Thesis
Reference
Spatiotemporal characterization of DNA replication in the C. elegans embryo
STROBINO, Maude
Abstract
Histone H3.3 is a variant of histone H3. Here we show that loss of H3.3 results in replication defects in Caenorhabditis elegans embryos at elevated temperatures. To characterize these defects, we adapt methods to determine replication timing, map replication origins, and examine replication fork progression. Our analysis shows that the genome is replicated from early and late firing origins and is partitioned into domains of early and late replication. We find that under temperature stress conditions, additional replication origins become activated.
Moreover, loss of H3.3 results in altered replication fork progression around origins. These replication defects are accompanied by replication checkpoint activation, a delayed cell cycle, and increased lethality in checkpoint-compromised embryos. Our comprehensive analysis of DNA replication in C. elegans reveals the genomic location of replication origins and the dynamics of their firing, and uncovers a role of H3.3 in the regulation of replication origins under stress conditions.
STROBINO, Maude. Spatiotemporal characterization of DNA replication in the C.
elegans embryo. Thèse de doctorat : Univ. Genève, 2020, no. Sc. Vie 60
DOI : 10.13097/archive-ouverte/unige:140702 URN : urn:nbn:ch:unige-1407021
Available at:
http://archive-ouverte.unige.ch/unige:140702
Disclaimer: layout of this document may differ from the published version.
UNIVERSITÉ DE GENÈVE FACULTÉ DES SCIENCES Section de Biologie
Département de Biologie Moléculaire Professeur Florian A. Steiner
__________________________________________________________________
SPATIOTEMPORAL CHARACTERIZATION OF DNA REPLICATION IN THE C. ELEGANS EMBRYO
THÈSE
présentée aux Facultés de médecine et des sciences de l’Université de Genève pour obtenir le grade de Docteur ès sciences en sciences de la vie, mention
Biosciences moléculaires
par
Maude STROBINO
de Genève (GE)
Thèse No 60
GENÈVE Repromail
2020
SUMMARY 5
RÉSUMÉ 7
ACKNOWLEDGEMENTS 9
INTRODUCTION 10
C. ELEGANS AS A MODEL ORGANISM 10
DNA REPLICATION 13
DNA REPLICATION INITIATION 15
REPLICATION ORIGIN MAPPING 18
DNA DAMAGE AND CHECKPOINTS 26
REPLICATION IN TIME AND SPACE 28
HISTONES AND HISTONE VARIANTS 31
AIM OF THE THESIS 34
REFERENCES 35
RESULTS 43
METHODS FOR THE ANALYSIS OF DNA REPLICATION IN C. ELEGANS 43
REPLI-SEQ 43
CHROMATIN ENDOGENOUS CLEAVAGE (CHEC-SEQ) 45
EDU-SEQ TO MAP EARLY ORIGINS 50
EDU-SEQ TO FOLLOW REPLICATION FORK PROGRESSION 54
REFERENCES 55
LOSS OF HISTONE H3.3 RESULTS IN DNA REPLICATION DEFECTS AND ALTERED ORIGIN DYNAMICS IN C.
ELEGANS. 57
AUTHOR CONTRIBUTIONS 57
INTRODUCTION 59
RESULTS 63
DISCUSSION 78
METHODS 85
DATA ACCESS 93
ACKNOWLEDGEMENTS 93
DISCLOSURE DECLARATION 94
SUPPLEMENTARY MATERIALS: 94
REFERENCES 101
DISCUSSION 109
THE GENOME OF C. ELEGANS IS PARTITIONED IN EARLY AND LATE REPLICATING DOMAINS 109 COMBINING TWO METHODS TO ROBUSTLY MAP REPLICATION ORIGINS 111
LOSS OF H3.3 IMPACTS FORK PROGRESSION AROUND ORIGINS 114
LOSS OF H3.3 CAUSES REPLICATION STRESS IN THE C. ELEGANS GENOME 115
CONCLUSION AND FUTURE DIRECTIONS 116
REFERENCES 121
Summary
During each cell cycle, the genomic information is copied exactly once with high fidelity, through a process called DNA replication. This essential and highly regulated process arises during the S phase of the cell cycle and starts at very precise loci called replication origins. Recent years have brought a lot of progress in understanding the mechanism of DNA replication, but still no DNA motifs have been found to define origins in metazoans, and it is now broadly accepted that the chromatin context strongly influences origins definition. Chromatin context is mainly determined by the presence of histone modifications and histone variants, such as H3.3. Histone H3.3 is a variant of histone H3 involved in differentiation, fertility and development. This histone was found to be enriched around replication origins in Drosophila and Arabidopsis, raising the possibility of a role of this variant in origin definition. In contrast to other organisms, total loss of H3.3 in C. elegans results only in subtle phenotypes under stress conditions offering the possibility to study the role of H3.3 and its implication in origins definition in vivo.
The aim of my PhD thesis was to determine the role of H3.3 in origin definition and replication fork progression in C. elegans.
In the first part of my thesis, I adapted molecular methods to C. elegans to determine replication timing, map replication origins, and follow DNA replication fork progression.
In the second part of my thesis, I used these new tools to analyze the spatiotemporal regulation of DNA replication in wild-type C. elegans in comparison with mutants lacking H3.3.
