loss of histone h3.3 results in dna replication …46 dna ahead of the replication fork (sonneville...
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Title: Loss of histone H3.3 results in DNA replication defects and 1
altered origin dynamics in C. elegans. 2
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Authors: Maude Strobino, Joanna M. Wenda and Florian A. Steiner 4
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Affiliations: 8
Department of Molecular Biology and Institute for Genetics and Genomics in Geneva, 9
Section of Biology, Faculty of Sciences, University of Geneva, 1211 Geneva, 10
Switzerland 11
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Correspondence to: [email protected] 13
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Abstract 15
Histone H3.3 is a replication-independent variant of histone H3 with important roles in 16
development, differentiation and fertility. Here we show that loss of H3.3 results in 17
replication defects in Caenorhabditis elegans embryos. To characterize these defects, 18
we adapt methods to determine replication timing, map replication origins, and examine 19
replication fork progression. Our analysis of the spatiotemporal regulation of DNA 20
replication shows that despite the very rapid embryonic cell cycle, the genome is 21
replicated from early and late firing origins and is partitioned into domains of early and 22
late replication. We find that under temperature stress conditions, additional replication 23
origins become activated. Moreover, loss of H3.3 results in impaired replication fork 24
progression around origins, which is particularly evident at stress-activated origins. 25
These replication defects are accompanied by replication checkpoint activation, a 26
prolonged cell cycle, and increased lethality in checkpoint-compromised embryos. Our 27
comprehensive analysis of DNA replication in C. elegans reveals the genomic location 28
of replication origins and the dynamics of their firing, and uncovers a role of H3.3 in the 29
regulation of replication origins under stress conditions. 30
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Keywords: Histone H3.3, replication, origins, Repli-seq, EdU-seq, ChEC-seq, DNA 32
combing, C. elegans 33
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Introduction 36
At every cell cycle, the entire genome is replicated exactly once for accurate transmission of 37
genetic information to the daughter cells. To ensure rapid and uniform duplication of the 38
genome, DNA replication is initiated in a bi-directional way at many different sites in the 39
genome, called replication origins. To prevent re-replication and to control the activity of origins 40
under replication stress, replication initiation is composed of two non-overlapping steps called 41
‘origin licensing’ and ‘origin firing’. Origin licensing occurs during G1 through the formation of the 42
pre-replicative complexes. During this process, the origin recognition complex (ORC) binds to 43
the origins and recruits the replication licensing factors Cdt1 and Cdc6, which together load the 44
complex of the minichromosome maintenance proteins 2-7 (Mcm2-7), a helicase that unwinds 45
DNA ahead of the replication fork (Sonneville et al., 2012). To ensure that each origin fires only 46
once per cell cycle, Cdt1 and Cdc6 are subsequently removed from the nucleus or degraded to 47
prevent the Mcm2-7 complex from rebinding origins that have already fired (Blow and Dutta, 48
2005; Feng and Kipreos, 2003; Kim and Kipreos, 2007). Origin firing occurs upon entry into S 49
phase, through binding of different initiation factors like Sld2 (RecQL4/RecQ4 in humans), Sld3 50
(Treslin/TICRR in humans) and Dpb11 (TopBP1 in humans), and phosphorylation of the Mcm2-51
7 complex (Bruck and Kaplan, 2015; Kanemaki and Labib, 2006; Labib and Diffley, 2001). 52
These firing factors are present in limiting amounts (Mantiero et al., 2011; Tanaka et al., 2011), 53
and all origins therefore do not fire synchronously at the start of S phase. Instead, some origins 54
fire early and some late, resulting in large domains of early and late replication (Desprat et al., 55
2009; Dileep and Gilbert, 2018; Farkash-Amar et al., 2008; MacAlpine et al., 2004; Takahashi et 56
al., 2019). These domains may change during development, and it has been shown in multiple 57
organisms that early replicating domains are associated with transcriptionally active regions 58
(Hiratani et al., 2010, 2008; Schübeler et al., 2002; Therizols et al., 2014; White et al., 2004; 59
Woodfine et al., 2004). 60
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Upon replication origin firing, the replication forks move bi-directionally until replication bubbles 62
of opposing directions meet and the replication of the corresponding stretch of chromosome is 63
complete. Under specific conditions, e.g. conflicts between transcription and replication forks, 64
depletion of nucleotides, or depletion of histones, replication forks can stall or collapse (Bester 65
et al., 2011; Groth et al., 2007; Macheret and Halazonetis, 2018). This replication stress can 66
lead to DNA damage (Macheret and Halazonetis, 2015; Técher et al., 2017), and triggers the 67
activation of the ATR checkpoint, which phosphorylates the Chk1 kinase, leading to cell cycle 68
arrest (Hyun et al., 2016; Liu et al., 2000; Moiseeva et al., 2019). As a response to moderate 69
replication stress, cells can activate additional origins (Técher et al., 2017). These origins, called 70
latent or dormant origins, are inactive and do not fire during the undisturbed S phase, but act as 71
backup origins in case of replication stress (Boos and Ferreira, 2019; Woodward et al., 2006). 72
How firing origins are selected over dormant origins, and under what conditions dormant origins 73
are activated is unclear (Fragkos et al., 2015). 74
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It is widely accepted that replication origins are not defined by DNA sequences in higher 76
eukaryotes and therefore cannot be predicted from genome sequences, making their 77
identification challenging. Numerous assays have been developed to map replication origins in 78
different organisms (Prioleau and MacAlpine, 2016), mostly by mapping different replication 79
elements such as nascent DNA strands (Besnard et al., 2012), replication bubbles (Mesner et 80
al., 2013), Okazaki fragments (Petryk et al., 2016), replication initiation sites (Langley et al., 81
2016), or initiation factors (Dellino et al., 2013; Miotto et al., 2016). Despite these efforts, the 82
number, location and dynamic regulation of replication origins remains controversial. For 83
example, two studies in C. elegans recently provided genome-wide maps of replication origins 84
using two different methods, and arrived at largely different conclusions (Pourkarimi et al., 2016; 85
Rodríguez-Martínez et al., 2017). The first study used Okazaki fragment sequencing to identify 86
~2’000 origins and concluded that replication origins are defined prior to the broad onset of 87
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zygotic transcription and maintained during development (Pourkarimi et al., 2016). The second 88
study applied mapping of small RNA-primed nascent DNA strands to identify ~16’000 origins 89
and described a major re-organization of replication origins after gastrulation (Rodríguez-90
Martínez et al., 2017). Additional approaches will therefore be required to understand how 91
origins are defined and dynamically regulated. 92
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The variant histone H3.3 has been linked to different steps of DNA replication. It is enriched at 94
