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Molecular and Cellular Biology, September 2000, p. 6308-6316, Vol. 20, No. 17
0270-7306/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
Replication of Heterochromatin and Structure of Polytene
Chromosomes
Thomas J.
Leach,1
Heather L.
Chotkowski,1
Michael G.
Wotring,1
Robert L.
Dilwith,1 and
Robert
L.
Glaser1,2,*
Laboratory of Developmental Genetics,
Wadsworth Center, New York State Department of
Health,1 and Department of
Biomedical Sciences, State University of New York at
Albany,2 Albany, New York 12201
Received 21 April 2000/Accepted 30 May 2000
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ABSTRACT |
Heterochromatin is characteristically the last portion of the
genome to be replicated. In polytene cells, heterochromatic sequences
are underreplicated because S phase ends before replication of
heterochromatin is completed. Truncated heterochromatic DNAs have been
identified in polytene cells of Drosophila and may be the
discontinuous molecules that form between fully replicated euchromatic
and underreplicated heterochromatic regions of the chromosome. In this
report, we characterize the temporal pattern of heterochromatic DNA
truncation during development of polytene cells. Underreplication
occurred during the first polytene S phase, yet DNA truncation, which
was found within heterochromatic sequences of all four
Drosophila chromosomes, did not occur until the second polytene S phase. DNA truncation was correlated with underreplication, since increasing the replication of satellite sequences with the cycE1672 mutation caused decreased production
of truncated DNAs. Finally, truncation of heterochromatic DNAs was
neither quantitatively nor qualitatively affected by modifiers of
position effect variegation including the Y chromosome,
Su(var)2052, parental origin, or temperature.
We propose that heterochromatic satellite sequences present a barrier
to DNA replication and that replication forks that transiently stall at
such barriers in late S phase of diploid cells are left unresolved in
the shortened S phase of polytene cells. DNA truncation then occurs in
the second polytene S phase, when new replication forks extend to the
position of forks left unresolved in the first polytene S phase.
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INTRODUCTION |
Eukaryotic chromosomes contain two
distinct chromatin domains, euchromatin and heterochromatin.
Euchromatin typically encompasses a majority of the chromosome and
contains most genes and other unique sequences. Heterochromatin
encompasses a smaller proportion of the chromosome, typically around
the centromere and telomeres, and is enriched in noncoding, highly
repetitive satellite sequences (22). The presence of
heterochromatin and its association with repetitive sequences is a
conserved feature of eukaryotic chromosomes and is found in most, if
not all, organisms including yeast, flies, and humans. Heterochromatin
is defined cytologically by its dark staining and condensed appearance
and genetically by its ability to suppress gene expression. Numerous
protein components of heterochromatin have now been identified,
including the SIR proteins in yeast (18); HP1, Su(var)3-7,
and ORC proteins in higher eukaryotes (58); specific
acetylated isoforms of histone H4 (3, 56); and variant forms
of histone H3 (34). Our understanding of how these and other
proteins contribute to the structure of heterochromatin and how that
structure relates to the dynamic functions of heterochromatin is
beginning to be broadened (4, 38).
Another conserved characteristic of heterochromatin is its late
replication during S phase; it is most often the last portion of the
genome to be replicated (31). The mechanism for late replication is not known, although the condensed structure of heterochromatin and its ability to suppress transcription suggest that
it might also suppress its own replication. If heterochromatin suppresses replication origins, for example, heterochromatic sequences might be late in replicating because initiation must occur at distant
euchromatic origins or because suppressed origins fire only in late S
(53). Alternatively, replication fork elongation might be
substantially slower through heterochromatin than through euchromatin,
resulting in longer and therefore later replication of heterochromatic
sequences (17, 47).
A particularly dramatic consequence of the late replication of
heterochromatic sequences is found in the polytene chromosomes of
Drosophila melanogaster. The copy number of euchromatic
sequences in the polytene chromosomes of the larval salivary gland
exceeds 1,000c, while heterochromatic satellite sequences within the
same chromosomes are severely underrepresented, remaining at or close to a copy number of 2c (11, 13, 43). Underrepresentation is
likely to be a consequence of the failure of satellite sequences to be
replicated during the polytene cell cycle because of a shortened S
phase that ends before the satellite sequences are fully replicated (30). Consistent with this model, specific mutations in
cyclin E, a regulator of S phase in both diploid and
polytene cells of Drosophila (46), can increase
the copy number of satellite sequences in polytene chromosomes by
increasing the length of S phase and allowing late replication of
satellite sequences to be completed (30).
DNA molecules of discontinuous structure must occur between the fully
replicated euchromatic and underreplicated heterochromatic regions of a
polytene chromosome. DNA structures consisting of nested replication
forks have been proposed to populate such regions (26);
however, truncated linear DNAs rather than fork-containing molecules
have been found in these regions, prompting reconsideration of the
underreplication model of heterochromatic underrepresentation (14,
15, 51). In this study, we characterized the temporal pattern of
heterochromatic DNA truncation during the development of polytene
chromosomes. The data suggest that replication barriers within
heterochromatin inhibit fork elongation through satellite sequences.
Replication forks stalled at such barriers fail to be resolved during
the shortened S phase of polytene cells, causing underreplication of
satellite sequences. Truncated linear DNAs initially form and then
accumulate as replication forks repeatedly extend to the same barriers
during the multiple S phases that generate polytene chromosomes.