I showed that despite the very rapid embryonic cell cycle, the genome is partitioned into domains of early and late replication. Early replication is correlated with high gene expression and the incorporation of the histone variant H3.3. I found that the genome-wide replication timing and origin firing is unchanged in H3.3 null mutants.
However, I observed an altered replication fork progression from origins upon loss of H3.3.
The results presented in my thesis reveal the genomic location of replication origins and the dynamics of their firing in C. elegans. They also uncover a role of H3.3 in the regulation of fork progression under stress conditions.
Résumé
Durant chaque cycle cellulaire, l’information génétique est copiée avec une haute fidélité exactement une fois à l’aide d’un processus appelé réplication de l’ADN. Ce processus, essentiel et hautement régulé, a lieu durant la phase S du cycle cellulaire et est initié à des endroits très spécifiques nommés origines de réplication. Ces dernières années, de nombreux progrès ont été faits dans la compréhension de ce mécanisme mais aucun motif dans l’ADN n’a été trouvé pour définir les origines chez les metazoaires et il est maintenant communément accepté que la conformation de la chromatine influence fortement la position des origines. La conformation de la chromatine est définie par la présence de modifications des histones et de variants d’histones, tels que H3.3. L’histone H3.3 est un variant de l’histone H3 important pour le développement, la différenciation cellulaire et la fertilité. Cet histone est enrichi autours des origines de réplication chez Drosophila et Arabidopsis, soulevant l’hypothèse d’un rôle de ce variant dans la détermination des origines. Au contraire des autres organismes, la perte totale de H3.3 chez C. elegans résulte en des phénotypes subtiles n’ayant lieu que dans des conditions de stress, permettant ainsi d’étudier le rôle possible de H3.3 et son implication dans la définition des origines in vivo.
Le but de ma thèse était de déterminer le rôle de H3.3 dans la définition des origines et la progression de la fourche de réplication chez C. elegans.
Dans la première partie de ma thèse, j’ai adapté différentes techniques moléculaires à C. elegans afin d’analyser la régulation temporelle de la réplication, cartographier les origines de réplication et suivre la progression de la fourche de réplication.
Dans la deuxième partie de ma thèse, j’ai utilisé ces nouveaux outils pour analyser la régulation spatio-temporelle de la réplication de l’ADN chez un vers de type sauvage en comparaison avec un vers ne possédant plus de H3.3.
J’ai montré que malgré le cycle cellulaire très rapide de l’embryon, le génome est tout de même séparé en domaines de réplication précoce et tardive.
Les domaines de réplication précoce sont corrélés avec la présence de gènes hautement exprimés et l’incorporation du variant d’histone H3.3. J’ai également montré que la temporalité de la réplication, lorsqu’elle est observée sur l’ensemble du génome, reste inchangée avec la perte de H3.3. Cependant, J’ai observé une altération de la progression des fourches de réplication autours des origines en absence de ce variant d’histone.
Les résultats présentés dans ma thèse révèlent ainsi la localisation génomique des origines de réplication ainsi que la dynamique de leur activation chez C. elegans. Ils identifient également un rôle pour H3.3 dans la régulation de la progression de la fourche de réplication en condition de stress.
Acknowledgements
First of all, I would like to thank Prof. Florian Steiner for giving me the opportunity to work on this project and for his supervision and support during all these years. I am grateful to Prof. Thanos Halazonetis and Prof. Conrad Nieduszynski, for accepting to be the members of my thesis committee. I also would like to thanks the members of my TAC committee, Prof. Thanos Halazonetis and Prof. François Karch, for their help, suggestions and advices during my entire thesis.
Thanks to all the members of the lab, present and past, for all the great time that we had together. You were all always very supportive during difficult time and thanks to you we always had a great atmosphere within the lab. I also would like to thank all the members of the depatment for your friendship and support and a special thanks to the members of the Halazonetis lab for accepting me on your lab half of the time.
More particularly, thanks to Laura Padayachy, Morgane Macheret, Jonathan Mailler and Giacomo Rossetti for your help, your support and all the great time during conferences and during our meetings at “Le petit Raphael”.
Merci aussi à Véronica Pinna Rodriguez et Cécile Bulut. Votre bonne humeur et votre gentillesse, lors de mes visites au bout du couloir, ont illuminé nombreuses de mes journées tout au long de ma thèse. Un grand merci également à Nicolas Roggli pour ton aide avec les analyses et les figures.
Je voudrais finalement remercier ma famille, et en particulier ma Maman, Jade, Naïla et Naïm pour votre soutien, votre patience, votre amour et tous ces merveilleux moments au fil des années. Naïm tu es à mes côtés depuis le Bachelor, tu m’aides,
tu me pousses et me soutiens. Merci.
Introduction
C. elegans as a model organism
Nematodes have played an important role in biology discoveries long before the establishment of Caenorhabditis elegans as a model organism by Sydney Brenner in 1974 (Brenner, 1974). Antonie van Leeuwenhoek, the inventor of the microscope, already mentioned the presence of such small worms in old wine in 1676 (Haidar, 2016; Nigon and Félix, 2017). In 1900, C. elegans was named and described for the first time by Maupas (Maupas, 1900; Nigon and Félix, 2017). In the 1940s, the groups of Nigon and Dougherty started using nematodes as model organisms. They established culture conditions and investigated the sex determination of C. elegans (Nigon and Félix, 2017). Nevertheless, the study of C. elegans was not widespread until Sydney Brenner advertised and popularized it as a model organism to study development and the nervous system (Brenner, 1974). Through the work of Brenner and his co-workers, the use of C. elegans gained the attention of many disciplines of biology. In 1998, it became the first animal to have its entire genome sequenced.