replication origins in Drosophila melanogaster and Arabidopsis thaliana (Deal et al., 2010; Eaton 95
et al., 2011; MacAlpine et al., 2010; Paranjape and Calvi, 2016; Stroud et al., 2012), and loss of 96
H3.3 results in a slower replication fork in chicken cells (Frey et al., 2014). The association of 97
H3.3 with early replication domains, and its recruitment to sites of DNA repair have also been 98
described in human cells (Adam et al., 2013; Clément et al., 2018). H3.3 is incorporated into 99
transcriptionally active regions as well as in centromeric and telomeric repeats (Buschbeck and 100
Hake, 2017; Henikoff and Smith, 2015), but the link between its genomic location and its 101
function is not always clear. Loss of H3.3 results in lethality or sterility in most organisms (Hödl 102
and Basler, 2009; Jang et al., 2015; Sakai et al., 2009; Tang et al., 2015; Wollmann et al., 103
2017). The C. elegans genome contains five genes encoding H3.3 homologues, and we 104
recently showed that mutation of all five genes resulted in viable worms with a reduced number 105
of viable offspring at higher temperatures (Delaney et al., 2018). 106
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Given the role of H3.3 in DNA replication in other organisms, we speculated that the reduced 108
brood size observed in C. elegans H3.3 null mutants is caused by defects in DNA replication. 109
Here we show that at higher temperatures, loss of H3.3 causes mild DNA replication defects 110
and activation of the embryonic DNA replication checkpoint. To investigate these defects, we 111
adapt ChEC-seq, Repli-seq, EdU-seq and DNA combing for use in C. elegans, which we apply 112
to define and characterize replication timing and replication origins. We find that during S phase, 113
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replication and the firing of replication origins are temporally regulated. H3.3 is enriched at 114
replication origins, but is not required for origin firing. Instead, loss of H3.3 results in delayed 115
fork progression around replication origins, in particular around origins that are only active under 116
temperature stress conditions. Together, our results uncover a role of H3.3 in DNA replication in 117
C. elegans and show that replication origin use can be adapted in response to stress. 118
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Results 121
Genome-wide replication timing in C. elegans embryos 122
Histone H3.3 has been linked to the regulation of DNA replication in a variety of organisms 123
(Adam et al., 2013; Clément et al., 2018; Deal et al., 2010; Eaton et al., 2011; Frey et al., 2014; 124
MacAlpine et al., 2010; Paranjape and Calvi, 2016; Stroud et al., 2012). We therefore 125
hypothesized that the reduced brood sizes observed in C. elegans H3.3 null mutants under mild 126
temperature stress are caused by defects in DNA replication (Delaney et al., 2018). Our first aim 127
was to determine whether loss of H3.3 causes global changes in the temporal program of 128
genome replication. To characterize replication timing during the rapid cell cycle in early C. 129
elegans embryogenesis, we adapted Repli-seq for use in this organism (Hansen et al., 2010). 130
This method relies on the incorporation of the thymidine analogue 5-ethynyl-2'-deoxyuridine 131
(EdU) into newly replicated regions of the genome (Figure 1A). To separate early and late 132
replicated genomic regions, we pulse-labelled asynchronous embryonic cell populations with 133
EdU, then sorted the cells according to their DNA content and sequenced EdU-containing 134
fragments. We obtained cells that just entered S phase (early replication), showed ongoing DNA 135
synthesis (mid replication), or were at or near completion of replicating the genome (late 136
replication) (Figure S1A). These Repli-seq experiments revealed that the genome is partitioned 137
into mutually exclusive domains of early and late replication (Figure 1B, C). Surprisingly, 138
different chromosomes display distinct overall replication timing patterns. Domains of early 139
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replication are enriched on chromosomes I, II and III, while domains of late replication are more 140
prevalent on chromosomes IV, V and X (Figure 1C and Figure S1B). We found that domains of 141
early replication correlate well with the presence of HIS-72, the most highly expressed C. 142
elegans H3.3 homologue (Figure 1B and Figure S1C). However, H3.3 null mutant worms 143
showed largely the same early and late replication domains as wild-type worms, indicating that 144
H3.3 is not a major driver of global DNA replication timing (Figure 1B, C and Figure S1B). 145
Genes in domains of early replication show higher levels of expression in embryo compared to 146
genes in domains of later replication, consistent with the observation from other organisms that 147
regions of open chromatin replicate early (Ekundayo and Bleichert, 2019; Prioleau and 148
MacAlpine, 2016) (Figure 1D). However, there is no enrichment of large genes in late replicating 149
domains that would suggest potential interference between transcription and replication under 150
stress conditions, as observed in human cells (Figure 1E) (Macheret and Halazonetis, 2018). 151
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Mapping replication origins by ChEC-seq and EdU-seq 153
Given the presence of domains of early and late replication, we reasoned that there should be 154
distinct populations of early and late firing replication origins. With the enrichment of H3.3 at 155
replication origins in other organisms, we speculated that loss of H3.3 may cause subtle defects 156
in the position or timing of origin firing in H3.3 null mutants. We therefore next aimed to define 157
replication origins and their timing during S phase. Previous origin mapping approaches in C. 158
elegans do not allow for the analysis of origin dynamics during the cell cycle (Pourkarimi et al., 159
2016; Rodríguez-Martínez et al., 2017). We therefore took an alternative approach that 160
combines the mapping of replication origin factors and the observation of nucleotide 161
incorporation at origins. We first profiled the genomic binding sites of proteins involved in 162
replication origin licensing and firing. We selected two conserved proteins, CDT-1 and TRES-1, 163
that are involved in different steps of origin activation (Figure 2A). CDT-1 is the homologue for 164
human Cdt1 and is implicated in the licensing of all origins during G1 phase (Zhong et al., 2003) 165
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(Figure 2A). TRES-1 is the homologue of human Treslin and is subsequently recruited 166
exclusively to the origins that are going to fire (Guo et al., 2015; Kumagai et al., 2010). 167
Immunofluorescence microscopy confirmed that CDT-1 is present on chromatin during 168
anaphase and telophase, but levels become undetectable during S-phase. TRES-1 only 169
appears on chromatin during telophase, but remains present during S phase (Figure 2B). These 170
observations are consistent with their roles in origin licensing and origin firing, respectively. 171
Typically, genomic protein binding sites are profiled by chromatin immunoprecipitation. 172
However, this approach depends on antibody availability and solubility of the chromatin 173
component of interest, all of which can be limiting factors. To circumvent these problems, we 174
adapted Chromatin Endogenous Cleavage (ChEC) to C. elegans. This method relies on the 175
fusion of the protein of interest with micrococcal nuclease (MNase), which cuts DNA around the 176
binding sites of the fusion protein and thereby releases DNA fragments that can be sequenced 177
and mapped (Schmid et al., 2004; Zentner et al., 2015) (Figure 2C). We generated MNase 178