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MATERIALS AND METHODS |
Drosophila strains and crosses.
Information
about Drosophila genes and nomenclature may be found at
Flybase (http://flybase.bio.indiana.edu). Flies were maintained at
23°C and 55% relative humidity on cornmeal-brewer's yeast-glucose medium. Strains CH(3)336 and CH(2)423, which
contain single heterochromatic P-element insertions, were described by
Zhang and Spradling (63). Ovaries used for cyclin E analysis
(see Fig. 3) were isolated from adult females of strain
cycE1672/CyO; CH(3)336. Homozygous
cycE1672/cycE1672 females are viable
but sterile and can be recovered from this stock (30).
Ovaries used for the analysis of Dp1187 (see Fig. 4) were
isolated from adult females of strain In(1)sc8
Df(1)sc8, y; Dp(1;f)1187, y+. Salivary
glands used for the Y chromosome modifier analysis (see Fig.
5A) were isolated from male larvae of genotype X/Y; CH(3)336/+, which were obtained from cross X/X;
CH(3)336 × X/Y, and from male larvae of genotype X/O;
CH(3)336/+, which were obtained from cross X/X;
CH(3)336 × X
Y. Salivary glands used for the
Y chromosome modifier analysis shown (Fig. 5B) were isolated
from male larvae of genotype X/Y, y; DP(1;f)1187,
y+, which were obtained from cross X/X, y; × X/Y, y, Dp(1;f)1187, y+, and from male larvae of
genotype X/O; Dp(1;f)1187, y+, which were
obtained from cross XX, y × X/Y, y,
Dp(1;f)1187, y+. Su(var)2052/+; CH(3)336/+
flies were obtained from cross Su(var)2052/CyO × CH(3)336. Female or male CH(3)336,
P[ry+] flies and female or male Dp(1;f)1187,
y+ flies were crossed to
y1;ry506 animals to obtain
CH(3)336 and Dp(1;f)1187 chromosomes,
respectively, transmitted from either the female or male parent.
DNA isolation and analysis.
Fluorescence-activated cell
sorter analysis of nuclei was done essentially as described by Lilly
and Spradling (30). Briefly, ovaries dissected from adult
females were homogenized, and nuclei were isolated on sucrose
gradients. After the nuclei were stained with
4',6-diamidino-2-phenylindole (DAPI), they were sorted on a Becton
Dickinson FACS Vantage using a Nova Enterprise UV laser at 15 mW.
Nuclei of specific ploidy classes were collected and prepared in
agarose inserts. Approximately 400 pairs of ovaries were dissected, and
106 nuclei were collected for each ploidy class (see Fig.
2B). DNAs used for the Y chromosome modifier analysis (see
Fig. 5) were isolated from whole salivary glands dissected from roaming
third-instar larvae.
DNA was prepared in agarose inserts by standard procedures as described
previously (16). The DNA was fractionated by contour-clamped homogeneous electric field pulsed-field gel electrophoresis on a
Bio-Rad DRII or CHEF Mapper apparatus. The two-dimensional agarose gel
electrophoresis technique used to separate the X and
Dp1187 hybridization signals in samples analyzed in the
experiment in Fig. 5B has been described previously (16).
Briefly, DNA was first digested with NotI and fractionated
by pulsed-field gel electrophoresis. A lane of the first-dimension gel
was excised, and the DNA in the gel was digested in situ with a second
enzyme. The DNA was then fractionated by conventional electrophoresis in a direction perpendicular to the direction of the first
electrophoresis. The restriction sites for the second enzyme are
polymorphic on the X and Dp1187 chromosomes;
therefore, the X and Dp1187-derived restriction
fragments differ in size and therefore are separated in the second
dimension. After pulsed-field gel or two-dimensional electrophoresis,
DNA was analyzed by Southern blot hybridization (45; also see
reference 16). In-gel hybridizations, which provide a more
sensitive and accurate method for detecting DNAs separated by
pulsed-field gel electrophoresis (28), were used for the
experiments in Fig. 4. Both Southern blot and in-gel hybridizations were done using the procedures of Church and Gilbert (6).
Probes were labeled with 32P to specific activities of
approximately 109 cpm/µg by random priming
(12). Hybridization signals were analyzed qualitatively
using Kodak XAR5 film and quantitatively using stored PhosphorImager
screens and ImageQuant software (Molecular Dynamics).
Clones of bacterial
lacZ sequences were used as
hybridization probes for analysis of
CH(3)336 and
CH(2)423. Subclones of
X chromosome genomic DNA
used as hybridization probes for analysis
of
Dp1187 have
been described previously (
15) and included a
2.6-kb
AseI-
HindIII fragment used for the
experiments in Fig.
4 and a 0.8-kb
BglII-
PstI
fragment used for the experiments in
Fig.
5, both from the
sc8 region of
Dp1187. A 3.6-kb
EcoRI fragment containing the
zfh-2
gene was used
for the analysis of the fourth chromosome (see Fig.
4) (
32).
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RESULTS |
Truncation of heterochromatic DNAs occurs after the second polytene
S phase.
Defined heterochromatic restriction fragments were
studied in Drosophila by using chromosomes
CH(3)336 and Dp(1;f)1187 (Dp1187) (Fig.