The sequence revealed that the C. elegans genome encodes approximately 19’000 genes, and that about 60% to 80% of the genes have homologues in the human genome (C. elegans Sequencing Consortium, 1998; Kaletta and Hengartner, 2006).
A large part of the success of C. elegans as a model organism is owed to its simplicity. This small, free-living soil nematode is easy to culture, and its transparent body allows the observation of cellular processes in vivo. The C. elegans life cycle is rapid, taking three to four days for the fertilized embryo to develop into an adult worm that will in turn produce about 300 embryos. The development of C. elegans can be
subdivided in 7 different stages: the embryonic stage, four different larval stages and the adult stage. An alternative stage, called dauer, arises during development under suboptimal growth conditions, such as in absence of food or at high temperature.
The nematode stays at this particular stage until optimal conditions are restored, where it resumes development into L4 stage (Fig. 1). C. elegans development follows a stereotypic pattern of cell divisions called an invariant lineage, with each cell dividing into predetermined daughter cells. The wild-type adult hermaphrodites contain exactly 959 somatic cells. The majority of the somatic cell divisions take place during embryonic development, while the germline develops mainly during L4 and adult stages (Hall et al., 2017)
Figure 1. C. elegans life cycle. The life cycle starts at 0 minute with fertilization. ~ 40 minutes post-fertilization the first cleavage occurs. After 150 minutes, embryos are laid and continue their development ex utero until hatching. Hatching is followed by four different larval stages before the worm becomes an adult. The dauer stage occurs in periods of food starvation or with high temperatures. Modified from Altun and Hall, 2006.
C. elegans are normally self-fertilizing hermaphrodites (XX), but in the rare case of meiotic nondisjunction of chromosome X, males are produced. These specific features allow easy maintenance of a strain while also making it possible to cross two different strains (Antoshechkin and Sternberg, 2007). Furthermore, due to its small genome and its invariant lineage, C. elegans is a perfect organism to study developmental questions. Because of its short life cycle, this nematode is a convenient model for investigating protein function through deletion of specific genes. In the past, this was achieved by random mutagenesis, but since 2013, genome editing using the CRISPR/Cas9 system has emerged as the method of choice (Dickinson et al., 2013). Alternatively, RNAi, which was discovered in C.
elegans (Lee et al., 1993), is commonly used in this model organism to modulate levels of gene expression. RNAi relies on the supply of exogenous double-stranded RNA (dsRNA) that is processed into small interfering RNAs, which target and degrade RNAs with sequence homology. The dsRNA can be transmitted to worms by different methods: injection of in-vitro transcribed dsRNA into the animals (Fire et al., 1998), feeding the animals with bacteria producing dsRNA (Timmons et al., 2001), soaking the animals with a solution containing dsRNA (Tabara et al., 1998) and production of the dsRNA in vivo from a transgene (Tavernarakis et al., 2000;
Wang and Barr, 2005).
C. elegans is now used as a model organism in numerous fields of biology: drug discovery (Carretero et al., 2017), host-microbiota interactions (Zhang et al., 2017), immunity (Ermolaeva and Schumacher, 2014), neurobiology (Ghosh-Roy and Chisholm, 2010; Portman, 2007), aging (Olsen et al., 2006), etc. In the field of DNA replication, this animal has been investigated mainly using genetic tools (Korzelius et al., 2011; Ossareh-Nazari et al., 2016; Sonneville et al., 2012), but a molecular characterization of DNA replication is lacking. The aim of my thesis work was to develop and adapt molecular tools to investigate the DNA replication in C. elegans, while at the same time take advantage of the C. elegans tools to study the molecular basis of DNA replication.
DNA replication
During each cell cycle, the genomic information has to be copied exactly once with high fidelity, through a process called DNA replication. The phase of the cell cycle during which the DNA is replicated is defined as the phase S. Once the DNA is copied, the cell transitions to the G2 phase and prepares itself to divide. The division happens in phase M, or mitosis, where duplicated DNA is segregated equally into the two daughter cells. Finally, the two daughter cells increase their size in phase G1 until they start a new cycle by replicating their DNA (Fig. 2A).
The progression through the cell cycle needs to be tightly regulated to ensure that the cell division happens only after sufficient growth of the cell and only once the DNA has been faithfully replicated. Cyclin-dependent kinases (CDKs) play a key role in this regulation and act as checkpoint factors during the cell cycle (Hochegger et al., 2008). CDKs are present at a stable concentration level during the entire cell
cycle, but their activation depends on the binding of specifics cyclins present at variable concentration all along the cell cycle (Fig. 2B) (Hochegger et al., 2008).
Figure 2. Regulation of the cell cycle progression. (A) The cell cycle contains three major transitions: the entry of the S phase (G1-S), the entry of mitosis (G2-M) and the transition between the metaphase and the anaphase. The two first transitions are controlled by the activation of CDKs that phosphorylate distinct sets of substrates. The transition between metaphase and anaphase is regulated by the degradation of cyclins and also other proteins as securin (Rahal and Amon, 2008).
