It is widely believed that chromatin, the nucleoprotein packaged state of
eukaryotic genomes, can carry epigenetic information and thus transmit gene
expression patterns to replicating cells. However, the inheritance of genomic
packaging status is subject to mechanistic challenges that do not confront the
inheritance of genomic DNA sequence. Most notably, histone proteins must at
least transiently dissociate from the maternal genome during replication, and it
is unknown whether or not maternal proteins re-associate with daughter genomes
near the sequence they originally occupied on the maternal genome. Here, we use
a novel method for tracking old proteins to determine where histone proteins
accumulate after 1, 3, or 6 generations of growth in yeast. To our surprise,
ancestral histones accumulate near the 5′ end of long, relatively inactive
genes. Using a mathematical model, we show that our results can be explained by
the combined effects of histone replacement, histone movement along genes from
3′ towards 5′ ends, and histone spreading during replication. Our
results show that old histones do move but stay relatively close to their
original location (within around 400 base-pairs), which places important
constraints on how chromatin could potentially carry epigenetic information. Our
findings also suggest that accumulation of the ancestral histones that are
inherited can influence histone modification patterns.
Replicating chromatin involves disruption of histone-DNA contacts and subsequent
reassembly of maternal histones on the new daughter genomes. In bulk, maternal
histones are randomly segregated to the two daughters, but little is known about
the fine details of this process: do maternal histones re-assemble at preferred
locations or close to their original loci? Here, we use a recently developed
method for swapping epitope tags to measure the disposition of ancestral histone
H3 across the yeast genome over six generations. We find that ancestral H3 is
preferentially retained at the 5′ ends of most genes, with strongest
retention at long, poorly transcribed genes. We recapitulate these observations
with a quantitative model in which the majority of maternal histones are
reincorporated within 400 bp of their pre-replication locus during replication,
with replication-independent replacement and transcription-related retrograde
nucleosome movement shaping the resulting distributions of ancestral histones.
We find a key role for Topoisomerase I in retrograde histone movement during
transcription, and we find that loss of Chromatin Assembly Factor-1 affects
replication-independent turnover. Together, these results show that specific
loci are enriched for histone proteins first synthesized several generations
beforehand, and that maternal histones re-associate close to their original
locations on daughter genomes after replication. Our findings further suggest
that accumulation of ancestral histones could play a role in shaping histone
modification patterns.
IntroductionIn addition to the information encoded in DNA sequence, replicating cells can inherit
epigenetic information, which refers to variable phenotypes that are heritable
without an underlying change in DNA sequence. It is widely accepted that chromatin,
the nucleoprotein packaging state of eukaryotic genomes, provides one potential
carrier of epigenetic information. Although definitive proof that chromatin per se
carries epigenetic information during replication exists in very few cases [1], genetic
studies in numerous organisms have identified key roles for chromatin regulators in
multiple epigenetic inheritance paradigms [2],[3].The idea that chromatin structure carries epigenetic information poses a central
mechanistic question—since chromosome replication involves dramatic
perturbations to chromatin structure ranging from old histone displacement to
widespread incorporation of newly synthesized histones, how can chromatin states be
stably maintained? To understand the mechanism by which chromatin states could be
inherited, it is necessary to understand the unique challenges posed by histone
protein dynamics during replication [4]–[7]. First, histones must at least
transiently dissociate from the genome during passage of the replication
fork—if old histones carrying epigenetic information do not re-associate with
daughter genomes at the location from which they came, this could lead to
“epimutation,” analogous to DNA bases moving relative to one another
during genomic replication. Second, it is unknown to what extent newly synthesized
histones deposited at different loci differ in their covalent modification patterns.
Finally, how old histones influence new histones, the basis for positive feedback,
can be considered analogous to asking what the equivalent of base-pairing is during
chromatin replication.Classic radioactive pulse-chase studies demonstrated that, in bulk, maternal histones
segregate equally to the two daughter cells [4],[8]–[10]. It is unknown, however,
whether maternal histones remain close to the locus from which they were evicted by
the replication fork or whether maternal histones are incorporated at preferred
genomic loci in the two daughter genomes [5],[7],[11]. The extent of maternal histone
dispersal affects the stability of epigenetic states in theoretical models of
chromatin inheritance [12], making experimental determination of this parameter a
key goal for epigenetics research.To address these fundamental questions, we carried out a genetic pulse-chase with
epitope-tagged histone H3 [13] to follow ancestral H3 for several cell divisions after
removal of the ancestral tag. We find that old histone proteins do not accumulate at
epigenetically regulated loci such as the subtelomeres but instead accumulate at the
5′ ends of long, poorly transcribed genes. As expected, old histones do not
accumulate at loci exhibiting rapid histone turnover, but we also find that 3′
to 5′ movement of old histones along coding regions and histone movement
during replication are required to explain the patterns of ancestral histone
retention we observe. We estimate that maternal histones stay within ∼400 bp of
their original location during replication, providing the first measure of this
crucial parameter. Finally, we identify a number of factors that affect old histone
localization, such as topoisomerase I and the H4 N-terminal tail, which both affect
the 5′ bias in localization patterns. In contrast, CAF-1 mostly affects
histone turnover at promoters. Together, these results provide a detailed overview
of the movement of ancestral histones across multiple cell generations and identify
a number of mechanisms that play a role in shaping the landscape of ancestral
histone retention.ResultsTo follow the movement of old histone proteins over multiple cell generations, we
utilized a novel pulse-chase technique [13] to follow ancestral
epitope-tagged histone H3 for several cell divisions after swapping epitope tags
from H3-HA to H3-T7 (Figure 1A,
B). We have previously described use of this technique to assay
replication-independent H3 turnover in arrested cells and have shown that prior to
recombination all cells carry the H3-HA, and that recombination is 98%
efficient in cells that are not dividing due to nutrient deprivation (Figure S1).
Unlike inducible pGAL-based systems for measuring
replication-independent histone dynamics [14]–[17], here the epitope-tagged
histone is under the control of its endogenous promoter, avoiding potential
artifacts of H3/H4 misexpression [18] on histone dynamics throughout the cell cycle.10.1371/journal.pbio.1001075.g001Figure 1Overview of system for tracking ancestral histone proteins.(A) Recombination-based swapping of epitope tags on histone H3. Histone H3 is
tagged at its endogenous locus with a C-terminal HA epitope tag surrounded
by LoxP sites. Upon induction of Cre recombinase with β-estradiol, the
HA tag is recombined out and H3 is left with a C-terminal T7 tag. (B)
Experimental overview. Yeast carrying HA-tagged H3 are arrested by nutrient
depletion, and the HA→T7 swap is induced by overnight incubation with
β-estradiol. After the tag swap, yeast are released from arrest and HA
and T7 tags are mapped across the genome at varying times post-release. (C)
Chromosome III overview. HA/T7 ratios are shown as a heatmap across
chromosome III at 3 generations after release. Notable in this view is a
lack of accumulation of H3-HA at TEL3L or the silent mating loci. (D)
Close-up views of two genomic loci. Data are shown as a heatmap for 3 and 6
generations after the tag swap.We used MNase-ChIP [15],[19] for the HA and T7 tags after recombination but before
release into the cell cycle, and 3 and 6 generations after releasing yeast into the
cell cycle [13]. This material was hybridized to tiling microarrays
covering 4% of the yeast genome [20], and HA/T7 ratios of normalized
HA and T7 signals were computed for the 3 and 6 generation data (Figure 1C, D). Since HA is
eliminated via recombination leaving new H3-T7, high HA/T7 ratios indicate loci
enriched for ancestral histone H3. Surprisingly, many of the highest HA/T7 levels
were associated with coding regions (discussed below).Overall, HA/T7 patterns are consistent at 3 and 6 generations, but the dynamic range
of HA/T7 enrichment diminished from 3 to 6 generations (Figure 1D, Figure S2).
This is an expected consequence of the fact that ∼1%–2% of
cells do not recombine the HA tag away (Figure S1)—since the amount of ancestral H3
is decreasing by at least 2-fold in each generation, the relative contribution of
the ∼2% of cells still expressing H3-HA will increase over time, with
this genomic background eventually competing with the real signal from increasingly
rare ancestral H3 (<2% of total H3 after 6 doublings).Ancestral Histones Are Retained Over Long, Poorly Transcribed GenesTo extend our analyses to the entire genome, we carried out deep sequencing of HA
and T7 libraries. HA- and T7-tagged H3 were immunoprecipitated after the tag
swap but before release from arrest (0 generations), after release into a G2/M
cell cycle block, and at 1, 3, and 6 generations after release. Sequencing reads
were mapped to the yeast genome, normalized for read count, and HA/T7 ratios
were computed genome-wide. These data correlated well with our microarray data,
and we further validated these measurements by q-PCR at SPA2
and BUD3, two genes which both exhibit high and low HA/T7
ratios at their 5′ and 3′ ends, respectively (Figure
S3).In previous work, we and others [13],[15]–[17],[21]–[23] showed that
there is a partial correlation between transcription levels and
replication-independent histone dynamics. To understand how transcription might
affect multigenerational histone retention in our system, we aligned all yeast
genes by their transcription start site (TSS) and clustered genes (K-means,
K = 5) based on the pattern of the 3-generation HA/T7
ratios along the gene body (Figures S4, S5, Table S1).