1). These chromosomes have been used
extensively to study the structure and function of
Drosophila heterochromatin (15, 24, 25, 37, 62,
63). CH(3)336 has a P element inserted into
pericentromeric heterochromatin in region h54-57 of chromosome 3. The P
element is inserted into an "island" of complex sequences consisting primarily of clustered transposable elements
(62). Interspersion of such islands among large tracks of
satellite repeats is the normal sequence structure of
Drosophila heterochromatin (8, 27).
Dp1187 was derived from the X chromosome first by a large pericentric inversion, In(1)sc8, which
directly juxtaposed centromere-proximal heterochromatin and distal
euchromatic sequences, followed by a larger internal deletion that
created the 1300-kb Dp1187 minichromosome (27). The euchromatic sequences of Dp1187 directly adjoin 1.688 satellite repeats (16), while on the normal X
chromosome, these euchromatic sequences are located approximately 15 Mb
from pericentromeric heterochromatin (27).
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FIG. 1.
Structure of CH(3)336, CH(2)423, and
Dp(1;f)1187 chromosomes. A portion of the
CH(3)336 or CH(2)423 chromosome is shown
containing a P-element transposon (triangle) inserted into
heterochromatic sequences (gray line) on the third
[CH(3)336] or second [CH(2)423] chromosome.
lacZ sequences within the P element were used as a probe
(dark line above the P element) to detect the 650-kb
[CH(3)336] or 185-kb [CH(2)423] full-length
NotI restriction fragment (dark line below the chromosome).
The entire Dp(1;f)1187 chromosome is shown, with
heterochromatic (gray line) and euchromatic (black line) sequences
indicated. The coordinate system used defines 0 kb as the position of
the sc8 euchromatic-heterochromatic breakpoint.
Euchromatic sequences used as a probe (fat black line) and full-length
restriction fragments (thin black lines) are indicated below the
chromosome. N, NotI; P, PmeI; Z,
lacZ.
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It has been demonstrated previously that restriction fragments that
extend from euchromatic into heterochromatic sequences
of a chromosome
are truncated in size in polytene cells (
15).
These
truncated molecules are likely to be the discontinuous DNAs
that were
predicted to form between regions of fully replicated
euchromatic and
underreplicated heterochromatic sequences of a
polytene chromosome. In
this earlier study, however, the presence
of truncated heterochromatic
DNAs was correlated only with polytene
tissues and not with the
specific developmental transition from
a diploid to a polytene cell
cycle (
15). To determine when DNA
truncation occurs relative
to the underreplication of heterochromatic
sequences, chromosomes were
analyzed during the transition from
a diploid to a polytene
structure.
Ovaries from adult flies contain two abundant polytene cell types,
follicular epithelial cells and nurse cells. Isolated nuclei
from whole
adult ovaries can be separated into specific ploidy
classes by flow
cytometry (Fig.
2A). The 2c to 16c nuclei
are
predominantly from the abundant follicular epithelial cells, and
the 32c to 512c nuclei are from the less abundant but higher-ploidy
nurse cells (
50). The 4c peak is a double peak that consists
of nuclei from diploid follicle cells that have replicated their
DNA
completely and nuclei from follicle cells that have committed
to
polytene development and underreplicated their heterochromatic
sequences during their first polytene S phase (Fig.
2A). Nuclei
larger
than 4c have only a single fluorescence maximum since all
polytene
cells underreplicate their DNA comparably with each S
phase.
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FIG. 2.
Truncation of heterochromatic DNAs occurs after the
second polytene S phase. (A) Nuclei from flies carrying the
CH(3)336 chromosome were isolated from ovaries and sorted by
flow cytometry into discrete ploidy classes. (B) Nuclei from each
ploidy class were collected, and their DNA was isolated, restriction
digested, fractionated by pulsed-field gel electrophoresis, and
analyzed by Southern blot hybridization. The ploidy of nuclei analyzed
is indicated above each autoradiogram. Nuclei of 2c to 32c ploidy are
predominantly from follicle cells, and nuclei of 64c to 512c ploidy are
predominantly from nurse cells. The sizes of the DNA molecules are
indicated in kilobase pairs, and the ratio of abundance of the 340-kb
molecules to the 650-kb full-length restriction fragment is indicated
below each autoradiogram. The well of each lane is indicated (w).
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Nuclei from ovaries of
CH(3)336 flies that contain a P
element inserted into third-chromosome heterochromatin were separated
by flow cytometry, and nuclei of each ploidy class were collected.
The
heterochromatic restriction fragment flanking the P element
was
analyzed by Southern blot hybridization from nuclei of each
ploidy
class (Fig.
2B). DNA from the 2c and 4c nuclei contained
only the
full-length restriction fragment of 650 kb. Truncated
DNAs appeared
abruptly in 8c nuclei and were present in all nuclei
of 8c and higher
ploidy. The truncated DNAs were heterogeneous
in size, ranging from 77 kb up to the size of the 650-kb full-length
restriction fragment,
although molecules of 340 kb and, to a lesser
extent, 77 kb were
prominent. Quantitative differences between
follicle cells and nurse
cells in the abundance of truncated DNAs
were observed: truncated DNAs
were more abundant in follicle cells
than in nurse cells. For example,
in follicle cells, DNAs truncated
to 340 kb were were always more
abundant than the full-length
restriction fragment and the 77-kb
truncated DNA was readily detected,
while in nurse cells, DNAs
truncated to 340 kb were not always
more abundant than the full-length
restriction fragment and the
77-kb DNA was not always detected (Fig.
2B). In both cell types,
the abundance of truncated DNAs appeared to
increase with increasing
ploidy, and this increase may occur more
rapidly in follicle cells
than in nurse
cells.
