When does histone synthesis occur in relation to DNA replication?

When does histone synthesis occur in relation to DNA replication?

We are searching data for your request:

Forums and discussions:
Manuals and reference books:
Data from registers:
Wait the end of the search in all databases.
Upon completion, a link will appear to access the found materials.

Do histones have to be synthesized before DNA is replicated to allow the DNA to coil around histones?

Yes, they have to. But that is just half of the story.

The (canonical) histones which are used in DNA replication are synthesized at the beginning of the S phase, and subsequently transported into the nucleus. Studies have shown that newly synthesized DNA is immediately packed into nucleosomes. Thus, it is necessary that these structures are available prior to (or at least just in time with) the replication.

However, there are different models on how the histones are inserted into new DNA. One model assumes that old and new nucleosomes are both incorporated into newly synthesized DNA after the replication fork. Others assume a semi-conservative approach where the old histones are disassembled into their subunits and then mixed together with the newly synthesized ones. For example, there is a model that proposes that the parental H2A/H2B and the H3/H4 dimers disassociate, while another assumes a disassociation of the H3/H4 dimers as well.

Figure: Nucleosome synthesis models for DNA replication


a Angélique Galvani and Christophe Thiriet (2013). Replicating - DNA in the Refractory Chromatin Environment, The Mechanisms of DNA Replication, Dr. David Stuart (Ed.), InTech, DOI: 10.5772/52656. Available from:

In Molecular Biology of the Cell (Chapter 4), it is written that

The major histones are synthesized primarily during the S phase of the cell cycle and assembled into nucleosomes on the daughter DNA helices just behind the replication fork (see Figure 5-32). In contrast, most histone variants are synthesized throughout interphase. They are often inserted into already-formed chromatin, which requires a histone-exchange process catalyzed by the ATP-dependent chromatin remodeling complexes discussed previously

DNA Replication Steps and Process

DNA is the genetic material that defines every cell. Before a cell duplicates and is divided into new daughter cells through either mitosis or meiosis, biomolecules and organelles must be copied to be distributed among the cells. DNA, found within the nucleus, must be replicated in order to ensure that each new cell receives the correct number of chromosomes. The process of DNA duplication is called DNA replication. Replication follows several steps that involve multiple proteins called replication enzymes and RNA. In eukaryotic cells, such as animal cells and plant cells, DNA replication occurs in the S phase of interphase during the cell cycle. The process of DNA replication is vital for cell growth, repair, and reproduction in organisms.

Key Takeaways

  • Deoxyribonucleic acid, commonly known as DNA, is a nucleic acid that has three main components: a deoxyribose sugar, a phosphate, and a nitrogenous base.
  • Since DNA contains the genetic material for an organism, it is important that it be copied when a cell divides into daughter cells. The process that copies DNA is called replication.
  • Replication involves the production of identical helices of DNA from one double-stranded molecule of DNA.
  • Enzymes are vital to DNA replication since they catalyze very important steps in the process.
  • The overall DNA replication process is extremely important for both cell growth and reproduction in organisms. It is also vital in the cell repair process.

DNA Metabolism: Synthesis, Replication and Degradation

a. The initiation of DNA synthesis requires priming by a short length of RNA (10-200 nucleotides long).

b. This priming process involves the nucleophilic attack by the 3′-hydroxyl group of the RNA primer on the α-phosphate of the deoxynucleoside triphosphate with splitting off pyrophosphate.

c. The 3′-hydroxyl group of the recently at­tached deoxyribonucleoside monophos­phate is then free to carry out a nucleophilic attack on the next entering deoxyribonucleoside triphosphate on its α-phosphate moiety with the splitting off pyrophosphate.

d. The selection of proper deoxyribonucleotide is dependent upon proper pairing with the other strand (template) of DNA molecule.

e. The template dictates in which dNTP is complementary and holds this by hydro­gen bonding.

f. The polymerization of deoxyribonucleotides takes place by such a process in a discontinuous phase of about 100 nucleotides in length.

g. This newly synthesized DNA strand at­tached to RNA primer is called okazaki fragments.

h. When many okazaki fragments are gener­ated, the replication complex begins to remove the RNA primers by DNA polymer­ase I and the gaps left by their removal are filled up by proper base paired deoxynucleotide. The enzyme DNA ligase seal the fragments of newly synthesized DNA.

2. Replication of DNA:

a. Due to unwinding of double helix of DNA, each strand acts as a template for the formation of a new strand. This process is called replication.

b. Types of Replication:

(i) Conservative replication:

The paren­tal strand is never completely sepa­rated. So, after one round of replica­tion, one daughter duplex contains only parental strands and the other only daughter strands.

(ii) Semiconservative replication:

The process of unwinding of the double helical daughter molecules, each of which is composed of a parental strand and a newly synthesized strand formed from the complementary strand, called semiconservative rep­lication.

b. Process of Replication:

(i) Initiation of DNA replication:

(1) Replication begins at a specific ini­tiation point and this is a unique se­quence of bases called Ori.

(2) In Ori there are two series of short re­peats such as three repeats of a 13- base pair sequence and four repeats of a 9-base pair sequence.

(3) In the initiation process about 20 Dna A protein molecule each with a bound ATP, bind at the four repeats of 9-base pair sequence, DNA is wrapped around the complex (initial).

(4) With the help of ATP and histone like protein HU, the three 13-base pair re­peats are denatured to give open com­plex.

(5) With the help of Dna C protein ATP Dna B protein binds to the open com­plex to form prep riming complex. As a result, unwinding of DNA occurs and priming replication starts.

In the elongation proc­ess of replication two operations oc­cur such as leading strand synthesis and lagging strand synthesis.

(1) Leading strand synthesis:

(a) Leading strand synthesis begins with the synthesis of RNA primer (10 to 60 nucleotides) by Dna G protein at the replication origin.

(b) Deoxyribonucleotides are then added to this primer by DNA polymerase III (it keeps pace with the replication fork).

(c) SSB molecules help to stabilize the separated strand.

(d) Helicases separate the two DNA strands at the fork.

(e) Topoisomerase II (DNA gyrase) acts to relieve the stress generated by helicases.

(f) This leading strand is replicated in a continuous manner in the 5′ to 3′ direction.

(2) Lagging strand synthesis:

(a) Lagging strand is replicated discontinuously and must be ac­complished in short fragments (okazaki fragments) synthesized in the direction opposite to fork movement.

(b) Synthesis of okazaki fragments:

The multi-protein primo-some complex travels in the same direc­tion as the replication fork.

At intervals, Dna G protein syn­thesizes an RNA primer for a new okazaki fragment. The synthesis proceeds in the direction opposite to fork movement. Each primer is extended by DNA polymerase III.

(c) When the new okazaki fragment is complete, the RNA primer is re­moved by DNA polymerase I. The remaining nick (a phosphodiester bond broken to leave a free 3′ OH and 5′ phosphate) is sealed by DNA ligase.

Very little is known about this process, it is assumed that DNA topoisomerase IV appears to be necessary for final separation of the two completed circular DNA molecules.

3. Degradation of DNA:

DNA damage may be classified into 4 forms:

(a) Single Base Alteration:

The one base damage includes the hydration of the cytosine residue by ultraviolet irra­diation.

(b) Two Base Alteration:

The two base damage includes Thymine-thymine dimer formation via a cyclobutane moiety.

Chain breaks may be created by irradiation such as X-ray exposure.

Cross-linkage agents which link bases of opposite strands also induce 2 base alterations. Cross­links can also occur between the DNA molecule and histones.

4. The DNA Polymerase Complex:

a. The different DNA polymerase molecules engage in DNA replication.

These are in­volved in three important properties:

b. Chain elongation shows the rate at which polymerization occurs. Processivity is an expression of the number of nucleotides added to the nascent chain before the polymerase disengages from the template. The proof-reading function identifies copying errors and corrects them.

c. InE. Coli, polymerase 111 (Pol 111) func­tions at the replication fork. It catalyzes the highest rate of chain elongation and is the most processive. It is capable of po­lymerizing 0.5 Mb of DNA during one cycle on the leading strand. It is the prod­uct of Dna E gene in E. Coli.

d. Polymerase 11 (Pol 11) is mostly con­cerned with proof-reading and DNA repair.

e. Polymerase 1 (Pol 1) completes chain syn­thesis between okazaki fragments on the lagging strand.

f. In mammalian cells, the polymerase is ca­pable of polymerizing about 100 nucleotides per second. This reduced rate is the result of interference by nucleosomes.

5. Enzymes Repair Damaged DNA:

a. It has been concluded that surviving spe­cies have the mechanisms for repairing DNA damage caused as a result of replica­tion errors or environmental insults.

b. Replication depends on the specific pair­ing of nucleotide bases. Proper pairing is dependent upon the presence of favoured tautomer’s of the purine and pyrimidine nucleotides. The proper base pairing can be assured by monitoring the base pairing twice.

Such double monitoring occurs in both bacterial and mammalian systems. This double monitoring does not produce errors of mis-pairing due to the presence of the un-favoured tautomer’s.

c. Replication errors lead to the accumula­tion of mutations.

d. DNA damages are replaced by mismatch repair, base excision repair, nucleotide excision repair, and double strand break repair. The defective in one strand can be returned to its original form by relying on the complementary information stored in the unaffected strand.

Some Repair Enzymes are Multifunctional:

i. Some repair enzymes are found as compo­nents of the large TF11H complex that plays central role in gene transcription. Another component of TF11H is involved in cell cycle regulation. Some repair enzymes are involved in gene rearrange­ment that normally occur.

ii. In patients with ataxia-telangiectasia, an autosomal recessive disease in humans, there exists an increased sensitivity to damage by x-ray. Patients with Fanconi’s anemia have defective repair of cross-link­ing damage.

iii. All these clinical syndromes are associ­ated with increased frequency of cancer.


