, the results were largely independent of the concentration used (Fig. 5). At all concentrations many cells divided in the expected area of the tx/ty
plots. Only the population of cells at the age of 0 to 2 h
was highly affected, pointing to a block in G1. In agreement with this conclusion, none of the cells shown in Fig.
5 divided a second time in the presence of the drug.
A number of cells that were older than 2 h at the time
of treatment reached the end of the video without dividing
(Fig. 6). From the TLV data obtained so far it was not possible to decide in which cell cycle phase these cells were
hit. To answer this question we analysed, by TLV, cells
that were pulse-labelled for one hour with BrdU before the
addition of α-amanitin (10.0 µg/ml). After recording, the
incorporated BrdU was detected immunohistochemically.
All cells that reached the end of recording without dividing were not labelled by BrdU (Fig. 7), which means that
these cells were either in G1 or in G2. However, 9 out of
10 cells (boxed area in Fig. 7) were younger than the minimum time required for progression through G1 and S
(approx. 8 h; see Table 1). Therefore, these cells were in
G1 when exposed to BrdU and α-amanitin. The stage of
119
Fig. 7. Simultaneous incorporation of BrdU and TLV. The
experiment was carried out as described in Figs 4 and 5, except
that the cells were exposed to BrdU for 1 h directly before
exposure to α-amanitin. Incorporation of BrdU was determined
after recording by immunostaining. Solid squares represent BrdUlabelled cells, open squares are unlabelled cells. Gray lines
indicate the approximate positions of the borders between the
G1/S and S/G2 compartments, as explained in Fig. 1.
the 10th cell, hit at the age of approx. 14 h, could not, however, be determined by this approach. Similar findings were
made in a number of other experiments. In total, 61 cells
were counted that reached the end of recording without
division; 56 out of these cells were younger than 8 h. These
observations strongly support our conclusion that G1 is by
far the most sensitive phase with respect to the action of
α-amanitin. In addition, the data in Fig. 7 indicate a normalsized pool of S-phase cells, suggesting that α-amanitin
treatment did not lead to a prolongation of S/G2 progression.
Finally, we wished to confirm the insensitivity of S/G2
cells to the inhibitory effect of α-amanitin by a second,
independent, approach. We determined, by the analysis of
metaphase chromosomes, the effect of α-amanitin and
cycloheximide on the progression through S and G2. Applying BrdU for different intervals before chromosome preparation allowed us to determine the frequencies of cells in
S or G2 at the time of BrdU addition. Table 3 summarises
these results. α-Amanitin had no effect on the frequencies
of S and G2 cells. The small differences seen in Table 3
clearly fall within the range of experimental variation. On
the other hand, even a low concentration of cycloheximide
(0.05 µg/ml) led to a decrease in the number of S-phase
cells, and at the higher cycloheximide concentration (0.1
µg/ml) the pool of G 2 cells was also clearly enlarged. These
independent experiments demonstrate again that inhibition
of protein biosynthesis by approximately 50% already has
a prolonging effect on S/G2. In contrast, even a high αamanitin concentration showed no significant effect on the
progression through S and G2.
120
S. Adolph, S. Brüsselbach and R. Müller
Table 3. Influence of -amanitin and cycloheximide on
the frequencies of S and G2 cells as determined by
cytogenetic analysis of metaphase chromosomes
Treatment
(µg/ml) (h)
S-phase
Number of
of exp.
GLL G 2-phase metaphases
BrdU
(h)
G
GL
2
3
4
5
6
7
13
33
4
24
35
35
2
23
44
31
26
98
73
24
16
8
7
7
10
5
6
826
788
899
969
395
3
4
5
6
10
1
27
44
29
33
22
70
34
24
1
1
1
109
110
123
5
6
5
6
3
-
23
33
1
7
34
40
23
26
43
23
77
67
1
1
1
1
69
106
151
134
Untreated
α-Amanitin
10.0
3
4
5
Cycloheximide
0.05 6
6
0.1
6
6
Cells were seeded on slides and incubated with BrdU and the drugs as
indicated. BrdU incorporation was analysed by the antibody technique.
Unlabelled chromosomes indicated that the cells were in G2 at the time of
BrdU application. Labelled cells were grouped according to their
chromosomal replication banding pattern in three groups. G, GL and GLL
represent normal, late and very late G-bands, respectively. Normal Gbands correspond to the standard idiogramme of G-banded chromosomes
(Evans, 1989).