Unexpectedly, initial truncation of heterochromatic DNAs did not
correlate temporally with the start of heterochromatic
underreplication.
Recall that half of the 4c follicle cell nuclei had
begun polytene
development and underreplicated their DNA (Fig.
2A), yet
no truncated
DNAs were detected in the 4c sample (Fig.
2B). Even if the
truncated
DNAs are the product of stalled replication forks that
subsequently
brake in vivo or in vitro, they should be present in the
underreplicated
4c nuclei. Lack of temporal correlation between the
appearance
of the truncated DNAs and the start of heterochromatic
underreplication
raised the possibility that truncation is not a
consequence of
underreplication but instead is caused by some other
cellular
process that occurs during polytene
development.
Truncation of heterochromatic DNAs is correlated with the extent of
underreplication.
If truncation of heterochromatic DNA is a
consequence of underreplication, the abundance of truncated DNAs should
correlate with the extent of underreplication. This hypothesis was
tested by characterizing truncated DNAs in flies containing a mutation in cyclin E that increases the extent of heterochromatic replication. Cyclin E is a regulator of S phase in both diploid and polytene cells
of Drosophila (46).
CycE1672 is a viable hypomorphic P-element
mutation of cyclin E that increases the replication of
satellite sequences, principally in the polytene chromosomes of ovarian
nurse cells, leading to a decrease in heterochromatic underrepresentation (30). CycE1672 is
proposed to lengthen S phase, allowing more time for late replication
of satellite sequences (30). Flow cytometry analysis showed
that the DNA content in nurse cell nuclei of
cycE1672 flies increased significantly, as
expected (Fig. 3A). The DNA content in
follicle cell nuclei also increased a slight but detectable amount
(Fig. 3A). The CH(3)336 chromosome was crossed into a
cycE1672 background, and the heterochromatic
restriction fragment flanking the P element was analyzed by Southern
blot hybridization in samples of pooled follicle cell nuclei (8c to
32c) and pooled nurse cell nuclei (64c to 512c) from wild-type and
cycE1672 mutant flies (Fig. 3B). Truncated DNAs
in mutant cycE1672 flies were significantly less
abundant in nurse cell nuclei and slightly less abundant in follicle
cell nuclei than were truncated DNAs in the wild-type flies (Fig. 3B).
Thus, truncation of heterochromatic DNAs does correlate with the extent
of heterochromatic underreplication. When more replication of satellite
sequences was allowed, fewer truncated DNAs were produced (Fig. 3). We
would predict that if replication of satellite sequences were complete,
no truncated DNAs would be formed. These results strongly support the
hypothesis that DNA truncation is a consequence of heterochromatic
underreplication (Fig. 3B), even though truncated DNAs do not appear
until the second cycle of underreplication (Fig. 2B).
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FIG. 3.
Truncation of heterochromatic DNAs is correlated with
the extent of underreplication. (A) Nuclei from flies carrying the
CH(3)336 chromosome that were either heterozygous (open) or
homozygous (solid) for the cycE1672 mutation
were isolated from ovaries and sorted by flow cytometry into discrete
ploidy classes. The increased fluorescence observed for nuclei from the
homozygous mutant flies is due to increased replication of
heterochromatic sequences (30). (B) Nuclei from follicle
cells (8c to 32c) and from nurse cells (64c to 512c) were collected and
pooled. DNA was isolated and analyzed as described for Fig. 2. The
18-kb DNA is the NotI fragment flanking the euchromatic P
element inserted into the cycE gene that creates the
cycE1672 mutation.
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Truncation of heterochromatic DNAs occurs on all chromosomes.
Truncation of heterochromatic DNA was also analyzed on the
Dp1187, second, and fourth chromosomes. Ovary nuclei were
isolated from flies carrying the Dp1187 minichromosome and
sorted by flow cytometry into three classes: diploid nuclei (2c to 4c),
follicle cell nuclei (8c to 32c), and nurse cell nuclei (64c to 512c). Truncated DNAs were analyzed in the three samples by in-gel
hybridization. Analysis was done on both undigested and restriction
enzyme-digested DNA. Consistent with what was observed for
CH(3)336, only full-length Dp1187 molecules were
detected in diploid nuclei while truncated DNAs were detected in the
polytene nuclei of both follicle cells and nurse cells (Fig.
4). Significant quantitative differences between follicle cells and nurse cells were observed in the abundance of truncated Dp1187 DNAs. Full-length Dp1187
chromosomes were abundant in follicle cell nuclei but were barely
detectable in nurse cell nuclei, suggesting that replication of
heterochromatic sequences of Dp1187 is more efficient in
follicle cells (Fig. 4). Qualitative differences were also observed.
The population of truncated Dp1187 DNAs formed in nurse cell
nuclei and revealed by NotI restriction digestion was 130 kb
larger, on average, than was the population of truncated DNAs formed in
follicle cell nuclei (Fig. 4, +1000 samples). This observation is
consistent with earlier experiments in which truncated DNAs of two
different sizes were observed in NotI-digested
Dp1187 DNA isolated from whole ovaries (15).
Finally, two distinct species of truncated Dp1187 molecules were detected in uncut DNA from both follicle cell and nurse cell nuclei (Fig. 4).
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FIG. 4.