We observed a striking enrichment of H3-HA just downstream of the 5′ ends
of genes (typically peaking around the +3 nucleosome). One exception to the
5′ pattern described is found in one cluster of short genes with uniformly
low H3-HA levels (Figure S4, Cluster 1), which is enriched for
GO categories (such as protein translation) related to high gene expression
levels. In contrast, long genes were generally associated with higher levels of
ancestral H3 (see for example Cluster 5).To better visualize these trends, we sorted genes by the extent of ancestral H3
retention after 3 generations (Figure 2A–B). Retention of ancestral histones correlates both
with low expression levels and with longer genes (Figure 2C–D, Figure S6).
While it is the case that longer genes tend to be expressed at lower levels than
short genes (Figure 2E),
these factors are partially independent here—even when we focus on genes
of 1–2 kb length, we still observe the correlations between ancestral
histone retention and low expression (Figure 2E–F, and see below).
Interestingly, in both microarray and sequencing datasets we found that
epigenetically repressed loci such as the silent mating loci and subtelomeres
[24],[25] did not preferentially accumulate ancestral histone
proteins (Figure 1C, Figure
S7)—analysis of both unique and repetitive subtelomeric genes
showed similar H3-HA retention patterns to euchromatic genes of similar length
and expression. This was not a consequence of silencing defects in our strains,
as they showed efficient mating (unpublished data).10.1371/journal.pbio.1001075.g002Figure 2Ancestral H3 molecules accumulate at the 5′ ends of long,
poorly transcribed genes.(A–B) Heatmap of sites of ancestral H3 accumulation. Genes are
aligned by TSS (indicated), and Log2 HA/T7 ratios are
indicated as a heatmap. Genes are ordered by the median HA/T7 ratio over
the 5′-most 1 kb at 3 generations. Grey over coding regions
indicates missing data; grey downstream of genes indicates sequence
downstream of the 3′ end of the gene to show gene length.
Accumulation of ancestral histones at the 5′ ends of genes peaks
around the +3 nucleosome, as expected given that the +1 and
+2 nucleosomes are generally subject to high rates of
replication-independent H3/H4 replacement [15],[17]. (C) An 80 gene
sliding window average of Pol2 ChIP levels [75] for genes ordered as
in (A–B), showing that genes with low levels of ancestral H3
retention are highly transcribed. (D) 80 gene sliding window average of
gene lengths, showing that genes with high levels of ancestral H3
retention tend to be long. (E) The median HA/T7 ratio over the 5′
end of genes (1 kb) was calculated for all genes, and median values of
this retention metric are shown for groups of genes ordered by
transcription rate (x-axis) and gene length
(y-axis). While these are not
independent—highly expressed genes tend to be short—for a
given gene length genes transcribed at higher levels exhibit low HA
retention levels. This is true mostly of genes shorter than 3 kb, which
encompasses the majority of yeast genes. (F) Average HA/T7 ratios (Log2)
for genes between 1 and 2 kb, broken into high (red), low (green), and
intermediate (blue) transcription rates.What properties of short or highly transcribed genes might lead to loss of
ancestral histones? Replication-independent histone replacement occurs most
rapidly over intergenic regions and over the coding regions of highly
transcribed genes [15],[17],[21],[23], the converse of the pattern of ancestral H3
retention we observe. Indeed, ancestral histone retention is broadly correlated
with “cold” regions of low H3/H4 turnover (Figure 3A). Importantly, however, for a given
level of H3/H4 turnover, ancestral H3 retention varied
significantly—retention at a given nucleosome was better correlated with
the average turnover rate of several surrounding nucleosomes than with the
immediate turnover rate (see, for example, Figure 3B–C). This observation suggests
that maternal histones preferentially re-associate with daughter genomes near
the location from which they originated—if old histones scattered randomly
at replication, ancestral H3 retention patterns should more precisely
anticorrelate with replication-independent turnover patterns, as is discussed in
more detail below.10.1371/journal.pbio.1001075.g003Figure 3H3 retention anticorrelates with replication-independent turnover in
a gene length-dependent manner.(A) Scatterplot of ancestral H3 retention (median Log2 HA/T7 for the
5′ 1 kb, y-axis) versus replication-independent
turnover (Dion et al. [15], Z score, x-axis). (B) HA
retention is plotted against 5′ H3 turnover as above but with
short and long genes plotted separately. For a given level of H3
turnover, ancestral retention is greater at longer genes. (C) Averages
of the 5′ HA/T7 retention parameter (median HA/T7 for the
5′-most 1 kb) are shown for genes broken into different length and
5′ turnover groups. For all turnover levels, longer genes retain
more H3-HA than do shorter genes.Accumulation of Ancestral Histones at 5′ Ends of GenesWhy do old histone proteins accumulate near the 5′ ends of genes? We
considered two alternative possibilities for classes of mechanisms causing this
pattern. In the first mechanism, we reason that if histone proteins tend to
maintain their locations along the genome, the 5′ enrichment of old
histones implies that old 3′ histones are evicted and replaced by new
histones during some phase of the cell cycle. However, previous measures of
turnover in G1- or G2/M-arrested yeast [15],[26] cannot explain the
5′/3′ ratios we observe. Furthermore, we found that mutations in
candidate 5′/3′-marking complexes such as cohesin [27],[28] or
H3K4/K36 methylases [29] did not affect 5′-biased retention of old
histones at target loci (Figure S8A).A second possible explanation for widespread 5′ accumulation of ancestral
histone proteins is that the histone proteins move from 3′ to 5′
over genes over time. This could result from RNA polymerase passage, because
some RNA polymerases pass histone octamers in a retrograde direction during
transcription [30],[31]. Although it is debatable whether this is true of
Pol2 in vitro [32],[33], in vivo we previously observed that inactivation of
Pol2 leads to a modest shift of nucleosomes from 5′ to 3′ [34], consistent
with the idea that Pol2 movement normally shifts nucleosomes in a 5′
direction. To test whether this movement was related to transcription, we asked
whether the 5′ peak of H3-HA accumulation shifted further 5′ with
increasing transcription rate. We normalized all gene lengths to one, then
plotted the HA/T7 ratio for all genes sorted by transcription rate (Figure S9).
Consistent with the prediction of transcription-dependent retrograde movement,
we did observe a subtle signal of H3-HA peaks shifting further 5′ at
higher transcription rates. While this analysis could be confounded by the
higher transcription rates seen over shorter genes, even when we focus on
1–2 kb genes, we observe that poorly transcribed genes exhibit a much
flatter profile than genes expressed at average levels (Figure 2F), as expected if Pol2 transit were
required for H3/H4 “passback.” Finally, we show below that per-gene
estimates of passback exhibit significant correlation with Pol2 levels.
Together, these results are most consistent with a model in which histone
proteins move from 3′ to 5′ over coding regions over time (further
detailed in the Discussion).Quantitative Estimation of Nucleosome Dynamics During ReplicationA key question we sought to address in this study is whether maternal histones
re-associate near their original positions after passage of the replication
fork. We reasoned that changes in HA/T7 patterns over the course of several
generations might provide insight into the effects of replication on nucleosome
dynamics. HA/T7 patterns change dramatically between arrest and 1 generation of
release (with or without G2/M arrest) and then are very similar between 1 and 3
generations, before the background of nonswitching cells starts to dominate the
profile at 6 generations (Figure
4). As expected, HA/T7 data at generation 0 exhibited widespread HA
loss/T7 gain at promoters and +1 nucleosomes as a result of the rapid
replication-independent turnover at these loci [13],[15],[17].
Importantly, to rule out the possibility that 5′ accumulation of H3-HA was
an effect of our arrest-release protocol, we also measured HA/T7 distributions 6
h after inducing recombination in actively growing midlog cultures of yeast
(Figures
S3 and S10). Despite heterogeneity in switching
times in this protocol (only 65% of yeast have switched from HA to T7 3 h
after switch induction, 85% after 6 h), we nonetheless observed that
HA/T7 distributions were remarkably similar in midlog-switching cells to HA/T7
patterns observed in cells undergoing the arrest/release protocol, with
preferential ancestral histone accumulation at the 5′ ends of long, poorly
transcribed genes.10.1371/journal.pbio.1001075.g004Figure 4Kinetic analysis of ancestral H3 retention.HA/T7 ratios were measured genome-wide after recombination but before
release (Gen 0), after release into nocodazole (G2/M), and after 1, 3,
or 6 generations of growth post-release. Data for all genes were
averaged and are plotted as indicated.We asked whether these dynamic observations could be used to quantitatively rule
out specific models concerning the mechanisms for segregation of maternal
histones to daughter genomes. However, the resolution of this question is
complicated by replication-independent processes we discuss above that can
remove or shift ancestral histones, and that cannot be fully removed
experimentally (for example, yeast will not proceed through the cell cycle in
the absence of RNA polymerase). To understand the relationship between these
issues, we designed an analytical model that accounts for three processes that
affect H3 molecules in coding sequences (Figure 5A) and then examined the effect of
removing any of the three. Briefly, our model includes a nucleosome-specific
term for H3 turnover taken from prior experimental results [15], with H3 turnover resulting
in loss of HA. In addition, it includes a gene-specific parameter accounting for
lateral movement of histones (“passback”). Further, the model also
includes a global parameter that describes the extent of histone
“spreading” via dissociation/re-association during replication.