(B) The level of CDK activity is variable among the different phases. These differences in concentration are a major regulator of the progression of the cell cycle. Once CDK activity reaches a specific threshold, the cell moves on to the next phase in the cell cycle. Modified from Hochegger et al., 2008.
It is imperative that the cell replicates its genome with high accuracy exactly once during each cell cycle to transmit precise genomic information to the daughter cells.
DNA replication is therefore a highly regulated and controlled process. The proteins involved in DNA replication and in replication checkpoints are intensively investigated, as their missfunctionement could lead to genomic instability, which is an underlying cause for cancer.
DNA replication initiation
DNA replication is initiated at specific genomic loci called replication origins.
Replication initiation is composed of two non-overlapping steps: origin licensing and origin firing. Origin licensing occurs during G1 phase when the origin recognition complex (ORC) is recruited to replication origins. In budding yeast, origins are sequence-specified (also called ARS for autonomous replicating sequences) and are recognized by ORC (Bolon and Bielinsky, 2006). In most other eukaryotes, the origins do not seem to be defined by their DNA sequence, and it remains unclear how ORC recognizes its binding sites (Parker et al., 2017; Vashee et al., 2003). In metazoans, origins are mainly present in GCs rich regions, such as CpG island (Cayrou et al., 2011; Delgado, 1998). It has been proposed that the binding of ORC is mediated by the local chromatin structure (Gilbert, 2001; Vashee et al., 2003). The discovery that H4K20me2 is recognized by the bromo-adjacent homology (BAH) domain of human ORC1 argues in favor of this hypothesis (Kuo et al., 2012).
Nevertheless, this histone mark is not sufficient to define origin, as this is one of the most abundant histone modifications and marks a number of sites that greatly exceeds the number of replication origins (Prioleau and MacAlpine, 2016). Also, it has been shown that Drosophila lacking H4K20me2 are viable (McKay et al., 2015).
Some other histone modifications, such as H3K9ac, H3K4me3, H4K20me1, and the variant histones H2AZ and H3.3, are enriched around early origins (Paranjape and Calvi, 2016; Smith et al., 2016). These marks and histone variants are representative of active chromatin and are mainly found on active promoters, enhancers or transcriptionally active sites and could indicate a correlation between transcription and replication origins (Picard et al., 2014). These two processes, transcription and replication, need to be tightly coordinated, in order to avoid conflicts between the two
machineries. Transcription may affect the DNA replication in two opposite ways. On one hand, the presence of RNA synthesis may inhibit the licensing of origins nearby, and on the other hand, transcription may create an open chromatin configuration that is more permissive for DNA replication (Fragkos et al., 2015).
Once loaded to the origins, ORC recruits the minichromosome 2-7 (MCM) complex with the help of cell division cycle 6 (Cdc6) and cycle 10-dependent transcript 1 (Cdt1) (Bleichert, 2019). MCM is a hexameric complex with helicase activity that, once activated, unwinds the DNA in front of the replication fork. Two copies of the MCM complex are loaded in a head-to-head conformation at replication origins to allow the bi-directional movement of the replication fork (Miller et al., 2019). The loading of the two hexamers is tightly coordinated, with the loading of the second MCM being driven by the loading of the first one (Miller et al., 2019).
Origin licensing outside of G1 phase can lead to re-replication of parts of the genome, leading to DNA replication stress (Alexander and Orr-Weaver, 2016). To avoid re-replication, the licensing factors Cdt1 and Cdc6 need to be tightly regulated.
Cdc6 is recruited by ORC, and they together mediate the loading of MCM, which is in complex with Cdt1 (Coster et al., 2014; Ticau et al., 2015). After MCM loading at origins, first Cdc6 and subsequently Cdt1 are released from the DNA (Ticau et al., 2015). The release of Cdt1 is concomitant with the closing of MCM around DNA and ATP hydrolysis by its helicase domain (Ticau et al., 2017). To avoid reloading, once released, Cdt1 is degraded (Pozo and Cook, 2016). Its activity is also inhibited by Geminin, a protein expressed during S and G2 phases (Wohlschlegel et al., 2000).
The majority of Cdc6 is also degraded at the beginning of S phase and the remaining Cdc6 is exported out of the nucleus during S and G2 phases (Kalfalah et al., 2015).
The second step of replication initiation, origin firing, occurs during S phase when the level of activation of the CDKs starts to increase, thus the MCM helicase activity. The MCM double hexamer is remodeled into two active helicases, named CMG (Cdc45 - MCM - GINS) by CDKs and DDKs (Dbf4-dependent kinases). The loading of Cdc45 onto MCM depends on the association of two proteins, Treslin and TopBP1, regulated themself by CDKs (Boos et al., 2011; Kumagai and Dunphy, 2017;
Kumagai et al., 2011). As Treslin is not essential for the movement of the replication fork, this protein is released once the origin is activated (Kanemaki and Labib, 2006).
The number of origins that are licensed in each cell cycle exceeds the number of origins that actually fire (Cayrou et al., 2011; Fragkos et al., 2015). This observation led to the creation of three categories of origins, depending their firing efficiency:
constitutive - origins that fire in each cell at every cell cycle, flexible - origins that are used in some cells, and dormant - origins that are normally inactive but can fire under specific circumstances to prevent genome instability (Fig. 3) (Fragkos et al., 2015).