Truncation of heterochromatic DNAs occurs on all
chromosomes. Nuclei from flies carrying the Dp(1;f)1187,
CH(2)423, or wild-type fourth chromosome (Chrom 4) were isolated
from ovaries and sorted by flow cytometry into discrete ploidy classes.
Nuclei from 2c to 4c follicle cells (dip), from 8c to 32c follicle
cells (Fc), and from 64c to 512c nurse cells (Nc) were collected and
pooled. DNA was isolated, restriction endonuclease digested,
fractionated by pulsed-field gel electrophoresis, and analyzed by
in-gel hybridization. Dp(1;f)1187 DNA was digested with
PmeI (+522) or NotI (+1000) or left uncut.
CH(2)423 DNA was digested with NotI, and
fourth-chromosome DNA was uncut. DNAs detected in 2c to 4c follicle
cells (dip) are full-length restriction fragments or chromosomes, and
the smaller DNAs detected in follicle (Fc) and nurse cells (Nc) are
truncated molecules. Autoradiograms of the follicle and nurse cell
samples for the fourth chromosome had to be manually aligned due to
severe skewing of the lanes during electrophoresis in the region of the
wells.
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Second-chromosome heterochromatin was analyzed using chromosome
CH(2)423, which has a P element inserted into
heterochromatic
sequences in pericentromeric region h42-43 (Fig.
1)
(
63). Ovary
nuclei were isolated from flies carrying the
CH(2)423 chromosome
and sorted by flow cytometry as
described above. Truncation of
the flanking heterochromatic restriction
fragment was analyzed
by Southern blot hybridization. Only the
full-length restriction
fragment was detected in diploid nuclei;
truncated DNAs were detected
only in the polytene nuclei of follicle
cells (Fig.
4). It is
interesting that truncated DNAs were most
abundant in follicle
cell nuclei for
CH(2)423 and
CH(3)336 (Fig.
2 and
4) but were
most abundant in nurse cell
nuclei for
Dp1187 (Fig.
4). Since
the abundance of truncated
DNAs is inversely correlated with the
extent of heterochromatic
replication (Fig.
3), these observations
suggest that the efficiency of
replicating heterochromatic sequences
varies between different regions
of heterochromatin in the same
cell and is not a genomewide property of
all heterochromatic sequences
in any single cell
type.
Truncation of heterochromatic DNAs was also analyzed on the wild-type
fourth chromosome, which is small enough to be directly
resolved by
pulsed-field gel electrophoresis (
32). Ovary nuclei
isolated
from flies carrying a wild-type fourth chromosome were
sorted by flow
cytometry, and their DNA was analyzed by in-gel
hybridization.
Full-length 4,125-kb chromosomes were detected
only in diploid nuclei
whereas truncated chromosomes of 2,595
and 1,025 kb were detected in
polytene nuclei of both follicle
cells and nurse cells (Fig.
4). No
full-length fourth chromosomes
were detected in either follicle or
nurse cell nuclei (Fig.
4).
Except for the difference in size, the
relative pattern of truncated
DNAs observed for the fourth chromosome
was strikingly similar
to the pattern observed for uncut
Dp1187 (Fig.
4). This suggests
that underreplication of
satellite sequences and the resulting
truncation of heterochromatic
DNAs occurs by the same mechanism
on both the
Dp1187 and
fourth chromosomes. A quantitative difference
between the
Dp1187 and fourth chromosomes was observed in follicle
cell
nuclei, where full-length
Dp1187 chromosomes were detected
but full-length fourth chromosomes were not (Fig.
4).
Truncation of heterochromatic DNA is not sensitive to modifiers of
PEV.
Since packaging of DNA into heterochromatin suppresses
transcription, we considered whether heterochromatin might play a
similar role in suppressing replication and thereby contribute to
heterochromatic underreplication in polytene cells. If the
heterochromatin structure contributes to underreplication, genetic
modifiers of that structure might alter the truncated DNAs that form as
a consequence of underreplication. Heterochromatin structure can be
altered using modifiers of position effect variegation (PEV), which
occurs when gene transcription becomes variably suppressed due to the
proximity of a gene to heterochromatin (21, 60). Several
modifiers of PEV, including the Y chromosome,
Su(var)2052, parental source, and temperature,
were tested, but none significantly influenced the formation of
truncated heterochromatic DNAs. The Y chromosome is a strong
suppressor of PEV (49) and suppresses PEV of the
rosy gene on CH(3)336 (63) and the
yellow gene on Dp1187 (25). Truncation
of heterochromatic DNA on the CH(3)336, Dp1187, and fourth
chromosomes was analyzed in salivary glands from X/O and
X/Y animals. The Y chromosome had no effect on
the truncation of CH(3)336 heterochromatin (Fig.
5A) or fourth-chromosome heterochromatin
(data not shown), but it did have a quantitative effect on the overall
replication of the Dp1187 chromosome (Fig. 5B). Total
Dp1187 DNA was less abundant in salivary glands from X/O animals than in those from X/Y animals (Fig.
5B). This effect appeared to be a general influence on total
Dp1187 copy number, consistent with earlier analysis of PEV
on Dp1187 (25), and not a specific effect on
truncation of heterochromatic DNAs. The truncated DNAs detected in
X/O animals, although less abundant, were the same size and
of the same abundance relative to the full-length restriction fragment
as those observed in X/Y animals (Fig. 5B). This suggests
that Dp1187 replication that occurred in X/O
salivary glands was subject to the same relative amount of
underreplication producing the same truncated DNAs.