fusion constructs at the endogenous cdt-1 and tres-1 loci using CRISPR/Cas9. MNase 179
insertions resulted in worms that are phenotypically wild-type. To generate genome-wide 180
binding maps of CDT-1 and TRES-1, we performed ChEC-seq experiment on purified 181
embryonic nuclei and identified peaks of CDT-1 and TRES-1 signal. We observed CDT-1 and 182
TRES-1 peaks that are distributed across the genome in both datasets. Moreover, as expected, 183
the CDT-1 peaks mostly overlap with TRES-1 peaks (licensed and firing), but some CDT-1 184
peaks show no corresponding TRES-1 signal (licensed only) (Figure 2D). 185
The CDT-1 and TRES-1 ChEC-seq data allowed us to discriminate between licensed and firing 186
origins, but we could not establish the timing of origin firing within S phase. To identify 187
replication origins that fire immediately after entry into S phase, we modified the Repli-seq 188
protocol. Instead of analyzing asynchronous cell populations, we used hydroxyurea (HU) to 189
synchronize cells at the entry of the S phase. Upon HU treatment, virtually all cells become 190
arrested at the entry of the S phase, and a large part of the cell population proceeds with DNA 191
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replication upon removal of HU (Figure S2A). We combined the release into S phase with a 192
second HU block in the presence of EdU, allowing the cells to replicate short stretches of DNA 193
around early replication origins before arresting. The EdU-containing DNA fragments were then 194
isolated and sequenced. These EdU-seq experiments revealed discrete peaks that are located 195
in close proximity to with a subset of the CDT-1 and TRES-1 peaks (licensed and firing with 196
EdU signal) (Figure 2D). The EdU-seq signal sometimes appear as double peaks flanking the 197
CDT-1 and TRES-1 peaks, indicating that we detect short fork movements away from the 198
replication origins. We conclude that the sites enriched for EdU-seq, CDT-1 and TRES-1 signal 199
correspond to replication origins that fire upon entry into S phase. 200
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Classifications of origins genome wide 202
To obtain a map of embryonic replication origins genome-wide, we combined the peak calls 203
from the CDT-1 ChEC-seq, TRES-1 ChEC-seq and EdU-seq experiments. In total, we obtained 204
a map of 1110 origins. Based on their signal in the three datasets, unsupervised clustering 205
distinguished three types of origins, which we classified as “early firing”, “late firing” and 206
“dormant” (Figure 3A). Early firing origins show enrichment in all three datasets, as they are 207
licensed (CDT-1) and fire at the onset of S phase (TRES-1 and EdU-seq). Late firing origins 208
show enrichment in CDT-1 and TRES-1 ChEC-seq datasets, but not EdU-seq, indicating that 209
they fire after the onset of S phase. Dormant origins show enrichment only in the CDT-1 ChEC-210
seq dataset, indicating that they are licensed, but do not fire under standard conditions (Figure 211
3A). While the enrichment of CDT-1 and TRES-1 overlaps within a narrow window at the 212
replication origins, the distribution of EdU-seq signal is more broad, which likely reflects short 213
stretches of fork progression away from the replication origins after the release from the initial 214
HU block. As expected, firing origins are found mostly outside of gene-coding regions, while 215
dormant origins are enriched within genes (Figure S2B). The genomic localization of the 216
replication origins identified here largely overlap with those previously identified (Figure S2C), 217
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but our mapping approach offers the advantage of capturing origin firing dynamics during the 218
cell cycle, and also includes dormant origins that may act as backup origins for use under 219
specific conditions. Similar to D. melanogaster and A. thaliana (Deal et al., 2010; Eaton et al., 220
2011; MacAlpine et al., 2010; Paranjape and Calvi, 2016; Stroud et al., 2012), H3.3 is enriched 221
around the origins of all three categories (Figure 3B), suggesting that it may play a role in origin 222
identity or activation. 223
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Altered origin dynamics in H3.3 null mutants 225
We next compared the distribution of firing origins in wild type and H3.3 null mutant worms. For 226
this, we repeated the TRES-1 ChEC-seq and the EdU-seq experiments in wild type and H3.3 227
null mutant strains at 25°C, where a difference in brood-size between the two strains is 228
observed. The TRES-1 ChEC-seq experiments showed that most origins licensed in wild type 229
worms were also utilized in absence of H3.3 (Figure 4A). In both strains, we also observed 230
weak TRES-1 signal at origins that are dormant at 20°C. These origins were initially identified 231
solely based on the presence of CDT-1 under normal conditions, and the presence of TRES-1 232
at 25°C shows that these are functional origins that become activated at higher temperatures. 233
This result confirms the presence of “backup” origins that are inactive during undisturbed S 234
phase, but are activated under replication stress (Boos and Ferreira, 2019; Woodward et al., 235
2006). We did not detect any difference between the firing of early origins in wild type and H3.3 236
null mutant based on EdU-seq signal (Figure 4C, 0min). These results indicate that the origin 237
distribution and the firing of early origins remains unchanged upon loss of H3.3. 238
Although the firing of early origins is not affected by the loss of H3.3, we were wondering if fork 239
progression following origin firing could be altered. The EdU-seq method not only allows one to 240
identify early firing origins, but can also be used to visualize fork progression around these 241
origins, by adding an EdU pulse at 0, 30 or 60 minutes after the release from the HU block 242
(Figure 4B-C). As expected, the EdU signal enrichment at the early firing origins is no longer 243
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present after 30 minutes, and regions occupied by early firing origins incorporate very little EdU 244
after 60 minutes, which reflects the bi-directional movement of the replication forks away from 245
the origins (Figure 4B-C, WT). This bi-directional fork movement is difficult to observe for late 246
origins, as we have no means to synchronize their firing. Nevertheless, the EdU-seq signal at 247
late origins appears depleted at 30 minutes, suggesting that they fire within the first 30 minutes 248
after S-phase onset. In H3.3 null mutants, as mentioned above, early origins appear to fire 249
normally, and we observe bi-directional fork movement. However, we found that the EdU signal 250
remains higher around early origins after 30 and 60 minutes, indicating that fork progression is 251
delayed (Figure 4B-C, ∆ H3.3). This delay in fork progression appears at origins of all classes 252
and results in an enrichment of EdU incorporation at late origins. The difference between wild 253
type and H3.3 null mutant cells is even more pronounced at origins that are dormant under 254
standard conditions. These origins fire in both strains at 25°C, as evidenced by the increased 255
presence of TRES-1 compared to 20°C (Figure 3A and Figure 4A), but fork progression from 256
the origins is impaired in H3.3 null mutants, resulting in an accumulation of EdU-seq signal at 257
later time points. The enrichment at late and dormant origins is observed within a very narrow 258
window, suggesting that the forks stall or collapse immediately after origins fire. The 259
coincidence of this EdU-seq signal enrichment with the center of late and dormant origins is 260