Figure 3. Three categories of origins. The constitutive origins (green circle) fire in each cell at every cell cycle. The flexible origins (blue circle) are used differently depending on the cell and the developmental stage. The dormant origins (star and red circle in the last panel) are activated under specific conditions such as DNA damage. Closed circles represent inactive origins and open circles active origins. Modified from Fragkos et al. 2015.
Replication origin mapping
As explained above, in metazoans, DNA replication origins are more likely not defined by an underlying DNA sequence, but by chromatin features that are not well understood. Therefore, the identification of replication origin location is challenging, and different methods have been developed to map them (Fig. 4).
Figure 4. Methods to map replication origins. Different methods have been developed to map replication origins. SNS-seq, OK-seq, Bubble-seq, ChIP-seq, EdUseq-HU and D-NAscent are discussed in this chapter. The cartoons show a replication bubble at a replication origin (black bar) bound by origin factors (gray circle), Okazaki fragments (gray dashed lines) and nascent strands (gray solid line). The elements used to map the origins in each method are highlighted in red.
Short nascent strand sequencing (SNS-seq)
The use of nascent strands to map replication origins was first developed in 1989 by Vassilev and Johnson and relies on the identification of short DNA molecules that are produced immediately after the initiation of DNA replication (Vassilev and Johnson, 1989). This method was further modified and adapted by different groups.
For example, in 2007, the group of Le Beau mapped replication origins in human genome-wide by hybridizing isolated short single-stranded DNA (nascent strands) to microarrays (Fig. 5) (Lucas et al., 2007). However, the use of a microarray
prevented them from detecting any signal within repetitive regions of the genome, and the result depended on the hybridization efficiency of the probes (Valenzuela, 2012).
Figure 5. Combining nascent strand isolation with microarrays to map replication origins. (A) Isolation of short single-strand DNA by a sucrose gradient. (B) Extraction of whole genomic DNA.
(C) Labelling of nascent strand and genomic DNA with different dyes and hybridization to a microarray. Active origins are defined as enriched in nascent strand samples in comparison to the whole genome. Ori, origin of replication; ssDNA, single stranded DNA. Modified from Lucas et al.
2007.
Later, this method was further improved by replacing microarrays with high throughput sequencing and the use of the lambda exonuclease, an enzyme that digests any broken DNA fragments, but not the nascent strands because they are
protected by a RNA-primed sequence (Bielinsky and Gerbi, 1998; Cayrou et al., 2012; Ganier et al., 2019; Prioleau and MacAlpine, 2016). The use of the lambda exonuclease increased the quality of the data by reducing background signal, but is prone to producing false positives because this nuclease has a sequence bias, e.g. it is inefficient in digesting G-quadruplexes (G4) (Foulk et al., 2015). Nascent strand sequencing is attractive because of its experimental simplicity, but enriches only for strong initiation events (Prioleau and MacAlpine, 2016).
Okazaki fragments sequencing (OK-seq)
The lagging strand of the DNA is replicated in a discontinuous manner, by ligating together small DNA segments called Okazaki fragments. These fragments are polymerized by DNA polymerase δ and joined by DNA ligase I. Okazaki fragments sequencing to map replication origins has been developed in S. cerevisiae in 2012 (McGuffee et al., 2013; Smith and Whitehouse, 2012). To enrich for Okazaki fragments, DNA ligase I is inactivated, and the small DNA fragments isolated and sequenced. OK-seq preserves the strand identity, allowing to distinguish between Okazaki fragments replicated at the Watson or Crick strand. The origins are detected as loci with transition in strandedness (McGuffee et al., 2013; Prioleau and MacAlpine, 2016). This method also has limitations, as it can not map weak initiation sites or resolve origins that are in close proximity to each other (Ganier et al., 2019).
Bubble trapping sequencing (bubble-seq)
In 2006, bubble trapping sequencing was introduced as a new method to detect replication origins (Mesner et al., 2006). This method takes advantage of the fact that
circular DNA can be separated from linear DNA by electrophoresis (Dean et al., 1973). The authors showed that DNA fragments containing origin have a circular bubble shape that can be isolated by gel electrophoresis (Mesner et al., 2006). They first mapped origins in Chinese Hamster Ovary (CHO) cells (Mesner et al., 2006), and later used this method to isolate fragments containing origins from a human cell line (GM06990) and identify their location in the human genome (Mesner et al., 2013). They also classified their origins according to the replication timing by comparing their origin map with data from another study that detected replication timing in the same cell line (Hansen et al., 2010). Bubble-seq method has the advantage of being able to detect origins with very low initiation frequency, but lacks precision in defining replication origin sites (Prioleau and MacAlpine, 2016).
Chromatin immunoprecipitation sequencing (ChIP-seq)
ChIP-seq is a common method used to map genomic binding sites of proteins. The chromatin is fragmented, and the protein of interest is targeted by a specific antibody which is linked to beads that allow immunoprecipitation of the protein-DNA complex.
The DNA bound by the protein is then isolated and sequenced. ChIP-seq has been used in many species for the mapping of replication origins e.g. yeast (Belsky et al., 2015; Wyrick et al., 2001), Drosophila (MacAlpine et al., 2010) and human (Dellino et al., 2013; Miotto et al., 2016). The majority of studies using the ChIP-seq approach for origin mapping determined the binding sites of subunits of ORC. For this method, the specificity of the antibodies used is critical to obtain reliable data. Furthermore, this method can be challenging for proteins that form insoluble complexes or bind to chromatin in a transient way.