Finally, the experimentally measured background of 2% nonswitching cells
(Figure
S1) was included. The free parameters of the model (describing global
histone spreading and gene-specific lateral movement per generation) were
estimated to maximize the likelihood of experimental observations (Text
S1).10.1371/journal.pbio.1001075.g005Figure 5Quantitative modeling reveals three distinct dynamic
processes.(A) Outline of quantitative model. From a given starting distribution,
histones are subject to turnover [15],
transcription-associated lateral movement (“passback”), and
replication-mediated spreading. Model is described in detail in Text
S1. (B) The model captures major features of the experimental
data. HA/T7 ratios for experimental data and model predictions are shown
for all genes as a heatmap. (C) Distribution of lateral passback
parameter (per generation) for all genes. Note that the vast majority
(92%) of genes were associated with retrograde 3′ to
5′ movement along coding regions. (D) Estimation of
replication-based spreading of maternal histones. Model likelihood
(Text S1) is plotted on the y-axis for
various width spreading distributions (defined as 1 standard deviation
of the Gaussian describing histone movement at replication—see
Text
S1 for model details). (E) Eliminating any of the three model
features worsens fit to data. Plotted are averages at 3 generations for
genes over 2 kb for data versus predictions of various models
(“STP” refer to replication-mediated Spreading,
replication-independent Turnover, and Passback). Note that the model
eliminating turnover underestimates turnover effects, as histones that
spread or are passed over the 5′ end of the gene are still
eliminated in this model (i.e., in this model we effectively only
eliminate turnover within CDS, not in intergenic regions), providing
another basis for high loss of 5′ histones.To account for any first-pass effects of Pol2 behavior during initial re-feeding
of nutrient-depleted yeast (Figures S3 and S10), we
examined this model with two starting conditions—the first started with a
uniform genomic distribution of H3-HA, while the second started with the
experimental distribution of HA/T7 observed after release into G2/M arrest
(Figure 4). Both model
variants predicted HA/T7 ratios with good correlations to the experimental data
(Figure 5B shows data
starting from a uniform distribution, Figure 5E and Figure S11
start from the G2/M distribution). Examination of estimated parameters revealed
expected behaviors. For instance, the distribution of lateral histone movement
estimates (Figure 5C) was
strongly biased towards retrograde 3′ to 5′ movement of histones,
consistent with the previously measured effects of rpb1-1
inactivation on nucleosome positioning [34]. Passback values were also
significantly (p = 9.6439e-19) correlated
with transcription rate (Figure S12).Our model allows us to estimate the extent of histone movement during
replication. Figure 5D shows
the likelihood of the full model plotted for various values for
replication-dependent histone spreading. The best fit model allowed histones to
spread ∼400 bp in either direction, or roughly two nucleosome widths, during
replication (more precisely, in this model two-thirds of histones stay within
400 bp of their original locations, as this value is the standard deviation of a
Gaussian function describing spreading; see Text S1).
Results from models with 800 and 1,600 bp spreading parameters are shown in
Figure 6 for comparison.
Our estimate of ±400 bp spreading is particularly interesting given
electron microscopy results demonstrating that nucleosomes are destabilized over
650–1,100 bp around the replication fork on replicating SV40
minichromosomes [35],[36].10.1371/journal.pbio.1001075.g006Figure 6Dependency of histone dynamics model on spreading parameter.(A–B) Parameters in the quantitative model described in Figure 5 were
re-optimized after fixing the spreading term to 400 bp (as in Figure 5), 800, or
1,600 bp. Data and simulations are shown averaged for genes over 2 kb
for models starting with a uniform H3-HA distribution (A) or starting
with the experimentally measured G2/M HA/T7 distribution (B).
(C–D) Examples of data and three models with different spreading
parameters. Genomic coordinates are chromosome 2 490–540 kb (C),
and chromosome 1 60–110 kb (D). Y-axis shows
measured (Data) or predicted HA/T7 values, in Log2.Elimination of any one or two of the three components of the
model—spreading, turnover, or passback—resulted in significantly
worse fits between model predictions and experimental data (Figure 5E). This can be intuited as follows.
First, in the absence of histone spreading, unmitigated histone movement from
3′ to 5′ results in a much tighter 5′ ancestral histone peak
and results in much more extensive change from one generation to the next than
we observe. Second, eliminating histone turnover shifts the 5′ ancestral
peak closer to the +1/+2 nucleosome. Third, preventing lateral histone
movement results in a 3′-shifted, flatter ancestral histone profile.While our model provided good quantitative fits of ancestral H3 patterns for many
genes, we nonetheless note that many genes were not perfectly fit by this model.
Generally, we found that the model poorly fit short genes, and overall the model
almost universally predicted lower HA/T7 at the +N nucleosome (the last
nucleosome in a gene) than was observed (Figures S11, S13). We
ascribe these failures to the fact that we considered each gene in isolation and
therefore did not model shifts of old nucleosomes from adjacent genes, which
would result in poor fits over short genes in particular. Interestingly, the
better fit at the +1 nucleosome than at the +N nucleosome is
consistent with rapid promoter turnover more effectively isolating genes from
one another at their 5′ ends in vivo.Overall, the strong correlation between our model and the experimental data
supports the hypothesis that at least three dynamic processes affect nucleosomes
and shape the landscape of ancestral histone retention and provide the first
quantitative estimate of maternal histone dynamics during replication.Topoisomerase I and the H4 N-Terminal Tail Play Roles in Establishing the
5′/3′ Gradient of Ancestral H3 MoleculesTo further investigate the mechanism of 5′ accumulation, we asked whether
gene-specific passback parameters were correlated with specific gene annotations
(Table
S2) [34],[37]. Interestingly, we find that the estimated passback
distance was much greater at TFIID-dominated (“growth”) genes than
at SAGA-dominated (“stress”) genes (Figure 7A) [38]. As a result, 5′
accumulation was much more pronounced at TFIID-dominated than at SAGA-dominated
genes (Figure 7B). Almost
every described aspect of chromatin structure and gene expression, from
nucleosome positioning to evolutionary lability (reviewed in [39],[40],[41]), differs
between these two broad types of genes. Mechanistically, one interesting
correlate is that TFIID recruitment has been proposed to be mediated in part by
acetylation of the N-terminal tail of histone H4 [38],[42].10.1371/journal.pbio.1001075.g007Figure 7Mutants affecting ancestral histone retention.(A) Distribution of lateral nucleosome distances from model (Figure 5). Shown are
the passback parameters for SAGA-dominated and TFIID-dominated genes as
defined in Huisinga et al. [38]. (B)
TFIID-dominated genes preferentially accumulate 5′ H3-HA. Averages
of 3 generation experimental data are shown for the indicated gene
classes. (C) H4 tail deletion dramatically reshapes the landscape of
ancestral histone retention. Yeast carrying an N-terminal H4 tail
deletion were processed as in Figure 1A–B, and averages for
all genes are plotted as indicated. We note that this strain has
retained a wild-type HHT2-HHF2 locus for viability, so
results must be interpreted with caution. However, we find similar but
less dramatic effects in an H4K5,12R mutant (Figure
S8B), supporting the observation here that passback is
affected by the H4 N-terminal tail. (D) H4 tail deletion preferentially
affects TFIID-dominated genes. Data for wild-type and
H4tailΔ yeast are plotted for the indicated
gene classes. (E) Topoisomerase I plays a role in 5′ accumulation
of ancestral histones. top1Δ yeast were processed
as in Figure
1A–B, and averages for all genes are plotted as
indicated. (F) TOP1 deletion affects 5′ passback
preferentially at long genes. Data for wild-type and
top1Δ yeast are plotted for the indicated gene
classes.To investigate this link experimentally, we examined whether mutations of the H4
tail influenced ancestral histone H3 retention. In an H4K5,12R mutant that
cannot be acetylated on these two tail residues, the 5′-biased HA/T7 was
partially lost (Figure S8B), consistent with the possibility that acetylation of H4
tail lysines may contribute to H3/H4 passback. We also deleted the H4 N-terminal
tail, although in this strain background this mutation proved lethal and so all
recovered strains retained a wild-type copy of the H4-H3 locus
(HHF2-HHT2). Thus, results with this strain must be
interpreted with extreme caution, as we do not know the effect of wild-type,
untagged nucleosomes on the behavior of the epitope-tagged histones.Nonetheless we present here results of mapping of HA and T7 3 generations after
release from the HA/T7 tag swap, since the H4 tail deletion has dramatic effects
on global nucleosome dynamics (Figure 7C), with low HA/T7 at 5′ ends followed by a nearly
flat profile over the remainder of coding regions. This profile suggests a
requirement for the H4 tail in H3/H4 passback, and possibly on
replication-mediated spreading (see Figure 5E). Interestingly, the effect of H4 tail deletion was much
more pronounced at TFIID-dominated genes (Figure 7D), suggesting that the exaggerated
H3/H4 passback inferred at these genes involves the H4 tail. The effects of the
H4 tail deletion were not simply due to the extensive changes in the
transcriptome [43], as we measured changes in genome-wide RNA Pol2
localization in our H4 mutant strains, finding that the relationship between
Pol2 levels and HA/T7 behavior qualitatively changed in this mutant (Figure
S14). While we must be cautious interpreting results obtained with the H4
tail deletion, the fact that H4K5,12R mutants (which were viable and did not
retain any wild-type H3/H4) also exhibit diminished 5′ bias in ancestral
H3 retention provides independent support for a key role for the H4 tail in
H3/H4 passback.We also explored the role of supercoiling in the 5′-biased retention of old
histones. Topoisomerases relax DNA supercoiling and thereby help to maintain
chromatin architecture. Transcription of DNA templates by Pol2 differentially
affects supercoiling in front of and behind the passing polymerase, thereby
differentially affecting 5′ and 3′ nucleosomes [44],[45]. To assess the role of this
activity in 5′ accumulation of old histones, we examined the consequences
of inactivation of the major topoisomerase Top1, which in vitro can resolve both
negative and positive supercoils [46],[47]. Cells
lacking Top1 showed reduced 5′ bias in ancestral nucleosome accumulation
(Figure 7E), indicating
that resolving DNA topology problems before or after passage of the
transcription or replication machinery influences the mobility and/or stability
of nucleosomes. Consistent with expectations of a greater buildup of supercoils
over longer transcription units, we confirmed a stronger effect of
TOP1 deletion at longer genes (Figure 7F).Replication Timing, Chaperones, and Ancestral Histone RetentionWe finally turn to the role of replication factors in ancestral histone
retention. We first asked whether replication timing affected H3-HA retention.