particularly remarkable, as these origins were identified solely based on a completely 261
independent method, namely the mapping of the CDT-1 and TRES-1 binding sites. The 262
aberrant fork progression in H3.3 null mutant worms is only seen under mild temperature stress 263
conditions, and is not observed at 20°C (Figure S3), indicating that H3.3 is required for 264
regulating origin dynamics under stress. 265
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Fork speed is unaffected by loss of H3.3 267
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To establish if loss of H3.3 resulted in a general slowing of the replication fork speed along the 268
chromatin fiber, we adapted DNA combing to C. elegans (Michalet et al., 1997). This method 269
relies on the pulse labeling of DNA by the subsequent incorporation of two thymidine analogues, 270
IdU and CldU. By visualizing the length of IdU and CldU incorporation on stretched individual 271
chromatin fibers, replication fork speed can be estimated (Figure 5A). We found that fork speed 272
is on average 1.4 kb/min (Figure 5B), but did not detect any significant difference between wild 273
type and H3.3 null mutant embryos, indicating that H3.3 is not required for normal progression 274
of the moving replication fork. 275
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Increased DNA damage in H3.3 null mutant embryos 277
DNA combing assesses the speed of the moving replication fork, but it does not measure the 278
average fork speed genome-wide, which can be affected by fork pausing, stalling or collapse. 279
Our EdU-seq time course analysis suggested that replication origins fire normally upon loss of 280
H3.3, but that the progression of forks is altered around origins. We therefore probed the 281
embryos for the presence of replication stress that might be indicative of fork stalling and 282
collapse. In C. elegans embryos, DNA replication defects are characterized by an increased 283
length of the first embryonic cell cycle (Brauchle et al., 2003; Holway et al., 2006; Stevens et al., 284
2016). We measured cell cycle timing from the appearance of a cleavage furrow in the P0 cell to 285
nuclear envelope breakdown of the AB cell (Figure 6A). We found that the cell cycle time was 286
significantly increased in H3.3 null mutant embryos compared to wild type (Figure 6B). 287
Removing the replication checkpoint through targeting the checkpoint kinase 1 (CHK-1) by RNAi 288
restored normal cell cycle timing, confirming that the increased cell cycle length is linked to a 289
replication defect (Figure 6B). Moreover, removal of CHK-1 resulted in an increase of the 290
embryonic lethality in H3.3 null mutants (Figure 6C). Together, our results indicate that loss of 291
H3.3 results in defects in DNA replication or DNA damage repair, and that these defects 292
become detrimental upon removal of CHK-1. 293
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Taken together, we find that replication of the C. elegans genome is partitioned into early and 295
late domains, and that the global timing of replication and replication origin firing is unchanged 296
in H3.3 null mutant worms. However, loss of H3.3 leads to defective fork progression around 297
origins and DNA replication defects at 25°C. Our results identify the replication dynamics during 298
the embryonic cell cycle and uncover a role of H3.3 in DNA replication in C. elegans embryos. 299
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Discussion 302
The variant histone H3.3 has previously been linked to DNA replication, but its mechanistic role 303
in this process, and how its function relates to its genomic localization are not well understood. 304
In most animal models, loss of H3.3 results in lethality or sterility, making the developmental 305
analysis of H3.3 mutant challenging (Couldrey et al., 1999; Hödl and Basler, 2009; Sakai et al., 306
2009; Tang et al., 2015). Our recent discovery that C. elegans H3.3 null mutants are viable 307
allowed us to analyze potential developmental defects in more detail (Delaney et al., 2018). 308
Here we describe that under stress conditions, H3.3 is required for DNA replication dynamics 309
around replication origins and protects the genome from replication stress. 310
311
For understanding the replication defects upon loss of H3.3 in detail, we first had to develop 312
tools to investigate DNA replication dynamics in C. elegans embryos. DNA replication timing in 313
C. elegans had not been analyzed in detail, and replication origins have only recently been 314
mapped by different techniques, with conflicting outcomes (Pourkarimi et al., 2016; Rodríguez-315
Martínez et al., 2017). We therefore adapted techniques that are based on the incorporation of 316
EdU to determine the genome-wide replication timing (Repli-seq), and to follow the fork 317
progression around replication origins (EdU-seq). These experiments revealed the surprising 318
finding that despite the very short cell cycle during embryogenesis, genomic regions of early 319
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and late replication are mutually exclusive, and distributed in a non-random fashion. Partitioning 320
of the genome into early and late replicating domains is very well described for mammalian 321
genomes, both in cell population averages (Desprat et al., 2009; Farkash-Amar et al., 2008; 322
Hiratani et al., 2008; MacAlpine et al., 2004) and single cells (Dileep and Gilbert, 2018; 323
Takahashi et al., 2019) but had not yet been observed in C. elegans. Replication timing has 324
been attributed to the 3D organization of chromatin into discrete domains (Marchal et al., 2019). 325
We also observed this robust spatial organization of replication timing in C. elegans, despite 326
absence of strong 3D compartmentalization (Crane et al., 2015). Chromosomes I-III are 327
enriched for early replication domains, while most of chromosomes IV, V and X, and the 328
telomeric regions of all chromosomes replicate late. We note that genes located in 329
chromosomes I-III are on average more highly expressed than genes on chromosomes IV-X, 330
and may therefore constitute an environment of open chromatin that is permissive for the 331
binding of replication initiation factors. The domains of early replication are also enriched for 332
H3.3. A similar correlation between early replication and H3.3 incorporation has been observed 333
in human cells (Clément et al., 2018). Given this conserved correlation, it is maybe surprising 334
that loss of H3.3 does not globally alter the domains of early and late replication. However, 335
replication timing has been found to be remarkably robust, and remains unaltered in a wide 336
range of mutants that affect chromatin organization (Marchal et al., 2019). Our results support 337
the notion that the concentrations of some factors implicated in DNA replication are limiting, and 338
that they will access the regions of open chromatin more readily (Mantiero et al., 2011). The 339
concentration of replication factors is tightly controlled during S phase, and failure to export or 340
degrade factors involved in licensing can lead to over-replication of the genome (Sonneville et 341
al., 2012). 342
343
To comprehensively identify replication origins throughout S phase, we mapped the genomic 344
binding sites of the origin licensing factor CDT-1 and the origin firing factor TRES-1. For 345
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determining their binding sites genome wide, we adapted the ChEC-seq method for use in C. 346
elegans (Schmid et al., 2004; Zentner et al., 2015). We then combined the results with the early 347
origins identified by EdU-seq. Our origin identification thus combines methods that detect 348
