© 2000 Oxford University Press
Nucleic Acids Research, 2000, Vol. 28, No. 21 4083–4089
Poly(dA·dT) sequences exist as rigid DNA structures in
nucleosome-free yeast promoters in vivo
Bernhard Suter, Georg Schnappauf and Fritz Thoma*
Institut für Zellbiologie, ETH-Zürich, Hönggerberg, CH-8093 Zürich, Switzerland
Received August 1, 2000; Revised and Accepted September 8, 2000
ABSTRACT
Poly(dA·dT) sequences (T-tracts) are abundant
genomic DNA elements with unusual properties
in vitro and an established role in transcriptional
regulation of yeast genes. In vitro T-tracts are rigid,
contribute to DNA bending, affect assembly in nucleosomes and generate a characteristic pattern of CPDs
(cyclobutane pyrimidine dimers) upon irradiation
with UV light (UV photofootprint). In eukaryotic cells,
where DNA is packaged in chromatin, the DNA structure
of T-tracts is unknown. Here we have used in vivo UV
photofootprinting and DNA repair by photolyase to
investigate the structure and accessibility of T-tracts
in yeast promoters (HIS3, URA3 and ILV1). The same
characteristic photofootprints were obtained in yeast
and in naked DNA, demonstrating that the unusual
T-tract structure exists in living cells. Rapid repair of
CPDs in the T-tracts demonstrates that these T-tracts
were not folded in nucleosomes. Moreover, neither
datin, a T-tract binding protein, nor Gcn5p, a histone
acetyltransferase involved in nucleosome remodelling,
showed an influence on the structure and accessibility
of T-tracts. The data support a contribution of this
unusual DNA structure to transcriptional regulation.
INTRODUCTION
Poly(dA·dT) sequences (called T-tracts) have been extensively
characterised in vitro. They adopt an unusual structure (here
referred to as T-tract structure) that is distinct from typical B-type
DNA. T-tracts are essentially straight and rigid (1–3) with a
compressed minor goove (4) and a short helical repeat of 10 bp
per turn, compared to 10.5 bp per turn determined for canonical
B-DNA (5,6). The structure of T-tracts is not sensitive to changes
in cations or salt concentration (7), but folding of T-tract DNA in
nucleosomes can disrupt the T-tract structure, indicating that
the structural constraints in nucleosomes dominate over those
of the T-tracts (8,9). Since the formation of DNA lesions
depends on the DNA structure (10,11), the unusual T-tract
structure generates a characteristic pattern of cyclobutane
pyrimidine dimers (CPDs) upon irradiation with UV light (UV
photofootprint). Tn-tracts in plasmid DNA with n ≥ 4 give high
yields of CPDs at the 3′-end, low yields in the middle and variable
yields towards the 5′-end, depending on the flanking base (12).
Because of their unusual structure, T-tracts influence the
conformation of longer stretches of free DNA and they interfere with the formation and arrangement of nucleosomes.
Phased runs of short T-tracts cause DNA bending (see for
example 13,14) and bending is abolished when the T-tract
structure is disrupted by a CPD lesion (15). Long poly(dA·dT)
sequences exclude nucleosome formation during in vitro
reconstitution (16–18) except at high temperatures (19). Short
T-tracts (∼20 bp) form nucleosomes (8,20–23), but appear to
be excluded from the centre (20,24). Whether T-tracts form
nucleosomes apparently depends on their length, the flanking
sequences and the reconstitution conditions. Thus, nucleosomes with different T-tracts may have differential stability
with respect to disruption and nucleosome positioning.
