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- Herausgeber: John Wiley & Sons
- Kategorie: Wissenschaft und neue Technologien
- Serie: ISTE Invoiced
- Sprache: Englisch
Archaea constitute a new branch of life alongside bacteria and eukaryotes. These microorganisms are unique in their cellular and molecular aspects. They have evolutionary links with the first eukaryotic cells and are now being used to elucidate fundamental biological questions.
Champions of extremophilicity, archaea are helping to lift the veil on the limits of life on Earth. Biology of Archaea 2 presents the diverse molecular mechanisms involved in the fundamental processes of genome maintenance and regulation of gene expression in archaeal cells.
This book analyzes the complex machinery involved in chromosome replication, DNA repair, RNA synthesis (transcription) and protein synthesis (translation), and explores the different classes of RNAs and enzymes involved in RNA maturation and degradation. These regulate the stability of messenger and regulatory non-coding RNAs, and contribute to the formation of the mature forms of ribosomal RNAs and transfer RNAs. These molecular mechanisms are closely related to those of eukaryotes.
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Veröffentlichungsjahr: 2025
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Table of Contents
Cover
Table of Contents
Title Page
Copyright Page
Preface
1 Replication of Archaeal Chromosomes
1.1. Replication initiation
1.2. The elongation phase of replication
1.3. Okazaki fragment maturation
1.4. Restarting replication forks
1.5. Replication termination
1.6. Chromosome segregation and decatenation
1.7. References
2 Archaeal DNA Repair
2.1. General introduction
2.2. Several different types of DNA damages occur in archaea
2.3. Different archaeal DNA repair pathways
2.4. Coordination of the different pathways by the replication clamp
2.5. Summary and conclusion
2.6. References
3 Transcription in Archaea
3.1. RNA polymerase
3.2. The three stages of transcription
3.3. Transcription regulation
3.4. References
4 RNA Classes and Their Maturation and Degradation Enzymes
4.1. Introduction
4.2. RNA classes in archaea
4.3. Ribonuclease families in archaea
4.4. Ubiquitous RNA-binding proteins in archaea
4.5. Conclusion
4.6. References
5 Ribosome and Transfer RNA Biogenesis
5.1. Ribosome biogenesis
5.2. Synthesis and maturation of tRNAs
5.3. Methodical toolboxes
5.4. References
6 The Diversity and Function of Noncoding RNAs in Archaea
6.1. Introduction
6.2. Noncoding RNA diversity
6.3. ncRNA identification
6.4. Circularization of noncoding RNAs
6.5. Conclusion
6.6. References
7 Translation in Archaea
7.1. Initiating the translation
7.2. Elongation peptide synthesis
7.3. Translation termination and ribosome recycling
7.4. Conclusion
7.5. References
List of Authors
Index
End User License Agreement
List of Tables
Chapter 2
Table 2.1. Experimental conditions used in the laboratory to mimic different D...
Table 2.2. Examples of functional modulation of archaeal DNA repair enzymes by...
Table 2.3. Simplified presentation of the phylogenetic distribution of the maj...
Chapter 3
Table 3.1. Conservation of RNA polymerase subunits and general transcription f...
Chapter 4
Table 4.1. Ribonuclease families in archaea. Occurrence, activities and known ...
Chapter 7
Table 7.1. Bacterial, eukaryotic and archaeal ribosomes. Ribosomal protein con...
Table 7.2. Main characteristics of translation initiation in the three domains...
Table 7.3. Players in translation elongation in the three domains of life
Table 7.4. Players in translation termination and ribosome recycling in the th...
List of Illustrations
Chapter 1
Figure 1.1. The different phases of replication initiation . Comment on Figu...
Figure 1.2. Diagram of the composition and organization of the replisome.
Figure 1.3. Maturation mechanism of archaeal Okazaki fragments. Comment on F...