Nucleosomes surrounding early-firing origins tended to lose H3-HA more rapidly
than late-firing origins (unpublished data), but this likely stems from the fact
that replication timing correlates with replication-independent turnover [23],[26]. Focusing
only on nearby coding regions (Figure S15), we found that late-replicating
genes were associated with slightly 5′ shifted ancestral H3 peaks relative
to genes near early origins (consistent with decreased spreading or turnover),
suggesting that different replication forks might affect chromatin in different
ways, although the modest effect precludes a stronger interpretation.To directly address the role of fork-associated chromatin proteins in histone
spreading at replication, we examined mutations of PCNA and Chromatin Assembly
Factor (CAF-1), which plays a key role in replication-coupled histone deposition
[48],[49]. Three different mutants of PCNA that disrupt
interactions with replication proteins or with replication-coupled
chromatin-assembly factors showed only minor effects on 5′ retention of
ancestral H3 at target genes SPA2 and BUD3
(Figure
S8C). In contrast, ancestral H3 retention at the 5′ ends of
these target genes was slightly increased upon deletion of the
CAF-1 subunit CAC1 (unpublished data). To further explore the
role of CAF-1 in histone retention patterns, we deep sequenced HA and T7 tags
from cac1Δ yeast 3 generations after release (Figure 8). These data show a
dramatic 5′ shift in the peak of ancestral H3 retention in these mutants.
This shift is most consistent with a decrease in histone turnover at the
5′ ends of genes in this mutant, which we have independently confirmed
using G1-arrested yeast expressing pGAL-driven Flag-H3 [50].
However, we cannot rule out the possibility that the role of CAF-1 in retention
of old histones in 5′ and promoter regions involves interactions with PCNA
during DNA replication. Interestingly, the 5′ accumulation observed in
wild-type yeast is otherwise little changed in the CAF-1 mutant over long genes
(Figure
S16), suggesting that 3′ to 5′ movement of histones is
normal, and that preferential retention of old histones at their maternal
locations may be carried out by alternative histone chaperones such as the Hir
complex or Asf1 in this mutant. Unfortunately, both hir and
asf1 mutants are lethal in our strain background (likely
because our strain carries only one copy of the H3/H4 gene pair [51]), preventing
us from testing this hypothesis.10.1371/journal.pbio.1001075.g008Figure 8Effects of Chromatin Assembly Factor-1 complex on ancestral H3
patterns.Yeast lacking CAF-1 subunit Cac1 were processed as in Figure 1A, and HA/T7
ratio averages are shown for all genes in wild-type and
cac1Δ mutants 3 generations after release.Consequences for Histone Modification PatternsOur results are most consistent with histone retrogression from 3′ to
5′ over genes, which raises the question of whether old histones carry
modifications associated with mid- and 3′ coding regions (e.g., H3K36 and
H3K79 methylation) towards the 5′ end of genes. Alternatively, there could
be active erasure of these modifications. We therefore compared genes exhibiting
high levels of ancestral H3 retention with prior genome-wide analyses of histone
modifications [52],[53]. Histone modification patterns generally conformed to
the patterns expected based on transcriptional behavior—genes that retain
high levels of ancestral histones are poorly transcribed (Figure 2D), and correspondingly exhibit low
levels of transcription-related marks H3K9ac, H3K14ac, H4ac, and H3K4
methylation (Figure S17 and unpublished data). However, these are all
5′-biased marks [19],[29],[52],[54], and based on retrograde movement of old histones are
therefore not expected to accumulate with age.More interestingly, we found that genes with high levels of old nucleosomes were
enriched for H3K79me3 throughout their coding regions, particularly at the
5′ end (Figure 9,
Figure
S17). The H3K79 methylase is nonprocessive, indicating that K79
methylation status should essentially act as a timer [55]. Further, analyses of
genome-wide H3K79me3 patterns show anticorrelation between this modification and
locations of high nucleosome turnover [15],[56], supporting the idea that
K79me3 identifies old H3 protein. We recently confirmed that old H3 protein is
enriched for H3K79me3 by mass spec analysis of old nucleosomes (D. DeVos, FvL et
al., submitted).10.1371/journal.pbio.1001075.g009Figure 9Ancestral H3 retention and histone modification patterns.(A) Scatterplot of previously measured H3K79me3 [52] levels averaged
over the 5′ CDS of genes versus the median HA/T7 for the 5′
1 kb of each gene. (B) As in (A), for K79me3 averaged over mid-CDS of
each gene.Finally, we also observed higher levels of H3K36me3 at 5′ and mid-CDS of
genes exhibiting elevated ancestral histone retention relative to genes with
intermediate H3-HA retention (Figure S17). This observation is consistent
with the above hypotheses that old histones move from 3′ to 5′ and
thus might carry typical mid-CDS and 3′-end histone modifications to the
5′ ends of genes (Figure S18). Together, these results provide
further evidence that our system accurately captures the behavior of old
histones.DiscussionThe fate of stable proteins in rapidly dividing cells is of great interest for fields
from protein damage to aging to epigenetic inheritance. In particular, models for
the inheritance of chromatin-based information [5]–[7],[12],[57],[58] require a quantitative
understanding of the fate of specific maternal histone proteins during the
disruptive replication process. Some models for epigenetic inheritance of chromatin
states require that old nucleosomes are retained near their original positions,
whereas other models (such as those based on replication timing; [59]) are less
sensitive to the fates of old histones. However, due to the lack of methods to
directly track histone dispersal during replication, these models have not been
experimentally tested in vivo. Here, by using a novel genetic pulse-chase assay, we
characterize ancestral histone retention patterns across the yeast genome. By
accounting for known replication-independent processes, we used these data to
estimate the effects of replication on histone movement, finding that H3/H4 are
retained close to their original locations during replication. We also identified a
number of mutants that affect various aspects of ancestral H3/H4 movement and
retention.Ancestral Histones Accumulate at the 5′ Ends of GenesMost surprising to us was the observation that ancestral H3 molecules accumulate
near the 5′ ends of coding regions, peaking around the +3 nucleosome.
The high HA/T7 ratio observed at the 5′ ends of genes is not an artifact
of the epitope tags used, as we have observed the converse behavior (high T7/HA)
when we switch the epitope tags used (unpublished data). Furthermore, this
unusual behavior is not an artifact of the conditions used for growth arrest and
release, as we observe a similar 5′/3′ gradient of H3-HA when yeast
are subjected to the epitope switch during active midlog growth (Figure
S10).What is the mechanistic basis for the 5′/3′ gradient of HA/T7 we
observe? We consider two classes of mechanisms—in one, histone proteins do
not move laterally and the 5′/3′ gradient results from preferential
loss of 3′ H3/H4, while in the other the gradient results from lateral
histone movement combined with loss at the 5′ end. While we cannot
definitively answer which mechanism explains our results, we strongly disfavor a
model with preferential 3′ nucleosome eviction and no lateral movement
based on the following observations. First, we tested a number of relevant
mutants for changes in the 5′/3′ HA/T7 bias (Figure S8).