The unusual structure of T-tracts and their potential to
destabilise or exclude nucleosomes has implications for their
functional role in eukaryotic cells. In the yeast Saccharomyces
cerevisiae T-tracts >10 bp occur at frequencies higher than
expected in the genome (25). In the HIS3, DED1 and URA3
genes, as well as in many other genes, T-tracts act as upstream
promoter elements necessary for wild-type levels of transcription (26–28). Since transcription activation depends on the
length of the T-tract and T-tracts can be replaced by
poly(dG·dC), another rigid structure, it was suggested that the
T-tracts operate not by recruitment of specific transcription
factors, but rather by their intrinsic DNA structure. The T-tracts
might affect nucleosome stability and enhance accessibility for
binding of factors to nearby sequences (28). This hypothesis is
consistent with the observations that several T-tract promoters
of yeast genes (including URA3, HIS3 and DED1) are not
packaged in stable nucleosomes as judged by enhanced
nuclease sensitivity (22,29–31) and by rapid repair of UV
lesions by photolyase (32). There are, however, examples
where T-tracts were reported in nucleosomal regions (33,34),
indicating that T-tracts alone are not sufficient to disrupt
nucleosomes, but they might lead to a local distortion in the
nucleosome (35). From these observations it is not clear
whether and how T-tracts contribute to chromatin organisation
and whether additional factors are required to disrupt nucleosomes. The only T-tract binding protein identified so far in
yeast is datin. It recognises T-tracts >9–11 bp in vitro (36,37),
but an effect of datin on nucleosome formation is not known. It
was reported that Gcn5p, which is a histone acetyltransferase
*To whom correspondence should be addressed. Tel: +41 1 633 33 23; Fax: +41 1 633 10 69; Email: [email protected]
Present address:
Bernhard Suter, Department of Molecular and Cell Biology, 222 Stanley Hall, Berkeley, CA 9472, USA
4084 Nucleic Acids Research, 2000, Vol. 28, No. 21
of the yeast SAGA complex, acts as a co-activator of HIS3
transcription (38) by acetylation of histones in the promoter
region (39). This implies that nucleosome remodelling is
involved in structural organisation of the HIS3 promoter.
The central questions addressed here are whether T-tracts in
yeast promoters adopt the unusual rigid structure in vivo and
whether those T-tracts are incorporated in nucleosomes.
MATERIALS AND METHODS
Yeast strains
JMY1 [MATa his3-∆1 trp1-289 rad1-∆ ura3-52 YRpTRURAP
(URA3 ARS1)] and FTY117 [MATa his3-∆1 trp1-289 rad1-∆
ura3-52 YRpCS1 (HIS3 TRP1 ARS1)] have been described
(32). GSY5 (MATa ade2-1 ura3-1,15 trp1-1 leu2-3, 112 can1-100
rad1∆ gcn5∆::LEU2) and GSY2 (MATa ade2-1 ura3-1,15
trp1-1 leu2-3, 112 can1-100 rad1∆) were generated from
W303-1a by restoration of the genomic HIS3 gene using a
1.6 kb BamHI fragment with the pet56-HIS3-ded1 sequence of
YRpCS1 for gene replacement and by disruption of RAD1 and
GCN5, respectively. EWY1002C-1 [MATa his3-∆200 leu2-3-112
ura3-52 lys2-801(am) trp1-1(am)] and EWY1002C-2 [MATa
his3-∆200 leu2-3-112 ura3-52 lys2-801(am) trp1-1(am)
dat1∆::LEU2] (36) were provided by Dr E. Winter and transformed with YRpCS1 (HIS3 TRP1 ARS1) to yield BSY1 and
BSY2, respectively.
UV irradiation of yeast cells
UV irradiation and photoreactivation were as described (32).
Yeast strains were grown at 30°C in minimal medium (2%
dextrose, 0.67% Yeast Nitrogen Base without amino acids)
supplemented with the required amino acids to a final density
of ∼1 × 107 cells/ml. GSY2 and GSY5 were grown for 6 h in
the presence of 3-aminotriazol to induce HIS3 transcription.