Cellular DNA is constantly altered by endogenous and exogenous factors, resulting in tens of thousands of lesions in a human cell every day (Lindahl, 1993). This damage may be classified into two types according to size: non-bulky DNA and bulky DNA. Non-bulky DNA lesions include base mismatches, abasic sites, and small base modifications, which in general are repaired by mismatch repair (MMR), base excision repair (BER), nucleotide incision repair (NIR), direct reversal repair (DRR), and translesion DNA synthesis (TLS) (Gros et al., 2004 Fortini and Dogliotti, 2007 Sharma et al., 2013 Yi and He, 2013 Ignatov et al., 2017). Bulky DNA lesions include, among other types of damage: double-strand breaks, DNA-protein cross-links (DPCs), and intra- and inter-strand DNA cross-links. The structural complexity of certain bulky DNA lesions requires the use of several DNA repair pathways acting in a coordinated manner, including homologous recombination (HR), non-homologous DNA end-joining (NHEJ), nucleotide excision repair (NER), TLS and BER Fanconi anemia (FA) signaling system and complex proteolytic machinery (Ishchenko et al., 2006 Ho and Schärer, 2010 Duxin et al., 2014 Tretyakova et al., 2015 Martin et al., 2017). Non-bulky DNA lesions cause limited and local DNA perturbations, whereas bulky ones induce significant distortions in the overall DNA helix structure (Ide et al., 2011). DNA-protein cross-links (DPCs) are formed when a protein covalently binds to DNA (Tretyakova et al., 2015). They are difficult to repair because of their super-bulky character compared with known voluminous, helix-distorting DNA lesions, such as UV-induced pyrimidine dimers. These super-bulky adducts can be generated by exposure of cells to endogenous and exogenous cross-linking agents (Stingele et al., 2017 Zhang et al., 2020). The presence of protein covalently attached to DNA strongly interferes with DNA replication, transcription, repair, and chromatin remodeling (Kuo et al., 2007 Klages-Mundt and Li, 2017 Yudkina et al., 2018 Ji et al., 2019). DPCs may be classified into five types, according to the nature of the covalent link in the DNA-protein complex and the presence of DNA strand breaks (Ide et al., 2015, 2018 Nakano et al., 2017). Type 1, the most common type of DPC, is formed when proteins covalently link to a nitrogenous base in undisrupted DNA. Type 2-4 cross-links occur when DNA-cleaving enzymes are trapped in a covalent intermediate with a DNA strand (Ide et al., 2015, 2018 Nakano et al., 2017). Type 2 is formed when bi-functional DNA glycosylases and repair enzymes containing β-lyase activity such as DNA polymerase β and Parp1 irreversibly bind to a cleaved apurinic/apyrimidinic (AP) site (Ide et al., 2015, 2018 Nakano et al., 2017). Type 3 is generated during abortive DNA strand cleavage by topoisomerase 1 (Top1) and formation of a covalent tyrosinyl–phosphodiester bond between the protein and the 3′-terminal DNA phosphate moiety of SSB (Ide et al., 2015, 2018 Nakano et al., 2017). The abortive action of topoisomerase 2 (Top2) generates type 4 DPC, in which tyrosine is linked to the 5′-terminal phosphates of double-strand breaks (DSB) (Ide et al., 2015, 2018 Nakano et al., 2017). Recently, a new type of DPC emerged after the discovery of HMCES, a 5-hydroxymethylcytosine (5hmC) binding protein which can recognize abasic sites in single stranded DNA (ssDNA) and form a covalent ssDNA-HMCES crosslink to prevent error-prone translesion synthesis past the lesion (Mohni et al., 2019). Because of the differences in structure and composition between these five groups, each type of DPC is processed by a distinct repair mechanism. It seems difficult to remove super-bulky Type 1 DPC in the canonical linear DNA excision repair pathways because the presence of a protein molecule blocks access to DNA. Nevertheless, recent studies have revealed that nucleotide excision repair (NER) and homologous recombination (HR) can remove certain types of DPCs in a nuclease-dependent manner (Zhang et al., 2020). However, it is still not clear whether these repair pathways could deal with other types of DPC. Stingele et al. (2017) have proposed that each constituent of DPC: DNA, protein, and the covalent linkage between them might be processed by three different repair mechanisms. A recent paper by Kühbacher and Duxin (2020) provides comprehensive review on the formation and repair of DPCs. In this review, we summarize the current knowledge regarding the repair mechanisms involved in removal of DHCs induced by various genotoxic agents. Covalent cross-linking to DNA occurs more often with DNA binding proteins, such as histones, transcription factors, and DNA metabolizing enzymes including repair factors and topoisomerases (Klages-Mundt and Li, 2017). In the cell nucleus, histones are assembled into an octamer forming the nucleosome core with 147 bp of DNA wrapped around and tightly bound to it (Luger et al., 1997, 2012). This basic chromatin structure makes histones primary targets of DNA cross-linking agents, leading to the formation of DNA-histone cross-links (DHC) (Solomon and Varshavsky, 1985). Currently, the repair mechanisms counteracting DHCs generated by various factors only started to unravel.

DNA-Histone Cross-Links (DHCs)

Nucleosomal DNA is packaged into compact units referred as chromosomes, in which core nucleosome particles are connected by stretches of linker DNA up to 80 bp length. A nucleosome core particle (NCP) is composed of two copies each of histones H2A, H2B, H3, and H4. The molecular weight of individual histones range from 11 to 22 KDa, whereas the molecular weight of histone octamer in NCP is 210 KDa (Eickbush and Moudrianakis, 1978 Luger et al., 1997). The stability of the nucleosome is based on various protein-protein interactions, and numerous non-covalent electrostatic and hydrogen bonds between histones and the DNA duplex (Luger et al., 1997, 2012 Davey et al., 2002 Rohs et al., 2009). The primary structure of chromatin can be depicted as a beads-on-a-string organization of individual nucleosomes, which can be further folded into compact secondary and tertiary structures, with the help of histone variants present in certain nucleosomes and post-translational modifications (PTMs) situated in disordered histone tails (Woodcock and Dimitrov, 2001 Luger et al., 2012). The folding of chromatin into primary, secondary, and tertiary structures is crucial for regulating the accessibility of DNA to complex multi-protein machinery involved in DNA replication, transcription, and repair. Non-covalent interactions between DNA and histones enable chromatin dynamics to switch between the closed and open conformations. DHCs impair chromatin flexibility, which may subsequently affect long-distance interactions in chromatin that would indirectly disturb DNA replication, transcription, and repair within a topologically associating domain (TAD) (Hinz et al., 2010 Todd and Lippard, 2010 Tretyakova et al., 2015 Hauer and Gasser, 2017 Nakano et al., 2017). DHCs belong to type 1, a non-enzymatic form of DPC, in which a protein is covalently attached to an undisrupted DNA (Ide et al., 2011). Several comprehensive studies describing the mechanisms of formation of DHCs have been published recently (Ming et al., 2017 Shang et al., 2019 Yang and Greenberg, 2019), nevertheless, it is not known whether specific repair mechanisms for the removal of DHCs exist. In this review, we focus mainly on the repair pathways of DHCs and briefly describe their formation.

Formation of DHCs

A water-soluble covalent complex of DNA and histones (H2A and H2B) was first identified in a UV cross-linking assay (Smith, 1966 Sperling and Sperling, 1978). With this finding, it became evident that UV irradiation can induce DHCs in addition to well-known pyrimidine dimers. It was then discovered that exogenous and endogenous aldehydes could also form DHCs in cells (Lam et al., 1985 Kuykendall and Bogdanffy, 1992). More than 10% of amino acid residues in histones are lysines, whereas, aldehydes preferentially react with ε-amino groups of lysine side-chains with the formation of a Schiff base, which further reacts with exocyclic amino groups of guanine, adenine, and cytosine DNA bases, creating methylene linkage. Many cross-linking agents, such as chromate, metal ions, and cisplatin (cis-diaminedichloroplatinum-II), also induce DHCs in cells (Zhitkovich and Costa, 1992). Platinum compounds not only cause DNA-DNA cross-links but also covalently link DNA-protein complexes. In the case of histones (Figure 1A), these compounds cross-link ε-amino-groups of lysines and N 7 atoms of guanosines (Tretyakova et al., 2015 Ming et al., 2017). Cross-links between DNA and methionine residues were also observed in an X-ray structure of nucleosomes treated with platinum compounds (Wu et al., 2008). Exposure of purified nucleosome to bi-functional alkylating agents (e.g., nitrogen mustards) also cross-links histones to guanosines in DNA (Shang et al., 2019) however, these types of cross-links in cells are much less abundant than DNA cross-links with cysteines and histidines of non-histone proteins (Loeber et al., 2009).

Figure 1. Mechanisms of histone-DNA cross-links formation. (A) Reaction mechanisms of DNA cross-linking agents. (B) Direct cross-linking of histones to modified DNA bases. (C) Abasic site mediated cross-linking of histones to DNA. NCP-NH2: N-terminal amine (Lysine) of histones in the nucleosome core.

Histones can also directly react with 5-formylcytosine, a naturally occurring modified DNA base, and 8-oxoguanine, a major oxidative DNA damage product. Lysine amino groups react with 5-formylcytosine (Figure 1B), with the formation of a reversible Schiff base (Li et al., 2017 Raiber et al., 2018). The reaction of lysine side-chains with 8-oxoguanosine produced a stable protein cross-linked spiroiminodihydantoin (Sp) adduct (Xu et al., 2008).

Finally, the majority of DHCs are produced by a reaction between histone lysines and an aldehyde form of the 2′-deoxyribose at apurinic/apyrimidinic (AP) sites (Figure 1C) that are either directly formed upon damage or generated during excision of damaged bases in the base excision repair pathway (Solomon and Varshavsky, 1985 Sczepanski et al., 2010). The resulting Schiff base often undergoes strand-breaking ß-elimination, followed by a reversal of a histone-DNA cross-link. Since histone emerges unaltered from the reaction, the whole process is sometimes referred to as histone-catalyzed strand cleavage at AP sites (Ren et al., 2019). It should be noted that histone PTMs and the chromatin state could have a significant impact on DHC formation at abasic sites and with DNA bases (Sczepanski et al., 2010 Bowman and Poirier, 2015).

Mechanisms of Repair of DHC

Although DPCs, especially DHCs, often occur in cells and present a constant threat to genome stability, it is presumed that, except for tyrosyl-DNA phosphodiesterases, there is no specialized DNA repair pathway dedicated to meet these super-bulky challenges. Instead, the cell employs several distinct DNA repair and protein degradation mechanisms to target cross-linked DNA and protein/histone components in a given DPC/DHC. The covalently bound protein could be detected and degraded to a small peptide by cell proteolytic machinery, such as the specialized proteases SPRTN/Wss1, Ddi1, and GCNA1, or by proteasome, an ATP-dependent multi-subunit protease complex, whereas the damaged DNA component is detected and repaired in the NER, BER, HR, NHEJ, and FA pathways.