Su(var)2052 is a suppressor of PEV and creates a
missense mutation in the HP1 protein, a known component of
heterochromatin (41). The Su(var)2052
mutation had no effect on the truncation of CH(3)336
heterochromatin in follicle cell, nurse cell, or salivary gland
chromosomes (data not shown). Transmission of a chromosome through the
female or male germ line influences the extent of PEV on that
chromosome in the subsequent generation, with passage through the male
germ line enhancing PEV (49). Parental source effects have
been demonstrated for the Dp1187 chromosome (25).
The parental source, however, had no effect on the truncation of
heterochromatic DNAs on the CH(3)336 or Dp1187
chromosomes analyzed in the follicle and nurse cells of the ovary (data
not shown). Finally, temperature influences PEV, with low temperatures
during development enhancing PEV and high temperatures suppressing PEV
(49). High temperatures suppress PEV of genes on
heterochromatic P elements like the one on CH(3)336 (63). Flies grown at 18 and 27°C showed no difference in
the truncation of heterochromatic DNA on the CH(3)336,
DP1187, or fourth chromosomes in follicle and nurse cells of the
ovary (data not shown).
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FIG. 5.
Truncation of heterochromatic DNA is not sensitive to
modifiers of PEV. (A) Salivary glands were dissected from
X/Y or X/O third-instar larvae carrying the
CH(3)336 chromosome. Total salivary gland DNA was isolated
and analyzed as described for Fig. 2. (B) Salivary glands were
dissected from X/Y or X/O third-instar larvae
carrying the Dp(1;f)1187 chromosome. Total salivary gland
DNA was isolated, digested with NotI, and analyzed using a
two-dimensional electrophoresis technique that physically separates DNA
molecules derived from the normal X chromosome and
Dp(1;f)1187 chromosome, both of which are present in these
animals. Without separation, hybridization to the X
chromosome NotI fragment would obstruct the hybridization
signal from truncated Dp(1;f)1187 DNAs.
|
|
| |
DISCUSSION |
Model for heterochromatic underreplication and the structure of
polytene chromosomes.
Heterochromatic underreplication in polytene
cells occurs when S phase ends before late replication of
heterochromatic DNA has been completed. This is probably due to changes
in cyclin E regulation (30). Premature termination of S
phase would produce unresolved replication forks between fully
replicated euchromatic and underreplicated heterochromatic sequences
(26). The truncated heterochromatic DNAs described in this
report, however, were not detected in the initial 4c nuclei that
underreplicate their DNA, suggesting that truncation is not caused by
simple breakage of unresolved replication forks (Fig. 2). Instead,
truncated DNAs were detected after the second underreplication S phase.
How could the abundance of truncated DNAs be dependent on the extent of underreplication yet require two underreplication S phases to form
(Fig. 2 and 3)? We propose that truncated DNAs are produced when
replication forks in the second polytene S phase extend to the position
where unresolved replication forks formed in the first polytene S phase
(Fig. 6). DNA synthesized during the
first S phase provides the template strands for DNA synthesis in the second S phase, and since these template strands simply end at unresolved replication forks, replication in the second S phase would
produce truncated linear DNAs (Fig. 6). This "replication runoff"
model not only explains why truncated DNAs would appear specifically in
the second S phase but also explains why their abundance would be
dependent on the degree of underreplication; specifically, the fraction
of replication forks left unresolved in the first S phase would
determine the abundance of truncated DNAs produced in the second S
phase. Furthermore, the gradual increase in the abundance of truncated
DNAs that is observed as nuclei increase in ploidy (Fig. 2) would be
caused by forks that complete replication in early S phases but are
left unresolved in later S phases, leading to the production of
additional truncated DNAs. While the replication runoff model explains
the origin of truncated linear DNAs, it does not preclude the
possibility that underreplication also creates DNA molecules with
structures of sufficient complexity that they are unable to migrate out
of the sample wells during electrophoresis (Fig. 2). Such molecules, if
they exist, would probably have structures more complex than simple
replication forks (14).
View larger version (34K):
[in this window]
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|
FIG. 6.
Model for heterochromatic underreplication and the
structure of polytene chromosomes. Double-stranded DNA in a
hypothetical region of Drosophila pericentromeric
heterochromatin is shown, with NotI restriction sites (N)
and a region of probe homology indicated. Satellite repeats inhibit
their own replication by blocking fork elongation. The barrier formed
by satellite repeats could reflect an intrinsic property of highly
repetitious sequences or could result from the compaction of satellite
repeats into a chromatin structure distinct from other heterochromatic
sequences. The first polytene S phase ends before replication forks
stalled at satellite barriers are resolved, causing heterochromatic
underreplication. Truncated linear DNAs are generated in the second
polytene S phase, when replication forks extend to the same barriers
where forks were left unresolved in the first polytene S phase. DNAs
are shown with blunt ends for clarity, but they would probably have
staggered ends, particularly on lagging strands. Once produced,
truncated DNAs would be replicated in each subsequent S phase.
|
|
In the replication runoff model, the sizes of truncated DNAs are
determined by the locations of unresolved replication forks
produced in
the first underreplication S phase (Fig.
6). Although
they formed a
heterogeneous population, most truncated DNAs were
of preferred sizes
(Fig.
2 and
4), suggesting that unresolved
forks were located at
preferred positions when S phase ended.
