Specific Aim:
Many human diseases are caused by mutations within specific genes. Examples
of such diseases include sickle cell anemia, Huntington’s disease, and Xeroderma
pigmentosa. It is unclear however, what makes these disease genes particularly
susceptible to genetic mutation. It has previously been demonstrated that several factors
can influence the mutability of regions of DNA. One of these factors is nuclear
replication timing. Late-replicating regions of DNA have been shown to be more prone
to mutation than early-replicating regions. Furthermore, replication timing has been
demonstrated to correlate with nuclear positioning. Late-replicating regions of DNA tend
to be located near the nuclear periphery while regions of DNA that replicate earlier in the
S phase of the cell cycle are more often located in the euchromatic regions of the nucleus.
It is therefore possible that nuclear position may place certain regions of DNA, such as
disease genes, at a higher risk of mutation.
The purpose of this research proposal is to identify the nuclear position, and
replication timing of the xeroderma pigmentosum variant gene in post-temporal decision
point nuclei. In this experiment, fluorescence in situ hybridization (FISH) will be used to
demonstrate the location of the XPV gene in the nucleus, as well as to verify the precise
replication timing of the locus. Additionally, the nuclear position of the XPV gene will
be shifted, and the gene sequenced after multiple cell cycles, to determine if nuclear
position directly influences the rate of accumulation of mutations. If shifting the nuclear
positioning and/or replication timing of the XPV gene alters the mutation rate we may
conclude that these are factors which put the XPV gene at a higher risk for mutation.
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Ultimately, the goal of this study is to further understanding of the factors which make
disease genes more prone to mutations during replication.
Preliminary Studies/Significance:
Elucidating the reasons for increased mutability of specific regions of DNA is the
first step in pinpointing the origin of inherited human diseases. One promising possibility
is that the replication-timing of different DNA regions may influence the rate at which
they sustain mutations. DNA is not a smooth, continuous process, but proceeds in spurts,
pausing occasionally as lagging-strand synthesis catches up with leading-strand. It is
during these stalls that DNA polymerases are most prone to make a mistake that will lead
to mutations in the DNA. Recent evidence suggests that particular regions of latereplicating DNA, or regions near the transition from early- to late-replicating may be
subject to a higher rate of mutation (Watanabe et al., 2002).
Unlike bacterial and viral genomes, eukaryotic genomes possess multiple origins.
Replication of the eukaryotic genome initiates at selected origins and proceeds in a
specific temporal order during S phase of the cell cycle. The region of DNA that is under
the control of a specific origin is known as a replicon. In many cases, groups of adjacent
origins fire simultaneously. Such groups of simultaneously active replicons are called
replicon clusters (Zink, 2006). The temporal order of replicon activation is carefully
controlled, resulting in the sequential replication of chromosomal subregions (Cimbora
and Groudine, 2001)
Inside the nucleus, DNA is not distributed uniformly, but assembled into distinct
formations called replication foci (RF). Recent studies have indicated that each RF is
actually a replicon cluster organized into a three-dimensional structure. The current view
2
is that chromosomes are organized into stable subunits known as sub-chromosomal foci
(SF), which are maintained throughout all phases of the cell cycle. During S-phase, all of
the proteins necessary for DNA replication assemble at the SF, thus converting the SF
into RF.
During DNA replication, RF are observed to localize at specific regions of the
nucleus during different temporal stages of S-phase. During the early stages of S-phase,
many small RF are found within the nuclear interior, but are absent from nucleoli or
perinuclear regions of the nucleus. The progression of S-phase is accompanied by a
disappearance of RF in the nuclear interior, and the appearance of a smaller number of
larger-sized RF focused around perinuclear and perinucleolar regions. More importantly,
these patterns of RF distribution are maintained in the organization of SF throughout
interphase, and are also observed in the nuclei of all daughter cells, suggesting that
regions of DNA with specific replication timing always occupy the same position in the
nucleus (Sadoni et al., 2004).