Proteasome-Dependent Proteolysis of Histones Cross-Linked to DNA

Proteasome-mediated proteolysis is the major pathway for the degradation of damaged proteins in a cell. A 26S proteasome consists of a cylindrical 20S core particle and one or two 19S regulatory particles (Ciechanover, 1998 Lecker et al., 2006). Although 20S core can bind to different regulatory particles, only the 19S particle confers the ability to degrade ubiquitylated proteins (Coux et al., 1996 Adams, 2004 Stadtmueller and Hill, 2011). Considering the vital role of the proteasome in the degradation of damaged protein, proteasome and ubiquitin involvement in the proteolysis of DHCs or DPCs remains a topic of debate. Inhibition of proteasome in Xenopus egg extracts did not stabilize the DPCs (Nakano et al., 2007 Duxin et al., 2014). However, many studies of the repair of DPCs in mammalian cells suggest proteasome participation (Adams, 2004 Baker et al., 2007 Zecevic et al., 2010 Larsen et al., 2019). Proteasome involvement in DHC removal surfaced for the first time in the research of Quievryn and Zhitkovich (2000), who discovered that proteasome inhibitors prevent the removal of DHCs and sensitize human cells to lower levels of formaldehyde. A study in Xenopus egg extracts found that DPCs are ubiquitylated by TRAIP E3 ubiquitin ligase and are subsequently degraded by the proteasome (Duxin et al., 2014 Larsen et al., 2019). However, an earlier study clearly demonstrated that DPCs are not marked with polyubiquitin chains, but are nevertheless subjected to proteasomal degradation by a mechanism that is not well understood (Nakano et al., 2009). The 26S proteasome can degrade purified non-ubiquitylated histones (Kisselev et al., 2006), raising the possibility of proteasomal degradation of non-ubiquitylated damaged histones in cells. A couple of studies have demonstrated that during replication stress induced by genotoxic agents, histones are hyperacetylated, and then specifically degraded in a ubiquitin-independent manner by a complex of 20S proteasome with PA200 proteasome activator, a distinct regulatory particle (Qian et al., 2013 Mandemaker et al., 2018). Although these studies have demonstrated that the ubiquitin-independent degradation of acetylated histones alleviates replication stress, the additional function of PA200-20S proteasome in DHC repair cannot be excluded. Moreover, PA200 was detected in nuclear speckles, and its role in DNA repair has been proposed (Ustrell et al., 2002). Thus, more detailed understanding of the role of proteasome in DHC repair requires further investigation.

The 20S proteasome is a hollow, barrel-shaped particle composed of 28 non-identical subunits arranged into four stacked rings. The active sites are sequestered inside an internal cavity separated from regulatory 19S and PA200 complexes by a gated channel. This 13Å channel is too narrow for a folded protein to enter (Löwe et al., 1995 Groll et al., 1997). For complete degradation of a DNA-cross-linked protein, the cross-linked DNA nucleotide itself would have to enter the proteolytic chamber, pulling a DNA strand inside. However, the DNA component of a DPC might be too bulky to enter the channel. Therefore, proteasome can remove only part of a cross-linked protein, converting DHC into a smaller DNA-peptide cross-link. Alternatively, traditional proteases, in which an active site is located in a cleft on the enzyme surface, could be involved in excision of the bulk of the non-cross-linked polypeptide chain, which can then be degraded by any of these proteases and by the proteasome.

Archaea usually have a single circular chromosome, the size of which may be as great as 5,751,492 base pairs in Methanosarcina acetivorans, which boasts the largest known archaean genome. One-tenth of this size is the tiny 490,885 base-pair genome of Nanoarchaeum equitans, which possesses the smallest archaean genome known it is estimated to contain only 537 protein-encoding genes. Smaller independent pieces of DNA, called plasmids, are also found in archaea. Plasmids may be transferred between cells by physical contact, in a process that may be similar to bacterial conjugation.

Figure: Archaea: Cluster of halobacterium (archaea)

Histone demethylases as emerging players in regulation of DNA replication and cell division

In addition to the central roles that histone lysine demethylases play in gene regulation, cell fate decisions and reprogramming, it has recently emerged that these enzymes are also involved in fundamental molecular processes that underpin DNA replication, cell cycle dynamics and cell division (Fig 3).

Figure 3. Histone demethylation is an integrated part of the cell cycle

Forming origins and DNA replication

The initiation of DNA replication and the copying of genetic information is a highly regulated and precisely controlled process. Establishing the correct chromatin environment is essential for proper formation of replication origins and replication itself 168 169 170 . Interestingly, recent studies have implicated histone demethylases in several aspects of DNA replication. For example, the H3K4me3 demethylase KDM5C appears to play an important role in forming origins and initiating replication at actively transcribed early-replicating genes 171 . This relies on an elevated expression of KDM5C during early S phase where it functions to actively remove H3K4me3 from replication origins, promoting the formation of the pre-initiation complex and driving occupancy of PCNA. In the absence of KDM5C, or its demethylase activity, H3K4me3 persists at these sites and early origins fail to efficiently initiate replication, leading to cell cycle arrest 171 172 . It is still unclear precisely how removal of H3K4me3 is involved in this process however, several proteins in the origin of replication complex are known to encode chromatin reader domains 173 . Perhaps components of the origin of replication complex or other replication-associated factors are responsive to the modification state of H3K4.

Once replication has been initiated, the process of ongoing replication is also regulated by the activity of histone demethylases. The levels of the H3K9me3 demethylase KDM4A/JMJD2A/JHDM3A are elevated at S phase, coincident with loss of H3K9me3 and an increase in H3K9me1/2 during replication 174 175 . H3K9me3 in chromatin is normally bound by the chromodomain-containing protein HP1γ, which contributes to the formation of condensed heterochromatic structures 176 . During S phase, KDM4A demethylase activity counteracts HP1γ binding at heterochromatic regions, creating more accessible chromatin required for passage of the DNA replication machinery 174 . This system appears to be tightly controlled through the cell cycle by regulating KDM4A protein levels which is required for accurate replication timing 83 174 . KDM4A-dependent effects on DNA replication are observed in mammalian cells and the model organism C. elegans, suggesting that this is an evolutionarily conserved function of the enzyme 174 . In keeping with a role for KDM4A activity in controlling DNA replication, overexpression of KDM4A leads to genomic instability in a demethylase-dependent manner, through driving re-replication and site-specific copy gain in genomic regions implicated in cancer 177 .

As we begin to understand more about the function of the histone demethylases, it seems likely that they have more widespread, conserved and even co-opted functions in the regulation of DNA replication. This is supported by observations in S. pombe demonstrating that KDM1A and KDM1B contribute to programmed replication fork pausing that promotes imprinting and mating-type switching 178 . Together, these observations suggest that there is likely an underappreciated role for histone demethylases in regulating the processes that initiate and regulate accurate replication of the genome.

Cell cycle transitions and organizing chromosomes

Control of cell cycle timing and dynamics is essential for proper cell division and recent work has demonstrated that histone demethylases play several distinct roles in controlling normal cell division (Fig 3) 10 . One specific way this is achieved is through their capacity to directly regulate the expression of genes required for normal cell cycle progression 72 81 179 180 181 182 . This is exemplified by the demethylase KDM7B, which binds to the promoters of several key cell cycle regulators, including E2F1 target genes, and is required for their transcriptional activation by removing the repressive H3K9me1/2 and H4K20me1 179 . In keeping with this role, KDM7B protein levels and its binding to chromatin are highly regulated during the cell cycle and this appears to play important roles in the G1/S and G2/M transitions 81 179 180 . Similarly, KDM1A positively regulates the expression of MAD2 and BUBR1, which are part of the mitotic checkpoint complex and are required for proper chromosome segregation during mitosis 181 . Transcriptional regulation by histone demethylases also ensures genomic stability during cell division 183 184 185 possibly by removing modifications associated with transcriptionally permissive chromatin states during mitosis 184 186 . For example, KDM8/JMJD5 is involved in repression of transcription at non-coding satellite repeat regions, possibly by removal of H3K36me2. In the absence of KDM8 activity, elevated H3K36me leads to defective spindle formation and causes abnormal cell division and genomic instability 184 . However, the mechanism by which KDM8 regulates H3K36me remains contentious as other studies failed to observe histone demethylase activity for KDM8 and, instead, suggest that KDM8 may act as a protein hydroxylase 187 188 189 .

Interestingly, during cell cycle transitions, histone demethylases can also function independently of their effects on gene transcription. As cells enter into prophase of mitosis, they need to deposit H4K20me1 on chromatin in order to load Condensin II, a structural protein complex required for chromosome condensation 179 190 . As chromatin-bound KDM7B would normally demethylate H4K20me1, its removal from chromatin is required to stabilize H4K20me1 and promote this transition. The cell achieves this through CDK1/cyclin B-dependant phosphorylation of KDM7B, which then leads to KDM7B dissociation from chromatin in prophase 179 . Although this dynamic engagement between KDM7B and chromatin is in fitting with its functions during the cell cycle, other histone demethylases appear to support normal chromosome segregation through alternative mechanisms. KDM4C/JMJD2C/JHDM3C remains associated with chromosomes throughout mitosis and is proposed to maintain low levels of H3K9me and regulate chromosome segregation 183 . However, deletion of KDM4C in mouse does not appear to overtly affect development, physiology or reproduction, suggesting that some of the effects observed in cell culture may not completely reflect an essential requirement in vivo 191 . Moving forward, a better understanding of how histone demethylases are involved in cell cycle progression and cell division in animals will be essential, given that misregulation of these enzymes appears to play roles in proliferation and cell division in cancer.

A point mutation is a single-letter swap – an exchange of two bases, adenine to cytosine, for example, at a single location in the DNA molecule. Since the sequence of letters in a gene determines the sequence of amino acids in the protein it encodes, a point mutation can change the amino acid sequence of the resulting protein. Sometimes a change in the protein's amino acid sequence can have dramatic results. For example, sickle cell disease occurs when a single-point mutation in the gene that encodes the hemoglobin molecule results in deformed red blood cells.

Sometimes, copying errors can insert or delete extra letters of the genetic code. Because these insertions and deletions, called indels, can make the protein produced by the gene much shorter or much longer, these errors can have a significant impact. Indels can have a dramatic effect on the protein's structure and function. Insertion or deletion of a single letter can sometimes cause a frameshift mutation, in which the entire amino acid sequence of the resulting protein is changed.


Abbas, T., Keaton, M. A. & Dutta, A. Genomic instability in cancer. Cold Spring Harb. Perspect. Biol. 5, a012914 (2013).

Flach, J. et al. Replication stress is a potent driver of functional decline in ageing haematopoietic stem cells. Nature 512, 198–202 (2014).

Marahrens, Y. & Stillman, B. A yeast chromosomal origin of DNA replication defined by multiple functional elements. Science 255, 817–823 (1992). This article defined the multipartite nature of a S. cerevisiae replication origin.

Leonard, A. C. & Méchali, M. DNA replication origins. Cold Spring Harb. Perspect. Biol. 5, a010116 (2013).

Cayrou, C. et al. Genome-scale analysis of metazoan replication origins reveals their organization in specific but flexible sites defined by conserved features. Genome Res. 21, 1438–1449 (2011).

Friedman, K. L., Brewer, B. J. & Fangman, W. L. Replication profile of Saccharomyces cerevisiae chromosome VI. Genes Cells 2, 667–678 (1997).