Loss of H3K4 methylation (a 5′-biased histone mark) or H3K36 methylation
(a mid and 3′-biased histone mark) did not affect HA/T7 patterns at
selected target genes. Similarly, 5′ retention of ancestral H3 was
unaffected by mutants of cohesin, whose loading is associated with regions of
high H3/H4 turnover and which accumulates at the 3′ ends of genes [27],[28]. Second,
direct measurements of H3/H4 turnover using a pGAL-driven
epitope tagged H3 do not provide evidence for ubiquitous 3′ histone
replacement during G1 arrest [15], during G2/M arrest [26], or in unsynchronized yeast
[15]. Thus,
while we cannot definitively rule out some cryptic 3′ replacement event in
this system, all direct tests have failed to support this hypothesis.Conversely, multiple observations support the hypothesis that H3/H4 proteins move
from 3′ to 5′ over protein-coding regions over time. First, seminal
in vitro studies on transcription of nucleosomal templates showed that several
RNA polymerases can transcribe through a nucleosome without displacing the H3/H4
tetramer. The proposed mechanism by which histones remain associated with the
DNA is a “bubble propagation” mechanism—DNA partially unwraps
from the histones, RNA polymerase enters, and DNA behind the polymerase
re-associates with the histone octamer, resulting in a net retrograde movement
of histones after the polymerase has passed. This mechanism is relatively well
established for SP6 polymerase and RNA Polymerase III [30],[31], whereas there is some
controversy regarding the effect of RNA Polymerase II on nucleosome positioning
[32],[33]. Of course, it is not unreasonable to expect that
nucleosome movement during transcription in vivo will also be affected by
polymerase-associated factors such as histone chaperones and ATP-dependent
remodelers that are not present in the in vitro systems. In any case, these
studies provide a plausible mechanism by which RNA polymerase transit results in
retrograde nucleosome movement.Second, we have previously found that inactivation of Pol2 using the
temperature-sensitive rpb1-1 allele results in a net 5′
to 3′ shift in the majority of coding region nucleosomes [34], consistent
with the hypothesis that polymerase transit normally shuttles nucleosomes from
3′ to 5′. Third, highly transcribed genes (such as those encoding
ribosomal proteins) in yeast paradoxically exhibit very tightly spaced coding
region nucleosomes (e.g., 155–160 bp between adjacent nucleosomes rather
than ∼165 bp), and this tight spacing relaxes upon Pol2 inactivation, again
consistent with nucleosomes being passed upstream during transcription [34]. Taken
together with the absence of any evidence for 3′ H3/H4 eviction, we
therefore argue that the most parsimonious explanation of the surprising
5′ accumulation of ancestral histones is retrograde movement of histones
over genes against the direction of transcription. Note that while we favor the
hypothesis that the act of RNA polymerase transit itself is the mechanism
linking transcription to H3/H4 passback, polymerase is not the only candidate
factor leading to retrograde histone movement. Notably, we found that
top1Δ mutants exhibit diminished signatures of H3/H4
passback (Figure 7), and
this decrease was stronger at longer genes, suggesting the possibility that some
aspect of cleavage and rotation of twisted DNA by Top1 contributes to the
passback observed. However, it is also possible that Top1 differentially affects
histone turnover in 5′ and 3′ regions or affects passback by
affecting Pol2 passage [60].We analytically assess several predictions of the “passback” model.
First of all, if RNA polymerase transit were the driver of retrograde histone
movement, then one might predict that passback should correlate with
transcription rate. We find the expected correlation to be statistically
significant (p = 9.6439e-19, Figure S12)
but weak nonetheless (R = 0.12). Importantly, we previously
observed that 5′ to 3′ nucleosome movement in
rpb1-1 mutants was also significantly but poorly correlated
with transcription rate [34]. The reason for the mediocre correlation between
polymerase abundance and passback is hinted at by the fact that TFIID-dominated
genes exhibit much greater passback values than do SAGA-dominated genes (Figure 7A). We have previously
noted that SAGA-dominated (“stress”) genes exhibit higher levels of
H3 turnover, per polymerase, than do TFIID-dominated genes
[15]. In
vitro, a single polymerase's transit displaces an H2A/H2B dimer from the
histone octamer, but a second polymerase encountering a histone hexamer will
displace the remaining histones [61],[62]. Coupled with the
observation that SAGA-dominated genes exhibit larger “bursts” of
polymerase, this suggests that closely spaced polymerases are required for H3/H4
eviction over coding regions, but evenly spaced polymerases leave time for dimer
replacement on damaged nucleosomes [54],[62]. We believe this model
also explains some of the behavior of ancestral histones in this
study—SAGA-dominated genes display little passback and overall diminished
levels of ancestral H3 (Figure
7A–B), an expected consequence of the loss of old histones via
turnover. Correlations between polymerase and passback are therefore expected to
be subtle—at increasing transcription rates, we expect an increased
likelihood of a closely spaced pair of polymerases, and the resulting H3/H4
eviction would eliminate any trace of the passback that had occurred to that
point.It is important to note that the transcription-dependent passback postulated here
cannot simply be interpreted as a model in which every round of polymerase
passage shifts the histone octamer upstream by one position (∼165 bp). In
Figure 7A, our estimates
of passback per cell cycle have a mean of ∼90 bp at TFIID-dominated genes,
less than the spacing between adjacent nucleosomes. If taken literally, these
values would be difficult to reconcile with the observation that the majority of
yeast nucleosomes are well positioned [20]. Instead, we interpret the
passback values in terms of probability that an octamer will be passed back in a
given cell cycle in each cell—a passback value of ∼80 bp suggests that
there is a 50% chance that histones on a given gene will be shifted back
one position towards the 5′ end in a single cell cycle. Physically, we
imagine that polymerase passage results in relatively short retrograde movement
of H3/H4, which then have some probability of returning to their original
position, and some probability of shifting to a new upstream location.Our results show a surprising pattern of ancestral histone retention in yeast,
with old histone proteins accumulating near the 5′ ends of genes—the
histone proteins located at the +3 nucleosome are the oldest histone
proteins over a typical yeast gene. These data are best explained by a model in
which H3/H4 proteins shuttle from 3′ to 5′ over coding regions over
time, with eventual loss of old histone proteins when they are eventually moved
into the +1 and +2 nucleosome positions.Maternal Histone Spreading During ReplicationThe process of genomic replication is enormously disruptive to chromatin
structure, as the melting of the DNA double helix is accompanied by histone
dissociation from the genome [4]–[7]. Thus,
understanding where maternal histones re-associate relative to the locus from
which they were evicted is a key constraint for understanding the potential of
chromatin as an epigenetic information carrier. The ideal experiment for
measuring this would be to epitope tag the histones at one specific locus (e.g.,
the +5 nucleosome over BUD3) in a large population of
yeast, allow replication to proceed, and measure the new locations of the tagged
histones. Despite numerous attempts, this type of tagging has proven technically
intractable to date. Here we measure instead the bulk distribution of ancestral
histones. Importantly, this still provides information on locus-specific histone
behavior—as turnover rates are not homogeneous across the genome, even
before we release yeast into the cell cycle the landscape of H3-HA exhibits
variability (Figure 4, see
generation 0), and so in effect only a subset of ancestral locations are
epitope-tagged before release. This enables us to infer the dynamic behavior of
histone proteins during replication via analysis of the evolution of the H3-HA
distributions over time.Two observations provide an intuition regarding the effects of replication on
histone locations. First, ancestral histone retention exhibits the expected
anticorrelation with replication-independent turnover (Figure 3A). However, old histones are more
efficiently retained at cold (low turnover) loci that occur in long cold
domains, whereas short domains of cold nucleosomes lose ancestral histones over
time. This observation is inconsistent with two extreme models for histone
behavior during replication—if old histones were to completely dissociate
from the genome during replication and randomly re-associate with the genome,
then ancestral histone retention should precisely recapitulate turnover
measurements. Conversely, if old histones were to reassociate precisely with
their original locations, then ancestral retention should essentially integrate
turnover for multiple generations. Thus, some process that shuffles histone
proteins locally must be invoked along with turnover to shape the ancestral
retention landscape.In principle, the preferential retention of old histones on longer genes could
simply result from passback—shorter genes will more quickly have all of
their histones passed “over a cliff” at the 5′ end. However,
we find relatively static 5′/3′ gradients of old histone retention
over time (Figure 4). While
it is the case that H3-HA domains gradually shorten over time as predicted by
the model that passback results in old histones being moved to promoters where
they are replaced (unpublished data), this effect is subtle and is
quantitatively much less dramatic than predicted from passback alone. This leads
to the second intuition regarding histone spreading during replication. Many
examples exist for relatively static gradients in biological systems being
established via a combination of directional active transport coupled with
passive diffusion. Most relevant in our opinion is the “pump leak”
model [63]
for membrane ion gradients—active transport of ions across membranes,
coupled with a passive leak of ions back into the cell, results in a static
gradient. Here, we envision transcription-related passback as the active
transport mechanism, with spreading during replication being somewhat analogous
to the leak that results in a steady gradient rather than a continuous 3′
to 5′ march of histone proteins.We present a quantitative model that recapitulates our experimental data with
only three dynamic processes—turnover, passback, and spreading.