proteins involved in origin identity with methods that rely on the identification of sites of 349
nucleotide incorporation during origin firing. This approach allows the identification of origin 350
firing dynamics during the cell cycle, and also includes dormant origins. C. elegans origins have 351
been mapped previously using Okazaki fragment sequencing or nascent strand sequencing, 352
both of which should identify firing origins (Pourkarimi et al., 2016; Rodríguez-Martínez et al., 353
2017). Our approach identified significantly fewer origins (~1’000 vs ~2’000 or ~16’000, 354
respectively), but the genomic location of the origins identified in this study largely overlap with 355
those previously identified (Figure S2C). Previous approaches utilized the depletion of DNase 356
lig-1 by RNAi (Pourkarimi et al., 2016) or liquid culture (Rodríguez-Martínez et al., 2017) for the 357
cultivation of worms for origin analysis. Both treatments result in worms that are superficially 358
wild type, as is the case for H3.3 null mutant worms. Given our observation that even mild 359
temperature stress can affect origin firing, and that liquid culture can affect gene expression 360
(Çelen et al., 2018), these treatments could alter origin use and contribute to the observed 361
differences in origin numbers. To independently estimate the number of origins required to 362
replicate the entire genome in our system, we measured the speed of the replication fork on 363
single molecules by DNA combing, and determined the time required to reach maximum EdU 364
incorporation within one cell cycle. We found that a plateau of EdU incorporation is reached 365
about 40 minutes after the first observed EdU incorporation, suggesting that cells require about 366
40 minutes to replicate the entire genome under these conditions (Figure S4). This is 367
substantially longer than the S-phase duration observed in developing embryos (Sonneville et 368
al., 2012), and we attribute the delay to the dissociation and the treatment with HU and EdU. 369
Based on this replication time and the fork speed measured in the DNA combing experiment, 370
we estimate the number of firing origins required for replicating the entire genome to be about 371
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924. This number is remarkably close to the number of firing origins identified by ChEC-seq and 372
EdU-seq (798), and we therefore think that our mapping strategy accurately reflects active 373
replication origins. While our methods allow for the detection of origins that only fire under 374
specific conditions, they still rely on averaging signal from very large populations of cells, and 375
will therefore be less sensitive to detect origins that are only licensed at specific developmental 376
time points or in specific cell types. 377
378
H3.3 is enriched at replication origins in D. melanogaster and A. thaliana, but the functional 379
significance of this enrichment remains unclear, and no defects in origin dynamics have been 380
detected upon loss of H3.3 (Deal et al., 2010; Eaton et al., 2011; MacAlpine et al., 2010; 381
Paranjape and Calvi, 2016; Stroud et al., 2012). We find that H3.3 is enriched at replication 382
origins, but consistent with the findings from D. melanogaster, we found no evidence that loss of 383
H3.3 inhibits replication origin firing (Paranjape and Calvi, 2016). The EdU-seq enrichments at 384
early origins remains unchanged, and H3.3 null mutant worms are able to activate dormant 385
origins similar to wild type under stress conditions, as evidenced by the TRES-1 signal observed 386
at 25°C. However, we found clear differences in EdU incorporation around origins as S phase 387
progresses. Fork progression from origins is impaired, and the EdU-seq signal remains enriched 388
at replication origins, in particular at those that only fire under stress conditions. The speed of 389
the replication fork along the chromatin fiber is not affected by the loss of H3.3, as determined 390
by DNA combing experiments. It is therefore likely that instead the frequency of fork stalling or 391
fork collapse is increased as the fork is progressing away from the replication origins, resulting 392
in the increased EdU-seq signal around replication origins. Our observation of the replication 393
checkpoint activation through CHK-1 is also consistent with increased replication stress rather 394
than a general slowing of fork progression. The defects are only observed at 25°C, and origin 395
dynamics in wild type and H3.3 null mutant worms are indistinguishable at 20°C, consistent with 396
the fertility defects of H3.3 null mutant worms that are only observed at higher temperatures. 397
17
The reasons behind the temperature sensitivity of the replication fork upon loss of H3.3 are 398
currently unclear. 399
400
The increased replication stress in the H3.3 null mutant could be caused directly by altering the 401
chromatin organization, or indirectly by affecting gene expression. Decreased nucleosome 402
density may affect replication fork progression. In D. melanogaster embryos, a decrease in 403
histone concentration results in an increased cell cycle length and CHK-1 activation, similar to 404
our observations in C. elegans embryos (Chari et al., 2019). However, only three genes 405
encoding for H3.3 are expressed during embryogenesis, compared to 16 genes encoding 406
canonical H3 (Boeck et al., 2016). It is therefore unlikely that loss of H3.3 results in a dramatic 407
drop in nucleosome density genome wide. However, replacing H3.3 nucleosomes with H3 408
nucleosomes may alter the chromatin dynamics and also contribute to the observed replication 409
stress. Absence of H3.3 may not cause replication stress directly, but may prevent fork restart 410
after fork pausing or collapse, similar to what has been observed after DNA damage in chicken 411
cells (Frey et al., 2014). Alternatively, loss of H3.3 could alter the transcriptional landscape 412
genome-wide, thus resulting in increased conflicts between transcription and DNA replication. 413
Indeed, the majority of the dormant origins, for which we find the strongest effects, are located 414
within genes (Figure S2B). However, we previously reported that loss of H3.3 only results in 415
minor changes in the transcriptome (Delaney et al., 2018). Loss of H3.3 could also alter the 416
histone marks around origins. C. elegans replication origins were found to be enriched for H3 417
acetylation, H3K4 methylation and H3K27 acetylation (Pourkarimi et al., 2016), and it is possible 418
that the presence of H3.3 aids the deposition of some of these marks. However, the causal 419
relationship between histone modifications and the identity of replication origins is not clear. 420
Many studies have accumulated evidence that chromatin context is important for the activity of 421
replication origins, and we now show that H3.3 plays an important role in the fork progression 422
following origin firing under stress condition. More research will be needed to establish a causal 423
18
relationship between the chromatin context and replication origin formation, to understand how 424
the timing of origin firing is regulated, and to determine how the replication fork interacts with 425
nucleosomes containing different histone modifications or variants. 426
427
428
Materials and Methods 429
Worm culture and strain generation 430
Worms were grown on NGM plates seeded with OP50 for maintenance, and on peptone-rich 431
NGM plates seeded with NA22 for large-scale experiments. The sequence coding for MNase 432
was inserted at the cdt-1 and tres-1 loci using CRISPR/Cas9 as described in (Arribere et al., 433