Heichinger, C., Penkett, C. J., Bahler, J. & Nurse, P. Genome-wide characterization of fission yeast DNA replication origins. EMBO J. 25, 5171–5179 (2006).

Méchali, M. Eukaryotic DNA replication origins: many choices for appropriate answers. Nature Rev. Mol. Cell Biol. 11, 728–738 (2010).

Yeeles, J. T., Deegan, T. D., Janska, A., Early, A. & Diffley, J. F. Regulated eukaryotic DNA replication origin firing with purified proteins. Nature 519, 431–435 (2015). The first in vitro reconstitution of regulated DNA replication from purified S. cerevisiae proteins.

Masai, H., Matsumoto, S., You, Z., Yoshizawa-Sugata, N. & Oda, M. Eukaryotic chromosome DNA replication: where, when, and how? Annu. Rev. Biochem. 79, 89–130 (2010).

Tanaka, S. & Diffley, J. F. Interdependent nuclear accumulation of budding yeast Cdt1 and Mcm2–7 during G1 phase. Nature Cell Biol. 4, 198–207 (2002).

Remus, D. et al. Concerted loading of Mcm2–7 double hexamers around DNA during DNA replication origin licensing. Cell 139, 719–730 (2009).

You, Z. & Masai, H. Cdt1 forms a complex with the minichromosome maintenance protein (MCM) and activates its helicase activity. J. Biol. Chem. 283, 24469–24477 (2008).

Maiorano, D., Moreau, J. & Méchali, M. XCDT1 is required for the assembly of pre-replicative complexes in Xenopus laevis. Nature 404, 622–625 (2000).

Maiorano, D., Rul, W. & Méchali, M. Cell cycle regulation of the licensing activity of Cdt1 in Xenopus laevis. Exp. Cell Res. 295, 138–149 (2004).

Blow, J. J. & Gillespie, P. J. Replication licensing and cancer — a fatal entanglement? Nature Rev. Cancer 8, 799–806 (2008).

Siddiqui, K., On, K. F. & Diffley, J. F. Regulating DNA replication in eukarya. Cold Spring Harb. Perspect. Biol. 5, a012930 (2013).

DePamphilis, M. L. Origins of DNA replication in metazoan chromosomes. J. Biol. Chem. 268, 1–4 (1993).

Taylor, J. H. Increase in DNA replication sites in cells held at the beginning of S phase. Chromosoma 62, 291–300 (1977).

Blumenthal, A. B., Kriegstein, H. J. & Hogness, D. S. The units of DNA replication in Drosophila melanogaster chromosomes. Cold Spring Harb. Symp. Quant. Biol. 38, 205–223 (1974).

Callan, H. G. DNA replication in the chromosomes of eukaryotes. Cold Spring Harb. Symp. Quant. Biol. 38, 195–203 (1974). References 20 and 21 were the first to describe the increased number of replication origins activated in early D. melanogaster and amphibian embryos.

Kang, S., Warner, M. D. & Bell, S. P. Multiple functions for Mcm2–7 ATPase motifs during replication initiation. Mol. Cell 55, 655–665 (2014).

Heller, R. C. et al. Eukaryotic origin-dependent DNA replication in vitro reveals sequential action of DDK and S-CDK kinases. Cell 146, 80–91 (2011).

Tanaka, S. et al. CDK-dependent phosphorylation of Sld2 and Sld3 initiates DNA replication in budding yeast. Nature 445, 328–332 (2007).

Jares, P. & Blow, J. J. Xenopus cdc7 function is dependent on licensing but not on XORC, XCdc6, or CDK activity and is required for XCdc45 loading. Genes Dev. 14, 1528–1540 (2000).

Masumoto, H., Muramatsu, S., Kamimura, Y. & Araki, H. S-Cdk-dependent phosphorylation of Sld2 essential for chromosomal DNA replication in budding yeast. Nature 415, 651–655 (2002).

Zegerman, P. & Diffley, J. F. Phosphorylation of Sld2 and Sld3 by cyclin-dependent kinases promotes DNA replication in budding yeast. Nature 445, 281–285 (2007). References 23–27 describe the phosphorylation events during the activation of replication origins.

Ilves, I., Petojevic, T., Pesavento, J. J. & Botchan, M. R. Activation of the MCM2–7 helicase by association with Cdc45 and GINS proteins. Mol. Cell 37, 247–258 (2010). This paper shows that the DNA helicase function of the MCM complex is activated by formation of a new complex.

On, K. F. et al. Prereplicative complexes assembled in vitro support origin-dependent and independent DNA replication. EMBO J. 33, 605–620 (2014).

Kumagai, A., Shevchenko, A. & Dunphy, W. G. Treslin collaborates with TopBP1 in triggering the initiation of DNA replication. Cell 140, 349–359 (2010).

Boos, D. et al. Regulation of DNA replication through Sld3–Dpb11 interaction is conserved from yeast to humans. Curr. Biol. 21, 1152–1157 (2011).

Kumagai, A., Shevchenko, A. & Dunphy, W. G. Direct regulation of treslin by cyclin-dependent kinase is essential for the onset of DNA replication. J. Cell Biol. 193, 995–1007 (2011).

Thu, Y. M. & Bielinsky, A. K. Enigmatic roles of Mcm10 in DNA replication. Trends Biochem. Sci. 38, 184–194 (2013).

Im, J. S. et al. Assembly of the Cdc45–Mcm2–7–GINS complex in human cells requires the Ctf4/And-1, RecQL4, and Mcm10 proteins. Proc. Natl Acad. Sci. USA 106, 15628–15632 (2009).

Gros, J., Devbhandari, S. & Remus, D. Origin plasticity during budding yeast DNA replication in vitro. EMBO J. 33, 621–636 (2014).

Delgado, S., Gomez, M., Bird, A. & Antequera, F. Initiation of DNA replication at CpG islands in mammalian chromosomes. EMBO J. 17, 2426–2435 (1998).

Cadoret, J. C. et al. Genome-wide studies highlight indirect links between human replication origins and gene regulation. Proc. Natl Acad. Sci. USA 105, 15837–15842 (2008).

Costas, C. et al. Genome-wide mapping of Arabidopsis thaliana origins of DNA replication and their associated epigenetic marks. Nature Struct. Mol. Biol. 18, 395–400 (2011).

Besnard, E. et al. Unraveling cell type-specific and reprogrammable human replication origin signatures associated with G-quadruplex consensus motifs. Nature Struct. Mol. Biol. 19, 837–844 (2012).

Smith, O. K. & Aladjem, M. I. Chromatin structure and replication origins: determinants of chromosome replication and nuclear organization. J. Mol. Biol. 426, 3330–3341 (2014).

Cayrou, C. et al. New insights into replication origin characteristics in metazoans. Cell Cycle 11, 658–667 (2012).

Valton, A. L. et al. G4 motifs affect origin positioning and efficiency in two vertebrate replicators. EMBO J. 33, 732–746 (2014).

Xu, J. et al. Genome-wide identification and characterization of replication origins by deep sequencing. Genome Biol. 13, R27 (2012).

Kohzaki, H. & Murakami, Y. Transcription factors and DNA replication origin selection. Bioessays 27, 1107–1116 (2005).

Goldman, M. A., Holmquist, G. P., Grey, M. C., Caston, L. A. & Nag, A. Replication timing of genes and middle repetitive sequences. Science 224, 686–692 (1984).

Martin, M. M. et al. Genome-wide depletion of replication initiation events in highly transcribed regions. Genome Res. 21, 1822–1832 (2011).

Sequeira-Mendes, J. et al. Transcription initiation activity sets replication origin efficiency in mammalian cells. PLoS Genet. 5, e1000446 (2009).

Lunyak, V. V., Ezrokhi, M., Smith, H. S. & Gerbi, S. A. Developmental changes in the sciara II/9A initiation zone for DNA replication. Mol. Cell. Biol. 22, 8426–8437 (2002).

Deaton, A. M. & Bird, A. CpG islands and the regulation of transcription. Genes Dev. 25, 1010–1022 (2011).

Danis, E. et al. Specification of a DNA replication origin by a transcription complex. Nature Cell Biol. 6, 721–730 (2004).

Knott, S. R. et al. Forkhead transcription factors establish origin timing and long-range clustering in S. cerevisiae. Cell 148, 99–111 (2012).

Belleli, R. et al. NCOA4 transcriptional coactivator inhibits activation of DNA replication origins. Mol. Cell 55, 123–137 (2014).

MacAlpine, H. K., Gordan, R., Powell, S. K., Hartemink, A. J. & MacAlpine, D. M. Drosophila ORC localizes to open chromatin and marks sites of cohesin complex loading. Genome Res. 20, 201–211 (2010).

Berbenetz, N. M., Nislow, C. & Brown, G. W. Diversity of eukaryotic DNA replication origins revealed by genome-wide analysis of chromatin structure. PLoS Genet. 6, e1001092 (2010).

Eaton, M. L., Galani, K., Kang, S., Bell, S. P. & MacAlpine, D. M. Conserved nucleosome positioning defines replication origins. Genes Dev. 24, 748–753 (2010). This study shows that the replication origin sequence is sufficient to establish a nucleosome-free region, but that ORC is necessary for precise nucleosome positioning at adjacent regions.

Lubelsky, Y. et al. Pre-replication complex proteins assemble at regions of low nucleosome occupancy within the Chinese hamster dihydrofolate reductase initiation zone. Nucleic Acids Res. 39, 3141–3155 (2011).

Givens, R. M. et al. Chromatin architectures at fission yeast transcriptional promoters and replication origins. Nucleic Acids Res. 40, 7176–7189 (2012).

Hizume, K., Yagura, M. & Araki, H. Concerted interaction between origin recognition complex (ORC), nucleosomes and replication origin DNA ensures stable ORC-origin binding. Genes Cells 18, 764–779 (2013).

Liu, J., McConnell, K., Dixon, M. & Calvi, B. R. Analysis of model replication origins in Drosophila reveals new aspects of the chromatin landscape and its relationship to origin activity and the prereplicative complex. Mol. Biol. Cell 23, 200–212 (2012).

Lombrana, R. et al. High-resolution analysis of DNA synthesis start sites and nucleosome architecture at efficient mammalian replication origins. EMBO J. 32, 2631–2644 (2013).

Iizuka, M., Matsui, T., Takisawa, H. & Smith, M. M. Regulation of replication licensing by acetyltransferase Hbo1. Mol. Cell. Biol. 26, 1098–1108 (2006).

Miotto, B. & Struhl, K. HBO1 histone acetylase activity is essential for DNA replication licensing and inhibited by geminin. Mol. Cell 37, 57–66 (2010).

