Replication origins experimentally identified in archaeal chromosomes
One fundamental challenge of cells is to accurately copy their genetic material for cell proliferation. This task is performed by core machineries considered conserved in all three domains of life: bacteria, archaea and eukaryotes .
For the vast majority of bacteria, the genome consists of one circular DNA molecule. Replication is initiated at a single replication origin from which two replication forks progress in the opposite direction. Replication termination takes place in the terminus region opposite the origin so that each replication fork has copied approximately one half of the genome. Studies of
Unlike bacterial genomes, eukaryotic chromosomes contain numerous replication origins that can be used as backup origins to rescue arrested forks. Consequently, the importance of replication restart pathways in eukaryotes has long been ignored. However, recent studies have demonstrated that fork restart pathways operate also in eukaryotic cells and are important for cell viability under replication stress conditions. Eukaryotic replication restart pathways described also involve recombination proteins, as in bacteria. Thus it appears that general rules regarding replication restart and the key role of recombination proteins in these processes are conserved in bacteria, yeast and higher eukaryotes, but little is known in archaea, the third domain of life. This is of interest as archaea appear to be evolutionary hybrids between bacteria and eukaryotes.
Three main archaeal phyla are currently recognized: Crenarchaeota, Euryarchaeota  and Thaumarchaeota . Similarly to most bacteria, archaeal genomes are also formed by a circular DNA molecule. However, unlike bacteria, some archaeal species have a single origin, whereas others have multiple origins per chromosome. Moreover, the ploidy of the genome in archaea varies considerably, with some species having one copy per cell whereas others have up to 25 copies of their genome in proliferating cells. As archaeal DNA replication consists both of evolutionary conserved as well as original features, understanding replication restart in these microorganisms will shed light on these fundamental but very complex pathways crucial to fulfill DNA replication. In this chapter we present recent advances on replication in archaea, followed by focused description of the Hef/XPF protein and its implication in replication restart in archaeal cells.
2. DNA replication origins in archaea
2.1. Multiple replication origins
Bacteria replicate their circular chromosome from a defined site called a replication origin. Two replication forks assemble at the replication origin and move in opposite directions. Each replication fork progresses at the same rate, and termination occurs at specific sites opposite the origin. Archaeal chromosomes are also circular, but whereas some archaea initiate replication from a single origin others replicate their chromosome from multiple replication origins, as observed for eukaryotic linear chromosomes (Table 1).
|At least 2|||
Replication from a single replication origin was reported experimentally in the euryarchaea
2.2. Archaeal replication initiator Orc1/Cdc6 proteins and origins recognition
How replication is regulated to allow a single circular DNA molecule to be replicated from uneven multiple origins is an ongoing question in archaea (Figure 1).
From each replication origin two replication forks are assembled and progress at the same rate so that termination of the replication is asynchronous. The origin region usually has a high content of adenine and thymine residues flanked by several conserved repeated motifs known as Origin Recognition Boxes (ORBs). In manycases archaeal replication origins are linked to replication genes [15, 18] and are located near genes coding initiator proteins. Despite the conservation of the replication origin-initiator structure, archaeal replication origins exhibit considerable diversity in terms of both ORB elements and their initiator genes [7, 11, 12]. Because replication origins can be dramatically diverse, it may facilitate differential usages by these microorganisms to adapt to various harsh environments.
All sequenced archaeal genomes encode proteins homologous to the eukaryotic initiator proteins Orc1 and Cdc6. Because the archaeal proteins are related both to the eukaryotic Orc1 subunit, involved in the replication origin recognition, and Cdc6, involved in the replicative helicase recruitment, they may combine both activities in a single polypeptide. Indeed, several studies have shown that the archaeal replicative helicase MCM is recruited by Cdc6/Orc1 proteins at replication origins [14, 19, 20]. Archaeal Cdc6/Orc1 proteins also share mechanistic similarities with the bacterial initiator protein DnaA.