Cells were harvested and resuspended in minimal medium to
yield ∼3 × 107 cells/ml. The cell suspension was transferred to
plastic trays to form thin layers and irradiated under four
Sylvania G15T8 germicidal lamps (predominantly 254 nm as
measured by an UVX radiometer; UVP Inc., CA). After irradiation the media were supplemented with the appropriate amino
acids and 3-aminotriazol (GSY2 and GSY5). Photoreactivation of
the cell suspension was done in plastic trays using Sylvania
Type F15 T8/BLB bulbs (peak emission at 375 nm) at ∼1.5 mW/
cm2 at 22–26°C. After different exposure times aliquots were
collected and chilled on ice. One UV-irradiated sample was
incubated at 23°C in the dark. DNA isolation was either done
by phenol extraction or using Qiagen Genomic-Tips. DNA was
dissolved in 10 mM Tris, 1 mM EDTA, pH 8.0 (TE). All postirradiation steps were carried out under yellow light to prevent
photoreactivation.
UV irradiation and photoreactivation of DNA
Purified DNA was exposed in droplets to UV light. For photoreactivation in vitro UV-irradiated DNA was mixed with
Escherichia coli photolyase (0.1 µg/µl) to a final concentration
of 0.1 µg photolyase/µg DNA and exposed in droplets to
photoreactivating light for 50 min at 2 mW/cm2.
Primer extension analysis
DNA was digested with EcoRI, purified and dissolved in TE.
Primer labelling and primer extension were done as described
(40,41) with modifications for detection of CPDs in genomic
DNA (42). The reaction products were analysed on 8% polyacrylamide gels. DNA damage at individual sites or at clusters
of sites was calculated from PhosphorImager scans as a
fraction of the total amount of signal per lane. The signal of
non-irradiated DNA was subtracted to yield total damage
[CPDs and pyrimidine (6–4) pyrimidone photoproducts (6–4PPs)].
The signal for DNA treated with E.coli photolyase displays
non-CPD lesions. For repair analysis, initial damage at each
site was set to 0% repair. The sequences of the primers were 5′TGTCATCTTTGCCTTCGTTTATCTTGC (HIS3 gene, top
strand), 5′-TGATTTATCTTCGTTTCCTGCA (URA3 gene,
top strand) and 5′-TATTATCTCTGCTATTTTGTGACG
(ILV1 gene, bottom strand).
RESULTS
UV photofootprinting was applied in yeast and to naked DNA to
identify the structures of T-tracts based on their characteristic
formation of CPD lesions (12). We focused on the promoter
regions of the yeast URA3 and HIS3 genes, since the T-tracts of
both genes act as transcriptional elements in vivo (26,27) and
are located in nuclease-sensitive regions (22,29).
Poly(dA·dT) sequences located in open promoter regions
adopt T-tract structure in vivo
The URA3 promoter was investigated in strain JMY1
harbouring the URA3 gene in a minichromosome
(YRpTRURAP; Fig. 1). For damage induction, protein-free
DNA and yeast cells in suspension were irradiated with UV
light (predominantly 254 nm) which generates preferentially
cis-syn CPDs and, to a much lesser extent, 6–4PPs (43). DNA was
purified and DNA lesions were assessed by primer extension with
Taq polymerase, which is efficiently blocked in front of the 3′-end
of CPDs and 6–4PPs (40,41). The intensities of the bands
reflect the yields of UV lesions. Irradiation conditions were
chosen that generated <0.1 CPD/molecule in the promoter
region.
Damage induction is shown for DNA irradiated in TE buffer
and in cellular chromatin (Fig. 1B, lanes 2 and 6). These
lesions were predominantly CPDs, since they could be
removed by incubation with E.coli photolyase in vitro (lanes 3
and 5) and by photoreactivation in vivo (lanes 7–12; see
below).
The URA3 promoter contains a T10-, a T8- and a T7-tract in
the top strand. In free DNA, damage induction in the T10- and
T7-tracts was highest at the 3′-end and lower towards the 5′-end
(Fig. 1B, lane 2 and enlargement, and C). Damage in the T8-tract
was also highest at the 3′-end, decreased towards the middle of
the tract, but slightly increased again towards the 5′-end. Both,
the T10- and T7-tracts are flanked by adenines at their 5′-ends,
whereas the T8-tract has a cytosine at this position (Fig. 1A).