While those positions could be
where actively elongating replication
forks happen to be "caught"
when S phase ends, the data suggest
that the size of truncated DNAs is
not directly determined by
the length of the S phase. For example, the
cycE1672 mutation reduced the abundance of
truncated DNAs by increasing
the length of S phase (Fig.
3)
(
30), yet the truncated DNAs
detected in
cycE1672 flies were not larger, as would be
expected if the length of
the S phase were determining the size of
truncated DNAs (Fig.
3). How does extending S phase allow more
heterochromatic replication
without changing the size of truncated
DNAs? If, during S phase,
replication forks stall for extended periods
at barriers to fork
elongation, the locations of such barriers would be
the preferred
positions at which replication forks would most often be
caught
when the S phase ends. By extending the length of the S phase,
the
cycE1672 mutation would increase the
probability that forks are able to
traverse these barriers and complete
replication before S phase
ends, but the forks still left unresolved
would most often be
located at the same barriers, producing the same
sized truncated
DNAs. Transient stalling of replication forks at fixed
barriers
would also explain the pattern of truncated DNAs produced in
CH(3)336 heterochromatin. The same 340- and 77-kb truncated
DNAs were detected
in all tissues analyzed, but their abundances
relative to the
full-length restriction fragment differed
significantly. In 64c
nurse cell nuclei, truncated DNAs were
approximately equal in
abundance to the full-length molecules (Fig.
2);
in 8c follicle
cell nuclei, truncated DNAs were about twofold more
abundant than
full-length molecules (Fig.
2); and in the 1,024c nuclei
of salivary
glands, only truncated DNAs, including a significant amount
of
77-kb DNAs, and no detectable full-length molecules were detected
(Fig.
5). These observations are consistent with stalling of
replication
forks at the same 340- and 77-kb barriers in all three
tissues
but with the probability of forks traversing these barriers
before
S phase ends differing, with forks being caught least often in
nurse cells and most often in salivary
glands.
Tissue-specific differences in the abundance of truncated DNAs, as
discussed above for
CH(3)336, appear to be a property of
a
given region of heterochromatin and not a consequence of
tissue-specific
differences in overall S-phase length, which would
affect all
regions of heterochromatin in the same way. For example,
truncated
DNAs were more abundant in follicle cells than nurse cells
for
heterochromatin assayed on the second and third chromosomes (Fig.
2
and
4) but the opposite was true for heterochromatin assayed
on the
Dp1187 chromosome, where truncated DNAs were more abundant
in nurse cells than in follicle cells (Fig.
4). Therefore, whether
a
replication fork is able to complete replication or is left
unresolved
when S phase ends must be determined, at least in part,
by more
regional parameters, such as the location of origins used
to replicate
specific segments of heterochromatin or when those
origins fire during
S
phase.
Underreplication in polytene chromosomes occurs primarily to satellite
sequences (
13,
19,
20). Complex sequences present
in
heterochromatin, such as islands of transposable elements,
can
replicate efficiently despite their heterochromatic location
(
1,
62). If replication barriers exist in heterochromatin,
satellite
sequences themselves could be the barriers that inhibit
their own
replication. At least some satellite sequences can have
strong
intrinsic curvature (
33,
42), and some microsatellite
repeats can cause stalling of DNA polymerases (
44). Such
properties
multiplied over hundreds of contiguous kilobases of
repetitious
DNA might present a barrier to fork elongation. It is also
possible
that compaction of satellite sequences into heterochromatin
could
inhibit fork elongation. While the inability of PEV modifiers
to
influence underreplication suggests that heterochromatin structure
does
not play a role (Fig.
5), PEV modifiers have been identified
based on
their effects on transcription, not replication, and
the components of
heterochromatin that suppress transcription
may not be the same as
those that suppress replication. Furthermore,
PEV modifiers have
usually been selected as mutations that influence
the ability of
heterochromatin to act at a distance, typically
across a
euchromatic-heterochromatic breakpoint. The heterochromatin
into which
satellite sequences are packaged may not be as sensitive
to PEV
modifiers as the heterochromatin that "spreads" into normally
euchromatic sequences. Finally, the size difference that was observed
between truncated
Dp1187 DNAs detected in follicle cells
versus
nurse cells (Fig.
4) is difficult to explain if the sizes of
truncated
DNAs were determined solely by the primary sequence, which is
the same in both cell types. Such size differences are more easily
explained if the barrier were a chromatin structure whose position
relative to underlying sequence could differ between cells. Thus,
satellite sequence, either directly or through their compaction
into
heterochromatin, may inhibit their own replication by blocking
fork
elongation.
The replication runoff model of polytene chromosome structure predicts
that DNA molecules with double-stranded free ends are
produced at sites
of stalled replication forks (Fig.
6). Since
polytene cells undergo
repeated S phases despite experiencing
incomplete DNA replication, as
well as skipping mitosis, they
clearly lack normal checkpoint controls
(
29,
52,
61) and
therefore might accumulate DNA containing
double-stranded free
ends without triggering the cell cycle arrest that
such structures
would induce in diploid cells. In fact, using
ligation-mediated
PCR, truncated
Dp1187 molecules have been
cloned from ovarian
nurse and follicle cells, and they have structures
consistent
with the presence in these polytene tissues of
Dp1187 DNAs containing
double-stranded free ends (B. Calvi
and A. Spradling, personal
communication). On occasion, polytene cells
might still attempt
to repair double-stranded free ends and, in so
doing, might create
ectopic fibers, a characteristic feature of
polytene chromosomes.