How multiple replication origins are regulated by Cdc6/Orc1 proteins in archaeal cells is a complex question. The number of Orc1/Cdc6 proteins varies between species, and recent genetic studies attempting to delete
For instance, four
Moreover, additional role of initiator proteins independent of replication origins has recently been suggested by serial deletions of
2.3. Are replication origins essential for viability in archaea?
The specific initiation sites, replication origins, on the chromosome of
How replication initiates in absence of replication origins? Because no evidence for activation of dormant origins has been found, authors favoured the hypothesis that replication initiation occurs randomly on the chromosome at recombination intermediates. Recombination-Dependent Replication (RDR) has first been observed in
But the deletion of
An alternative explanation for RDR is activation of dormant origins randomly in cells so that no preferential origin emerged at the level of a cell population [29, 30]. In this scenario the essentiality of RadA could imply that randomly-initiated replication forks more often collapse and have to be restarted. This notion would be consistent with an organization of archaeal genes on the genome preventing collision of replication machinery with transcription machinery  and physical connections recently suggested between replication and transcription machineries .
In conclusion, this study by Hawkins
3. Archaeal Hef/XPF proteins from the XPF/MUS81/FANCM family
Proteins belonging to the XPF/MUS81/FANCM endonuclease family act on 3’-flap DNA structures that are formed during DNA repair or replication restart. They are found throughout eukarya and archaea but to date have not been identified in bacteria. Eukaryotes have several XPF/MUS81/FANCM family members that all share a conserved nuclease domain  whereas MUS81 proteins possess only an active nuclease domain. In XPF, an active nuclease domain is fused to a SF2-helicase domain that is degenerated and appears to be inactive . By contrast, FANCM consists of a helicase:nuclease fusion in which the nuclease domain is degenerated [36, 37]. Other members can be found that have a degenerated nuclease and/or helicase domain. They assemble into heterodimeric complexes with MUS81, XPF or FANCM proteins to form distinct active complexes involved in DNA repair, meiotic recombination and replication restart  (Figure 3).
All archaea encode a protein of the XPF/MUS81/FANCM family of endonucleases. It exists in two forms. The long form, referred as Hef, consists of an N-terminal helicase fused to a C-terminal nuclease and is specific to the euryarchaea. The short form, referred as XPF, lacks the helicase domain and is specific to the crenarchaea and the thaumarchaea (Figure 3).
The long-formed Hef protein was first identified in
In vitrostudies of crenarchaeal XPF proteins
The crystallographic structure of XPF from the crenarchaea
The nuclease activity of XPF from
In vitrocharacterization of euryarchaeal Hef proteins
As mentioned previously Hef was identified in
These biochemical studies have indicated that both creanarchaeal XPF and euryarchaeal Hef proteins interact with PCNA and display biochemical activities consistent for being proteins involved in DNA repair and/or replication restart. Is this hypothesis supported by
3.3. What have we learned deleting
hefgene in euryarchaea
In contrast, the deletion of
|UV irradiation||Up to 150 J/m2||on plate||-||2 or 5 J/m2||on plate||+|
|MMS||0,04%||1 hour in suspension||-||0,05%||4 hours in suspension||++|
|γ-rays||Up to 1000 Gy||on plate||-||1700 Gy||in suspension||++|
|Phleomycin||1 or 2 mg/ml||1 hour in suspension||-|
|Mitomycin C||0,02 µg/ml||On plate||slow-growing||100 µg/ml||4 hours in suspension||+++|
A possible explanation for these phenotypic differences is that NER proteins in
To further investigate the role of Hef in
4. Dynamic localization of Hef proteins fused to the Green Fluorescent Protein (GFP) in living
The Green Fluorescent Protein (GFP) was originally isolated from the jellyfish
4.1. Fusion of the green fluorescent protein to the C-terminal end of Hef protein in
GFP has been used to investigate proteasome-dependant proteolysis and protein levels in
Whether GFP-fused Hef proteins remained functional was then tested by comparing cells deleted for
4.2. Localising the fluorescence signal in
H. volcaniiliving cells
We then observed the localisation of Hef::GFP proteins by fluorescence microscopy, comparing cells exposed to APD to non-treated control samples. Towards this goal, a drop of cells was spotted on an agarose slice placed on a glass slide. After allowing this drop to dry, the agarose pad was covered with a cover-slip for cell imaging studies using a wield-field microscope to visualize a large number of individual cells. Differential Interference Contrast (DIC) [also known as Nomarski Interference Contrast (NIC)], was first used to visualized the cells as it enhances the contrast in unstained, transparent samples. Then fluorescence imaging was performed (exciting at 474 nm and collecting at 527-554 nm). Note however that due to the small cell size (around 1 to 2 µm) and the use of the soft agarose, not all cells were in the same focal plane. In order not to lose any information, fluorescence images were acquired at different focal planes on the z-axis. Consecutive slices of cells in focus were then selected and used to perform a maximum intensity z-projection. At each pixel, the highest fluorescence signal was kept when comparing the selected images. This maximum intensity z-projection resulted into a two-dimensional picture where the maximal fluorescence signals from different focal planes were recorded (Figure 4).