2014), using plasmids with 1 kb homology arms as repair templates. A list of the strains used in 434
this study is given in Table S1. RNAi experiments were carried out by feeding, using clones 435
from the Ahringer library (Kamath et al., 2003). For experiments measuring effects at 25°C, 436
worms were grown at this temperature for at least 5 generations. 437
438
Phenotype analysis 439
For determining embryonic lethality at 25°C, L4 worms were placed on plates containing RNAi 440
or control food overnight. Single adults were transferred to new plates for 3h and then removed. 441
The number of eggs, hatched larvae and adults was counted on subsequent days to determine 442
the penetrance of embryonic lethality. Plates with 0 or 1 embryos were excluded from the final 443
count. Data from three independent experiments with 10 plates per experiment and condition 444
were used. Total number of embryos: WT control = 445, ∆ H3.3 control = 240, WT chk-1 RNAi = 445
270, ∆ H3.3 chk-1 RNAi = 251. 446
19
For determining the cell cycle length during early embryonic cell divisions at 25°C, L4 worms 447
were placed on plates containing RNAi or control food overnight. Adults were cut in egg buffer 448
(118mM NaCl, 48 mM KCL, 2mM CaCl2, 2mM MgCl2 and 25mM Hepes pH7,5) to release 449
young embryos. These embryos were mounted on 2% agarose pads and recorded every 10 s 450
with DIC settings on a Leica DMI8 microscope, with the temperature maintained at 25°C. Cell 451
cycle length was measured from the appearance of the cleavage furrow of the P0 cell to nuclear 452
envelope breakdown of the AB cell. WT control = 3, ∆ H3.3 control = 5, WT RNAi = 4, ∆ H3.3 453
RNAi = 4. 454
For determining the subcellular localization of CDT-1 and TRES-1, gravid adult worms were 455
washed in PBS containing 0.1% Triton X-100, anesthetized in DB buffer (50 mM sucrose, 75 456
mM HEPES pH 6.5, 60 mM NaCl, 5 mM KCl, 2 mM MgCl2, 10 mM EGTA pH 7.5, 0.1% NaN3) 457
and cut in half to release embryos. The embryos were transferred to poly-L-lysine-coated slides, 458
freeze-cracked and fixed in cold methanol for 5 min. Slides were washed two times in PBS and 459
incubated with anti-HA antibody (mAb 42F13, 1:60) for CDT-1 staining or FLAG antibody 460
(Sigma-Aldrich, F7425-.2MG, 1:200) for TRES-1 staining, overnight at 4°C. After two washes in 461
PBS, slides were incubated with secondary antibodies (Alexa Fluor 488 goat anti-mouse or 462
Alexa Fluor 488 goat anti-rabbit; 1:700) for 1 h at room temperature, washed once in PBS and 463
counterstained with DAPI (2 µg/ml) for 15 minutes. After a final PBS wash, the slides were 464
mounted in VECTASHIELD Antifade Mounting Medium. Images were acquired with Leica SP8 465
confocal microscope. For all the images Z-sections with 0.3 µm step were acquired, and 466
processed with Fiji software (maximum Z-projection, contrast adjustment, Gaussian blur filter 467
r=0.5) (Schindelin et al., 2012). 468
469
Chromatin Endogenous Cleavage (ChEC) 470
20
Worm cultures were synchronized by embryo isolation with sodium hypochlorite for 4 471
generations. Embryos were treated with chitinase (1h, RT , 2 U/ml), washed twice in buffer A 472
(15 mM Tris pH 7.5, 0.1 mM EGTA, 0.34 M sucrose, 0.2 mM spermine, 0.5 mM spermidine, 0.5 473
mM PMSF) and then resuspend in buffer A supplemented with detergents (0.25% NP-40 and 474
0.1% Tx-100). Nuclei were released with a glass dounce homogenizer and 15 stokes using the 475
loose inserting pestle and 15 stokes using the tight inserting pestle. Debris were spun down at 476
100 g for 2 minutes and the supernatant, containing nuclei, was saved. The debris were 477
resuspended in buffer A supplemented with detergents and the nuclei isolation step was 478
repeated. The supernatants containing the nuclei were pooled, and the nuclei were pelleted at 479
1000 g for 10 min. Nuclei were washed once in buffer A, once in buffer B (15 mM Tris pH 7.5, 480
80 mM KCl, 0.1 mM EGTA, 0.2 mM spermine, 0.5 mM spermidine, 1 mM PMSF), and 481
resuspended in buffer B. MNase was activated by the addition of CaCl2 to a final concentration 482
of 2 mM at 30°C. Samples were incubated for 5 to 10 minutes (for CDT-1 ChEC), 5 to 7 minutes 483
(for TRES-1 ChEC) or for 1 h (free MNase control). For the control sample, 0.3 U/ml of MNase 484
(Biolabs, Cat. No. M0247S) was added at the same time as the CaCl2. MNase was inactivated 485
by adding an equal volume of 2x stop buffer (400 mM NaCl, 20 mM EGTA). Samples were 486
treated overnight with Proteinase K and SDS at 55°C. Phenol-chloroform-isoamyl treatment was 487
used to remove Proteinase K prior the RNase treatment (Roche, Cat. No. 11119915001) for 488
1h30 minutes at 37°C. After chloroform extraction, DNA was precipitated overnight with ethanol 489
and 0.25 M NaCl. DNA was resuspended overnight in TE prior to constructing the libraries using 490
NEBNext® Ultra™ II DNA Library Prep. Fifty base pair paired-end read sequencing reactions 491
were then performed on an Illumina Hi-Seq 2500 sequencer at the Genomics Platform of the 492
University of Geneva, except for TRES-1 ChEC-seq samples from 25°C, which were sequenced 493
with one hundred base pair single-end read sequencing reactions. 494
495
Embryonic cell extraction for EdU incorporation or DNA combing 496
21
Cells were extracted from embryos as described in (Bianchi and Driscoll, 2006). Briefly, 497
embryos were isolated using sodium hypochlorite, treated with chitinase (1h, RT, 2U/ml), 498
washed once in M9 and resuspend in 10ml of L-15 medium (supplemented with 10% FBS (heat 499
inactivated, Gibco), 41 mM sucrose and 1% pen/strep (1:100)). Cells were dissociated with 500
three passes through a syringe with a 22 gauge needle and two passes through a syringe with a 501
26 gauge needle. Debris were pelleted by spinning at 80 g for 1 min, and the supernatant, 502
containing cells, was saved. Debris were resuspended in supplemented L-15 medium and cell 503
extraction was repeated twice more. Supernatants were pooled together and filtered using 5 504
micron filters. Cells were pelleted at 500 g for 15 min and resuspended in supplemented L-15 505
medium. 506
507
EdU-seq 508
EdU-seq was carried out as described (Macheret and Halazonetis, 2019) with minor 509
modifications. Embryonic cells were synchronized by addition of 20 mM of HU (Sigma, Cat. No. 510
H8627) during 1h at 20°C or 25°C. Cells were washed twice with PBS to remove HU and 511
resuspended in L-15 medium. At desired time points, 25 µM of EdU and 20 mM of HU were 512
added for 10 minutes. Cells were washed twice with PBS, fixed with 90% methanol and stored 513
at -20°C. Cells were washed with ice-cold PBS and permeabilized with PBS containing 0.2% 514
Triton X-100 for 30 minutes at room temperature in the dark. EdU was coupled to a cleavable 515
biotin-azide linker (Azide-PEG(3+3)-S-S-biotin) (Jena Biosciences, Cat. No. CLK-A2112-10) 516
using the reagents of the Click-it Kit (Invitrogen, Cat. No. C-10424). Cells were washed with 517
PBS and treated overnight at 50°C with proteinase K in the lysis buffer. DNA was extracted by 518
phenol/chloroform/isoamyl extraction followed by chloroform extraction. DNA was precipitated 519
overnight at -20°C with presence of ethanol and 0.25 M NaCl and resuspended overnight in TE. 520
For an estimation of the time required to replicate the entire genome, cells were isolated, 521
synchronized using HU, and released in the presence of EdU as described above. Cell aliquots 522
22
were harvested between 0 and 130 min. Cell permeabilization and click-it reaction were done as 523
described above to couple Alexa Fluor™ 647 Azide (Thermofisher Cat. No. A10277) to the 524
EdU-labeled DNA. Cells were washed twice with PBS and treated with RNase (Roche, Cat. No. 525
11119915001) for 30 minutes at 37°C. PI (Sigma, Cat. No. 81845) was added, and samples 526