DNA combing
DNA combing does not allow genome-wide mapping of replication origins, but is used to determine fork speed, distance between origins, or fork asymmetry, on single molecules of DNA (Bianco et al., 2012; Urban et al., 2015). In 1994, Bensimon et al. set up a new method called “molecular combing”, where DNA molecules are stretched on a silanized glass coverslips (Bensimon et al., 1994). This technique was then used to monitor origin firing and fork progression. For this purpose, cells are synchronized so they enter in S phase at the same time. At this moment, a first thymidine analogue (for example chloro-deoxyuridine (CldU)) is added for a specific time to label newly-synthesized DNA. This first thymidine analogue is then replaced by another one (for example iododeoxyuridine (IdU)) to detect the progression of the fork. DNA is purified in agarose plugs to avoid any DNA shearing, and stretched in silanized coverslips using a specific device, generating extension of 2 kb/µm. After denaturation, primary and secondary antibodies are used to visualize the thymidine analogues by fluorescence microscopy (Fig. 6) (Bianco et al., 2012). It is also possible to analyze origins at specific loci by combining DNA combing with fluorescence in situ hybridization (FISH) and using hybridization probe that is specific to a replication origin (Urban et al., 2015).
Figure 6. DNA combing. Cells are incubated with CldU and IdU and harvested. DNA is extracted in an agarose plug and combed on silanized coverslips. CldU and IdU are detected by immunofluorescence. Modified from Gali et al., 2019.
Isolation of nascent DNA in synchronized cell population (EdUseq-HU)
EdUseq-HU is a method to detect origins of replication by monitoring the incorporation of thymidine analogue around the origins. It is based on initiation site sequencing (Ini-seq), a method developed to map the initiation sites of DNA replication (Fig. 7) (Langley et al., 2016).
Figure 7. Ini-seq. Cells are synchronized in order to initiate DNA replication at the same time. By the use of thymidine analogues, small DNA fragments around origins are isolated and sequenced.
Modified from Langley et al., 2016.
This method was then improved by changing the synchronization method, the use of hydroxyurea (HU) to restrict the extent of thymidine analogue incorporation, and the replacement of antibodies with click-it chemistry in order to reduce the background.
Briefly, culture cells are synchronized using nocodazole and cells in mitosis are selected by a mitotic shake-off in order to have the entire population initiating DNA replication at the same time. The cells are then released from the nocodazole block and allowed to enter into S phase in presence of HU and 5-ethynyl-2′-deoxyuridine (EdU). HU inhibits the production of nucleotides and therefore limits fork progression.
EdU is an analogue of thymidine and is used to label nascent DNA. After DNA
isolation and sonication, click-it chemistry is performed to bind a cleavable biotin to the incorporated EdU, allowing the enrichment of nascent DNA (Macheret and Halazonetis, 2019). Due to these improvements, the method was re-named EdUseq- HU and is used to map early replication origins, and to detect the fork movement away from origins (Macheret and Halazonetis, 2018). A limitation of this method is that it cannot be applied for mapping mid, late or dormant origins but only early ones.
D-NAscent
All the methods presented so far (with the exception of DNA combing) map replication origins in populations of cells. The use of replication origins is thought to be variable in between cells, and for this reason it would be desirable to map replication origins in single cells. The Nieduszynski group recently analyzed genome replication on individual DNA molecules, introducing a method called D-NAscent, which is inspired by DNA combing but applied to a genomic scale (Müller et al., 2019). The strengths of this method relie on the fact that the sequencing technology is able to distinguish between natural bases and base analogues, combined with the ability to sequence long DNA molecules (> 160 kb) using the Oxford Nanopore Technologies’ MinION (Müller et al., 2019). This instrument detects changes in the electrical signal as a single molecule of DNA passes through a protein pore to determine the base sequence (Müller et al., 2019). It had previously been shown that this sequencing technologie is able to discriminate between methylated and unmodified bases (Rand et al., 2017; Simpson et al., 2017), and now the authors showed that it can also discriminate between thymidine and thymidine analogues such as BrdU (Müller et al., 2019). This allowed the identification of replication
Replication origin mapping in C. elegans
Investigation of DNA replication on a genomic scale has only recently been established in C. elegans. Two groups mapped replication origins using two different methods, Okazaki fragments sequencing (Pourkarimi et al., 2016) and nascent strand sequencing (Rodríguez-Martínez et al., 2017). The two studies came to contradictory conclusions about the number and genomic location of replication origins: Okazaki fragments sequencing identified about 2’000 origins which are defined prior to the broad onset of zygotic transcription and maintained during development, whereas nascent strand sequencing identified about 16’000 origins and a complete reorganization of origins after gastrulation.
DNA damage and checkpoints
Genomic sequence information has to be successfully transmitted from one cell generation to the next, without any mutation of the sequence. However, DNA is a reactive molecule with a high rate of chemical modifications when exposed to reactive oxygen species (ROS), ultraviolet radiation (UV), ionizing radiation (IR) etc.
