Department of Biological Sciences, State University of New York at Buffalo, Buffalo, NY 14260, USA
* Author for correspondence (e-mail: berezney{at}acsu.buffalo.edu)
Accepted 1 August 2002
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Summary |
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Key words: DNA replication patterns, Primary and transformed mammalian cells, Senescence, Retinoblastoma protein, Minichromosome maintenance proteins, Nuclear lamin proteins
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Introduction |
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The integrity of the nuclear organization is compromised in a variety of
diseases. Major structural components and functional domains of the nucleus
undergo dramatic changes during viral infection
(Maul, 1998;
Monier et al., 2000
;
Wilcock and Lane, 1991
),
malignant transformation, or in organisms with genetic syndromes. Examples of
altered nuclear components in transformed cells include the nuclear matrix
(Davido and Getzenberg, 2000
;
Pienta et al., 1989
),
chromatin (Chadee et al.,
1999
; Chadee et al.,
1995
; de Campos Vidal et al.,
1998
; Herrera et al.,
1996
; Vassilev et al.,
1995
), nuclear bodies and speckles
(Gordon et al., 2000
;
Manuelidis, 1984
;
Spector et al., 1992
) and
replication proteins (Bechtel et al.,
1998
). Regulatory activities play a key role in establishing
nuclear architecture that favors tumor suppression
(Chuang et al., 1997
;
Linares-Cruz et al., 1998
).
Genetic disorders have been traced to mutations in genes encoding for
components of chromatin [Rett syndrome
(Amir et al., 1999
)], of the
DNA replication, recombination and repair machinery [e.g. ICF syndrome
(Hansen et al., 1999
)], and
nuclear structural components, such as the nuclear lamins
(Wilson et al., 2001
).
We set out to investigate whether differentiation or malignant
transformation of mammalian cells is accompanied by significant changes in
nuclear architecture and the number and distribution of DNA replication,
transcription and RNA processing sites. In this study, we have compared the
spatio-temporal patterns of DNA replication sites within normal, immortalized
and transformed mammalian cells of various origin. Unlike a recent study which
reports a fundamental difference in the organization of DNA replication sites
in primary versus immortalized cell lines
(Kennedy et al., 2000), we
find that the number and distribution of these sites throughout the S-phase is
strikingly similar in all cell lines examined and is independent of the
technique used to visualize the replication sites. We further report that
neither the nuclear lamin proteins nor the retinoblastoma protein (pRb) are
significantly associated at any stage in S-phase with replication sites and
that pRb does not colocalize with minichromosome maintenance (Mcm) protein
family members.
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Materials and Methods |
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Visualization of DNA replication sites
The immunofluorescent labeling of DNA replication sites was performed in 4
different ways. (1) 5-bromo-2'-deoxyuridine (BrdU)-substituted DNA was
immunolabeled by the acid-depurination protocol as described
(Dimitrova et al., 1999),
using a monoclonal anti-BrdU antibody (Becton Dickinson) and FITC- or Texas
Red-conjugated donkey anti-mouse secondary antibodies (Jackson ImmunoResearch
Laboratories). (2) Detection of BrdU using the nuclease cocktail (BrdU
labeling and detection kit, cat. no. 1296736, Roche) was performed exactly as
recommended by the manufacturer. (3) Sites of DNA synthesis within
digitonin-permeabilized cells were labeled by in situ run-on reaction in a
replication cocktail (30 mM HEPES, pH 7.6, 7 mM MgCl2, 1 mM
dithiothreitol, 100 µM each dATP, dGTP and dCTP, 25 µM biotin-11-dUTP,
400 µM each GTP, CTP and UTP, 4 mM ATP, 40 mM creatine phosphate, 20
µg/ml creatine phosphokinase). Biotin-dUTP was detected with Texas
Red-conjugated streptavidin (Amersham). The digitonin permeabilization of
cells under conditions that preserve intact nuclear membranes has been
described (Dimitrova and Gilbert,
1998
). (4) PCNA and RPA were immunolabeled as described
(Dimitrova et al., 1999
).
Nuclear lamin proteins were detected using rabbit polyclonal antibodies (gift
of Dr G. Blobel, Rockefeller University).
In the triple labeling experiments in Figs
7,8,9,
nascent DNA was labeled with 5-chloro-2'-deoxyuridine (CldU) and
visualized using a rat anti-BrdU antibody (Harlan/SeraLab) as described
(Dimitrova et al., 1999).
Nucleolin was detected with a mouse monoclonal anti-nucleolin antibody (Santa
Cruz), the retinoblastoma protein with either a rabbit polyclonal
(NeoMarkers; a gift from Dr D. Goodrich, Roswell Park Cancer Institute) or a
mouse monoclonal antibody (a gift from Dr T. Melendy, SUNY Buffalo), and Mcm7
with a mouse monoclonal antibody (Santa Cruz).