Burke, T. W., Cook, J. G., Asano, M. & Nevins, J. R. Replication factors MCM2 and ORC1 interact with the histone acetyltransferase HBO1. J. Biol. Chem. 276, 15397–15408 (2001).

Tardat, M. et al. The histone H4 Lys 20 methyltransferase PR-Set7 regulates replication origins in mammalian cells. Nature Cell Biol. 12, 1086–1093 (2010).

Beck, D. B. et al. The role of PR-Set7 in replication licensing depends on Suv4–20h. Genes Dev. 26, 2580–2589 (2012).

Kuo, A. J. et al. The BAH domain of ORC1 links H4K20me2 to DNA replication licensing and Meier–Gorlin syndrome. Nature 484, 115–119 (2012).

Picard, F. et al. The spatiotemporal program of DNA replication is associated with specific combinations of chromatin marks in human cells. PLoS Genet. 10, e1004282 (2014).

Yu, Y. et al. Histone H3 lysine 56 methylation regulates DNA replication through its interaction with PCNA. Mol. Cell 46, 7–17 (2012).

Fu, H. et al. Methylation of histone H3 on lysine 79 associates with a group of replication origins and helps limit DNA replication once per cell cycle. PLoS Genet. 9, e1003542 (2013).

Stroud, H. et al. Genome-wide analysis of histone H3.1 and H3.3 variants in Arabidopsis thaliana. Proc. Natl Acad. Sci. USA 109, 5370–5375 (2012).

Pak, D. T. et al. Association of the origin recognition complex with heterochromatin and HP1 in higher eukaryotes. Cell 91, 311–323 (1997).

Prasanth, S. G., Shen, Z., Prasanth, K. V. & Stillman, B. Human origin recognition complex is essential for HP1 binding to chromatin and heterochromatin organization. Proc. Natl Acad. Sci. USA 107, 15093–15098 (2010). References 71 and 72 show the relationships between ORC and HP1.

Nagano, T. & Fraser, P. No-nonsense functions for long noncoding RNAs. Cell 145, 178–181 (2011).

Mohammad, M. M., Donti, T. R., Sebastian Yakisich, J., Smith, A. G. & Kapler, G. M. Tetrahymena ORC contains a ribosomal RNA fragment that participates in rDNA origin recognition. EMBO J. 26, 5048–5060 (2007).

Norseen, J. et al. RNA-dependent recruitment of the origin recognition complex. EMBO J. 27, 3024–3035 (2008).

Norseen, J., Johnson, F. B. & Lieberman, P. M. Role for G-quadruplex RNA binding by Epstein–Barr virus nuclear antigen 1 in DNA replication and metaphase chromosome attachment. J. Virol. 83, 10336–10346 (2009).

Hoshina, S. et al. Human origin recognition complex binds preferentially to G-quadruplex-preferable RNA and single-stranded DNA. J. Biol. Chem. 288, 30161–30171 (2013).

Christov, C. P., Gardiner, T. J., Szuts, D. & Krude, T. Functional requirement of noncoding Y RNAs for human chromosomal DNA replication. Mol. Cell. Biol. 26, 6993–7004 (2006).

Collart, C., Christov, C. P., Smith, J. C. & Krude, T. The midblastula transition defines the onset of Y RNA-dependent DNA replication in Xenopus laevis. Mol. Cell. Biol. 31, 3857–3870 (2011).

Newport, J. & Spann, T. Disassembly of the nucleus in mitotic extracts: membrane vesicularization, lamin disassembly, and chromosome condensation are independent processes. Cell 48, 219–230 (1987).

Sheehan, M. A., Mills, A. D., Sleeman, A. M., Laskey, R. A. & Blow, J. J. Steps in the assembly of replication-competent nuclei in a cell-free system from Xenopus eggs. J. Cell Biol. 106, 1–12 (1988).

Coue, M., Kearsey, S. E. & Méchali, M. Chromatin binding, nuclear localization and phosphorylation of Xenopus cdc21 are cell-cycle dependent and associated with the control of initiation of DNA replication. EMBO J. 15, 1085–1097 (1996).

Walter, J., Sun, L. & Newport, J. Regulated chromosomal DNA replication in the absence of a nucleus. Mol. Cell 1, 519–529 (1998).

Guelen, L. et al. Domain organization of human chromosomes revealed by mapping of nuclear lamina interactions. Nature 453, 948–951 (2008).

Goldberg, M., Jenkins, H., Allen, T., Whitfield, W. G. & Hutchison, C. J. Xenopus lamin B3 has a direct role in the assembly of a replication competent nucleus: evidence from cell-free egg extracts. J. Cell Sci. 108, 3451–3461 (1995).

Wilson, R. H. & Coverley, D. Relationship between DNA replication and the nuclear matrix. Genes Cells 18, 17–31 (2013).

Huberman, J. A. & Riggs, A. D. On the mechanism of DNA replication in mammalian chromosomes. J. Mol. Biol. 32, 327–341 (1968). This is a classical paper describing the replication unit clusters.

Nagano, T. et al. Single-cell Hi-C reveals cell-to-cell variability in chromosome structure. Nature 502, 59–64 (2013).

Brewer, B. J. & Fangman, W. L. Initiation at closely spaced replication origins in a yeast chromosome. Science 262, 1728–1731 (1993). This paper describes replication origin interference in S. cerevisiae.

Lebofsky, R., Heilig, R., Sonnleitner, M., Weissenbach, J. & Bensimon, A. DNA replication origin interference increases the spacing between initiation events in human cells. Mol. Biol. Cell 17, 5337–5345 (2006).

Marheineke, K. & Hyrien, O. Control of replication origin density and firing time in Xenopus egg extracts: role of a caffeine-sensitive, ATR-dependent checkpoint. J. Biol. Chem. 279, 28071–28081 (2004).

Buongiorno-Nardelli, M., Micheli, G., Carri, M. T. & Marilley, M. A relationship between replicon size and supercoiled loop domains in the eukaryotic genome. Nature 298, 100–102 (1982).

Lemaitre, J. M., Danis, E., Pasero, P., Vassetzky, Y. & Méchali, M. Mitotic remodeling of the replicon and chromosome structure. Cell 123, 1–15 (2005).

Courbet, S. et al. Replication fork movement sets chromatin loop size and origin choice in mammalian cells. Nature 455, 557–560 (2008).

Berezney, R., Dubey, D. D. & Huberman, J. A. Heterogeneity of eukaryotic replicons, replicon clusters, and replication foci. Chromosoma 108, 471–484 (2000).

Cseresnyes, Z., Schwarz, U. & Green, C. M. Analysis of replication factories in human cells by super-resolution light microscopy. BMC Cell Biol. 10, 88 (2009).

Cardoso, M. C., Schneider, K., Martin, R. M. & Leonhardt, H. Structure, function and dynamics of nuclear subcompartments. Curr. Opin. Cell Biol. 24, 79–85 (2012).

Takahashi, T. S., Yiu, P., Chou, M. F., Gygi, S. & Walter, J. C. Recruitment of Xenopus Scc2 and cohesin to chromatin requires the pre-replication complex. Nature Cell Biol. 6, 991–996 (2004).

Guillou, E. et al. Cohesin organizes chromatin loops at DNA replication factories. Genes Dev. 24, 2812–2822 (2010). References 53 and 99 report a link between cohesin recruitment and formation of the pre-RCs.

Jackson, D. A. & Pombo, A. Replicon clusters are stable units of chromosome structure: evidence that nuclear organization contributes to the efficient activation and propagation of S phase in human cells. J. Cell Biol. 140, 1285–1295 (1998). This is an extensive fluorescence microscopy study of replicon clusters and replication foci.

Moir, R. D., Montaglowy, M. & Goldman, R. D. Dynamic properties of nuclear lamins: lamin B is associated with sites of DNA replication. J. Cell Biol. 125, 1201–1212 (1994).

Moir, R. D., Spann, T. P., Herrmann, H. & Goldman, R. D. Disruption of nuclear lamin organization blocks the elongation phase of DNA replication. J. Cell Biol. 149, 1179–1192 (2000).

Lebofsky, R., van Oijen, A. M. & Walter, J. C. DNA is a co-factor for its own replication in Xenopus egg extracts. Nucleic Acids Res. 39, 545–555 (2011).

Cox, L. S. & Laskey, R. A. DNA replication occurs at discrete sites in pseudonuclei assembled from purified DNA in vitro. Cell 66, 271–275 (1991).

Rhind, N. & Gilbert, D. M. DNA replication timing. Cold Spring Harb. Perspect. Med. 3, 1–26 (2013).

Yoshida, K., Poveda, A. & Pasero, P. Time to be versatile: regulation of the replication timing program in budding yeast. J. Mol. Biol. 425, 4696–4705 (2013).

Eaton, M. L. et al. Chromatin signatures of the Drosophila replication program. Genome Res. 21, 164–174 (2011).

Dellino, G. I. et al. Genome-wide mapping of human DNA-replication origins: levels of transcription at ORC1 sites regulate origin selection and replication timing. Genome Res. 23, 1–11 (2013).

Letessier, A. et al. Cell-type-specific replication initiation programs set fragility of the FRA3B fragile site. Nature 470, 120–123 (2011).

Barlow, J. H. et al. Identification of early replicating fragile sites that contribute to genome instability. Cell 152, 620–632 (2013).

Hansen, R. S. et al. Sequencing newly replicated DNA reveals widespread plasticity in human replication timing. Proc. Natl Acad. Sci. USA 107, 139–144 (2010).

Ryba, T. et al. Evolutionarily conserved replication timing profiles predict long-range chromatin interactions and distinguish closely related cell types. Genome Res. 20, 761–770 (2010). This study reports the close relationship between replication domains and chromosome domains.

Dixon, J. R. et al. Topological domains in mammalian genomes identified by analysis of chromatin interactions. Nature 485, 376–380 (2012).

Pope, B. D. et al. Topologically associating domains are stable units of replication-timing regulation. Nature 515, 402–405 (2014).

Hayano, M. et al. Rif1 is a global regulator of timing of replication origin firing in fission yeast. Genes Dev. 26, 137–150 (2012).

Cornacchia, D. et al. Mouse Rif1 is a key regulator of the replication-timing programme in mammalian cells. EMBO J. 31, 3678–3690 (2012).

Yamazaki, S. et al. Rif1 regulates the replication timing domains on the human genome. EMBO J. 31, 3667–3677 (2012). References 115–117 are the first reports on the role of RIF1 in replication timing.

Silverman, J., Takai, H., Buonomo, S. B., Eisenhaber, F. & de Lange, T. Human Rif1, ortholog of a yeast telomeric protein, is regulated by ATM and 53BP1 and functions in the S-phase checkpoint. Genes Dev. 18, 2108–2119 (2004).