Figure 1.4. Restarting arrested replication forks . Comment on Figure 1.4.–
Chapter 2
Figure 2.1. The different types of abnormal sites including replication errors...
Figure 2.2. Early evolutionary genomics analysis of archaeal genomes indicated...
Figure 2.3. Different mechanisms ensuring high fidelity of DNA replication in ...
Figure 2.4. Two different strategies for repairing mismatches that occur durin...
Figure 2.5. An overview of a global genomic NER (GG-NER) where various enzymes...
Figure 2.6. During the short-patch BER, a monofunctional DNA glycosylase cleav...
Figure 2.7. In archaea, two major pathways repairing DSBs are homologous recom...
Figure 2.8. Two orthogonal views of the archaeal PCNA in complex with DNA. The...
Chapter 3
Figure 3.1. Fundamental theory of molecular biology. DNA, the stable, transmis...
Figure 3.2. RNAP structure. For a color version of this figure, see www.iste.c...
Figure 3.3. Transcription cycle in archaea. For a color version of this figure...
Figure 3.4. Transcription initiation. For a color version of this figure, see ...
Figure 3.5. Schematic representation of the archaeal transcription elongation ...
Figure 3.6. Structural elements essential for maintaining DNA–RNA hybrid integ...
Figure 3.7. Transcription termination and RNA release. For a color version of ...
Figure 3.8. Transcription regulation. (a) Transcription repressors bind to the...
Chapter 4
Figure 4.1. Metabolism in archaea: a mosaic of bacterial and eukaryotic charac...
Figure 4.2. RNA classes in archaea and their degradation and maturation enzyme...
Figure 4.3. RNA classes in archaea and their degradation and maturation enzyme...
Figure 4.4. Taxonomic distribution of ribonuclease families in archaeal phylog...
Chapter 5
Figure 5.1. Representative ribosomal subunit structures across the tree of l...
Figure 5.2. Ribosome and ribosome biogenesis key features overview across the ...
Figure 5.3. Phylogenetic distribution of ribosomal proteins across the tree of...
Figure 5.4. Unlinked rRNA genes in archaea (reproduced from Jüttner and Ferrei...
Figure 5.5. Ribosomal RNA processing in the Euryarchaeon Haloferax volcanii. ...
Figure 5.6. Conservation between archaea and eukaryotes of the late steps of r...
Figure 5.7. tRNA structure. Comment on Figure 5.7.– Cloverleaf structure of ...
Figure 5.8. Examples of unusual archaeal tRNA genes (figure reproduced from He...
Figure 5.9. Overview of the tRNA maturation process in archaea and associated ...
Figure 5.10. RNaseP in the three domains of life (figure reprinted from Wu et ...
Figure 5.11. Mechanism of CCA tail addition by an archaeal tRNA terminal trans...
Figure 5.12. Intron splicing in archaea (left panel adapted from Clouet-d’Orva...
Figure 5.13. Occurrence of modified nucleosides in tRNA (images courtesy of th...
Figure 5.14. Modified nucleosides found in tRNA and their distribution among l...
Figure 5.15. tRNA modifications identified in the archaeon H. volcanii and pre...
Figure 5.16. tRNA-derived fragments in archaea (figure adapted from Clouet-d’O...
Figure 5.17. Aminoacylation by aminoacyl tRNA synthetases (figure adapted from...
Figure 5.18. Classification of aminoacyl-tRNA synthetases (figure reproduced f...
Figure 5.19. Incorporation of non-canonical amino acids into proteins (figure ...
Chapter 6
Figure 6.1. Types of noncoding RNA-related regulatory mechanisms in archaea. P...
Figure 6.2. Schematic representation of ribonucleic complexes involving (a) bo...
Figure 6.3. Noncoding RNA diversity in archaea and mechanisms of interaction. ...
Figure 6.4. CRISPR system. In three steps, immunization of the organism is pro...
Figure 6.5. Classification of tRNA-derived small RNAs (according to Xie et al....