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The cells were imaged on a Zeiss epifluorescent microscope equipped with a RTE CCD camera (Princeton Instruments), controlled by IP Lab software (Scanalytics), or on an Olympus BX51 epifluorescent microscope equipped with a Sensicam QE CCD camera (The Cooke Corporation), controlled by Image-Pro Plus software (Media Cybernetics). The images were assembled in an Apple G3 Powerbook using Adobe Photoshop 5.0.2 and Claris Draw 1.0v4 software.
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Results and Discussion |
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Whereas a significant amount of information has accumulated regarding the
changes in cell morphology, chromosome structure, enzymatic activities and
cell cycle regulatory activities that distinguish normal proliferating cells,
senescent cells or immortal/transformed cells, very little is known about the
relationship between senescence or immortalization and the spatio-temporal
regulation of DNA replication. Scattered reports have been published on the
appearance and sequence of DNA replication patterns within normal or tumor
cells of various derivation (Fox et al.,
1991; Humbert et al.,
1992
; Humbert and Usson,
1992
; Kill et al.,
1991
; O'Keefe et al.,
1992
; van Dierendonck et al.,
1989
), but few attempts have been made to conduct a comprehensive
study of these patterns within aging cells at different passage levels or
within immortalized cells at different stages of transformation. Therefore, we
set out to investigate whether there are differences in the replication
patterns between transformed, immortal and young or aging primary mammalian
cells.
We collected a number of rodent and human cell lines that belong to these groups. Cells were synchronized at the beginning of S-phase and then released for various times to prepare cell populations at different stages of S-phase (Fig. 1). To test whether the cell synchronization procedures might influence the distribution of the replication sites, replication patterns were also analyzed in exponentially growing cells (Fig. 2). Furthermore, to ensure that the method employed to visualize the replication patterns does not produce artifactual results and lead to inaccurate conclusions, we used four different techniques to reveal sites of DNA replication.
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|
Visualization of BrdU-labeled DNA replication sites through the use
of nuclease/anti-BrdU antibody cocktail
In the first series of experiments, the cell cultures were synchronized at
the G1/S border by aphidicolin arrest. Aliquots of each cell culture were
released from the block for different time periods (5 minutes, 1 hour, 2
hours, 4 hours, 6 hours, 8 hours, 10 hours or 12 hours) and pulse-labeled for
5 minutes with BrdU just prior to fixation and immunolabeling of the sites of
BrdU incorporation with fluorescent anti-BrdU antibodies. Since the
immunodetection of BrdU is possible only within single-stranded DNA regions,
two methods were used to create such regions. First, a nuclease cocktail was
used to generate short single-stranded tracts accessible for binding of the
anti-BrdU antibodies (Dolbeare and Gray,
1988; Fox et al.,
1991
). Although this technique is not frequently used for labeling
of replication foci, it is considered to be the one that best preserves the
nuclear architecture (Dolbeare and Gray,
1988
; Kennedy et al.,
2000
). Fig. 1 shows
a gallery of immunofluorescent images representative of different stages of
S-phase within a subset of the cell lines analyzed, including primary (WI38,
NHF1 and NRK), immortal (CHOC 400 and 3T3) and transformed (JH1, HeLa, MCF7,
TE671 and HT1080) cells. We and others have previously reported three major
types of replication patterns in several mammalian cell lines, corresponding
to early (type I), middle (type II) and late (type III) stages of S phase
(Ma et al., 1998
;
Nakayasu and Berezney, 1989
).
A finer discrimination of the replication patterns, however, led to their
classification into five major types
(Dimitrova and Gilbert, 1999
;
Humbert and Usson, 1992
;
O'Keefe et al., 1992
). Based
on the number, size, shape and distribution of the fluorescent foci, the
replication patterns in the cell lines analyzed in this study were similarly
classified into five types.
The type IA pattern is detectable for a brief period of time (30
minutes) at the onset of S-phase. It consists of a relatively low number
(several dozens to few hundred) of small discrete foci scattered throughout
the nuclear interior, but excluded from the peripheral, nucleolar or
heterochromatic regions. Importantly, this pattern was observed in all cell
lines (Fig. 1), indicating that
DNA replication initiates at the beginning of S-phase within nuclear sites
that are similarly distributed in the different cell lines.
It has been suggested that the arrest of cells at the G1/S border leads to
the artificial activation of additional replication origin sites that are not
normally utilized in unperturbed cell cycles
(Li et al., 2000;
Taylor, 1977
). The results
presented in Fig. 1, as well as
previously published reports (Dimitrova
and Gilbert, 1999
; Dimitrova
and Gilbert, 2000b
; Fox et
al., 1991
; Jackson,
1995
), demonstrate that this does not apply to a tight aphidicolin
block administered for limited time intervals. Our observation of type IA
patterns within exponentially growing cell cultures
(Fig. 2) at a frequency
5-10% of the BrdU-positive nuclei is consistent with the duration of this
pattern observed in our synchronized cell studies (
30 minutes within a
typical
10-hour S-phase). We conclude that limited exposure to
aphidicolin does not change significantly the number of early-S-phase
replication sites or induce artifacts in the overall replication patterns
observed from early to late S-phase.