incubated overnight at room temperature in the dark. Levels of EdU incorporation were 527
assessed by flow cytometry (Gallios, Beckman Coulter). 528
529
Repli-seq 530
Asynchronous embryonic cells extracted from worms grown at 25°C were incubated for 5 531
minutes with 25 µM EdU (Invitrogen, Cat. No. A10044), washed twice with PBS and fixed with 532
90% methanol overnight. After fixation and click-it reaction as described in the EdU-seq section, 533
cells were washed twice with PBS and treated with RNase (Roche, Cat. No. 11119915001) for 534
30 minutes at 37°C. Propidium iodide (Sigma, Cat. No. 81845) was added, and samples were 535
incubated overnight at room temperature in the dark. Cells were sorted according to their DNA 536
content using a MoFlo Astrios flow sorter (Beckman Coulter) at the Flow Cytometry platform of 537
the Medical Faculty of the University of Geneva. DNA was extracted as described in the EdU-538
seq section. 539
540
Isolation and sequencing of EdU-labelled DNA from EdU-seq and Repli-seq 541
Five to eight µg of DNA were sonicated with a bioruptor sonicator (Diagenode) to obtain 542
fragments of 100-500 bp. EdU-labelled fragments were isolated using Dynabeads MyOne 543
Streptavidine C1 (Invitrogen, Cat. No. 65001) as previously described (Macheret and 544
Halazonetis, 2019). DNA was eluted by the addition of 2% of B-mercaptoethanol (Sigma, Cat. 545
No. M6250) and incubation for 1h at room temperature. The eluted DNA, as well as fragmented 546
total DNA, was used for library preparation using the TruSeq ChIP Sample Prep Kit (Illumina, 547
Cat. No. IP-202-1012). One hundred base pair single-end read sequencing reactions were 548
23
performed on an Illumina Hi-Seq 2500 sequencer at by the Genomics Platform of the University 549
of Geneva. 550
551
Data processing, domain annotation and peak calling 552
Sequencing reads were mapped to the C. elegans reference genome WBcel215 using 553
Novoalign software (default parameters, producing SAM format files). Reads were transformed 554
into 1’000 bins size (ChEC-seq and EdU-seq) or 10’000 bins size (Repli-seq) (JSON format) 555
with a custom script. Biological replicates that passed the quality control were merged (Repli-556
seq : WT early (2), WT mid (1), WT late (3), ∆ H3.3 early (2), ∆ H3.3 mid (2), ∆ H3.3 late (2), 557
ChEC-seq : CDT-1 20°C (5), TRES-1 20°C (4), TRES-1 25°C (3), TRES-1 25°C ∆ H3.3 (3), 558
EdU-seq : WT 0 min 20°C (4), all the other EdU-seq data (2)). A custom R script was used to 559
transform JSON files into bedgraphs. All files were normalized to the same number of reads. 560
Experimental samples were then normalized by subtracting the number of reads present in the 561
corresponding bin in the control file. The control samples were DNA from nuclei treated with 562
Free MNase (ChEC-seq), genomic DNA from cells treated with HU (EdU-seq), or genomic DNA 563
from cells without treatment (Repli-seq). Finally, the data were smoothed with a sliding window 564
averaging 5 consecutive bins using a custom R script. Genome Browser images were obtained 565
by using the Sushi Bioconductor package in R. 566
Domains of early and late replication were defined according to the signal enrichment present in 567
early and late Repli-seq in WT samples. Bins with positive signal in early and negative signal in 568
late datasets were defined as early domains, and vice versa for the ones with negative signal in 569
early and positive signal in late as late domain. The bins with either positive signal in early and 570
late repli-seq or negative signal in both datasets were not assigned to a domain. Domains 571
smaller than 5kb were merged with the previous domain. 572
24
For peak calling, normalized and smoothed data was converted to SGA files using the “ChIP-573
convert” tool in the Vital-it platform (https://ccg.epfl.ch//chipseq/). The output SGA files were 574
used as input for peak calling with the tool “ChIP-peak” of the Vital-it platform with these specific 575
parameters: Cut-off = 1000, Read counts = 10, Window = 3000, Vicinity = 5000, strand = any, 576
Repeat Masker = on, Refine peak position = off, Relative Enrichment 325 (CDT-1), 450 (TRES-577
1) or 350 (EdU). Peak lists from the ChEC-seq and EdU-seq experiments were merged, and 578
duplicate peaks removed. Peaks are considered duplicate if two datasets contains a peak within 579
the same 5kb window. To avoid sequencing artefacts, peaks that were also enriched in the 580
genomic DNA sample were removed. The final list of peaks was then clustered in 15 using 581
deepTools (Ramírez et al., 2016). Clusters with similar enrichment were merged together and 582
one cluster, containing signal only in the EdU-seq sample, was deleted, as the sites in this 583
cluster did not show fork movement after release from the HU block. Heatmaps were created 584
using deepTools (Ramírez et al., 2016). The genomic annotation of early, late and dormant 585
origins was determined using PAVIS website (https://manticore.niehs.nih.gov/pavis2/) (Huang et 586
al., 2013). Sequencing data has been deposited at the Gene Expression Omnibus (GEO) under 587
accession number GSE140804. 588
589
DNA combing and staining 590
DNA combing was performed as described in (Michalet et al., 1997). Embryonic cells were 591
incubated with 20 mM HU for 1h at 20°C or 25°C, washed twice with ice-cold PBS, and 592
incubated with 40 µM CldU for 20 minutes at the desired temperature. Cells were washed twice 593
with L-15 and then incubated in presence of 400 µM IdU for 20 minutes at the desired 594
temperature. Cells were washed with PBS and frozen at -20°C. Cells were embedded into 595
agarose plugs, and DNA was processed using the FiberPrep® DNA extraction kit (Genomic 596
Vision). The combing and antibody incubation was done as described in (Costantino et al., 597
2014). The DNA combing slides were imaged on a Leica DM5000B microscope, and the images 598
25
were analyzed with Fiji software (Schindelin et al., 2012). The analysis was carried out with data 599
from six independent experiments with a total of 49 measurements (WT, 20°C), 32 600
measurements (WT, 25°C), 28 measurements (∆ H3.3, 20°C) and 49 measurements (∆ H3.3, 601
25°C). 602
603
604
Acknowledgements 605
We thank Morgane Macheret and Thanos Halazonetis for sharing unpublished protocols, 606
Jonathan Mailler and Laura Padayachy for discussion and help with the DNA combing 607
experiments, Dario Menendez for help with strain generation, Caroline Gabus for reagent 608
preparation, Nicolas Roggli for help with data analysis and image processing, Florian Huber and 609
Ioannis Xenarios for help with the data analysis and Uli Laemmli and Gabriel Zentner for 610
reagents and advice when establishing the ChEC method. We thank all the Steiner lab 611
members for discussion and comments on the manuscript. We thank Thanos Halazonetis, 612
Damien Hermand, Jonathan Mailler, Laura Padayachy, Reinier Prosee, David Shore and 613
Maksym Shyian for comments on the manuscript. Some strains were provided by the CGC, 614
which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440). We are 615
grateful to the Genomics Platform of the iGE3, the Bioimaging Center of the Faculty of Sciences 616
and the Flow Cytometry platform of the Medical Faculty at the University of Geneva. The work 617
was funded by the Swiss National Science Foundation (Grants 31003A_156774 and 618
31003A_175606) and the Republic and Canton of Geneva. 619
620
621
Competing interests 622
The authors declare no competing interests. 623
26
624
625
Author contributions 626
FAS and MS conceived the study, MS carried out genetics, genomics and DNA combing 627
experiments, JMW carried out staining experiments, MS analyzed the data, MS and FAS 628