Figure 6.6. Development of sequencing techniques adapted to each RNA subcatego...
Figure 6.7. Classification of bioinformatics approaches for identifying noncod...
Figure 6.8. Circular RNA biogenesis. For a color version of this figure, see w...
Chapter 7
Figure 7.1. Schematic representation of translation initiation stages in (a) b...
Figure 7.2. Structural organization of common initiation factors in archaea an...
Figure 7.3. Translation initiation stages in P. abyssi.
Figure 7.4. Key stages and players in translation elongation.
Figure 7.5. Schematic representation of transfer RNA rearrangement during the ...
Figure 7.6. GTPase recruitment platform or L12/P stalk.
Figure 7.7. Key stages and players in translation termination and mRNA recycli...
Guide
Cover Page
Table of Contents
Title Page
Copyright Page
Preface
Begin Reading
List of Authors
Index
Wiley End User License Agreement
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SCIENCES
Biology, Field Director – Marie-Christine Maurel
Microbiology, Subject Head – Marie-Christine Maurel
Biology of Archaea 2
Molecular Biology of Archaea, From Genome Maintenance to the Regulation of Gene Expression
Coordinated by
Béatrice Clouet-d’Orval
Bruno Franzetti
Philippe Oger
First published 2025 in Great Britain and the United States by ISTE Ltd and John Wiley & Sons, Inc.
Apart from any fair dealing for the purposes of research or private study, or criticism or review, as permitted under the Copyright, Designs and Patents Act 1988, this publication may only be reproduced, stored or transmitted, in any form or by any means, with the prior permission in writing of the publishers, or in the case of reprographic reproduction in accordance with the terms and licenses issued by the CLA. Enquiries concerning reproduction outside these terms should be sent to the publishers at the undermentioned address:
ISTE Ltd27-37 St George’s RoadLondon SW19 4EUUK
www.iste.co.uk
John Wiley & Sons, Inc.111 River StreetHoboken, NJ 07030USA
www.wiley.com
© ISTE Ltd 2025The rights of Béatrice Clouet-d’Orval, Bruno Franzetti and Philippe Oger to be identified as the authors of this work have been asserted by them in accordance with the Copyright, Designs and Patents Act 1988.
Any opinions, findings, and conclusions or recommendations expressed in this material are those of the author(s), contributor(s) or editor(s) and do not necessarily reflect the views of ISTE Group.
Library of Congress Control Number: 2024949125
British Library Cataloguing-in-Publication DataA CIP record for this book is available from the British LibraryISBN 978-1-78945-169-6
ERC code:LS8 Ecology, Evolution and Environmental Biology LS8_10 Microbial ecology and evolution
Preface
Béatrice Clouet-d’Orval1, Bruno Franzetti2 and Philippe Oger3
1 MCD-CBI, CNRS, Université de Toulouse, France
2 IBS, CNRS, Université Grenoble Alpes, France
3 MAP, CNRS, INSA Lyon, Villeurbanne, France
More than 40 years ago, archaea were proposed as a new domain of life alongside bacteria and eukaryotes. It is now accepted that these organisms constitute a distinct group, unique in many cellular and molecular aspects. The discovery of these universally important organisms represents one of the greatest advances in modern biology. Archaea have evolutionary links with early eukaryotic cells and are now used to elucidate fundamental biological questions. As champions of extremophilia, archaea have also shed light on the limits of life on Earth. Over the years, our understanding of archaeal microorganisms has evolved considerably. Far from being exotic forms of microbial life, archaea are found to be omnipresent in all terrestrial sites (>20% of marine microbial biomass) and are notably present in the human microbiota. As new genomes are sequenced, it becomes evident that archaea represent a complex, extremely diverse world and also possess a largely unexplored virosphere. In soils and oceans, archaea play a key role in the planet’s major geochemical cycles. In particular, methanogens, exclusively represented by archaea, are central to issues of climate change and energy challenges. To understand their impact on the environment and human health, it is imperative to decode their unique features at a molecular level. The molecular machinery deciphered in archaea will provide key paradigms for understanding fundamental biological processes conserved across life forms. Notably, the molecular bases of genetic information processing in archaea are often very similar to their counterparts in eukaryotes (translation, transcription, replication, recombination and DNA repair), even though archaea have a cellular structure similar to bacteria. Moreover, the regulatory systems for gene expression through post-transcriptional mechanisms remain poorly understood.