The number and fluorescent intensity of foci rapidly increased as the cells
were allowed to progress further into S-phase (1-5 hours), until the nuclear
interior appears virtually filled with foci, including the regions close to
the nuclear periphery. This pattern, consisting of small and numerous discrete
granules [several hundreds to one- to two-thousand
(Ma et al., 1998)], is here
classified as type IB (Fig. 1).
It is the major replication pattern observed until the middle of S-phase.
Electron microscopy and biochemical data accumulated by different labs have
demonstrated that the euchromatic regions of the genome are involved in the
DNA replication process during this time
(Nakayasu and Berezney, 1989
;
O'Keefe et al., 1992
;
van Dierendonck et al., 1989
;
Williams and Ockey,
1970
).
A striking change in the distribution of the replication foci marks the transition from early to middle S-phase. Although the size and intensity of the individual foci did not change as compared to early-S, the number of the DNA replication sites dropped drastically and their localization switched from the internal euchromatic to predominantly perinuclear, perinucleolar and/or intranucleolar regions of the genome, known to accommodate mostly heterochromatin. This replication pattern, classified here as type II, persisted for 2-3 hours during mid-S (5-8 hours after release from aphidicolin arrest) in all cell lines (Fig. 1).
During late S-phase, the size of the replication sites increased and their
shape became irregular. Within type IIIA (1-2 hours), the number of
replication sites (several hundred) was intermediate between types IB and II.
The fluorescent foci were both at the nuclear periphery and scattered
throughout the interior and, consistent with previous observations
(Nakayasu and Berezney, 1989
;
O'Keefe et al., 1992
;
van Dierendonck et al.,
1989
), they often had a ring-, chain- or horseshoe-like appearance
(Fig. 1). The number of
replication foci decreased again at the very end of S-phase and their
morphology changed to more compact granules (this is especially pronounced in
the rodent cell lines see the NRK, CHOC 400, JH1 panels in
Fig. 1). This pattern,
classified as type IIIB (
1 hour), consisted of a small number of
heterogeneously sized (often extremely large) sites over peripheral or
interior nuclear regions. They stained intensely with DAPI, which is
characteristic of the constitutive heterochromatin.
In contrast to the early- and mid-S-phase patterns, not all cell lines showed identical morphology, number and distribution of replication sites within mid/late-S types II, IIIA and IIIB. For example, the late-S sites in the TE671, HCT116 and HT1080 human tumor cell lines remained relatively small in size and, in TE671 and HCT116, they occupied predominantly perinuclear regions with few sites in the nuclear interior (Fig. 1). In addition, whereas the rat and Chinese hamster cell lines showed a very small number (often just one) of large, homogeneously stained and often multi-lobed type IIIB sites, the human cell lines contained a higher number (a dozen or more) of late-S sites with a more heterogeneous morphology. The observed differences in the late-S replication patterns were not specific for the primary vs. the transformed cells, but rather reflected features characteristic of a certain cell type. For example, the replication patterns were indistinguishable between the primary NRK, the immortal CHO AA8 (not shown) and CHOC 400 cells, and the SV40-transformed JH1 cells. Likewise, the replication patterns were similar between the primary human cell lines (WI38 and NHF1) and the transformed HeLa cells.
When the replication patterns were analyzed within young (starting at passage 18), ageing (passage 35-45) or presenescent (passage 50) primary human fibroblasts (WI38), no major differences were found, with the exception of a marked decrease in the percentage of cells incorporating BrdU during the late passages, as expected for a senescent cell culture (data not shown). We conclude that there are no fundamental differences in the distribution of DNA replication sites and the spatio-temporal sequence of replication patterns between primary, immortal or transformed mammalian cells.
Visualization of BrdU-labeled DNA replication sites after mild HCl
hydrolysis
Our observations illustrated in Fig.
1 are in sharp contrast with the conclusions reached in a recent
study on the distribution of replication sites in primary mammalian
fibroblasts (Kennedy et al.,
2000). These investigators reported that primary cell lines
exhibited early-S replication pattern that lasted for approximately 3 hours
into S-phase and consisted of 5 to 20 perinucleolar foci. This unusual
early-S-phase pattern was observed in asynchronous cell cultures, as well as
in synchronized cell populations irrespective of the synchronization method
(i.e., contact inhibition with or without an aphidicolin or hydroxyurea
block). In contrast, immortalized cells exhibited numerous early-S replication
sites. Kennedy and colleagues (Kennedy et
al., 2000
), further suggested that previous observations of
numerous replication sites in early-S phase in primary cells are incorrect due
to the use of immunolabeling protocols (namely HCl hydrolysis) that destroy
nuclear structure. It was suggested that the use of DNase I to expose
single-stranded DNA regions for binding by the anti-BrdU antibodies better
preserves nuclear organization. Since we agree that HCl treatment (used in the
majority of the previous studies) is destructive for many nuclear antigens,
and, when used inappropriately, could produce artifacts, we applied in this
present study four different techniques for immunolabeling of DNA replication
sites to aliquots of the same cell cultures. This enabled a direct comparison
among the results obtained with each technique.