This damage pattern observed in the T-tracts of free DNA is
characteristic for the unusual structure of T-tracts (12).
Dimethyl sulphoxide (DMSO) unwinds DNA by dehydration
(44), perturbs the unusual T-tract structure and generates a
more even CPD pattern upon UV irradiation (12). When naked
Nucleic Acids Research, 2000, Vol. 28, No. 21 4085
Figure 1. UV photofootprints and repair analysis of T-tracts in the URA3 promoter of strain JMY1 [rad1∆; YRpTRURAP (URA3, ARS1)]. (A) Schematic drawing
of the chromatin structure and DNA elements in the top strand including nucleosome positions (ovals; overlapping positions at the TATA box) and T-tracts (T10, T8
and T7) (29,30). (B) Primer extension products showing damage formation and repair. UV damage generated by UV light (preferentially CPDs and 6–4PPs) in
naked DNA: lane 1, in 50% DMSO; lane 2, in TE buffer; lane 6, in living cells (chromatin). Lanes 7–12, UV photoproducts after photoreactivation for 3–120 min
in vivo. Lanes 3 and 5, non-CPDs (preferentially 6–4PPs) revealed by treatment with E.coli photolyase. Lane 4, DNA of non-irradiated cells. A, G, C and T,
sequencing lanes. The T-tract regions of lanes 1, 2 and 6 are enlarged in the bottom panel. (C) Quantitative analysis. The graphs show the percentage of total DNA
damage generated at individual dipyrimidine sequences. (D) DNA repair curves. Repair of CPDs is plotted versus incubation time (min) in the presence of
photoreactivating light. The pyrimidine clusters 562 and 516–537 are located in the flanking nucleosome.
DNA was irradiated in 50% DMSO the damage yields were
similar at all positions within the T-tracts, demonstrating a loss
of the T-tract structure (Fig. 1B and C). We therefore conclude
that the T7-, T8- and T10-tracts of the natural URA3 promoter
adopt typical T-tract structures in aqueous solution.
When DNA damage was generated in yeast cells, the CPD
patterns in all T-tracts were very similar to those induced in
free DNA (Fig. 1B, lanes 6 and 2, and C). These data are direct
evidence that the T-tracts of the URA3 promoter adopt the
same rigid DNA structure in chromatin in vivo as in free DNA.
To test whether this observation also applies to other T-tracts,
we analysed the HIS3 promoter region in the minichromosome
YRpCS1 of strain FTY117. This region, which includes the
divergent HIS3 and PET56 promoters (45), is nuclease sensitive
and flanked by positioned nucleosomes (Fig. 2A). All T-tracts
in free DNA as well as in chromatin showed a CPD pattern
characteristic of T-tract structures, with enhanced damage
formation at the 3′-end and disruption of the structure upon
incubation in DMSO (Fig. 2B, lanes 1, 2 and 6). The enhanced
yields of lesions at the 5′-end of the T9-tract are attributable to
the flanking cytosine, as in the T8-tract of URA3. Thus, the
HIS3 promoter in the minichromosome is a second example
where T-tracts in vivo maintain the same rigid structure as in
isolated DNA.
4086 Nucleic Acids Research, 2000, Vol. 28, No. 21
constraints on the T-tracts in genomic chromatin which would
affect their structure.
Efficient CPD repair in poly(dA·dT) sequences by photolyase
Figure 2. UV photofootprints and repair analysis of T-tracts in the extrachromosomal HIS3 promoter of strain FTY117 [rad1∆; YRpCS1 (HIS3 TRP1
ARS1)]. (A) Schematic drawing of chromatin structure and DNA elements
(top strand; according to 22). (B) Primer extension products. For space
reasons, only the lower part of the gel is shown.