DISCUSSION
This study was undertaken to investigate the role of transcriptional regulation in the control of cell cycle progression. To this end, we have performed time-lapse video
recording (TLV) and cytogenetic analyses to determine the
sensitivity of NIH3T3 cells at different stages in the cell
cycle to the RNA polymerase II inhibitor α-amanitin. The
results of this study were also compared to observations
made with the protein biosynthesis inhibitor cycloheximide
in analogous experiments. The most intriguing finding of
this study is the observation that G1 is highly sensitive to
both types of inhibitors, while S and G2 progression is
blocked to a significant extent only by higher concentrations of cycloheximide.
In the first part of this investigation, we determined the
effect of α-amanitin and cycloheximide on intermitotic
times, i.e. the total cell cycle lengths (tc). The results of
these experiments are shown in Fig. 3. It is obvious that in
both cases the fraction of cells with higher tc values was
increased. In the case of α-amanitin, at least 3 different
populations of cells were detectable: cells with largely
normal tc values (top part of the curve); cells with greatly
increased intermitotic times; and cells that had not divided
by the end of recording. The important question of where
in the cell cycle these cells were blocked was analysed in
the experiment shown in Fig. 5 (compare to control in Fig.
4a). Here, the influence of cell age on the inhibitory effect
of α-amanitin was studied. The data indicate that the majority of the cells at an age of <8 h were clearly affected, particularly those cells at ≤2 h post-mitosis. This is evident
from the fact that the respective area in the plots was
depleted of cells younger than 2 h, and that the fraction of
undivided cells dramatically increased (boxed areas in Fig.
5). This is different from the effects seen with cycloheximide, which at a low concentration also showed an effect
on young cells (<5 h), but at higher concentrations also led
to a dramatic prolongation and block of cell cycle progression in older cells.
The next important task was then to determine in which
cell cycle phases the inhibitors exerted their effects, i.e. to
relate the cell’s age to its position in the cell division cycle.
For this purpose we combined the results obtained by two
different techniques, the TLV, which allowed for the precise measurement of total cell cycle times, and a cytogenetic approach, which gave us accurate information about
the length of S+G2 and G2. Based on the assumption that
a prolonged cell cycle time can be correlated with a longer
duration of the different cell cycle phases we established
the data evaluation plot shown in Fig. 1. The validity of
this technique for relating cell age and the position in the
cell cycle was tested by identifying S-phase cells through
BrdU labelling within a cell population followed by TLV
(Fig. 7 and data not shown). In the experiment shown in
Fig. 7, 16 cells were BrdU-labelled and 14 out of these
were found in the predicted area. Conversely, 31 cells
remained unlabelled and 29 out of these appeared in the
correct area of the plot in Fig. 7. Similar results were
obtained in other experiments (data not shown). The fraction of ‘incorrectly’ assigned cells was therefore 2/16
(12.5%) and 2/31 (6.4%), respectively, i.e. there are some
cells with a long G1 and a short S/G2-phase The value of
approximately 90% correctly positioned cells can be considered sufficiently high to allow a conclusive interpretation of the data. On this basis we were able to assign cell
cycle phases to the effects seen in Fig. 5; α-amanitin has
a dramatic effect on cells in early G1 (≤2 h post-mitosis),
but clearly also affects cells at later stages in G1, while cells
in S and G2 are largely resistant to the inhibitory effect of
α-amanitin. This is supported by the data in Fig. 6, where
the number of cells blocked in cell cycle progression by αamanitin was plotted against the cell age. These data also
show the sensitivity of the cells to α-amanitin within the
first 8 h post-mitosis, in that >90% of the cells that did not
divide until the end of recording were 8 h or younger.
However, the conclusive interpretation of the data
required the clarification of another question, i.e. the kinetics of α-amanitin action and a quantitative assessment of
its effect on RNA polymerase II-dependent transcription.
Unfortunately, such a measurement is not as easy as in the
case of protein biosynthesis inhibitors, because radioactive
RNA precursors are not exclusively incorporated into
mRNA by RNA polymerase II, but to much higher extent
into rRNA and tRNA by the other RNA polymerases. We
therefore decided to measure the effect of α-amanitin on
the transcription of a single gene. This was possible, since
α-amanitin should affect the transcription of all genes to a
similar extent due to its inhibition of RNA polymerase II
rather than of a gene specific factor. For this purpose we
chose the c-fos gene whose transcription is induced within
minutes after mitogen stimulation. In addition, the c-fos
gene product, c-Fos, is rapidly synthesised as well, so that
Restriction of cell cycle progression
gene induction and its inhibition by α-amanitin could be
followed at the single cell level. The results of this experiment (Fig. 2a) showed that α-amanitin at a dose of 10
µg/ml inhibits c-Fos induction by 50% within 80 min. The
kinetics of α-amanitin mediated inhibition on RNA polymerase II are therefore fast enough to conclude that the
majority of the cells were in G1 at the time of α-amanitin
action. In addition, in none of the experiments we performed did we find a single cell in S-phase (identified by
BrdU labelling) whose cell cycle progression was visibly
prolonged or even blocked by α-amanitin.