To optimize the comparison, cells from the same synchronized cell populations employed in the nuclease labeling method (Fig. 1) were used for the HCl method. The same five S-stage-dependent replication patterns were observed as found following nuclease treatment (compare Figs 1 and 3). In addition, a comprehensive analysis of the percent of each replication pattern in exponentially growing cells is shown in Fig. 2. Fourteen different cell lines ranging from primary to immortalized to transformed cells were compared following application of the nuclease or HCl labeling protocols. No significant differences were observed in the distribution of the five replication patterns as a function of the labeling method.
|
While we cannot definitively explain the discrepancies between our results
and those of Kennedy and colleagues
(Kennedy et al., 2000), one
possibility is that the nuclease labeling procedure used in their study failed
to efficiently immunolabel the BrdU sites within the primary cell lines. In
this regard, Dolbeare and Gray demonstrated that the sole use of an
endonuclease may not be sufficient to expose enough single-stranded DNA for
significant immunodetection. Addition of exonuclease III to the
nuclease/antibody cocktail, however, resulted in the generation of more
extensive single-stranded DNA regions, which were very efficiently recognized
by the anti-BrdU antibodies (Dolbeare and
Gray, 1988
). It is, therefore, conceivable that, under the
conditions used by Kennedy and colleagues
(Kennedy et al., 2000
), DNase
I (an endonuclease) generated mostly nicks in the chromosomal DNA of the
primary fibroblasts (as testified by the TUNEL staining performed in that
study), but failed to create sufficient number of single-stranded DNA regions
for good immunolabeling. It is interesting to consider that the late-S-phase
primary cells and the immortalized and transformed cells in general might have
been more sensitive to the DNase I digestion and, therefore, were more
efficiently stained with the anti-BrdU antibodies.
This possibility is corroborated by the published observation that
chromatin of pRb-deficient primary mouse fibroblasts exhibits higher
sensitivity to micrococcal nuclease digestion than chromatin of pRb-positive
primary mouse or human fibroblasts
(Herrera et al., 1996).
Intriguingly, this higher sensitivity is especially pronounced during the
progression of the cells from late-G1-phase into early-S-phase. Higher
nuclease sensitivity is associated with relaxed chromatin structure, which
potentially results from the elevated levels of histone H1 and H3
phosphorylation (Hohmann,
1983
; Lu et al.,
1994
) within pRb-deficient cells at this time of the cell cycle
due to deregulated cyclin-dependent kinase 2 (Cdk2) activity
(Chadee et al., 1999
;
Herrera et al., 1996
). Similar
to the pRb-deficient primary fibroblasts, many transformed cell lines either
lack pRb, or express mutant inactive pRb
(Nevins, 2001
;
Niculescu et al., 1998
). Thus,
pRb-negative primary and transformed cells, as well as tumor cells expressing
various oncogenes may be more susceptible to BrdU labeling via the nuclease
cocktail technique due to the higher accessibility of chromatin to nuclease
attack (Chadee et al., 1995
;
Herrera et al., 1996
;
Taylor et al., 1995
). The
possibility of different chromatin/chromosome organization (and different
nuclease sensitivity, respectively) between primary and transformed cells
(de Campos Vidal et al., 1998
;
Takaha et al., 2002
) and of
chromatin rearrangements, taking place during the progress of S-phase, is an
exciting direction to pursue.
We emphasize that, unlike Kennedy et al.
(Kennedy et al., 2000) who
used post-microscopy deconvolution image processing, we have not applied any
computer processing to our raw microscopic images. Deconvolution algorithms,
when applied inappropriately, can lead to a significant loss of relevant
immunofluorescent signal. Weakly fluorescing replication foci, typical of
nuclei at the onset of S-phase, would be especially susceptible to elimination
by harsh deconvolution processing after inefficient immunolabeling, thus
leading to in silico generation of artifactual results.
Visualization of DNA replication sites without generation of
single-stranded DNA regions
To further test our conclusions, we also applied two protocols that do not
require the generation of single-stranded DNA regions to visualize the
location of replication sites. First, biotinylated dUTP, incorporated into
DNA, can be directly immunolabeled with fluorescent streptavidin without the
need of DNA denaturation. The cells have to be permeabilized to allow access
of the biotin-dUTP to the nucleus. Following permeabilization, DNA synthesis
continues at the same sites that were active in vivo
(Krude, 1995;
Mills et al., 2000
).
Unfortunately, most previous studies were performed with cells permeabilized
with the non-ionic detergents Triton X-100 or Nonidet P40. These detergents
severely compromise both the overall structure and the replication capacity of
mammalian nuclei (Dimitrova and Gilbert,
1998
), which raises legitimate concerns about the degree of
preservation of nuclear organization
(Kennedy et al., 2000
). To
optimize comparisons, we used aliquots of the same cell cultures from
Fig. 1 to prepare permeabilized
cells with intact nuclei (Fig.