Locus-specific turnover rates were previously measured [15] and are not fit by the model.
Passback is estimated for each gene separately, while spreading is a single
global parameter affecting all histones. Thus, our model has 4,811 free
parameters, which are used to fit over 100,000 HA/T7 ratios. This model does not
overfit the data, and this can best be appreciated by the fact that eliminating
a single parameter (spreading) greatly diminishes the agreement between model
and data.Using this model, we estimate that maternal histones spread little
(∼1–2 nucleosomes) during replication. This value has not been
measured before but is consistent with several related observations. First,
electron microscopy studies on replicating chromatin show a stretch of
∼650–1,100 bp of nucleosome-free DNA surrounding replication forks
[35],[36], consistent
with histone movement of ±400 bp we estimate here. Second, histone
proteins are retained in cis during in vitro replication even in the presence of
competitor DNA [64]–[67], indicating that histones do not freely diffuse away
from replication forks but likely are retained locally. Finally, we previously
observed that upon gene repression, loss of the active chromatin mark H3K4me3
occurs during S phase, but at very highly methylated nucleosomes H3K4me3 does
not return to baseline levels immediately, with methylation levels falling
little more than the 2-fold predicted by a dilution-based mechanism (see Figure S5
in [68]). This final result indicates that
“overmethylated” old histone proteins are retained near their
original location, since extensive spreading of old histone proteins would
enable a greater than 2-fold drop in methylation levels during S phase.Together, these results support the prospect of chromatin as a
“sloppy” epigenetic information carrier (“sloppy” in the
sense that some spreading of histones will preclude mononucleosome-resolution
information passage) [69], even if chromatin-based inheritance occurs
infrequently [1]. Thus, chromatin states are unlikely to be inherited
with mononucleosome precision, a view consistent with the fact that most or all
proposed epigenetic chromatin domains are associated with long (>1 kb) blocks
of histone modifications such as H3K9me3 or deacetylated H4K16 (reviewed in
[54]).Mutant StudiesTo further investigate the mechanisms underlying the patterns of ancestral H3
retention, we assessed HA/T7 ratios at target genes in 12 mutants and further
characterized HA/T7 genome wide for three of these mutants. Interestingly, a
number of histone modifying factors, including Swd1, Swd3, Rtt109, Nhp6, and
Set2, had either no effect or subtle effects (e.g., Rtt109) on the 5′
accumulation of old histones at our target genes (Figure S8).
These results suggest either that these mutants will have subtle global effects
on ancestral H3 retention or that they have more localized roles that do not
extend to the two target genes on which we focused.The three mutants we characterized at full genome coverage each had a distinct
effect on ancestral H3 retention. Most dramatically, loss of the H4 N-terminal
tail abolished the 5′ accumulation of ancestral histones—while the
H4 tail deletion results are complicated by the retention of wild-type H3/H4 in
this strain, the fact that similar results were obtained with clean H4K5,12R
mutants (Figure
S8B) provides independent support for observations obtained with the
H4 tail deletion. The mechanistic basis for the loss of 5′ H3-HA retention
is unknown to us—a flat HA/T7 profile is of course consistent with
complete loss of passback. Alternatively, the observed profile in this mutant
could be consistent with complete shuffling of maternal histones every
generation, which as described above would be expected to more closely
recapitulate a turnover-dominated profile. Importantly, loss of the H4 tail also
affects H3/H4 turnover—in Figure 7D, the increased HA/T7 ratio at the 5′ ends of
SAGA-dominated genes suggests a decrease in histone turnover in this mutant, and
we have independently confirmed a decrease in replication-independent turnover
in this mutant (F.v.L., manuscript in preparation). Analysis of Pol2 ChIP in H4
tail deletions shows that the effects of the H4 tail do not simply reflect
altered transcription but instead reflect a change in the relationship between
RNA Polymerase and histone dynamics over genes in this mutant (Figure
S14).We also observe a similar, albeit muted, effect of Topoisomerase I on the
5′ accumulation of ancestral histones. Interestingly, loss of both
topoisomerase I and II affects nucleosome occupancy and dynamics in S.
pombe, indicating that topoisomerases play key roles in histone
dynamics [60]. Here, we find that top1Δ
mutants exhibit diminished 5′ accumulation of ancestral H3 and that this
effect is stronger at longer genes than at shorter genes. As RNA polymerase
passage will cause greater changes in supercoiling over longer genes, the
preferential effects of Top1 on longer genes is consistent with the observation
in S. pombe that topoisomerase mutants show evidence of stalled
or slowed RNA polymerase over longer genes [60]. In addition to its
role in transcription, topoisomerase I plays a key role in replication [45]. We note
that the profile of top1Δ mutants here most closely mimics
the predictions of our analytical model with both passback and spreading being
compromised, but since neither of these is likely to be completely eliminated in
top1Δ mutants, more detailed kinetic analyses will be
required to make a quantitative statement about the role of Top1 in
replication-related movement of histones.Finally, we assessed the role of the histone chaperone CAF-1 in ancestral H3
retention. To our surprise, we found that H3-HA exhibited even stronger 5′
accumulation in this mutant, with the 5′ peak of HA/T7 occurring closer to
the +1 or +2 nucleosome (compared to the +3 peak location for
wild-type strains). This result most closely matches the predictions of a model
in which H3/H4 turnover has been slowed without loss of passback or spreading
(Figure 5E). We recently
tested this prediction using an alternative system for measuring
replication-independent turnover (pGAL-driven Flag-H3) and
confirmed the prediction that caf mutants affect
replication-independent histone replacement [50]. As CAF-1 and the Hir
complex are known to complement one another in yeast, we predict that a
caf hir double mutant would be necessary to uncover effects
of replication-coupled spreading. Unfortunately, since both hir
and asf mutants are lethal in our strain background, this
prediction cannot be tested at present.PerspectiveTaken together, our results provide a surprising view of histone dynamics over
multiple generations, with 5′ accumulation of ancestral histone proteins
over coding regions and little evidence for preferential histone retention at
epigenetically regulated loci such as subtelomeric genes. One unanticipated
implication of this observation is that 3′ histone marks are expected to
move towards the 5′ ends of genes over time, thereby shaping histone
modification profiles (as we document in Figure 9 and Figure
S17). This potentially necessitates mechanisms for erasure of these
inappropriate marks in order to maintain accurate encoding of gene polarity.
However, we note that active erasure of H3K4me3 after gene repression occurs
most efficiently at 5′ ends of genes, whereas nucleosomes over coding
regions mostly lose H3K4me3 by passive dilution ([68], see Figure S9).