Nonhomologous end-joining reactions between
free-ended DNAs at
different loci would produce ectopic ligations
between regions
of the genome that were underreplicated but not
necessarily homologous
by sequences, both of which are characteristic
features of ectopic
fibers (
64). Ectopic ligations would
also create chimeric molecules
more complex in structure than the
simple truncated DNAs proposed
in Fig.
6 and would complicate attempts
to localize replication
barriers, since the size of truncated DNAs
would be determined
not only by the position of fork arrest but also by
structural
changes introduced by ectopic ligation. Mapping of a series
of
truncated
Dp1187 DNAs onto the known heterochromatic
sequence
structure of
Dp1187 and assuming a simple
truncation of DNA resulted
in differing positions of putative
replication barriers depending
on the restriction enzyme used for the
analysis (
15). Perhaps
the source of this ambiguity is, in
part, a consequence of ectopic
ligations.
Heterochromatic replication origins.
Sequences flanking the
CH(3)336 P element were fully replicated in polytene
chromosomes despite their heterochromatic location, and the level of
replication was unaffected by the Y chromosome (Fig. 5)
(62). Even though replication is not suppressed, sequences flanking the P element are likely to be compacted into heterochromatin, since expression of the rosy gene present on the
CH(3)336 P element is subject to strong
heterochromatin-induced suppression (63). This contrasts to
the Dp1187 chromosome, whose replication is both suppressed
by heterochromatin and sensitive to modulation by the Y
chromosome (Fig. 5) (25). Why should replication in these
two chromosomes differ with respect to their sensitivity to
heterochromatin? The answer may be that sequences flanking the
CH(3)336 P element occur normally within heterochromatin
(63), while sequences of Dp1187 contain an
abnormal euchromatic-heterochromatic junction (27). Origins
that replicate the heterochromatic sequences flanking the
CH(3)336 P element may be specifically adapted to function
in heterochromatin, analogous to heterochromatic genes, which must be
located in heterochromatin for proper expression (60). On
the other hand, Dp1187 may be replicated by euchromatic origins that are suppressed by their proximity to heterochromatin on
the rearranged chromosome, analogous to the suppression of euchromatic
genes by heterochromatin-induced PEV. The existence of replication
origins specifically adapted to a heterochromatic environment is
supported by the replication behavior of the light gene.
light is fully replicated in polytene chromosomes despite its location within pericentromeric heterochromatin, but when it is
placed into a euchromatic environment by chromosome inversion, its
replication is suppressed consistent with the presence near light of a replication origin requiring a heterochromatic
environment for function (21a).
Implications for diploid cells.
If replication barriers exist
in diploid cells such that stalled replication forks occur normally
when cells replicate heterochromatic sequences, this would have
interesting implications for the biology of heterochromatin. For
example, heterochromatic sequences are characteristically the last
portion of the genome to be replicated (31). The mechanism
of late replication is not known, but origins responsible for
replicating heterochromatin might be intrinsically late firing or might
be located so far into euchromatin that passive replication of
heterochromatic sequences does not occur until late S phase
(53). If replication barriers exist in heterochromatin, however, inhibited fork elongation could also cause late replication. Elongating replication forks might arrive at heterochromatin in early S
or mid-S phase but might be unable to elongate through heterochromatic
barriers until late S phase, when a stochastic process or a specific
cellular event finally allows replication to be completed. In this
scenario, such barriers would have a significant impact on the
replication timing of specific regions of heterochromatin, and
replication timing may play an important role in centromere formation,
particularly in relation to the temporal expression pattern of the
centromere-specific histone variant CENP-A/Cse4 (9, 23, 48).
By their nature, replication forks cause dramatic changes in the
structure of both DNA and chromatin. Stalled forks may even
create
transient double-stranded free ends via isomerization of
the
replication fork and annealing of newly synthesized DNA strands
(
2,
10,
35). If stalled forks are common within
heterochromatin,
the structural perturbations they create could make
heterochromatin
prone to DNA rearrangements. For example, stalled forks
could
create preferred insertion sites for transposable elements
(
5,
36,
55,
59) and could explain, in part, why
heterochromatic
domains characteristically contain a high density of
transposons
(
7,
39,
40,
54). Structural polymorphisms
are also characteristic
of heterochromatic domains (
57) and
might arise by unequal exchange
within satellite repeats when a
cell misinterprets a fork stalled
at a heterochromatic barrier as
evidence of DNA damage and attempts
to repair the "lesion" by
homologous
recombination.
Given the unique characteristics of heterochromatin, it is perhaps not
unexpected that the way in which cells replicate heterochromatin
might
differ in important aspects from the way in which they replicate
euchromatin. Studying those differences will provide important
insights
into the biology of
heterochromatin.
| |
ACKNOWLEDGMENTS |
We are grateful to A. Spradling, P. Zhang, and G. Karpen for fly
strains and to J. Lock for zfh-2 DNA. We acknowledge the Wadsworth Center Cellular Immunology and Molecular Genetics Core Facilities for assistance with flow cytometry and contour-clamped homogeneous electric field electrophoresis, respectively.
This work was supported by grant GM53476 from the National Institute of
General Medical Sciences.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Wadsworth
Center, NYS-DOH, P.O. Box 22002, Albany, NY 12201-2002. Phone: (518)
473-4201. Fax: (518) 474-3181. E-mail: glaser{at}wadsworth.org.
| |
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Abstract
Introduction