One of the most surprising results of the present study
was the finding that α-amanitin has no detectable effect on
cells in S or G2. This conclusion is based on the evaluation of a large number of cells by TLV (375 cells) and was
fully confirmed by cytogenetic analyses (Table 3). This
indicates that the transcription of a large number of genes,
including many cell cycle genes, can be blocked by >90%
without any significant effect on the completion of an ongoing cell cycle. This is surprising as such genes include, for
example, cdc2 and cyclin B, which have been reported to
be induced in late S and G2 (Dalton, 1992; Pines and
Hunter, 1989). The question thus arises how these cells
manage to complete their cell cycle with a proper mitosis.
Our own and other authors’ observations have shown that
cells need an active protein synthesis machinery to proceed
through S and G2 (Zetterberg and Larsson, 1985; Figs 3
and 4). This leads to the conclusion that the α-amanitin
treated cells must contain sufficiently high levels of mRNA
to be able to synthesise the required amounts of protein.
Several possibilities exist to explain this situation: (i) the
basal transcription of these genes may be high enough to
allow for the accumulation of sufficiently high levels before
entry into S or G2; (ii) the mRNA may not be subject to a
rapid and complete degradation so that enough RNA is left
from preceding cell cycles; and (iii) the genes are not subject to stringent transcriptional regulation during a normal
cell cycle. It has to be borne in mind that all experiments
addressing cell cycle phase-specific expression of cyclins
and cdc2/cdk genes have been performed with cells that
were either stimulated in a state of quiescence, chemically
synchronised or sorted by elutriation. All these procedures
represent artificial conditions or some kind of unphysiological stress that may affect the pattern of gene expression.
To clarify these questions it is therefore necessary to
analyse the expression of these genes in normally cycling
‘untouched’ cells, e.g. by combining TLV and in situ staining procedures. Such experiments are in progress in our laboratory for a number of selected cell cycle genes.
Another interesting question concerns the action of αamanitin in the G1-phase. It seems clear from our results
that the only sensitive phase of the cell cycle is G1, suggesting that α-amanitin exposure interferes with progression across the R point (Pardee, 1989; see Introduction for
details). Which, however, are the genes that are crucial for
the progression into S and are blocked in α-amanitin treated
cells? This question is not easy to answer because not much
is known about G1-specific genes. Most genes induced
before entry into S are mitogen-inducible genes whose
expression is elevated upon stimulation of quiescent cells
(Bravo, 1990; Hershman, 1991). This however does not
121
mean that these genes behave in a similar way or even have
a function during G1 progression in normally cycling cells.
One example of this kind is the c-fos gene, which is induced
to very high levels by mitogen stimulation but which is
expressed only at very low levels throughout the cell cycle
(Bravo et al., 1986) and apparently has no crucial function
in normally cycling cells (Kovary and Bravo, 1991, 1992).
One might, however, speculate that genes whose transcription is not regulated to a significant extent during G1→S
progression, but whose transcripts have a short half-life are
the critical targets for the action of α-amanitin. Among such
genes may be the recently identified cyclins C, D and E
(Lew and Reed, 1992), members of the cdk family (Meyerson et al., 1992) and the transcription factor E2F (Nevins,
1992). Future experiments making use of anti-sense RNA
approaches or targeted gene disruption by homologous
recombination will have to clarify the relevance of these
genes for the progression of cells through the G1-phase and
their entry into S.
This work was supported by the Deutsche Forschungsgemeinschaft (Mu601/5-2, Mu601/5-3 and Mu601/7-1) and the Dr Mildred Scheel-Stiftung für Krebsforschung. S.B. is the recipient of
a fellowship from the Graduiertenkolleg ‘Zell- und Tumor-biologie’ at the Philipps-Universität Marburg.
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Restriction of cell cycle progression
(Received 15 February 1993 - Accepted 10 March 1993)
123