Furthermore, although DNA is mainly replicated by high fidelity DNA polymerases such as δ and ε, other DNA polymerases such as α, β, telomerase etc. that are more prone to induce mistakes also contribute to DNA replication and DNA repair (Chatterjee and Walker, 2017). To avoid the propagation of mutations by copying altered DNA, replication initiation is inhibited when damage to the DNA is detected (Boos and Ferreira, 2019). The inhibition of DNA replication is mediated by checkpoint proteins that are regulated by CDKs and DDKs. For example, in the presence of DNA damage, Rad53 phosphorylates Treslin, making the binding of this
protein to Topbp1 impossible, thereby blocking origin firing (Boos and Ferreira, 2019;
Zegerman and Diffley, 2010).
Checkpoints were initially discovered in yeast (S. cerevisiae), where Weinert and Hartwell observed that after UV irradiation wild-type strain stops to proliferate, while strain lacking Rad9 continues to proliferate and then die. In the same experiment, they also discovered that the phenotype observed in the strain lacking Rad9 could be rescued by arresting the cell cycle for 4 hours after irradiation (Weinert and Hartwell, 1988). To confirm that this time was needed to repair DNA damage caused by UV, they removed repair proteins such as Rad52 or Rad18. Under these conditions, yeast cells remained sensitive to irradiation even when the cell cycle was blocked for 4 hours. They concluded that Rad9 is a protein that is able to detect DNA damage and was blocking the cell cycle to give time for other proteins to repair the DNA damage before progressing in the cell cycle (Weinert and Hartwell, 1988).
DNA damage can be detected by two pathways: ATM (Ataxia telangiectasia mutated) or ATR (Ataxia telangiectasia and Rad53 related). The ATM pathway is activated in presence of double strand breaks, where p53 and the checkpoint kinase 2 (CHK2) inhibit cell cycle progression in G1, S or G2 phases (Paull, 2015). In the ATR pathway, DNA replication stress and single-stranded DNA are sensed by the checkpoint protein kinase 1 (CHK1) that inhibits the cell cycle in S or G2 phases (Awasthi et al., 2015).
There are different sources of DNA damage, both exogenous and endogenous. For example, even though DNA is copied by high fidelity polymerases, some nucleotide substitutions, insertions or deletions are introduced with a frequency of >10-6 per cell per generation (Kunkel, 2009). In the next round of DNA replication, these mistakes become fixed mutations. Another source of endogenous DNA damage is ROS. They
are produced during cellular respiration and can give raise to oxidative DNA lesions that can lead to double-strand breaks (Chatterjee and Walker, 2017; Sedelnikova et al., 2010). Exogenous agents responsible for DNA damage are e.g. IR or UV, both causing DNA breaks.
Replication in time and space
It has been observed in different organisms that not all replication origins fire at the same time, but that the genome possesses a replication timing program, with some regions of the genome always being replicated before the others (Concia et al., 2018; Dileep and Gilbert, 2018; Hansen et al., 2010). How replication timing is regulated is not entirely understood. One hypothesis proposes that some factors required for origin firing are present in limiting amounts in each cell (Mantiero et al., 2011; Tanaka et al., 2011). Hence, not all origins can fire synchronously at the entry of the S phase, and origin firing needs to be spread along the entire S phase (Boos and Ferreira, 2019). Large inter-origin distance can cause DNA damage, because long gaps between origins may not be completely replicated during the time of the S phase (Boos and Ferreira, 2019; Mantiero et al., 2011; Tanaka et al., 2011). To avoid such damage, origins fire in clusters, creating domains of early, mid or late replication. These domains are replicated according to a “domino model” (Boos and Ferreira, 2019), where once a domain finishes to replicate, its neighbouring domain starts (Maya-Mendoza et al., 2010). Chromatin organization influences which regions of the genome are first accessed by the limiting initiation factors and determine the identity of replication domains (Boos and Ferreira, 2019). Euchromatin, which is transcriptionally active, is preferentially replicated in early S phase, whereas
later during S phase (Boos and Ferreira, 2019). The segments of the chromosomes that replicate at a similar time are called constant timing regions (CTR) and are separated by timing transition regions (TTR). CTR are composed of one or several replication domains (Fig.8) (Marchal et al., 2019).
Figure 8. Replication timing along the chromosome. Early and late constant timing regions (CTR) are delimited by timing transition regions (TTR). CTR are composed of several replication domains (RD) that fire at similar replication timings (RT). Modified from Marchal et al. 2019.
This separation of the genome into early, mid and late replicating domains is very robust, and until now, no chemical perturbation or gene mutation that changes this timing has been identified (Marchal et al., 2019). Replication timing is not limited to segments of DNA but also applies to entire chromosomes. It has been proposed that replication timing along chromosomes is regulated by long non-coding RNAs (lncRNAs) (Donley et al., 2015; Marchal et al., 2019).
Replication domains are reorganized during development, and it has been shown that there are fewer and larger CTR in differentiated cells compared to undifferentiated cells (Hiratani et al., 2008; Marchal et al., 2019).
As described above, the firing of origins is determined by the activity of CDKs and DDKs that oscillate during the cell cycle. This raises the question how replication
domains with different timing of replication form during S phase, when CDKs and DDKs are active similarly everywhere in the nucleus. Two models to explain this observation have been recently proposed (Boos and Ferreira, 2019). In the first model, the timing is directly linked to chromatin organization, where the firing factors have different accessibility to the chromatin depending the compaction of the DNA (Fig. 9A). This model fits with the observation that firing factors are present in limiting amounts, and more accessible chromatin will have a higher probability of attracting some of these factors. The second model proposes that origins present within the same cluster are regulatorily coupled. Through a yet to be defined mechanism, the origins present in the same cluster are either inhibited and kept inactive until a specific signal releases the inhibition, or activated, with the firing of an origin stimulating the firing of neighboring origins (Fig. 9B). These two models are not mutually exclusive, and could both be relevant for regulating the replication timing (Boos and Ferreira, 2019).