The organization of chromatin is not inherited through mitosis however, but
established de novo shortly after cells enter G1 (Thomson, 2004). After associating with
its specific nuclear position, a particular region is more or less stationary and undergoes
only locally confined movements (Chubb, 2002). Several studies have demonstrated that
the replication machinery of DNA does not move uniformly through the nucleus but
rather assembles and disassembles, following established paths between neighboring SF
(Sporbert et al., 2002; Sadoni et al., 2004). This is important for two reasons. First, it
helps explain the observed temporal organization of RF; and second, it implies that
something about the presence of replication machinery at one origin is sufficient to
3
activate adjacent origins. While the precise nature of this activation is uncertain, it is
likely that activation of adjacent origins is due to the destabilization of local chromatin
structure from the incoming of a replication fork, or replication of adjacent DNA (Zink,
2006). Whatever the case, nuclear positioning appears to play a critical role in the
control of replication timing.
This introduces one of the key aspects of the temporal control of replication,
chromatin structure. Several studies have indicated the importance of local chromatin
structure in the determination of replication timing. According to Heun et al. (2001),
there are three types of late-firing origins. The first includes origins directly adjacent to
telomeres. In this case, late-firing seems to be maintained by SIR complexes on
nucleosomal fiber, which block large enzymes from access to the DNA. The second type
are still telomere-proximal, but their replication timing is not affected by sir mutations.
The third type are far from telomeres, yet appear to have patterns of histone
modifications that mark them for late initiation or contact with nuclear periphery (Heun
et al., 2001). In each case, the highly-condensed nature of local chromatin is implicated
in the delay of activation.
The importance of chromatin condensation in control of replication timing was
further demonstrated by Engelman et al. (2005), who demonstrated that by
hyperacetylating inactive, highly-condensed chromatin near the nuclear periphery it is
possible to shift a gene in this region into the euchromatic interior compartment, and
switch to earlier replication timing. This, along with previously mentioned data (Sporbert
et al., 2002; Sadoni et al., 2004) suggests that both nuclear position and local chromatin
structure have important roles in replication timing. Higher-order folding of local
4
chromatin could sterically hinder assembly of the replication machinery at an origin,
while the fact that replication machinery moves sequentially between adjacent SF, and
the observation that loci nearly always adopt the timing of their corresponding nuclear
compartment highlight the importance of nuclear positioning.
Studies have been performed that demonstrate the effect of nuclear positioning, as
well chromatin environment on the determination of replication timing (Sadoni et al.,
2004; Engelman et al., 2005). It has also been demonstrated that SNP’s tend to occur with
greater frequency in regions of late-replicating DNA, and that a very high proportion of
cancer-related genes are found in early/late transition regions (Watanabe et al., 2002)
What remains to be shown is whether the mutation rate of a known late-replicating gene
can be reduced by shifting the gene out of a peripherally-located, late-replicating region
of DNA and into a euchromatic region that replicates early in S-phase. Such a shift can
be induced through treatment with a histone deacetylase inhibitor (Engelman et al.,
2005), and it is likely that once the shift is made, the gene will remain in its new location
throughout future cell cycles (Gilbert, 2002). Once this is accomplished, the rate of
mutation can be determined by comparing the sequence of the target gene after a
determined number of generations in a cell line with the shift in nuclear positioning, and
a control cell line. Information acquired through the indicated study will contribute to
further understanding of the structure and function of genome organization, particularly
in relation to replication timing. Furthermore it will provide insight into the reasons for
increased mutability of certain DNA regions, with particular relevance to the
pathogenesis of diseases.
5
Experimental Design and Methods:
To study the effect of nuclear positioning and replication timing on the
mutability of specific gene regions within the chromatin, procedures described by several
previous studies are followed (Zink et al., 1998; Heun et al., 2001; Zink et al., 2004;
Englmann et al., 2005). These studies utilize fluorescence in situ hybridization (FISH)
analysis to pinpoint the nuclear positioning of specific genes. They also demonstrate that
treatment with trichostatin-A (TSA) can shift a given gene from the nuclear periphery
into the nuclear interior.