It is essential to establish whether T-tracts are incorporated in
nucleosomes, modified nucleosomes or complexed with other
proteins. We used CPD repair by photolyase (photoreactivation)
as a direct in vivo approach, in contrast to chromatin digestion
with nucleases, which is an in vitro assay. Photoreactivation is a
major pathway to remove UV-induced CPDs and it is modulated
by nucleosomes and proteins bound to DNA (32,48–50).
Nucleotide excision repair, which is the alternative pathway to
remove CPDs, was inactivated by disruption of the RAD1 gene
(rad1∆). Immediately after damage induction the cell suspension
was exposed to photoreactivating light for different times, the
DNA was extracted and the remaining lesions were analysed.
In a low resolution repair study we previously showed that
linker DNA between nucleosomes and whole open promoter
regions of the URA3 and HIS3 genes were repaired in 15–30
min, while repair of nucleosomes required ∼2 h (32). Thus,
photolyase is a new tool to measure whether a DNA lesion is in
a nucleosome in vivo. The high resolution data presented here
show rapid repair of all T-tracts in the URA3 and HIS3
promoters in minichromosomes (Figs 1B, lanes 7–12, and 2B,
lanes 7 and 8) and in the genome (Fig. 3A, lanes 6–11). After
30 min >90% of CPDs were removed at all positions within the
different T-tracts (T7, T10, T8 and CTTTC 618 in Fig. 1A and
D), while DNA lesions which map in the flanking nucleosome
were repaired at 40–50% (CTTT 562 and 516–537). Repair
was exclusively by photolyase. No repair occurred in the
absence of light (Figs 1B, lane 13, 2, lane 9, 3A, lane 12, and 3B,
lane 10). This efficient repair demonstrates that these T-tracts
were not folded in nucleosomes.
We noticed a subtle repair heterogeneity within T-tracts.
While all sites in the T9-tract of HIS3 were repaired at similar
rates, CPDs at the 5′-end of the T11-tract (excluding the TC dimer)
were more rapidly repaired than CPDs towards the 5′-end. This
was observed in the minichromosome and in the genome and
might indicate unknown factors which interfere with repair
(dots and arrowhead in Figs 2 and 3).
T-tract structure of poly(dA·dT) sequences in genomic
chromatin
The T-tract structures in the HIS3 promoter are
independent of Gcn5p
Chromatin folding involves several levels of DNA packaging
from nucleosomes to higher order structures (46). It has been
demonstrated that the arrangement of nucleosomes on the
DNA sequence (nucleosome positioning) depends on a
combination of DNA sequence, protein–DNA interactions,
boundaries and chromatin folding (47). It is therefore conceivable
that the location of a gene in chromosomes or small circular
minichromosomes affects the structure of T-tracts due to local
constraints imposed by packaging of DNA in chromatin. We
therefore tested whether the T-tract structure also occurs in the
genomic HIS3 gene. The HIS3 promoter has the same chromatin
structure in the genome as in the minichromosome, namely
with the T-tracts located in a nuclease-sensitive region (not
shown). The UV damage pattern in chromatin was very similar
to that observed in free DNA (Fig. 3A, lanes 1 and 5) and no
difference was observed between the genomic copy and the
HIS3 gene in the minichromosome. We conclude that the T-tract
structure is maintained in the genome and that there are no
Gcn5p is a histone acetyltransferase of the SAGA complex and
contributes to transcriptional activity of the HIS3 promoter
(38), possibly by nucleosome remodelling. Thus, it is conceivable
that Gcn5p activity is required to destabilise the nucleosomes
in the promoter and thereby allow T-tracts to adopt their rigid
structure. Figure 3B shows the results obtained in a strain with
a deleted GCN5 gene (gcn5∆). The photofootprints were
identical to those obtained in the GCN5 wild-type strain GSY2.