Our knowledge of archaea is rapidly evolving thanks to the advent of high-throughput sequencing of entire genomes. It is time to introduce a wide audience to the advances that illuminate our understanding of biology and reveal the originality of this domain of life.
The 14 chapters, divided into two volumes, review the discovery and evolution of the domain of archaea and summarize our current knowledge of cellular and molecular processes to better integrate this domain into the broader understanding of life.
January 2025
1Replication of Archaeal Chromosomes
Ghislaine Henneke1, Roxane Lestini2, Marc Nadal3 and Didier Flament1
1BEEP, CNRS, Ifremer, Université de Bretagne Occidentale, Plouzané, France
2Laboratoire d’optique et biosciences, Inserm, CNRS, Institut Polytechnique de Paris, Palaiseau, France
3IBENS, CNRS, Inserm, PSL University, Paris, France
It was only when Watson and Crick proposed the double helix structure of DNA, in the early 1950s, that the scientific community finally accepted that DNA could be the carrier of heredity. Since then, the study of DNA replication has become one of the key themes of molecular biology. First developed in bacteria in the 1960s, then in eukaryotes in the 1980s, it was only after Carl Woese and George Fox clarified the phylogenetic position of archaea that the study of the molecular mechanisms of DNA replication began in archaea, in the early 1980s. This new scientific venture has proved exciting in several ways. First, although the composition of the replication machinery shows a high degree of diversity in archaea, it is similar to that of eukaryotes, indicating a common evolutionary history (see Volume 1, Chapters 2 and 3).
Thus, archaeal proteins and the metabolic pathways involved in this process represent excellent models for study. Furthermore, the study of replication initiation in these organisms has recently challenged one of the dogmas of molecular biology, according to which an origin of replication is required to initiate the autonomous replication of a DNA fragment. Indeed, in some archaea, the origin(s) of replication (of which there can be several) can be genetically suppressed without the cell exhibiting growth defects. This suggests the presence of an alternative replication initiation mechanism in archaea, which remains to be characterized.
The general process of DNA replication is conserved throughout all cellular organisms. DNA replication can be divided into four main stages. The first stage involves initiation, during which specific proteins recognize and bind to the origin of replication. This DNA–protein complex recruits a helicase to unwind the double-stranded DNA at the origin. DNA primase and DNA polymerases are then recruited to the replication bubble generated by the helicase activity. The second stage is elongation, which corresponds to the duplication of both DNA strands simultaneously and bidirectionally. Due to the antiparallel structure of DNA and the direction of nucleotide polymerization from 5′ to 3′, the leading strand is synthesized continuously, while the lagging strand is replicated discontinuously in the form of Okazaki fragments. In the third stage, the RNA primers on the lagging strand are removed by the coordinated action of ribonucleases, flap endonucleases and DNA ligases. Finally, the last stage, known as termination, corresponds to the end of synthesis, when the replication forks meet or when they encounter a termination signal.
In this chapter, we will summarize what we know about DNA replication in archaea through the different phases of this process.
1.1. Replication initiation
1.1.1. Identifying the origins of replication
Although we now know the complete genome of some 420 cultivated isolates, our knowledge of the replication of these organisms is limited to <20 species, some of which are very closely related, and do not cover all the phyla of the archaea domain. In terms of genome structure, very different situations coexist, since some archaea have a single circular chromosome, like most bacteria, while halophilic archaea (Haloarchaea) have several chromosomes. Furthermore, while some archaea are haploid, others are polyploid. This genomic diversity implies different couplings with the cell cycle and different regulations of replication initiation.