For the first experiment, HeLa cells (or another immortal human cell line) are
cultivated and separated into several treatment groups. In order to establish conclusively
whether the nuclear positioning and replication timing of a particular region of DNA
increases the likelihood that the region will incur mutations, the XPV gene is shifted from
the nuclear periphery to the nuclear interior. This is accomplished by treating one group
of cells with trichostatin A (TSA) for 10 hours. A second, untreated group of cells serves
as a negative control. TSA is a histone-deacetylase inhibitor. By preventing histonedeacetylation, which is associated with chromatin condensation, TSA has been shown to
shift genes associated with late-replicating heterochromatin into euchromatic regions of
the nucleus (Zink et al., 2004; Englmann et al., 2005). It appears likely that once these
patterns of histone acetylation are established, the system becomes self-propagating and
the shifted gene will remain early-replicating and associated with euchromatin throughout
all future cell cycles (Gilbert, 2002; Zink, 2006).
6
To verify the shifting of the XPV gene into the nuclear interior, TSA-treated and
–untreated cells are fixed with formaldehyde for 10 minutes, to prepare for FISH
analysis. A short probe complementary to the coding region of the XPV gene is used.
The probe is specific to the 3’ region of the Pol eta domain (Figure 1). The probe for
XPV is labeled with biotin-UTP, by PCR. PCR has been found to be more efficient than
nick translation for the labeling of relatively short probes (Henegariu et al., 2001).
Additionally, a painting probe for chromosome 6 is used as a control for the specificity of
the XPV gene signals. As the XPV gene is located within chromosome 6 (Figure 2), it is
expected that the XPV signal should localize within, or at the periphery of, the
chromosome 6 territory, as defined by the FISH signal of the painting probe. The
painting probe is labeled with digoxigenin-dUTP, also by PCR. The probes are
denatured and applied to the slide containing formaldehyde-fixed cells. Hybridization is
allowed to proceed for 12-18 hours, and then the slides are washed in SSC/SDS to
remove any non-bound probe.
Figure 1: Coding domain of the Pol eta gene (XPV).
7
XPV
Figure 2: Location of XPV gene on chromosome 6 (6p21.1-p12)
Cells are also immunostained with rabbit antibodies for H4Ac8 or mouse
antibodies for LAP2β. H4Ac8 (histone H4 acetylated at Lys 8) is known to be enriched
in regions of early-replicating and transcriptionally active chromatin in the nuclear
interior, and depleted in regions of later-replicating and transcriptionally inactive
heterochromatin (Sadoni et al., 1999). LAP2β (lamina-associated polypeptide 2 β) is an
integral protein in the nuclear membrane, which binds to chromatin at the nuclear
periphery (Nili et al., 2001). Relative localization of these signals with the XPV gene
signal gives information about the chromatin environment in which the XPV gene is
located. If XPV is located near the periphery, it is expected that the XPV signal will
appear in a region depleted of H4Ac8 signal, but strongly enriched in LAP2β signal (Zink
et al., 2004). As a control, cells are also treated with a labeled probe for α-globin, a
known early-replicating gene that associates with euchromatin in the nuclear interior (Li
et al., 2005). The signal for this probe will appear in a region highly-enriched in H4Ac8
signal, but isolated from LAP2β.
Biotinylated gene-specific probes (XPV) are detected with avidin-Cy3, and the
signal is enhanced with a biotinylated anti-avidin antibody and a second layer of avidinCy3. Digoxigenin-labeled probes (chromosome 6) are detected with FITC-conjugated
anti-digoxigenin antibodies. FITC is the reactive isothiocyanate form of fluorescein.
8
Primary antibodies for H4Ac8 and LAP2β are detected with Cy3.5-conjugated goat antirabbit antibodies, and Cy5-conjugated goat anti-mouse antibodies, respectively.
Confocal imaging is used to detect fluorescent signals from the tagged probes.
This technique scans across the fixed cells with a single point of excitation light, rather
than uniformly illuminating the entire specimen at once, as with epifluoresence
microscopy. This allows for much better resolution for the detection of fluorescent
signals.