Figure 9. Two models for explaining replication timing. (A) Model 1. Replication timing is determined by chromatin organization. Chromatin organization regulates the accessibility to origins for the firing factors. (B) Model 2. Regulation of replication timing is coupled for origins in the same cluster. An unknown mechanism controls the firing of origins within a cluster either by inhibition or activation of origin firing. Modified from Boos and Ferreira, 2019.
As demonstrated in this section, DNA replication is an essential and highly regulated process. Recent years have brought a lot of progress in understanding the mechanism of DNA replication but a lot of questions remained. The regulation of the timing of origins firing is very precise and robust and more work is needed to understand the effect of a disruption of this timing. Furthermore, it is now broadly accepted that replication origins in higher eukaryotes are not defined by DNA sequence, but strongly influenced by the chromatin context. Chromatin context is mainly determined by the presence of histone modifications and histone variants, but we still lack understanding on their precise role.
Histones and histone variants
Chromatin structure is a major determinant of replication origins. Chromatin is organized into basic units called nucleosomes. These structures are composed of an octamer of histone proteins, two copies each of H2A, H2B, H3 and H4, wrapped by 147 bp of DNA (Fig. 10) (Luger et al., 1997).
Figure 10. The nucleosome: DNA wrapped around a histone octamer. A nucleosome is composed of 147 bp of DNA (brown and turquoise) and an octamer of histone proteins (blue: H3;
green: H4; yellow: H2A; red: H2B). Modified from Luger et al., 1997.
Histone proteins contain an α-helical histone-fold domain that is embedded in the nucleosome core and an N-terminal tail that is unstructured. The N-terminal tail is often subject to post-translational modifications (PTMs) such as acetylation, phosphorylation or methylation and plays an important role in the regulation of nucleosome function. The functional properties of nucleosomes can also be modified by the incorporation of histone variants such as H3.3, H2A.Z, CENP-A or H2A.X.
These histone variants have histone-fold domains that are very similar to those of canonical histones, and their tail sequences range from being almost identical to canonical histones (e.g. H3.3) to being completely different (e.g. CENP-A) (Shi et al., 2017; Xiong et al., 2016). H3.3, which is most relevant for the results presented in
this thesis, is a major variant of histone H3, and the two proteins differ only by few amino acids (Xiong et al., 2016). The substitution of few amino acids in H3.3 allow the recognition of this variant by the H3.3-specific histone chaperones HIRA and ATRX/DAXX that deposit H3.3 onto chromatin in a replication-independent manner (Delaney et al., 2018; Shi et al., 2017; Tagami et al., 2004). The differences between H3 and H3.3 are subtle, and it is therefore unlikely that incorporation of H3.3 changes the overall structure of the nucleosome. The presence of H3.3 may impact chromatin folding by creating a more open conformation (Chen et al., 2013; Shi et al., 2017). This assumption is supported by the observation that H3.3 is enriched at promoters and gene bodies (Henikoff and Smith, 2015). H3.3 is also deposited into transcriptionally inactive regions such as telomeres and centromeres (Shi et al., 2017). H3.3 is implicated in different processes, such as embryonic stem cell differentiation and epigenetic reprogramming (Banaszynski et al., 2013; Martire et al., 2019; Wen et al., 2014), neuronal plasticity (Maze et al., 2015) and centromere maintenance (Verreault and Rajan, 2012). A number of genome-wide studies have shown in different organisms, such as Drosophila melanogaster and Arabidopsis thaliana, that H3.3 is enriched around replication origins (Deal et al., 2010;
MacAlpine et al., 2010; Stroud et al., 2012). Furthermore, in chicken cells, a slower replication fork has been observed following the loss of H3.3 (Frey et al., 2014). H3.3 has also been associated with domains of early replication and sites of DNA repair in human cells (Adam et al., 2013; Clément et al., 2018) raising the possibility of a role of this variant in origin definition. For the majority of the organisms, mutation of the genes coding for H3.3 results in lethality or sterility (Hödl and Basler, 2009; Jang et al., 2015; Sakai et al., 2009; Tang et al., 2015). However, total loss of H3.3 in C.
elegans results only in subtle phenotypes under stress conditions (Delaney et al.,
2018). This particularity offered us the possibility to study the roles of H3.3 and its implication in origins definition in vivo.
Aim of the thesis
As explained in this chapter, DNA replication is an essential process that needs to be tightly regulated to avoid DNA damage and genomic instability. This process is studied since numerous years but the exact locations of these origins in metazoans and the relationship between replication and chromatin organization still needs to be investigated in detail. A number of studies have shown that the H3 variant histone H3.3 is linked to DNA replication, but little is known about its specific role. With the work described in this thesis, I aim to investigate the consequence of the loss of H3.3 for DNA replication. To study the role of H3.3 in DNA replication, we use C.
elegans, a model organism that allows to study these mechanisms in vivo. To be able to study the relationship between chromatin context and origins firing or fork progression we first adapted molecular methods to map the origins, follow the progression of the replication fork and study the timing of origins firing.
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