Buonomo, S. B., Wu, Y., Ferguson, D. & de Lange, T. Mammalian Rif1 contributes to replication stress survival and homology-directed repair. J. Cell Biol. 187, 385–398 (2009).

Hiraga, S. et al. Rif1 controls DNA replication by directing protein phosphatase 1 to reverse Cdc7-mediated phosphorylation of the MCM complex. Genes Dev. 28, 372–383 (2014).

Mattarocci, S. et al. Rif1 controls DNA replication timing in yeast through the PP1 phosphatase Glc7. Cell Rep. 7, 62–69 (2014).

Dave, A., Cooley, C., Garg, M. & Bianchi, A. Protein phosphatase 1 recruitment by Rif1 regulates DNA replication origin firing by counteracting DDK activity. Cell Rep. 7, 53–61 (2014).

Schwaiger, M., Kohler, H., Oakeley, E. J., Stadler, M. B. & Schubeler, D. Heterochromatin protein 1 (HP1) modulates replication timing of the Drosophila genome. Genome Res. 20, 771–780 (2010).

Hayashi, M. T., Takahashi, T. S., Nakagawa, T., Nakayama, J. & Masukata, H. The heterochromatin protein Swi6/HP1 activates replication origins at the pericentromeric region and silent mating-type locus. Nature Cell Biol. 11, 357–362 (2009).

Figueiredo, M. L., Philip, P., Stenberg, P. & Larsson, J. HP1a recruitment to promoters is independent of H3K9 methylation in Drosophila melanogaster. PLoS Genet. 8, e1003061 (2012).

Kim, S. M., Dubey, D. D. & Huberman, J. A. Early-replicating heterochromatin. Genes Dev. 17, 330–335 (2003).

Casas-Delucchi, C. S. et al. Histone hypoacetylation is required to maintain late replication timing of constitutive heterochromatin. Nucleic Acids Res. 40, 159–169 (2012).

Tazumi, A. et al. Telomere-binding protein Taz1 controls global replication timing through its localization near late replication origins in fission yeast. Genes Dev. 26, 2050–2062 (2012).

Cooper, J. P., Nimmo, E. R., Allshire, R. C. & Cech, T. R. Regulation of telomere length and function by a Myb-domain protein in fission yeast. Nature 385, 744–747 (1997).

Wu, P. Y. & Nurse, P. Establishing the program of origin firing during S phase in fission yeast. Cell 136, 852–864 (2009).

Wong, P. G. et al. Cdc45 limits replicon usage from a low density of preRCs in mammalian cells. PLoS ONE 6, e17533 (2011).

Patel, P. K. et al. The Hsk1(Cdc7) replication kinase regulates origin efficiency. Mol. Biol. Cell 19, 5550–5558 (2008).

Mantiero, D., Mackenzie, A., Donaldson, A. & Zegerman, P. Limiting replication initiation factors execute the temporal programme of origin firing in budding yeast. EMBO J. 30, 4805–4814 (2011).

Kwan, E. X. et al. A natural polymorphism in rDNA replication origins links origin activation with calorie restriction and lifespan. PLoS Genet. 9, e1003329 (2013).

Yoshida, K. et al. The histone deacetylases Sir2 and Rpd3 act on ribosomal DNA to control the replication program in budding yeast. Mol. Cell 54, 691–697 (2014).

Koren, A. et al. Genetic variation in human DNA replication timing. Cell 159, 1015–1026 (2014).

Hiratani, I. et al. Global reorganization of replication domains during embryonic stem cell differentiation. PLoS Biol. 6, e245 (2008).

Hiratani, I. et al. Genome-wide dynamics of replication timing revealed by in vitro models of mouse embryogenesis. Genome Res. 20, 155–169 (2010).

Hyrien, O., Maric, C. & Méchali, M. Transition in specification of embryonic metazoan DNA replication origins. Science 270, 994–997 (1995).

Sasaki, T., Sawado, T., Yamaguchi, M. & Shinomiya, T. Specification of regions of DNA replication initiation during embryogenesis in the 65-kilobase DNApolα-dE2F locus of Drosophila melanogaster. Mol. Cell. Biol. 19, 547–555 (1999).

Norio, P. et al. Progressive activation of DNA replication initiation in large domains of the immunoglobulin heavy chain locus during B cell development. Mol. Cell 20, 575–587 (2005).

Errico, A. & Costanzo, V. Mechanisms of replication fork protection: a safeguard for genome stability. Crit. Rev. Biochem. Mol. Biol. 47, 222–235 (2012).

Tercero, J. A. & Diffley, J. F. Regulation of DNA replication fork progression through damaged DNA by the Mec1/Rad53 checkpoint. Nature 412, 553–557 (2001).

Yekezare, M., Gomez-Gonzalez, B. & Diffley, J. F. Controlling DNA replication origins in response to DNA damage — inhibit globally, activate locally. J. Cell Sci. 126, 1297–1306 (2013).

McIntosh, D. & Blow, J. J. Dormant origins, the licensing checkpoint, and the response to replicative stresses. Cold Spring Harb. Perspect. Biol. 4, a012955 (2012).

Bartek, J., Lukas, C. & Lukas, J. Checking on DNA damage in S phase. Nature Rev. Mol. Cell Biol. 5, 792–804 (2004).

Donzelli, M. & Draetta, G. F. Regulating mammalian checkpoints through Cdc25 inactivation. EMBO Rep. 4, 671–677 (2003).

Takizawa, C. G. & Morgan, D. O. Control of mitosis by changes in the subcellular location of cyclin-B1–Cdk1 and Cdc25C. Curr. Opin. Cell Biol. 12, 658–665 (2000).

Karnani, N. & Dutta, A. The effect of the intra-S-phase checkpoint on origins of replication in human cells. Genes Dev. 25, 621–633 (2011).

Liu, H. et al. Phosphorylation of MLL by ATR is required for execution of mammalian S-phase checkpoint. Nature 467, 343–346 (2010).

Boos, D., Yekezare, M. & Diffley, J. F. Identification of a heteromeric complex that promotes DNA replication origin firing in human cells. Science 340, 981–984 (2013).

Lopez-Mosqueda, J. et al. Damage-induced phosphorylation of Sld3 is important to block late origin firing. Nature 467, 479–483 (2010).

Zegerman, P. & Diffley, J. F. Checkpoint-dependent inhibition of DNA replication initiation by Sld3 and Dbf4 phosphorylation. Nature 467, 474–478 (2010).

Anglana, M., Apiou, F., Bensimon, A. & Debatisse, M. Dynamics of DNA replication in mammalian somatic cells: nucleotide pool modulates origin choice and interorigin spacing. Cell 114, 385–394 (2003).

Syljuasen, R. G. et al. Inhibition of human Chk1 causes increased initiation of DNA replication, phosphorylation of ATR targets, and DNA breakage. Mol. Cell. Biol. 25, 3553–3562 (2005).

Allen, J. B., Zhou, Z., Siede, W., Friedberg, E. C. & Elledge, S. J. The SAD1/RAD53 protein kinase controls multiple checkpoints and DNA damage-induced transcription in yeast. Genes Dev. 8, 2401–2415 (1994).

Kato, R. & Ogawa, H. An essential gene, ESR1, is required for mitotic cell growth, DNA repair and meiotic recombination in Saccharomyces cerevisiae. Nucleic Acids Res. 22, 3104–3112 (1994).

Liu, Q. et al. Chk1 is an essential kinase that is regulated by Atr and required for the G2/M DNA damage checkpoint. Genes Dev. 14, 1448–1459 (2000).

Takai, H. et al. Aberrant cell cycle checkpoint function and early embryonic death in Chk1 −/− mice. Genes Dev. 14, 1439–1447 (2000).

Brown, E. J. & Baltimore, D. ATR disruption leads to chromosomal fragmentation and early embryonic lethality. Genes Dev. 14, 397–402 (2000).

Maya-Mendoza, A., Petermann, E., Gillespie, D. A., Caldecott, K. W. & Jackson, D. A. Chk1 regulates the density of active replication origins during the vertebrate S phase. EMBO J. 26, 2719–2731 (2007).

Zhao, H., Watkins, J. L. & Piwnica-Worms, H. Disruption of the checkpoint kinase 1/cell division cycle 25A pathway abrogates ionizing radiation-induced S and G2 checkpoints. Proc. Natl Acad. Sci. USA 99, 14795–14800 (2002).

Sorensen, C. S. et al. Chk1 regulates the S phase checkpoint by coupling the physiological turnover and ionizing radiation-induced accelerated proteolysis of Cdc25A. Cancer Cell 3, 247–258 (2003).

Maya-Mendoza, A., Olivares-Chauvet, P., Shaw, A. & Jackson, D. A. S phase progression in human cells is dictated by the genetic continuity of DNA foci. PLoS Genet. 6, e1000900 (2010).

Ge, X. Q. & Blow, J. J. Chk1 inhibits replication factory activation but allows dormant origin firing in existing factories. J. Cell Biol. 191, 1285–1297 (2010).

Vaziri, C. et al. A p53-dependent checkpoint pathway prevents rereplication. Mol. Cell 11, 997–1008 (2003).

Neelsen, K. J. et al. Deregulated origin licensing leads to chromosomal breaks by rereplication of a gapped DNA template. Genes Dev. 27, 2537–2542 (2013).

Melixetian, M. et al. Loss of geminin induces rereplication in the presence of functional p53. J. Cell Biol. 165, 473–482 (2004).

Zhu, W., Chen, Y. & Dutta, A. Rereplication by depletion of geminin is seen regardless of p53 status and activates a G2/M checkpoint. Mol. Cell. Biol. 24, 7140–7150 (2004).

Tada, S., Li, A., Maiorano, D., Méchali, M. & Blow, J. J. Repression of origin assembly in metaphase depends on inhibition of RLF-B/Cdt1 by geminin. Nature Cell Biol. 3, 107–113 (2001).

Wohlschlegel, J. A. et al. Inhibition of eukaryotic DNA replication by geminin binding to Cdt1. Science 290, 2309–2312 (2000).

Nguyen, V. Q., Co, C., Irie, K. & Li, J. J. Clb/Cdc28 kinases promote nuclear export of the replication initiator proteins Mcm2–7. Curr. Biol. 10, 195–205 (2000).

Saha, T., Ghosh, S., Vassilev, A. & DePamphilis, M. L. Ubiquitylation, phosphorylation and Orc2 modulate the subcellular location of Orc1 and prevent it from inducing apoptosis. J. Cell Sci. 119, 1371–1382 (2006).

Petersen, B. O., Lukas, J., Sorensen, C. S., Bartek, J. & Helin, K. Phosphorylation of mammalian CDC6 by cyclin A/CDK2 regulates its subcellular localization. EMBO J. 18, 396–410 (1999).