Now that the nuclear positioning of the XPV gene before and after treatment with
TSA has been determined, it is necessary to verify whether the replication timing is
shifted as well. In order to determine the replication timing of the XPV gene locus, one
group of cells is treated with TSA for 10 hrs, while a second control group of cells
remains untreated. All cells are pulse-labeled with 5’-bromo-2’-deoxyuridine (BrdU)
shortly prior to fixation. Incorporation of BrdU results in sister chromatid labeling at the
second mitosis after the labeling event (Latt, 1973). In other words, BrdU is integrated in
regions of DNA that are being replicated, and labeling patterns can be used to determine
cell stage progression from early to late S-phase. BrdU labeling patterns are commonly
assigned to five different stages, I-V (Sadoni et al., 1999). In early S-phase, BrdU is
localized throughout the euchromatic regions of the nucleus, and gradually shifts
outwards towards the periphery as the cell proceeds through S-phase (Englmann et al.,
2005).
After pulse-labelling, TSA-treated and untreated cells are fixed either with
formaldehyde, or methanol/acetic acid, for 10 minutes to prepare for FISH analysis.
Fixation with methanol/acetic acid of one group of TSA-treated, and one group of
9
untreated cells, acts as a control to ensure that prolonged sister chromatid cohesion is not
mistaken for late-replication. It has been demonstrated that for some gene loci,
prolonged sister chromatid cohesion following replication of the locus can lead to
incorrect determination of replication timing (Azuara et al., 2003). Nonetheless, fixation
with methanol/acetic acid, rather than formaldehyde, disrupts association between sister
chromatids. By comparing the results obtained from formaldehyde fixation with the
results from methanol/acetic acid fixation, it is possible to determine whether sister
chromatid adhesion affects the appearance of doublet FISH signals (Englmann et al.,
2005). After cell fixation, probes for the XPV gene are denatured and applied to the
slides containing the four different groups of cells (Table 1). Hybridization is allowed to
proceed for 12-18 hours, and then the slides are washed in SSC/SDS to remove any nonbound probe. BrdU-labelled chromosome territories are detected with rat anti-BrdU
antibodies. These primary antibodies are detected with FITC-conjugated anti-rat IgG
antibodies. XPV is detected as described previously. Replication of the XPV gene locus
is identified by several different markers. First, the appearance of doublet XPV gene
signals indicates that the locus has replicated. Additionally, it is only when the XPV
gene is undergoing active replication that BrdU and XPV FISH signals colocalize
(Englmann et al., 2005).
Confocal imaging is once again used to detect the fluorescent signals associated
with XPV and BrdU. Replication of the XPV gene is correlated with the temporal stage
of S-phase to which cells were assigned according to BrdU-labeling patterns. Finally, the
stage of S-phase in which XPV replicates will be compared between the four groups of
cells.
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Table 1: Groups of cells to act as controls for the effect of TSA treatment on replication
timing, and for the influence of sister chromatid cohesion on recognition of locus
replication.
Grp.
TSA
Fixation
1
2
3
4
+
+
-
Formaldehyde
Methanol/Acetic
Formaldehyde
Methanol/Acetic
The final step is to determine whether shifting nuclear positioning and replication
timing can directly reduce the rate of accumulation of mutations in the XPV gene. To
accomplish this, one group of cells is treated with TSA for ten minutes, and another
group is left untreated. Both groups of cells are allowed to proceed through multiple
rounds of the cell cycle. After a specific number of replications, cells from both groups
are transferred to separate PCR tubes, and PCR amplification of the XPV gene is
performed (sequence from NCBI: Accession_NM006502). The XPV gene is sequenced
for both groups, and the sequences are compared with the wild type XPV sequence. If
association with late-replicating, highly condensed regions of DNA does lead to
increased mutation rate, then the XPV gene from untreated cells should have sustained a
greater number of changes than the XPV gene from TSA-treated cells.
Based on findings from these studies, it should be possible to determine whether
nuclear positioning and replication timing directly influence the risk of mutation for a
specific region of DNA. This in turn will allow recognition of genes or DNA regions
which are particularly at risk for mutation. This would be of great benefit to the scientific
and medical communities, particularly in terms of the identification of cancer or other
disease-causing mutations. Future research should address whether the nuclear
positioning of chromosome regions and specific genes varies between individuals or
11
ethnic groups. If this variation does exist, recognition of the variation could allow for
improved genetic screening techniques, and/or greater specialization of
phamarcogenetics.
12
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