The first origin of replication observed in archaea was that of Pyrococcus abyssi. This Euryarcheota, of the order Thermococcales, has a single chromosome containing a single bidirectional origin of replication (Matsunaga et al. 2001). In methanogens, in silico studies indicate the presence of a single origin. Allele frequency analysis in Nitrosopumilus maritimus, a Thaumarchaeota, reveals the presence of a single origin of replication. However, multiple origins of replication per chromosome have been found in Halobacteria. In Crenarchaeota, there seems to be the presence, on a single circular chromosome, of up to four multiple origins of replication, this being unrelated to genome size. Surprisingly, in 2013, Thorsten Allers’ team at the University of Nottingham demonstrated that all the origins of replication in the halophilic archaea Haloferax volcanii could be deleted without preventing the mutant strain from dividing and continuing to grow (Hawkins et al. 2013). This same finding was then established in a hyperthermophilic archaea of the order Thermococcales (Gehring et al. 2017), raising the question of the role of these origins of replication in certain archaea, as well as the alternative mechanism enabling chromosome replication in the absence of the origin of replication. Although these studies still only concern a small number of archaea, it seems that the vast majority of archaea have multiple origins of replication, with Thermococcales and Thaumarchaeota being the exceptions. To date, all observed origins of replication allow bidirectional replication of the associated replicon.
1.1.2. Recognizing the origins of replication
Replication initiation takes place at specific sites, composed of at least two short repetitive sequences, the ORBs (origin recognition boxes) (Matsunaga et al. 2001; Robinson et al. 2007), located near an AT-rich DNA sequence enabling local separation of the two complementary strands, known as DNA unwinding element (DUE) (Figure 1.1(a)). These sequences are recognized by specific proteins, the Orc/Cdc6 proteins. These proteins belong to the same family as the Orc (origin recognition complex) proteins involved in the initiation of replication in eukaryotes. Their binding induces local DNA unwinding (Matsunaga et al. 2009). In eukaryotes, the six Orc proteins (Orc1–6) associate at replication origins with Cdc6 and Cdt1 proteins, so as to enable initiation. In archaea, the initiator proteins are both homologous to the Orc protein family, essentially Orc1, and to Cdc6 proteins. It was surprising to find these proteins in multiple copies, with more than 14 in H. volcanii (Norais et al. 2007). Furthermore, genes are most often localized in the immediate vicinity of the origin to which they bind, reflecting the localization of the bacterial initiator protein DnaA (Figure 1.1). Sulfolobus islandicus has been shown to possess three replication origins, OriC1, OriC2 and OriC3, which bind Orc1-1, Orc1-3 and WhiP proteins, respectively. Unlike the other initiation proteins, WhiP is related to Cdt1, a protein involved in origin recognition in eukaryotes. Although the order Sulfolobales has an Orc1-2 protein, it is WhiP that binds to the OriC3 region.
As in eukaryotes, expression of the genes encoding Orc/Cdc6 proteins is finely regulated during the cell cycle. In Sulfolobales, genes encoding Orc1-1 and Orc1-3 proteins are expressed at the start of the G1 phase (Lundgren and Bernander 2007), leading to the binding of these initiator proteins to replication origin sequences at the start of the G1 phase. The Orc1/Cdc6 proteins have an ATP binding and hydrolysis site, and conformational changes take place during the ATP binding/hydrolysis cycle. ATP binding is necessary for the activation of replication origins, but ATP hydrolysis is not essential for replication. For the moment, the precise signals leading to replication induction are not known. However, the third Orc1/Cdc6-like protein, Orc1-2, which is not involved in replication initiation, could act as a replication repressor. The sequence recognized by Orc1-2 overlaps with those recognized by Orc1-1 and Orc1-3 (at OriC1 and OriC2 origins, respectively). Consequently, its binding would competitively block these two origins, which is consistent with the fact that this protein is specifically expressed in the G2 phase.