Coulombe, P., Gregoire, D., Tsanov, N. & Méchali, M. A spontaneous Cdt1 mutation in 129 mouse strains reveals a regulatory domain restraining replication licensing. Nature Commun. 4, 2065 (2013).

Sugimoto, N. et al. Cdt1 phosphorylation by cyclin A-dependent kinases negatively regulates its function without affecting geminin binding. J. Biol. Chem. 279, 19691–19697 (2004).

Mendez, J. et al. Human origin recognition complex large subunit is degraded by ubiquitin-mediated proteolysis after initiation of DNA replication. Mol. Cell 9, 481–491 (2002).

Li, X., Zhao, Q., Liao, R., Sun, P. & Wu, X. The SCF Skp2 ubiquitin ligase complex interacts with the human replication licensing factor Cdt1 and regulates Cdt1 degradation. J. Biol. Chem. 278, 30854–30858 (2003).

Higa, L. A., Mihaylov, I. S., Banks, D. P., Zheng, J. & Zhang, H. Radiation-mediated proteolysis of CDT1 by CUL4–ROC1 and CSN complexes constitutes a new checkpoint. Nature Cell Biol. 5, 1008–1015 (2003).

Arias, E. E. & Walter, J. C. PCNA functions as a molecular platform to trigger Cdt1 destruction and prevent re-replication. Nature Cell Biol. 8, 84–90 (2006).

McGarry, T. J. & Kirschner, M. W. Geminin, an inhibitor of DNA replication, is degraded during mitosis. Cell 93, 1043–1053 (1998).

Petersen, B. O. et al. Cell cycle- and cell growth-regulated proteolysis of mammalian CDC6 is dependent on APC–CDH1. Genes Dev. 14, 2330–2343 (2000).

Sugimoto, N. et al. Identification of novel human Cdt1-binding proteins by a proteomics approach: proteolytic regulation by APC/CCdh1. Mol. Biol. Cell 19, 1007–1021 (2008).

Liu, E. et al. The ATR-mediated S phase checkpoint prevents rereplication in mammalian cells when licensing control is disrupted. J. Cell Biol. 179, 643–657 (2007).

Zielke, N., Edgar, B. A. & DePamphilis, M. L. Endoreplication. Cold Spring Harb. Perspect. Biol. 5, a012948 (2013).

Kim, J. C. et al. Integrative analysis of gene amplification in Drosophila follicle cells: parameters of origin activation and repression. Genes Dev. 25, 1384–1398 (2011).

Aggarwal, B. D. & Calvi, B. R. Chromatin regulates origin activity in Drosophila follicle cells. Nature 430, 372–376 (2004).

Gonzalez, S. et al. Oncogenic activity of Cdc6 through repression of the INK4/ARF locus. Nature 440, 702–706 (2006).

Seo, J. et al. Cdt1 transgenic mice develop lymphoblastic lymphoma in the absence of p53. Oncogene 24, 8176–8186 (2005).

Zhu, W. & Depamphilis, M. L. Selective killing of cancer cells by suppression of geminin activity. Cancer Res. 69, 4870–4877 (2009).

Ge, X. Q., Jackson, D. A. & Blow, J. J. Dormant origins licensed by excess Mcm2–7 are required for human cells to survive replicative stress. Genes Dev. 21, 3331–3341 (2007). This study shows that during replication stress, the excess of the MCM complex can be used to activate DNA replication origins that are not used in a normal cell cycle.

Cayrou, C., Coulombe, P. & Méchali, M. Programming DNA replication origins and chromosome organization. Chromosome Res. 18, 137–145 (2010).

Conclusion and outlook

A decade on from the initial discovery of histone lysine demethylases, our understanding of how these fascinating enzymes function in cells has progressed at an immensely rapid pace. During this time, the emergence of genome-wide technologies has allowed us to examine the function of these enzymes on chromatin with unprecedented breadth and precision. This has provided a surprisingly detailed understanding of the fundamental roles that these enzymes play in controlling gene expression, cell fate decisions during development, and the reprogramming of chromatin states. Furthermore, new functions for histone demethylases as critical regulators of other important cellular processes, including DNA replication, cell cycle dynamics and the repair of DNA damage, have been identified that clearly warrant further investigation.

Perhaps not surprisingly given their discovery as histone demethylases, these enzymes and their cellular functions have been studied within the guise of histone demethylation. However, it is now increasingly clear that these proteins also catalyse other hydroxylation reactions that regulate both protein and nucleic acid based processes. A clear challenge for the future will be to understand the primary molecular determinants that underpin the phenotypes that result from perturbing demethylase enzymes. Does this rely on histone demethylase activity, protein demethylase activity or the hydroxylation of other cellular substrates? Alternatively, are these outcomes driven independently of enzymatic activity all together? Addressing these important questions, particularly within the context of developmental transitions where these proteins appear to be of central importance, will inevitably rely on the generation of new animal models, where specific activities can be disrupted to study and define the molecular principles that underpin the function of these fascinating proteins in normal biology and, ultimately, disease.


Tetrasome formation

Formation of the H3–H4 tetramer–DNA complex, known as the tetrasome, does not require ATP hydrolysis, but is instead driven by the high affinity of (H3–H4)2 tetramers for DNA in a mechanism guided by CAF-1. The finding that CAF-1 binds to a single H3–H4 dimer and cooperatively binds to DNA in a length-dependent manner suggests a mechanism for tetrasome formation. Results from two groups suggest that the deposition step requires the association of two CAF-1•H3–H4 complexes, that co-assemble on DNA, followed by the concerted deposition of one H3–H4 tetramer (Figure 3) ( 55, 58).

Model for CAF-1 mediated nucleosome assembly. In absence of histones, the C-terminal WHD is inaccessible due to sequestration by the ED domain. ASF1 transfers a single H3–H4 dimer to CAF-1, resulting in the liberation of the WHD. Two CAF-1•H3–H4 complexes associate in close proximity to each other, mediated by PCNA–CAF-1 and DNA–CAF-1 contacts. The transient association of two CAF-1•H3–H4 complexes on DNA allows for H3-H3 contacts to form and the two histone chaperone complexes concertedly deposit one (H3–H4)2 tetramer onto the DNA prior to being released from the DNA. During the second assembly step, H2A–H2B histone chaperones mediate H2A–H2B deposition onto the preexisting tetramer forming full nucleosomes. For details see text.

Model for CAF-1 mediated nucleosome assembly. In absence of histones, the C-terminal WHD is inaccessible due to sequestration by the ED domain. ASF1 transfers a single H3–H4 dimer to CAF-1, resulting in the liberation of the WHD. Two CAF-1•H3–H4 complexes associate in close proximity to each other, mediated by PCNA–CAF-1 and DNA–CAF-1 contacts. The transient association of two CAF-1•H3–H4 complexes on DNA allows for H3-H3 contacts to form and the two histone chaperone complexes concertedly deposit one (H3–H4)2 tetramer onto the DNA prior to being released from the DNA. During the second assembly step, H2A–H2B histone chaperones mediate H2A–H2B deposition onto the preexisting tetramer forming full nucleosomes. For details see text.

Using DNA fragments of varying lengths, the Luger lab was able to reveal intermediate steps in the mechanism of CAF-1-mediated H3–H4 deposition. In-solution cross-linking studies show that the association of two CAF-1•H3–H4 complexes is dependent on the DNA binding capacity of the large subunit ( 58). The WHD plays an important role in the mechanism that enables H3–H4 deposition. In the absence of the H3–H4 cargo, the positively charged DNA binding surface of the WHD engages the acidic ED domain, thus auto-inhibiting potential WHD–DNA interactions. Upon H3–H4 binding to the ED domain, this interaction is destabilized and the WHD becomes available for DNA engagement. Because the WHD binds DNA in a cooperative manner, this greatly enhances the subsequent association of two CAF-1•H3–H4 complexes, and mutations in the WHD that interfere with DNA binding abolish dimerization of CAF-1•H3–H4 complexes ( 58). The tetramerization of H3–H4 requires the interaction of H3 α3 helices. Notably, mutation of the H3 α3 tetramerization interface still allows H3–H4 interaction with CAF-1 or DNA but perturbs the concerted deposition of H3–H4 dimers and tetrasome assembly ( 55, 58). This finding, together with crosslinking experiments, suggests that the two α3 helices of the H3–H4 dimers are positioned in close proximity to each other prior to deposition ( 58). In the final step, CAF-1 releases the histones once a tetrasome has successfully formed.

Regulated dimerization of histone chaperones guides nucleosome assembly

Histone chaperone dimerization is likely a means to control the oligomerization state of H3–H4 itself: presumably, maintenance of H3–H4 as dimers represents a response to the need to control and restrict the histone tetramerization reaction during critical tasks. As DNA is being replicated, the assembly of chromatin impacts the speed at which the replication fork progresses ( 13, 98–100). Rapid deposition of new (H3–H4)2 tetramers is therefore critical to prevent replication fork stalling and genomic instability. The concerted DNA-mediated association of histone-bound CAF-1 ensures a timely and controlled mechanism and reduces the possibility for unproductive interactions.

Thermodynamically, the free energy associated with H3–H4 tetramerization onto DNA is the sum of partial reactions, which exhibit opposing energetic expenditures: overall, the DNA must be substantially deformed at a high energetic cost. This must be offset by favorable energy from the establishment of contacts between H3–H4 and DNA, formation of the H3-H3 four-helix bundle and the hydrophobic effect. The presence of all of these would be required to provide the entire assembly pathway with the necessary free energy to proceed in an ordered fashion. Much like an enzyme, CAF-1 promotes an optimal micro-environment, which allows formation of these contacts. In addition, histone tetramerization and concerted DNA deposition could establish directionality of the reaction and explain why histone chaperones like CAF-1 do not catalyze nucleosome disassembly reactions - the energetic cost of tetrasome splitting and DNA unwinding is simply too high and can only be accomplished with the help of ATP-dependent remodelers or helicases. In conclusion, the exploitation of the directed DNA binding energy of H3–H4 by histone chaperones supports the mechanism that renders them independent of ATP hydrolysis, while still providing a high degree of directionality for the H3–H4 deposition process.