Thus, the T-tract structure in the HIS3 promoter is independent
of Gcn5p. Moreover, there was no obvious difference in photoreactivation of T-tracts, indicating that the T-tracts remained
accessible to photolyase in the absence of Gcn5p. Thus, both
the photofootprints and rapid repair data argue against Gcn5pdependent nucleosome remodelling in the T-tract region.
T-tract structures in vivo are independent of datin
Datin is the only known yeast protein that binds poly(dA·dT)
sequences of 9–11 bp (36,37). Datin behaves as a repressor and
Nucleic Acids Research, 2000, Vol. 28, No. 21 4087
Figure 3. UV photofootprints and repair analysis in the genomic HIS3 promoter in the presence and absence of Gcn5p. (A) Primer extension products of strain
GSY2 (rad1∆ GCN5). (B) Primer extension products of strain GSY5 (rad1∆ gcn5∆). Dots and the arrowhead indicate fast and more slowly repaired sites within
T11, respectively.
deletion of the datin gene. We conclude that the T-tract
structure as it occurs in chromatin in vivo is independent of
datin.
DISCUSSION
Figure 4. T-tract structure is independent of datin. (A) Primer extension
analysis of UV photoproducts in the HIS3 promoter of strains BSY1 (DAT1,
YRpCS1) and BSY2 (dat1∆, YRpCS1). (B) Primer extension analysis of UV
photoproducts in the ILV1 promoter of the same strains.
activator through poly(dA·dT) sequences of the HIS3 and ILV1
promoters, respectively (28,51). To investigate whether datin
affects the structure of T-tracts in vivo, yeast strains containing
the wild-type gene (DAT1) or a disruption (dat1∆) were transformed with YRpCS1 and the UV photofootprints in the HIS3
promoter were analysed (Fig. 4). There were no obvious differences in the CPD patterns of the T-tracts between the dat1∆
and wild-type (DAT1) strains and the patterns described above
(Figs 2 and 3). Similarly, the promoter of the genomic ILV1
gene showed typical T-tract patterns that were not affected by
A wide variety of experiments established that T-tract DNA
adopts an unusual rigid B-form structure in vitro and it was
inferred that this particular structure serves a functional role in
the genome. Here we show direct evidence that this unusual
structure exists in chromatin of living cells. We identified these
structures in promoters which are known to be transcriptionally
regulated by T-tracts. Moreover, we show that these T-tracts
are not folded in nucleosomes nor affected by datin or Gcn5p.
This suggests that the T-tract structure per se is involved in
transcriptional regulation of these genes.
Very recent work reported T-tract structures in artificial
minichromosome constructs (52). Here we show T-tract structures
in natural promoters. The T-tract structures did not depend on
whether the gene (HIS3) was located in a small circular minichromosome (3.2 kb) or in the genome, demonstrating that
there were no structural constraints in chromosomes or minichromosomes which would compromise these structures.
Moreover, T-tract structure was observed in tracts which
varied in length from 4 to 11 bp. We also observed T-tract
structures in the nuclease-sensitive 3′-end of the URA3 gene
and in the DED1 promoter (not shown). Therefore, the unusual
structure of T-tracts appears to be a dominant feature in yeast
chromosomes and further suggests a functional role of T-tract
structures. It was estimated that there are ∼1500 yeast genes
whose promoters contain T-tracts >10 bp (28). It is reasonable
to extrapolate that many of these promoters will reveal T-tract
structures when analysed.
4088 Nucleic Acids Research, 2000, Vol. 28, No. 21
Our results shed light on how chromatin is organised under
conditions where T-tract structures were observed in the HIS3
and URA3 promoters. Nuclease digestion of isolated chromatin
in vitro previously established that the T-tracts are located in
nuclease-sensitive regions flanked by positioned nucleosomes
(22,29,30). However, another report proposed that a ‘perturbed’
nucleosome includes the T-tracts of a modified HIS3 promoter
(28). Here we observed that the UV damage patterns of the
promoter regions were identical in vivo and in naked DNA.