4A). We previously demonstrated that controlled permeabilization
with digitonin produces nuclei with uncompromised nuclear structure, fully
preserved replication capacity and unaltered distribution of replication
proteins and origin sites (Dimitrova and
Gilbert, 1998
; Dimitrova and
Gilbert, 1999
; Dimitrova et
al., 1999
). As illustrated in
Fig. 4B,C, the labeling of
nascent DNA in situ by run-on replication in the presence of biotin-11-dUTP
generates the same replication patterns as the in vivo labeling with BrdU.
This is in agreement with previous reports on biotin-dUTP labeling of
mammalian nuclei (Kill et al.,
1991
; Mills et al.,
2000
; Nakayasu and Berezney,
1989
).
|
Finally, in order to visualize replication sites by a method that does not rely on the detection of nascent DNA, we stained aliquots of the same cell cultures as in Fig. 1 with antibodies specific for protein components of mamalian replication forks. This was done either independently, or in combination with BrdU staining. As expected, the immunolabeling of PCNA and RPA, two of the best characterized replication proteins, produced replication patterns identical to those generated by the immunolabeling of nascent DNA (not shown). The two proteins colocalized at discrete nuclear sites (Fig. 5I), whose distribution dynamically changed during S-phase.
|
To verify that these are active DNA replication sites, we performed
double-labeling experiments to demonstrate that protein components of
replication forks colocalize with BrdU-labeled sites. We emphasize that this
double-labeling must be performed in a specific order to avoid artifactual
results. As previously discussed, the mild HCl hydrolysis required for
immunolabeling of BrdU sites is destructive for many nuclear proteins
(Humbert et al., 1992). To
illustrate this point, we immunolabeled the lamin B proteins, whose
established localization at the nuclear boundary can be used as a reliable
reference point. The immunolabeling of the lamin B proteins was done either
before, or following HCl treatment.
As evident from Fig. 5A-C,
prior HCl hydrolysis destroyed the integrity of the nuclear lamina and
resulted in a dispersed distribution of the lamin B epitopes, in agreement
with the results reported by Kennedy et al.
(Kennedy et al., 2000).
However, when the lamin B staining was performed first, and the fluorescent
anti-lamin antibodies were covalently fixed to their specific sites before the
cells were treated with HCl (Fig.
5D-F), the exclusive localization of the lamin B antibodies to the
nuclear envelope was preserved [most of the few internal lamin B foci could be
traced to invaginations of the nuclear envelope (results not shown)].
Identical distribution is obtained when lamin B proteins are stained without
any HCl treatment. We did not detect colocalization of internal lamin B foci
and BrdU sites during any stage of S-phase (examples of early-S and mid-S
nuclei are shown in Fig. 5A-F and
G, respectively), and neither before
(Fig. 5D-G), nor after
(Fig. 5A-C) HCl hydrolysis.
Identical results were obtained with lamin A/C-specific antibodies
(Fig. 5J-Q). In contrast, PCNA
(Fig. 5H) or RPA sites (not
shown) colocalized with the BrdU sites, as expected for replication fork
proteins.
These results are in agreement with many previous studies, but differ
strikingly from the observations on the cell cycle distribution of these
proteins reported by Kennedy and colleagues
(Kennedy et al., 2000). They
report that abundant DNA replication-related proteins, such as PCNA and CAF-1
localize to only a few discrete perinucleolar foci during G1- and early
S-phase. Such observations are inconsistent with previous studies of these
proteins in both primary and immortalized mammalian cells. For example, both
RPA and PCNA are present in a soluble form during the entire cell cycle, thus
generating strong uniform nuclear staining. A chromatin-bound fraction of
these proteins, which localizes to the DNA replication sites, is detected only
during S-phase, after a prior extraction of the soluble fraction
(Bravo and Macdonald-Bravo,
1987
; Dimitrova and Gilbert,
2000a
; Dimitrova et al.,
1999
). Therefore, the very low number (5 to 20) of discrete foci
of total nuclear PCNA and CAF-1 during G1-phase and the first 2-3 hours of
S-phase in formaldehyde-fixed cells described by Kennedy and colleagues
(Kennedy et al., 2000
) is
bizarre and is not consistent with the known behavior of these proteins
(Bravo and Macdonald-Bravo,
1987
; Dimitrova et al.,
1999
; Humbert et al.,
1992
; Krude, 1995
;
Marheineke and Krude, 1998
;
Murzina et al., 1999
;
Shibahara and Stillman,
1999
).