If other old histone marks are not erased over coding regions, then we speculate
that the accumulation of old histone proteins at the +3 nucleosome could
potentially provide a mechanism by which a gene's transcriptional history
could be integrated to play a role in regulation of the transition from
transcriptional initiation to elongation.Most importantly, we find that old histones do not re-associate with daughter
genomes at precisely the locus from which they dissociated. Thus, any
inheritance of chromatin states must occur at the scale of ∼5–10
nucleosome domains rather than at single nucleosome resolution. These results
therefore constrain the maximum amount of information theoretically carried by
chromatin between generations. It will be of great interest in future studies to
identify mutants that affect histone movement during replication and to measure
their effects on the stability of epigenetic inheritance and to measure how
maternal histone incorporation differs between leading and lagging strand
daughter genomes.Materials and MethodsYeast Strains and Growth ConditionsFor tag switch experiments, yeast cells were grown overnight in YPD in the
presence of Hygromycin B (200 µg/mL, Invitrogen). The cells were then
diluted 1∶10 into fresh YPD and incubated for 30–36 h. Recombination
was induced by the addition of 1 µM β-estradiol (E-8875,
Sigma-Aldrich). Subsequently, cells were diluted 1∶25 in fresh YPD media
to release the cells back into the cell cycle and kept in log phase by 1∶2
dilutions into fresh media after each population doubling. Samples were taken
after 1, 2, 3, and 6 cell divisions or after 5 h of G2/M arrest. The number of
population doublings was determined by microscopy and OD. G2/M arrest was
induced by addition of 15 µg/ml Nocodazole (Sigma-Aldrich) and confirmed
by FACS analysis. Strains are listed in Table S3. Gene deletion mutants isogenic to
strains NKI2048, NKI2148, and NKI2048 were made by homologous recombination
using KanMX and/or NatMX selection markers. Gene deletion mutants isogenic to
NKI4128 were made by crossing NKI4114 with gene deletion mutants from the
MATa yeast knock-out collection using Synthetic Genetic Array
methods. Histone mutants were made by transformation of strain NKI2148 with a
HHF2-HHT2 CEN plasmid (pMP9), subsequent deletion of the
tagged HHF2-HHT2 locus, followed by transformation with a PCR
fragment encoding wild-type or mutated HHF2 in combination with
tagged HHT2. Deletion of the wild-type locus was confirmed in
H4K5,12R mutants, whereas all surviving H4 tail deletion mutants retained a copy
of the wild-type HHT2-HHF2 locus.Chromatin Immunoprecipitation (ChIP)ChIP was performed as described previously [13],[70] with the
following modifications. All steps were done at 4°C unless otherwise
indicated. Following cell lysis by bead beating the insoluble chromatin of
1×109 cells was washed, resuspended in 400 µl FA
lysis buffer (50 mM HEPES-KOH [pH 7.6], 150 mM NaCl, 1 mM EDTA,
1% Triton X-100, 0.1% sodium deoxycholate), and sheared using a
Bioruptor (Diagenode) for 6 min with 30 s intervals at high. The soluble
fraction was diluted 3-fold in buffer 15 mM Tris-HCl pH 7.4, 50 mM NaCl, 1.5 mM
CaCl2, 5 mM β-meracptoethanol, 5 mM MgCl2, after
which 25 units of micrococcal nuclease (Worthington) were added. The digestion
reaction was incubated 20 min at 37°C and stopped by the addition of 10 mM
EDTA and 10 mM EGTA; tubes were placed on ice. The majority of obtained
fragments was around 150 bp, as determined on a 2% TAE agarose gel
stained with ethidium bromide. The isolated chromatin of the equivalent of
3×108 cells was immunoprecipitated overnight at 4°C
using magnetic Dynabeads (Invitrogen), which were previously incubated with
antibody O/N at 4°C.Real-Time PCRChIP DNA was quantified by real-time quantitative PCR using the SYBR Green PCR
Master Mix (Applied Biosystems) and the ABI PRISM 7500. An input sample was used
to make a standard curve, which was then used to calculate the IP samples, all
performed in the 7500 fast system software. Primers used for qPCR are listed in
Table
S4.Linear amplification of DNAThe samples were amplified, with a starting amount of up to 75 ng for ChIP
samples, using the DNA linear amplification method described previously
[19].Microarray hybridization3 µg of aRNA produced from the linear amplification were used to label
probe via the amino-allyl method as described on www.microarrays.org. Labeled probes were hybridized onto a
yeast tiled oligonucleotide microarray [20] at 65°C for 16 h and
washed as described on www.microarrays.org.
The arrays were scanned at 5 micron resolution with an Axon Laboratories
GenePix 4000B scanner running GenePix 5.1. Image analysis and data
normalization were performed as previously described [19].Deep Sequencing Library ConstructionChIP DNA was treated with CIP (calf alkaline phosphatase NEB; in 1× NEB
buffer 3, 0.25 U/µl CIP; 45 min at 37°C, reaction clean up with Qiagen
MinElute spin columns). 20–150 ng of CIP treated ChIP DNA fragments were
blunt ended and phosphorylated with the EPICENTRE End-it-Repair kit (1×
buffer, 0.25 mM dNTPs,1 mM ATP, 1 µl/50 µl reaction of Enzyme mix)
for 1 h at RT and cleaned up with Qiagen MinElute spin columns. Adenosine
nucleotide overhangs were added using EPICENTRE exo-Klenow for 45 min at RT
(with 0.2 mM dATP). Illumina genome sequencing adaptors were then ligated using
the EPICENTRE Fast-Link ligation kit: 11.5 µl A tailed DNA eluted from a
MinElute column was mixed with 1.5 µl 10× ligation buffer, 0.75
µl 10 mM ATP, 0.5 µl Illumina DNA adaptors, and 1 µl Ligase.
The reaction was incubated for 1 h at RT and subsequently supplemented with 7.5
µl water, 1 µl 10× buffer, 0.5 µl 10 mM ATP, and 1
µl ligase, and incubated overnight at 16°C.The ligation reaction was cleaned up with MinElute columns (with an additional
wash step to eliminate all the excess adaptors) and the adaptor ligated
fragments were amplified by PCR as follows: 0.5 µl of each Illumina
genomic DNA sequencing primers, 10 µl 10× Pfx buffer 3 µl 10
mM dNTPs, 2 µl 50 mM MgSO4, and 1 µl Pfx DNA polymerase (Invitrogen)
were added to 30 µl DNA template in a 100 µl reaction. The cycling
parameters were: (1) 94°C, 2′; (2) 94°C, 15″; (3) 65°C,
1′; (4) 68°C, 30″; (5) repeat from (2) 17 times; (6) 68°C,
5′. The PCR product (200 to 300 bp in size) was gel purified from a
2% TAE agarose gel using the Freeze'N Squeeze columns (BioRad). Gel
purified fragments were finally precipitated with Sodium acetate and Ethanol and
pellets were resuspended (25 nM final concentration) in TE buffer and sent for
SOLEXA sequencing at the UMass Worcester core deep sequencing facility.RNA Pol II ChIP and Microarray HybridizationCells were grown as described above. Cell pellets (∼109 cells)
were flash frozen after formaldehyde crosslinking (1%) and kept at
−80°C overnight. Frozen cell pellets were resuspended in 300 µl
cell braking buffer (100 mM Tris pH 7.9, 20% glycerol, 1× Sigma
Protease inhibitors cocktail) and cell walls were broken down by bead beating
using 400 µl of 0.5 mm zirconia/silica beads (BioSpec Products) in the
BioSpec Mini-BeadBeater Model 8 three times for 1 min with 1 min pauses in
between. Cell pellets (5 min max speed spin in refrigerated microcentrifuge)
were then washed once and resuspended in 800 µl FA lysis buffer (with
1× Sigma Protease inhibitors cocktail). Chromatin was sheared by
sonication in a cup sonicator (Branson, 50% pulse at strength 7 for 3.5
min) to 250–400 bp fragments.The sheared chromatin suspension was pre-cleared with 100 µl Protein
A-agarose slurry (IPA 400 HC RepliGen) at 4°C for 1 h. 100 µl of the
pre-cleared solution was saved for the ChIP input sample and 7 µl of RNA
Pol II antibody (abcam ab81859, lot #: 933570 and GR6094-1) was added to the
rest and incubated overnight at 4°C with rotation. ChIP DNA isolation and
amplification by TLAD was done as described previously [19].2.5 μg of aRNA produced from the linear amplification were used to label
probes via the amino-allyl method as described on www.microarrays.org.
Labeled probes were hybridized onto a 4X44K yeast whole genome array (Agilent)
at 65°C for 16 h. The arrays were scanned with the Agilent microarray
scanner.Data availabilityData are downloadable at http://www.umassmed.edu/bmp/faculty/rando.cfm and have been
deposited in GEO (Accession # GSE28269).AnalysisRaw sequencing data of HA and T7 libraries after the tag swap before release
from arrest (0 generations), and at 1, 3, and 6 generations after release
were uniquely mapped to the S. cerevisiae genome.
Nucleosome positions were called from aggregated HA and T7 sequencing of the
3 generation sample using Template-Filtering [34]. For each nucleosome,
we counted the number of supporting reads for each sample separately and
calculated the ratio of HA reads to T7 reads at each nucleosome for each
time point. Note, for the H4tailΔ data the T7 data quality was poor, so
we used wild-type T7 sequences for this comparison.For aggregated analyses such as those shown in Figure 2E or Figure 3, we calculated the median of the
Log2 HA/T7 over the 1 kb starting at a given gene's transcription start
site to provide a summary retention score per gene.ModelDescribed in Text S1.Supporting InformationFigure S1Recombination efficiency. Yeast were plated onto nonselective media and onto
media selecting for the HA tag (linked to Hygro), before
(t = 0) and after (t = o/n)
inducing recombination. Roughly 2% of yeast fail to swap out the
HA-Hygro insert.(PDF)Click here for additional data file.Figure S2HA/T7 at 3 and 6 generations after release. HA/T7 ratios (Log2) for
individual nucleosomes are scatterplotted as indicated, showing good
correlation but a slope <1 (red line), consistent with the background of
nonswitching cells observed in Figure S1.(PDF)Click here for additional data file.Figure S3Validation of target genes. (A, C) Deep sequencing data (3 generations) for
SPA2 (A) and BUD3 (C). (B, D) qPCR
shown for the 5′ and 3′ ends of SPA2 (B) and
BUD3 (D) at the indicated number of generations after
tag-swap and release. Midlog refers to samples taken 3 h after the tag-swap
was induced in exponentially growing cells that had not undergone a recent
arrest. Note that only a fraction of all the cells had recombined out the
old tag during the 3 h. qPCR amplicon locations are indicated under the gene
annotation.(PDF)Click here for additional data file.Figure S4K means clustering of HA/T7 ratios. Log(2) HA/T7 ratios at 3 generations
after release are shown as a heatmap for all genes, aligned by transcription
start site (TSS) and clustered (K means, K = 5).