prepared the manuscript, FAS acquired funding. 629
630
631
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848
Figures and legends: 849
850
35
851
Figure 1. Replication timing is unchanged in absence of H3.3 (A) Schematic description of 852
Repli-seq. Embryonic cells were dissociated, exposed to an EdU pulse for 10 min and sorted 853
36
according to their DNA content. EdU-labelled DNA was sequenced and mapped to the genome. 854
(B) Representative genome browser views of Repli-seq and H3.3 ChIP-seq. Repli-seq signal is 855
shown for wild type (WT) and H3.3 null mutant (∆ H3.3) worms on regions of chromosomes I 856
and V. Early S phase is shown in black and late S phase in red. HIS-72 (H3.3) ChIP-seq signal 857
is shown in green for the same regions. ChIP-seq data from (Delaney et al., 2019). (C) Color-858
coded replication timing for each chromosome. Repli-seq signal from early (black), mid (orange) 859
and late (red) S phase for WT and ∆ H3.3. Data for each chromosome was sorted according to 860
the signal of early S phase in WT. (D) Gene expression levels for the genes present in domains 861
of early and late replication. RNA-seq data from (Kramer et al., 2015). (E) Genes sizes for the 862
genes present in domains of early and late replication. 863
864
37
865
Figure 2. Identification of replication origins in C. elegans embryos. (A) Schematic 866
description of the roles of CDT-1 and TRES-1 in replication origin firing. CDT-1 is required for 867
the licensing of all origins. TRES-1 is recruited only to origins that fire. (B) Localization of 868
38
HA::CDT-1 and FLAG::TRES-1 during the cell cycle. Immunofluorescence images using anti-HA 869
and anti-FLAG antibodies and DAPI are shown. Scale bar represents 5 µm. (C) Schematic 870
description of ChEC-seq. The protein of interest is fused with MNase. Upon activation with 871
calcium in purified nuclei, MNase cleaves and releases DNA fragments at the binding sites of 872
the fusion proteins. These small fragments are isolated and sequenced. (D) Representative 873
genome browser views of CDT-1 ChEC-seq (blue), TRES-1 ChEC-seq (violet) and EdU-seq 874
(pink) signal. Identified origins are highlighted in gray (licensed only) or dark and light green 875
(licensed and firing with and without EdU signal, respectively). Domains of late (red) and early 876
(gray) replication and positions of replication origins identified in previous studies are shown 877
below the genome browser. 878
879
39
880
Figure 3. Classification of replication origins in C. elegans embryos. (A) Heatmaps (top) 881
and average plots (bottom) of CDT-1 ChEC-seq (blue), TRES-1 ChEC-seq (violet) and EdU-seq 882
(pink) signal at replication origins. Origins were identified by peak calling using the CDT-1 883
ChEC-seq, TRES-1 ChEC-seq and EdU-seq datasets obtained from worms grown at 20°C and 884
separated into early, late and dormant origins through unsupervised clustering. Signal is shown 885
for a 50 kb window around each origin. (B) Heatmaps (top) and average plots (bottom) of HIS-886
72 (H3.3) ChIP-seq signal (Delaney et al., 2019) at replication origins, as in (A). 887
888
40
889
Figure 4. Replication origin dynamics is altered H3.3 null mutants. (A) Heatmaps (top) and 890
average plots (bottom) of TRES-1 ChEC-seq signal at replication origins, for wildtype (WT, 891
41
violet) and H3.3 null mutant (∆ H3.3, gray) worms grown at 25°C. Signal is shown for a 50 kb 892
window around each origin. (B, C) EdU-seq time course, for WT (pink) and ∆ H3.3 (gray) worms 893
grown at 25°C. (B) Genome browser views of EdU-seq signal at 0, 30 and 60 minutes, showing 894
fork progression at representative examples of early, late and dormant origins. (C) Heatmaps 895
(top) and average plots (bottom) of EdU-seq signal at 0, 30 and 60 minutes, at all origins. Signal 896
is shown for a 50 kb window around each origin. 897
898
42
899
Figure 5. Replication fork speed is not affected by loss of H3.3.. (A) Representative 900
examples of DNA combing images used to measure replication fork speed for wildtype (WT) 901
and H3.3 null mutant (∆ H3.3). IdU incorporation is shown in green and CldU incorporation in 902
magenta. Scale bar represents 10µm. (B) Fork speed determined by DNA combing at 20°C and 903
25°C for the WT and ∆ H3.3. 904
905
43
906
Figure 6. Loss of H3.3 results cell cycle delays and replication checkpoint activation. (A) 907
Determination of cell cycle length, from the onset of the cleavage furrow of the first cell division 908
(arrow) to nuclear envelope breakdown of the AB cell (star). Representative still images from a 909
movie of a developing H3.3 null mutant (∆ H3.3) embryo with and without depletion of chk-1 by 910
RNAi are shown. Scale bar represents 10µm. (B) Cell cycle timing for wild type (WT) and ∆ 911
H3.3 worms, with and without depletion of chk-1 by RNAi. (C) Embryonic lethality for WT and ∆ 912
H3.3, with and without depletion of chk-1 by RNAi. 913
914
44
915
Figure S1. Characteristics of early and late replicating domains. (A) Boundaries used to 916
sort embryonic cells according to their DNA content. Plot showing the number of cells counted 917
(cell counts) as a function of the amount of PI incorporation (PI-Area). Cells were sorted 918
according to their DNA content into early replication (gray), mid replication (yellow) and late 919
45
replication (red). (B) Color-coded replication timing for each chromosome. Repli-seq signal from 920
early (black), mid (orange) and late (red) S phase for wild type (WT) and H3.3 null mutant (∆ 921
H3.3) worms. Data is shown according to the chromosomal location. (C) Correlation plots 922
between Repli-seq data for early (left panel) or late (right panel) replication with HIS-72 ChIP-923
seq data (Delaney et al., 2019), in 10kb bins. The Spearman's rank correlation coefficient (r) is 924
shown. 925
926
46
927
Figure S2. Characteristics of replication origins. (A) FACS profiles from HU block 928
experiments for EdU-seq. Plots show distribution of EdU incorporation across cell populations in 929
absence of HU (left panel), upon HU block (center panel) and 30 minutes after release from the 930
HU block (right panel). (B) Classification of early, late and dormant origins found in this study 931
based on genomic location within (genic) or outside (non-genic) gene-coding regions. (C) Venn 932
diagrams representing the number of origins identified in this study that overlap with the origins 933
found in (Pourkarimi et al., 2016) (left) and in (Rodríguez-Martínez et al., 2017) (right). 934
935
47
936
Figure S3. Replication origin dynamics under non-stress conditions. EdU-seq time course, 937
for wild type (WT, pink) and H3.3 null mutant (∆ H3.3, gray) worms grown at 20°C. Heatmaps 938
(top) and average plots (bottom) of EdU-seq signal at 0, 30 and 60 minutes, at all origins. Signal 939
is shown for a 50 kb window around each origin. 940
941
48
942
Figure S4. Estimation of S-phase duration. Time course of EdU incorporation after HU 943
release at 20°C. Relative EdU incorporation is shown as a function of the time of incubation in 944
presence of EdU. The curve was obtained using a bell-shaped model with GraphPad Prism 945
version 8.0.0. 946
947
49
Table S1. Overview of C. elegans strains used in this study 948
949
Strain Genotype Source Description
N2 CGC Wild type
FAS43 his-69&his-70(uge44) III; his-72(tm2066) III; his-74(uge18) V; his-71(ok2289) X
Delaney et al., 2018
H3.3 null mutant
FAS50 cdt-1(uge34 [MNase::cdt-1]) I This study CDT-1 ChEC in WT background
FAS137 tres-1(uge102[MNase::tres-1]) I This study TRES-1 ChEC in WT background
FAS154 tres-1(uge104[MNase::tres-1]) I; his-69&his-70(uge44) III; his-72(tm2066) III; his-74(uge18) V; his-71(ok2289) X
This study TRES-1 ChEC in ∆H3.3 background
950