The retinoblastoma protein does not colocalize with DNA replication
sites or mammalian Mcm proteins
pRb is a key regulator of cell cycle progression
(Zheng and Lee, 2001) and is
present in a functionally active, hypophosphorylated form during late
mitosis/G1-phase, or in a functionally inactive, unphosphorylated or
hyperphosphorylated forms during G0- and throughout late-G1/S/G2-phases,
respectively (Ezhevsky et al.,
2001
; Ho and Dowdy,
2002
; Moberg et al.,
1996
). The estimated number of pRb molecules per human cell is of
the order of 1 million (Goodrich et al.,
1991
) and the levels of this protein do not vary excessively
during the cell cycle (Buchkovich et al.,
1989
; Mihara et al.,
1989
; Muller et al.,
1997
). It is difficult to imagine that a million molecules of an
important cell cycle and differentiation regulator known to have numerous
genomic targets (Dyson, 1998
;
Wells et al., 2000
;
Zheng and Lee, 2001
) would
coalesce into only 5-20 perinucleolar foci within mammalian nuclei, as
observed by Kennedy et al. (Kennedy et
al., 2000
). Moreover, this immunofluorescent pattern differs from
the findings on the subnuclear localization of pRb family members in both
primary, immortalized and transformed cells reported by several other groups
(Bartek et al., 1992
;
Cinti et al., 2000
;
Fortunato and Spector, 1998
;
Mittnacht and Weinberg, 1991
;
Szekely et al., 1991
;
Zini et al., 2001
). Most of
these studies found numerous pRb-positive foci and/or speckles scattered
throughout the nucleus. However, differences have also been reported, mostly
concerning the size, number and intensity of the pRb granules
(Szekely et al., 1991
).
Whereas some of these differences could be explained by the different
functional status of pRb in primary vs. transformed cells (i.e. normal vs.
mutant pRb forms) (Cinti et al.,
2000
; Szekely et al.,
1991
), others clearly derive from the use of different fixation
protocols (Szekely et al.,
1991
) or from the utilization of unrelated anti-pRb antibodies
(Bartek et al., 1992
;
Mittnacht and Weinberg, 1991
;
Szekely et al., 1991
), which
recognize different epitopes and, potentially, different nuclear
subpopulations of pRb.
Since, to our knowledge, Kennedy et al.
(Kennedy et al., 2000) are the
only investigators who have directly examined the relative distribution of pRb
and nuclear DNA replication sites and in view of the apparent discrepancies,
we decided to reinvestigate this important issue using our collection of
anti-pRb antibodies. From the eight anti-pRb antibodies, which we tested, only
two (see Materials and Methods) gave positive immunofluorescent signal with
primary human WI38 and NHF1 fibroblasts. In agreement with the observations
reported by Szekely et al. (Szekely et
al., 1991
), we found that formaldehyde fixation of the cells
resulted in a more uniformly granular staining pattern (e.g. see
Fig. 6), whereas
methanol/acetone fixation resulted in a smaller number of heterogeneously
sized pRb granules (not shown). Since methanol treatment is known to both
extract and fix proteins through precipitation, it is likely that these
differences result from the loss and/or aggregation of pRb molecules in cells
fixed with organic solvents. We, therefore, used formaldehyde fixation in the
remaining experiments.
|
Under our experimental conditions, pRb exhibited exclusively nuclear
localization and all interphase nuclei within asynchronously growing primary
fibroblast cell cultures were positive. In agreement with previous reports in
the literature (Bartek et al.,
1992; Mittnacht and Weinberg,
1991
), we observed some variability in the intensity of the pRb
immunofluorescent signal (Fig.
6). The weaker-staining nuclei were generally, albeit not
exclusively, BrdU-negative (e.g. Fig.
6A-C) and, thus, it is likely that most of them are G1-phase
nuclei, since the levels of pRb are known to increase
twofold in the
course of the cell cycle (Buchkovich et
al., 1989
; Mihara et al.,
1989
; Szekely et al.,
1991
). In all nuclei the pRb labeling appeared as hundreds of
fluorescent foci scattered throughout the nucleus and often showed a bias
towards nuclear regions with lower DNA density
[Fig. 6G-I
(Szekely et al., 1991
)].
Simultaneous immunolabeling of pRb and the major nucleolar protein
nucleolin (Ginisty et al.,
1999) demonstrated that the nucleolar regions contain numerous pRb
foci (Fig. 7), consistent with
a role for the Rb family of proteins in the regulation of rDNA transcription
(Ciarmatori et al., 2001
;
Hannan et al., 2000
). We,
however, did not find any cells in the entire exponentially growing population
(Figs 6,
7), which exhibited the limited
perinucleolar pRb staining pattern described by Kennedy et al.
(Kennedy et al., 2000
).
Additionally, merged images of pRb and DNA replication sites following triple
labeling experiments revealed that pRb was present at hundreds of
extranucleolar sites, which showed little, if any, colocalization with
CldU-substituted nascent DNA during all stages of S-phase (Figs
7,8,9).
The absence of pRb within nuclear DNA replication sites argues against a
direct role for pRb in this process. This view is supported by the notion that
DNA replicates during S-phase when pRb is normally rendered inactive by
hyperphosphorylation (Ezhevsky et al.,
2001; Lundberg and Weinberg,
1998
). Furthermore, our immunocytochemical results are in
agreement with the observations reported by Goodrich et al.