Resulting images contained hundreds of cells that were analysed by quantitative image analyses using IMARIS software. Different imaging parameters were optimized to detect cells and fluorescence foci within cells using automatic thresholds to avoid user-bias. This approch allowed thousands of cells to be analysed in each condition tested, providing extremely high statistical power.
4.3. Hef::GFP molecules are recruited at arrested replication forks
Using such approach, we have shown that Hef::GFP proteins formed fluorescence foci even under normal growth condition, in the absence of any DNA damaging agents. The number of these foci was significantly increased from two to four foci
We have also observed that cell size was increased from 28 to 45 µm² in response to replication arrest (i.e. APD exposure). We have shown using other DNA damaging agents that increased cell size and number of foci were specific to APD treatment, suggesting that indeed
4.4. Diffusing pattern of Hef::GFP molecule upon replication arrests
To investigate the diffusion of Hef::GFP molecules inside and outside fluorescence foci, we performed Fluorescence Recovery After Photobleaching (FRAP) and Number and Brightness (N&B) experiments. These experiments were performed using a confocal microscope on cells immobilized on a poly-D-lysine coated cover-slip.
In FRAP experiments a region of interest was photobleached in a cell. The speed of fluorescence recovery in that region was then measured, reflecting the diffusion of Hef::GFP fluorescent molecules arriving from the non-photobleached region of the cell. In control cells (no aphidicolin), one major population of Hef::GFP diffusing molecules was observed. From the fit of the recovery curve we obtained the recovery constant, allowing then the apparent two-dimensional diffusion rate of Hef::GFP to be estimated at 0.8 to 2.3 µm² per second. This appeared markedly lower than expected for Hef dimer, as revealed by analytical ultracentrifugation experiments on purified
Such changes in the diffusion pattern of Hef::GFP molecules upon replication arrests were also observed using N&B technique that measures fluctuation of fluorescence intensity in each analysed pixel . These analyses were performed on one hundred images taken every 2 seconds, and cell regions including and excluding fluorescence foci were compared. Fluctuation of fluorescence intensity per pixel was then used to determine the number of diffusing molecules and their brightness. This information can then be used to deduce changes in the oligomeric state of the fluorescent molecules. When N&B technique was applied to diffusing Hef::GFP molecules, changes in the oligomeric state (i.e. higher brightness) were observed upon APD treatment. This observation revealed oligomerization and/or co-localisation of several Hef::GFP molecules induced by APD exposure (Figure 7), and provided a feasible explanation for the slow diffusion in APD treated cells revealed by FRAP experiments.
Overall, the results obtained from FRAP and N&B experiments were consistent with the notion that Hef::GFP molecules are actively recruited at arrested replication forks. Whether the slow-diffusion pattern of Hef::GFP molecules reflects their recruitment directly on DNA and/or as part of protein complexes at arrested replication forks are questions that remain to be addressed. Interestingly,
Moreover, our work has also shown that replication forks arrest spontaneously occured in
5. Concluding remarks
In conclusion, as archaea possess hallmarks of both bacterial and eukaryotic replication systems we believe that continuation of studies underlined will shed light on the evolutionary history of replication restart mechanisms and its complex machinery that we are just starting to unravel in eukaryotes and now archaea.
We thank A. Lestini for figures 1, 2, 3 and 4. Figures 5, 6, 7 and 8 were reproduced with permission from Lestini
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