Selected Gene Ontology (GO) enrichments for the various clusters are
indicated to the right of the clusters.(TIF)Click here for additional data file.Figure S5Histone retention anticorrelates with turnover. (A) Average profiles for the
5 clusters from Figure S4 are plotted relative to
TSS-aligned coding regions. (B) Replication-independent turnover (Z score,
[15])
was averaged for 5′ CDS, mid-CDS, and 3′ CDS for all genes in
each cluster. Note that Cluster 2, which exhibits a somewhat
3′-shifted peak of HA/T7 relative to Clusters 3–5 (see A),
consists of genes with relatively high 5′ turnover, which presumably
explains the downstream location of the HA/T7 peak in this cluster.(PDF)Click here for additional data file.Figure S6Histone retention anticorrelates with transcription frequency. (A–B) As
in Figure 2B–C.
(C) As in (B), but using “transcription frequency” defined in
Holstege et al. [71] rather than Pol2 ChIP. (D) As in Figure 2F, but using
Holstege et al. data rather than Pol2 ChIP data.(TIF)Click here for additional data file.Figure S7H3 retention at subtelomeric genes. Median HA/T7 over the 5′ 1 kb of
all genes is plotted versus distance from the closest telomere, with an 80
gene running window average shown in red. No specific enrichment of H3-HA is
observed near telomeres. Similar results are found for repetitive
subtelomeric genes (unpublished data).(PDF)Click here for additional data file.Figure S8Mutant analysis of ancestral H3 retention. (A) 5′/3′ ratio at
SPA2 or BUD3 were measured by q-PCR
for the various mutants 3 generations after release or after one round of
replication arrested in G2/M. 5′/3′ ratio relative to wild-type
level is plotted on the y-axis. Mutants are as indicated,
with swd1, swd3 referring to an average of
single replicates with each individual mutant and scc1
referring to one experiment using a pGAL1-SCC1 allele that
was shut off by release into glucose media after the tag switch (leading to
a G2/M arrest). Average of mutant/wt, ± S.E.M.
(n = 2). Swd1 and Swd3 are components
of the Set1 complex, which methylates H3K4. Set2 is the H3K36 methylase and
Scc1 is part of the cohesin complex. (B) 5′/3′ ratio at
SPA2 or BUD3 were measured by q-PCR
for the various mutants after one round of replication arrested in G2/M.
5′/3′ ratio is plotted on the y-axis with
wild-type set to 1. Average ± S.E.M.
(n = 2). H4K5,12R is a mutant in which
two of the acetylatable lysines of the H4 tail have been replaced by
arginine, mimicking the unacetylated state. Rtt109 is a histone
acetyltransferase that binds to Asf1 and acetylates new histone H3 on K56
[72].
Nhp6a/b are non-essential HMGB proteins [73] that are required for
FACT activity. The FACT core subunits Spt16 and Pob3 are essential,
precluding us from testing their functions in ancestral histone inheritance.
(C) As in (A–B), for the indicated PCNA (POL30) point
mutants. Average of mutant/wt, ± S.E.M.
(n = 2).(PDF)Click here for additional data file.Figure S9HA retention on length-normalized genes. (A) All genes were normalized to a
length of 1, and genes are ordered by Pol2 ChIP. Log2 HA/T7 ratios are shown
as a heatmap. (B) Running window average of data from (A). Note the 5′
shift of the downstream edge of the HA/T7 peak with increasing transcription
rates. (C) Pol2 ChIP for genes as ordered in (A–B). (D) Averages for
all length-normalized genes grouped into 6 bins of transcription level.(PDF)Click here for additional data file.Figure S105′ accumulation of ancestral histones occurs in the absence of nutrient
stress. (A) Yeast carrying the HA/T7 recombination cassette were grown
continuously in YPD, then were treated with β-estradiol for 6 h to
induce recombination. HA and T7 ChIPs were carried out after 6 h and deep
sequenced, and normalized HA/T7 ratios were calculated. Here, genes are
ordered in 5 clusters as in Figure S4. (B) Averaged data for cells
arrested, switched, and released for 3 generations (“3 gen”) are
shown alongside data from the midlog switch. Note that 5′ accumulation
occurs in both conditions but to a lesser extent in the midlog swap. This is
an expected result of the heterogeneity of switch timing in midlog
cells—only 65% of yeast have completed recombination after 3 h,
with 85% complete by 6 h (unpublished data), meaning that the midlog
switch represents a mixture of cells that have recently swapped tags with
those that swapped tags ∼1–3 generations prior. (C) As in (B), for
intermediate and long genes.(TIF)Click here for additional data file.Figure S11A quantitative model accurately captures ancestral H3 retention patterns.
Model predictions (red lines) and data (blue lines) for HA/T7 ratios at 1,
3, and 6 generations after tag swap are shown for 1–2 kb genes (A) and
>2 kb genes (B). The model performs better on longer genes than on short
genes.(PDF)Click here for additional data file.Figure S12Passback correlates with transcription rate. Estimated passback parameters
for each gene were compared to Pol2 ChIP values for each gene. Scatterplot
is colored by density of points—red indicates greater density of
points. White line indicates linear fit to dataset,
R = 0.12.(PDF)Click here for additional data file.Figure S13Short genes and 3′ ends are poorly predicted by the model. Genes are
ordered by length, and the difference between model predictions for 3
generations and actual data are shown as a heatmap—yellow indicates
the model predicts excessive old nucleosome loss, or lower HA/T7 ratios than
measured. Notably, the +N nucleosome is universally predicted to lose
more H3-HA than is measured. This is almost certainly a consequence of the
fact that our model considers all genes in isolation—there is no way
for histones to spread onto the 3′ end of a gene from adjacent genomic
loci in this model, although this likely occurs in vivo.(PDF)Click here for additional data file.Figure S14The H4 N-terminal tail qualitatively changes the relationship between
transcription and ancestral H3 retention. (A) Tag swap strains carrying an
H4 N-terminal tail deletion were processed as in Figure 2B. Genes are ordered by the
median HA/T7 over the 5′ 1 kb. (B) Pol2 ChIP was carried out in the H4
tail deletion strain 2 generations after release from arrest. Data here show
an 80 gene running window average of Pol2 ChIP level per gene. (C) As in
Figure 2D. Genes
with high HA/T7 ratios in the H4 tail deletion mutant actually tend to be
slightly more enriched for Pol2 than those with low HA/T7
levels, the opposite of what is seen in wild-type (although it is important
to note that the correlation with Pol2 levels in this mutant is very
weak—note that the scale bar for Pol2 ChIP here ranges from −0.1
to 0.1, whereas the scale bar in Figure 2D ranges from −0.2 to 1).
Thus, we can conclude with confidence that the effects of the H4 tail
deletion do not simply result from extensive transcription reprogramming in
these mutants, since the relationship between Pol2 and H3-HA retention
qualitatively changes in this mutant.(TIF)Click here for additional data file.Figure S15Replication time has subtle effects of ancestral H3 patterns. (A) Data for
the 20% earliest and 20% latest-replicating [74]
genes is averaged as indicated. (B) As in (A), with gene lengths normalized
to one.(PDF)Click here for additional data file.Figure S16CAF-1 mutation affects far-5′ end levels of ancestral H3. Averaged data
for clusters 4 and 5 (Figure S4) are shown for wild-type and
cac1Δ.(PDF)Click here for additional data file.Figure S17Ancestral H3 retention and histone modification patterns. Modification levels
[52]
compared to ancestral H3 retention. For each modification, genes were
grouped into high, middle, and low HA/T7 (based on the 5′-most 1 kb
median HA/T7), and for each group of genes modifications were averaged for
5-CDS (“5”), mid-CDS (“m”), and 3′-CDS
(“3”) as previously described [15],[19]. Groupings are indicated
for H3K9ac and for H3K79me3 and are the same for the other three
modifications.(PDF)Click here for additional data file.Figure S18Role for histone movement in shaping modification landscapes. Schematic for
retrograde histone movement in shaping histone modification landscapes.
Initial 5′ (green) and 3′ (purple) histone modification states
could, in the absence of erasing enzymes, eventually give rise to skewed
distributions via retrograde motion of old histones bearing 3′
modifications such as H3K36me3 (purple). Importantly, after a few cell
divisions old histones on average constitute only a minor fraction of all
histones at any given locus (e.g., see Figure
S3B, D). Modifications on ancestral histones will therefore make
subtle contributions to overall average modification patterns.(PDF)Click here for additional data file.Table S1HA/T7 ratios for 3 generations. Log2(HA/T7) for all genes is shown at varying
distances with respect to the TSS. Genes are sorted according to the median
HA/T7 over the 5′ 1 kb.(XLSX)Click here for additional data file.Table S2Enrichments of genesets for high or low retrograde passback values. A variety
of gene sets were searched for enrichment of relatively high or low values
for model estimates of lateral nucleosome movement. Negative KS values
indicate large 3′ to 5′ lateral movements; positive values
indicate the converse.(XLSX)Click here for additional data file.Table S3Strain list. Genotypes of strains used in this study.(XLS)Click here for additional data file.Table S4Primers used. Primer sequences.(XLS)Click here for additional data file.Text S1Quantitative model for multigenerational histone dynamics. Detailed
description of model shown in Figures 5 and 6.(PDF)Click here for additional data file.
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