These in vivo photofootprints argue against a nucleosome,
since examples are known where T-tract structure was lost
upon incorporation in nucleosomes (8,9). In addition, we found
that all T-tracts were repaired within <30 min and repair was
much faster than repair of CPDs in nucleosomes (32). Thus,
three lines of evidence (nuclease digestion, photofootprinting
and DNA repair data) are consistent with the absence of nucleosomes in the T-tract regions of the HIS3 and URA3 promoters.
A difficult question to address is whether the T-tracts structures
in vivo are stabilised by proteins or whether they are maintained by their intrinsic properties. Since T-tract size varied
between T4 and T11, one would have to propose a class of
different T-tract stabilising proteins. All the photofootprints
were very similar in naked DNA and in chromatin, in small and
long T-tracts, in minichromosomes and chromosomes and in the
presence and absence of datin and Gcn5p. With the exception of
the mild repair delay in the T11-tract of HIS3, there was no
indication for proteins in these regions. If proteins bind to T-tracts
in vivo, they do not disturb the T-tract structure and they do not
inhibit repair or they are displaced by damage induction. All
those observations together make it more likely that T-tracts
are not tightly associated with proteins and therefore participate in
transcription regulation by virtue of their intrinsic DNA structure,
as previously proposed (28).
We still lack a clear explanation as to why these T-tract
regions are not packaged in nucleosomes. Based on controversial
results on how T-tracts can be incorporated in nucleosomes
and that T-tracts were found in nucleosomal as well as in nonnucleosomal regions (see Introduction), T-tracts per se appear
not to be sufficient to exclude nucleosome formation in vivo.
However, under some conditions they may destabilise nucleosomes or affect their positions, as reported for long T-tracts
artificially inserted in minichromosome constructs (52). To
establish a non-nucleosomal region, additional activities might
be required. A sequence-specific protein, Abf1, and a T-rich
element appear to be required to create a nucleosome-free
region in the yeast RPS28A gene (31). Other obvious candidates
are nucleosome remodelling activities which contain histone
acetyltransferases, are brought to promoter regions by transcription factors and destabilise nucleosomes by acetylation of
histones (53). Gcn5p is a histone acetyltransferase of the
SAGA complex and contributes to transcriptional activity of
the HIS3 promoter (38). Disruption of the GCN5 gene was
reported to lead to invasion of the HIS3 promoter by a nucleosome (54). Our photofootprints and repair results do not support
that observation. Using a chromatin immunoprecipitation
approach it was reported that the promoter region contains
acetylated histones (39). The probe used in that study was
0.6 kb long, which, according to our data, includes the nucleosomal regions flanking the promoters of the HIS3 and PET56
genes (22). Therefore, the observation of acetylated histones is
compatible with non-nucleosomal T-tracts and modified
nucleosomes in the flanking regions.
With respect to DNA repair, we also need to consider that
chromatin structures are dynamic and might react on damage
induction, e.g. by rearranging nucleosome positions. If the T-tract
structure affects the stability or positions of nucleosomes, one
would expect that disruption of the T-tract structure by CPD
formation (15) would allow regeneration of nucleosomes in
these regions. The rapid repair by photolyase shown here is
strong evidence against such a chromatin rearrangement after
damage formation.
Having shown that T-tract structure exists in T-tracts of
different length in vivo, it will be interesting to use UV photofootprinting and photolyase to understand the structure and
role of T-tracts in other regions of the genome and in other organisms. Of particular interest is the characterisation of T-tracts in
nucleosomal arrays as well as phased T-tracts that induce DNA
bending in vitro.
ACKNOWLEDGEMENTS
We thank Dr E. Winter for yeast strains and Dr U. Suter for
continuous support. This work was supported by grants from
the Swiss National Science Foundation and by the Swiss
Federal Institute of Technology (ETH) (grants to F.T.)
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