(Goodrich et al., 1991
) on the
in vivo physiological effect of active pRb on DNA replication. In these
experiments, the microinjection of purified functional pRb into pRb-negative
cultured mammalian cells prevented G1 cells from entry into S-phase, but did
not have any major effect on those cells, which were already in S-phase. We
conclude that pRb does not have a direct role in mammalian DNA synthesis
during S-phase.
Our results, however, do not rule out the possibility that pRb might have
an indirect role in certain aspects of S-phase regulation in normal cells or
under conditions of stress (Knudsen et
al., 2000; Sever-Chroneos et
al., 2001
). For example, it has been shown that pRb, independent
of its phosphorylation status, associates with
(Pradhan and Kim, 2002
;
Robertson et al., 2000
) and
inhibits the activity of the maintenance DNA methyltransferase (Dnmt1)
(Pradhan and Kim, 2002
). Since
active Dnmt1, but not pRb, is present at DNA replication sites
(Leonhardt et al., 1992
;
Rountree et al., 2000
), it is
possible that pRb and Dnmt1 associate in the nucleosolic compartment. Thus,
pRb might modulate Dnmt1 activity by controlling the levels of chromatin-bound
functional Dnmt1. The importance of properly regulated Dnmt1 is underscored by
the observation that deregulated methyltransferase activity accompanies
neoplastic transformation of mammalian cells
(El-Deiry et al., 1991
;
Kautiainen and Jones,
1986
).
An alternative role for pRb in the regulation of genome replication is
suggested by the finding that human pRb interacts in vitro with a member of
the Mcm family of proteins, Mcm7 (Sterner
et al., 1998). Mammalian Mcm proteins begin to load onto chromatin
in late telophase (Dimitrova et al.,
2002
; Dimitrova et al.,
1999
) in a process known as replication licensing
(Blow, 2001
). It is generally
believed that Mcm-s, together with ORC, Cdc6 and Cdt1, assemble into
pre-replication complexes (pre-RCs) at chromosomal replication origins. The
loading of Mcm-s during late telophase completes the assembly of fully
functional pre-RCs and the licensing capacity of mammalian nuclei remains
unchanged throughout G1-phase (Dimitrova
et al., 2002
). By analogy with the role of Rb family members in
transcriptional repression during G1-phase, which at least in part is mediated
by association with promoter-bound E2F protein family members
(Dyson, 1998
;
Wells et al., 2000
;
Zheng and Lee, 2001
),
association of pRb with origin-bound Mcm family members could represent a
potential mechanism for keeping the origins dormant until a protein kinase
nuclear environment favoring activation of the origins has been established at
the G1/S-phase transition. To date, evidence for the existence in vivo of
pRb-Mcm7 complexes, however, has not been presented. Hence, we decided to
investigate whether pRb and Mcm proteins are found in close proximity within
nuclei of primary and transformed mammalian cells.
Mcm proteins bind tightly to licensed chromatin and this interaction is
resistant to mild detergent extraction
(Dimitrova et al., 2002;
Dimitrova et al., 1999
).
Therefore, in order to look at the active, possibly origin-bound form of
Mcm-s, we subjected exponentially growing cultures of WI38 and HeLa cells to a
Triton X100 extraction procedure prior to fixation, which removes all soluble
or loosely bound cytoplasmic and nuclear proteins. To mark DNA replication
sites, the cells were pulse-labeled with CldU immediately before extraction.
After fixation with formaldehyde, the cells were triple labeled with
antibodies specific for pRb, CldU and one Mcm family member (Mcm2, 3 or 7).
Representative results of WI38 cells with anti-Mcm7 antibodies are shown in
Fig. 8. Identical results were
obtained with HeLa cells and with Mcm2- and Mcm3-specific antibodies.
Even though we used asynchronous cell cultures, it is possible to classify
precisely the cells according to the cell cycle stage. G1 nuclei are
CldU-negative and Mcm-positive, since the Mcm proteins are tightly bound to
chromatin and, therefore, resistant to Triton extraction during G1-phase.
S-phase nuclei are CldU-positive and can further be categorized into early-,
mid- or late-S subgroups based on the characteristic replication patterns and
on the decreasing amount of chromatin-bound Mcm-s. Finally, G2-phase nuclei
(not shown) are both CldU- and Mcm-negative. The mild extraction procedure,
which we applied, removed also a significant fraction of pRb. Nevertheless,
most nuclei, including those in S-phase, remained positive for pRb, even
though the fluorescence intensity varied between individual nuclei and was
generally low in CldU-positive nuclei. pRb is known to associate tightly with
the nuclear matrix only during G1-phase when it is hypophosphorylated and
active (Mancini et al., 1994).