Regulation of CAF-1 recruitment through post-translational modifications

Histone deposition by CAF-1 is likely regulated not only by other proteins but also by post-translational modifications on the histone chaperone as well as on the histones. While phosphorylation regulates recruitment of CAF-1 to the replication fork in a cell cycle dependent manner (see introduction) ( 40), posttranslational modifications of the histones have the potential to directly affect the deposition mechanism. In yeast, H3–H4 dimers that are directed towards incorporation at the replication fork are acetylated by the acetyltransferase Rtt109 on lysine 56 of H3 (H3K56 ac ) this modification serves as one of the marks for newly synthesized histones ( 101, 102). In contrast, acetylation of H3K56 in humans only appears to be required for nucleosome assembly associated with DNA repair, while like yeast, histones targeted to the replication fork are acetylated by the cytosolic HAT1 on H4K5 and H4K12 ( 77, 103–108) and by HAT4 on H4K91 ( 109). CAF-1 is thought to recognize the H3K56 ac modification, based on biochemical studies reporting that CAF-1 binds to H3K56 ac -H4 with higher affinity than to unmodified histones ( 101, 105) and XL-MS showing numerous crosslinks between CAF-1 and the H3 N-helix containing K56 ( 63). The N-terminal tails of both histones are not required for nucleosome assembly by CAF-1 even though they are essential for chromatin formation in a physiological context ( 70, 77, 110). Therefore, if and how CAF-1 specifically recognizes modifications on H4 is not clear and future work should address how histone modifications contribute mechanistically to CAF-1 function.

Mechanistic implications for propagation of chromatin structure

During DNA replication, (H3–H4)2 tetramers are propagated as intact units from the parental DNA randomly to one of the two daughter strands ( 19, 111). This finding raised questions about a possible copying mechanism to re-establish chromatin marks on newly deposited histones when the template histone is not located within the same nucleosome ( 112–115). In relation to that, the histone chaperones not only perform assembly of (H3–H4)2 tetramers onto the newly-synthesized DNA, but could also regulate the initial tetrasome formation in response to certain histone modifications. Candidate chaperones involved in recycling of such modified H3–H4 are MCM2 and ASF1, as outlined in the introduction. Thus, the recent advances outlined here are compatible with a model in which CAF-1 acts as an acceptor of modified or native parental H3–H4 dimers for histone recycling during replication. Because of the coordinated deposition mechanism, the biochemical properties of CAF-1 could help to ensure that simultaneously transferred histone dimers are rapidly reassembled on DNA. In such a case, CAF-1 could fulfill a dual role, namely de novo assembly as well as recycling of parental (H3–H4)2 tetramers.

When does histone synthesis occur in relation to DNA replication? - Biology

Elongation synthesizes pre-mRNA in a 5′ to 3′ direction, and termination occurs in response to termination sequences and signals.

Learning Objectives

Describe what is happening during transcription elongation and termination

Key Takeaways

Key Points

  • RNA polymerase II (RNAPII) transcribes the major share of eukaryotic genes.
  • During elongation, the transcription machinery needs to move histones out of the way every time it encounters a nucleosome.
  • Transcription elongation occurs in a bubble of unwound DNA, where the RNA Polymerase uses one strand of DNA as a template to catalyze the synthesis of a new RNA strand in the 5′ to 3′ direction.
  • RNA Polymerase I and RNA Polymerase III terminate transcription in response to specific termination sequences in either the DNA being transcribed (RNA Polymerase I) or in the newly-synthesized RNA (RNA Polymerase III).
  • RNA Polymerase II terminates transcription at random locations past the end of the gene being transcribed. The newly-synthesized RNA is cleaved at a sequence-specified location and released before transcription terminates.

Key Terms

  • nucleosome: any of the subunits that repeat in chromatin a coil of DNA surrounding a histone core
  • histone: any of various simple water-soluble proteins that are rich in the basic amino acids lysine and arginine and are complexed with DNA in the nucleosomes of eukaryotic chromatin
  • chromatin: a complex of DNA, RNA, and proteins within the cell nucleus out of which chromosomes condense during cell division

Transcription through Nucleosomes

Following the formation of the pre-initiation complex, the polymerase is released from the other transcription factors, and elongation is allowed to proceed with the polymerase synthesizing RNA in the 5′ to 3′ direction. RNA Polymerase II (RNAPII) transcribes the major share of eukaryotic genes, so this section will mainly focus on how this specific polymerase accomplishes elongation and termination.

Although the enzymatic process of elongation is essentially the same in eukaryotes and prokaryotes, the eukaryotic DNA template is more complex. When eukaryotic cells are not dividing, their genes exist as a diffuse, but still extensively packaged and compacted mass of DNA and proteins called chromatin. The DNA is tightly packaged around charged histone proteins at repeated intervals. These DNA–histone complexes, collectively called nucleosomes, are regularly spaced and include 146 nucleotides of DNA wound twice around the eight histones in a nucleosome like thread around a spool.

For polynucleotide synthesis to occur, the transcription machinery needs to move histones out of the way every time it encounters a nucleosome. This is accomplished by a special protein dimer called FACT, which stands for “facilitates chromatin transcription.” FACT partially disassembles the nucleosome immediately ahead (upstream) of a transcribing RNA Polymerase II by removing two of the eight histones (a single dimer of H2A and H2B histones is removed.) This presumably sufficiently loosens the DNA wrapped around that nucleosome so that RNA Polymerase II can transcribe through it. FACT reassembles the nucleosome behind the RNA Polymerase II by returning the missing histones to it. RNA Polymerase II will continue to elongate the newly-synthesized RNA until transcription terminates.

The FACT protein dimer allows RNA Polymerase II to transcribe through packaged DNA: DNA in eukaryotes is packaged in nucleosomes, which consist of an octomer of 4 different histone proteins. When DNA is tightly wound twice around a nucleosome, RNA Polymerase II cannot access it for transcription. FACT removes two of the histones from the nucleosome immediately ahead of RNA Polymerase, loosening the packaging so that RNA Polymerase II can continue transcription. FACT also reassembles the nucleosome immediately behindd the RNA Polymerase by returning the missing histones.


RNA Polymerase II is a complex of 12 protein subunits. Specific subunits within the protein allow RNA Polymerase II to act as its own helicase, sliding clamp, single-stranded DNA binding protein, as well as carry out other functions. Consequently, RNA Polymerase II does not need as many accessory proteins to catalyze the synthesis of new RNA strands during transcription elongation as DNA Polymerase does to catalyze the synthesis of new DNA strands during replication elongation.

However, RNA Polymerase II does need a large collection of accessory proteins to initiate transcription at gene promoters, but once the double-stranded DNA in the transcription start region has been unwound, the RNA Polymerase II has been positioned at the +1 initiation nucleotide, and has started catalyzing new RNA strand synthesis, RNA Polymerase II clears or “escapes” the promoter region and leaves most of the transcription initiation proteins behind.

All RNA Polymerases travel along the template DNA strand in the 3′ to 5′ direction and catalyze the synthesis of new RNA strands in the 5′ to 3′ direction, adding new nucleotides to the 3′ end of the growing RNA strand.

RNA Polymerases unwind the double stranded DNA ahead of them and allow the unwound DNA behind them to rewind. As a result, RNA strand synthesis occurs in a transcription bubble of about 25 unwound DNA basebairs. Only about 8 nucleotides of newly-synthesized RNA remain basepaired to the template DNA. The rest of the RNA molecules falls off the template to allow the DNA behind it to rewind.

RNA Polymerases use the DNA strand below them as a template to direct which nucleotide to add to the 3′ end of the growing RNA strand at each point in the sequence. The RNA Polymerase travels along the template DNA one nucleotide at at time. Whichever RNA nucleotide is capable of basepairing to the template nucleotide below the RNA Polymerase is the next nucleotide to be added. Once the addition of a new nucleotide to the 3′ end of the growing strand has been catalyzed, the RNA Polymerase moves to the next DNA nucleotide on the template below it. This process continues until transcription termination occurs.


Transcription termination by RNA Polymerase II on a protein-encoding gene.: RNA Polymerase II has no specific signals that terminate its transcription. In the case of protein-encoding genes, a protein complex will bind to two locations on the growing pre-mRNA once the RNA Polymerase has transcribed past the end of the gene. CPSF in the complex will bind a AAUAAA sequence, and CstF in the complex will bind a GU-rich sequence (top figure). CPSF in the complex will cleave the pre-mRNA at a site between the two bound sequences, releasing the pre-mRNA (middle figure). Poly(A) Polymerase is a part of the same complex and will begin to add a poly-A tail to the pre-mRNA. At the same time, Xrn2 protein, which is an exonuclease, attacks the 5′ end of the RNA strand still associated with the RNA Polymerase. Xrn2 will start digesting the non-released portion of the newly synthesized RNA until Xrn2 reaches the RNA Polymerase, where it aids in displacing the RNA Polymerase from the template DNA strand. This terminates transcription at some random location downstream from the true end of the gene (bottom figure).

The termination of transcription is different for the three different eukaryotic RNA polymerases.

The ribosomal rRNA genes transcribed by RNA Polymerase I contain a specific sequence of basepairs (11 bp long in humans 18 bp in mice) that is recognized by a termination protein called TTF-1 (Transcription Termination Factor for RNA Polymerase I.) This protein binds the DNA at its recognition sequence and blocks further transcription, causing the RNA Polymerase I to disengage from the template DNA strand and to release its newly-synthesized RNA.

The protein-encoding, structural RNA, and regulatory RNA genes transcribed by RNA Polymerse II lack any specific signals or sequences that direct RNA Polymerase II to terminate at specific locations. RNA Polymerase II can continue to transcribe RNA anywhere from a few bp to thousands of bp past the actual end of the gene. However, the transcript is cleaved at an internal site before RNA Polymerase II finishes transcribing. This releases the upstream portion of the transcript, which will serve as the initial RNA prior to further processing (the pre-mRNA in the case of protein-encoding genes.) This cleavage site is considered the “end” of the gene. The remainder of the transcript is digested by a 5′-exonuclease (called Xrn2 in humans) while it is still being transcribed by the RNA Polymerase II. When the 5′-exonulease “catches up” to RNA Polymerase II by digesting away all the overhanging RNA, it helps disengage the polymerase from its DNA template strand, finally terminating that round of transcription.

In the case of protein-encoding genes, the cleavage site which determines the “end” of the emerging pre-mRNA occurs between an upstream AAUAAA sequence and a downstream GU-rich sequence separated by about 40-60 nucleotides in the emerging RNA. Once both of these sequences have been transcribed, a protein called CPSF in humans binds the AAUAAA sequence and a protein called CstF in humans binds the GU-rich sequence. These two proteins form the base of a complicated protein complex that forms in this region before CPSF cleaves the nascent pre-mRNA at a site 10-30 nucleotides downstream from the AAUAAA site. The Poly(A) Polymerase enzyme which catalyzes the addition of a 3′ poly-A tail on the pre-mRNA is part of the complex that forms with CPSF and CstF.

The tRNA, 5S rRNA, and structural RNAs genes transcribed by RNA Polymerase III have a not-entirely-understood termination signal. The RNAs transcribed by RNA Polymerase III have a short stretch of four to seven U’s at their 3′ end. This somehow triggers RNA Polymerase III to both release the nascent RNA and disengage from the template DNA strand.