Unlike nuclear matrix results obtained through the application of high salt
extraction procedures, our experiments show that a small fraction of pRb
resists low salt/detergent extraction also during S-phase. This observation
seems to contradict previously published observations on the low-salt
extractability of pRb (Mittnacht and
Weinberg, 1991
). We do not know the exact reason for this
discrepancy, but we believe that differences in buffer composition and
extraction procedures may provide a potential answer. We note that hypotonic
treatment used in previous studies
(Mittnacht and Weinberg, 1991
)
is known to disrupt nuclear architecture
(Zatsepina et al., 1997
),
whereas the cytoskeleton buffer used by us may preserve molecular interactions
better. Even though unexpected, our observations that small amounts of pRb
resist low-salt extraction during S-phase are corroborated by a recently
published report (Wells et al.,
2000
), which found that, surprisingly, Rb family proteins are
bound in vivo to a number of gene promoters at this cell cycle time. The
implications of these discoveries remain to be unveiled.
Consistent with previous reports on the lack of colocalization between Mcm
proteins and DNA replication sites
(Dimitrova et al., 1999;
Krude et al., 1996
;
Todorov et al., 1995
), we did
not observe spatial coincidence between chromatin bound Mcm-s and CldU-labeled
nascent DNA during any stage of S-phase
(Fig. 8). Importantly, pRb and
Mcm7 also did not significantly overlap
(Fig. 8). Identical results
were obtained with Mcm2- and Mcm3-specific antibodies (data not shown). To
address the possibility that Mcm-s and pRb might interact within the soluble
nucleoplasmic fraction, we also conducted experiments where the cells were
fixed directly after CldU pulse-labeling without prior detergent extraction
(Fig. 9). Again, no significant
overlap was detected between the pRb- and Mcm-specific immunofluorescent
signals. Thus, the lack of spatial proximity between pRb and Mcm proteins
makes it unlikely that extensive interactions between these proteins take
place in vivo within mammalian nuclei. This notion is further strengthened by
the observation that microinjection of a functional pRb mutant with deleted
N-terminal part (which mediates the interaction with Mcm7) is sufficient to
block cultured human cells in G1-phase and to prevent their entry into S-phase
(Goodrich et al., 1991
).
Furthermore, expression of a constitutively active form of pRb resistant to
Cdk phosphorylation has no effect on the establishment or maintenance of
mammalian pre-RCs (Sever-Chroneos et al.,
2001
). The significance of the biochemically detected interactions
between pRb and Mcm7 remains unclear at present
(Sterner et al., 1998
).
In conclusion, the results presented in this report are in good agreement
with a plethora of previous reports in the literature, in which the
replication patterns during S-phase were studied individually in primary,
immortal or transformed mammalian cells. We conducted a comprehensive study of
the distribution of DNA replication sites in over a dozen mammalian cell lines
with different proliferation capacities. Through the use of four independent
approaches for visualization of these sites, we have provided an explanation
for some puzzling discrepancies in the literature and have convincingly
demonstrated that there are no fundamental differences in the distribution of
DNA replication sites and the spatio-temporal sequence of replication patterns
between normal, immortalized or transformed mammalian cells. These studies
further demonstrate that pRb and nuclear lamin proteins A, B and C are not
significantly associated with replication sites in either early-S or any other
stage of S-phase. Moreover, no evidence was found for limited perinucleolar
foci containing labeled replicating DNA, nuclear lamins and/or pRb as reported
by Kennedy et al. (Kennedy et al.,
2000). Simultaneous examination of Mcm proteins and pRb enabled us
to extend these studies to determine whether pRb, together with Mcm-s, while
not associating with active replication sites, might associate with
replication origins and thus play a potential role in origin regulation or in
the transformation of pre-RCs into active replication complexes. Our findings
show for the first time the lack of association of pRb with Mcm-labeled sites
in G1-phase, as well as throughout S-phase.
The validity of the major replication patterns described in this and many
past studies is further substantiated by recent in vivo studies, whereby the
dynamics of DNA replication sites are visualized directly in live cells
through the microinjection of fluorescent nucleotides
(Manders et al., 1999;
Pepperkok and Ansorge, 1995
)
or through the stable transfection of cells with GFP-tagged variants of
protein components of the replication machinery
(Leonhardt et al., 2000
;
Somanathan et al., 2001
).
Taken together with the findings of similar distributions of replication sites
in other eukaryotes, including plant cells
(Fuchs et al., 1998
;
Fujishige and Taniguchi, 1998
;
Lafontaine and Lord, 1974
;
Sparvoli et al., 1976
), we
conclude that replication patterns are generally conserved in at least the
higher eukaryotic organisms. Observed differences occur during the mid/late
stages of S-phase and possibly reflect differences in DNA sequence content and
unique chromatin organization in different cell types, rather than the stage
of transformation or the replicative capacity of the cells. Deciphering the
mechanistic basis for this highly conserved spatio-temporal programming for
the replication of the genome and its associated replication factors within
the context of a global nuclear architecture is an exciting challenge for
future investigation.
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Acknowledgments |
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References |
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