While the main mode of replication in archaea seems to be the replicon hypothesis proposed by Jacob et al. (1963) – in other words, involving the attachment of an inducible protein transacting on a particular sequence – the presence of these origins, and the proteins capable of recognizing them, is not essential in archaea. In fact, in H. volcanii, deletion of the four replication origins of the largest chromosome enables faster growth of these cells without it being possible to identify a dormant origin, as is the case in Haloferax mediterranei (Yang et al. 2015). In the H. volcanii mutant without the origin sequences, the presence of the RadA protein, the homologue of the RecA/Rad51 homologous recombination proteins, is required to allow for replication (Hawkins et al. 2013). The same is true for Thermococcus kodakarensis, in which deletion of the single origin of replication and/or the Cdc6 protein does not alter the growth of this archaea.
This type of replication, known as RDR (recombination-dependent DNA replication), makes use of homologous recombination, as already described in bacteria (Ogawa et al. 1984), but with the consequence of reduced growth in bacteria. The precise mechanisms leading to the formation of a complex-enabling genome replication through this pathway could be involved in the polyploidy observed, but they are, as yet, unknown to us, and we will therefore concentrate hereafter on replication, starting from the sequences of replication origin.
Figure 1.1.The different phases of replication initiation .
Comment on Figure 1.1.– (a) A common feature of the archaeal origins of replication is the presence of a DUE (DNA unwinding element) sequence flanked by adjacent Orc-cdc6 (mORB) protein binding sites. The gene specifying the Orc-cdc6 initiator protein is found close to the origin. The Orc-cdc6 initiator protein binds to the mORB sites and then facilitates the recruitment of MCM replicative helicases; (b) by interacting preferentially with the open forms of MCM, Orc-cdc6 would thus enable their positions, which would probably take place on double-stranded DNA; (c) similar to the situation in eukaryotes, a transition would enable MCM to bind to single-stranded DNA in a targeted way and would cause the opening of the DUE region; (d) the CG complex ((Gins14-Gin23-Cdc45)2) binds to and activates the MCM helicase, enabling it to move by hydrolyzing ATP, which leads to dissociation of the Orc-cdc6 proteins, thus allowing primase binding; (e) primase (PriSLX) then binds the first nucleotide triphosphate (in red), and then synthesizes a short RNA chain forming the primer.
1.1.3. Pre-initiation complex formation
As in eukaryotes, it is the ORB-Orc1/Cdc6 complex that serves as the platform for the replisome. A critical step is the positioning of the replicative helicases. In archaea, the replicative helicase belongs to the MCM family of proteins, but unlike in eukaryotes, where it forms a ring consisting of six different subunits (MCM2-7), it is a homohexamer in archaea. Two helicases are positioned head-to-tail on the DNA at the ORB-Orc1/Cdc6 complex, each hexamer surrounding a double-stranded DNA (Figure 1.1(b)). Archaeal MCMs naturally exist in two forms, closed and open. The open form of the ring allows the double-stranded DNA to be positioned centrally, without the need for a helicase loader, unlike in bacteria and eukaryotes. The Orc1/Cdc6 proteins have two roles. On the one hand, they enable correct positioning of the two MCMs at the region of origin, possibly by facilitating the opening of the MCM hexamer; on the other hand, they generate a single-stranded DNA region at the DUE region. A rearrangement of the complex (that is yet-to-be-understood) would allow the positioning of a single-stranded DNA in the center of the ring. It is in this form that MCMs acquire their helicase activities, moving onto this single strand and ejecting the other strand of the DNA duplex (Figure 1.1(c)). These helicases become fully active after association with the (Gins23-Gins15)2 complex, forming a homolog of the eukaryotic GINS complex and two copies of the eukaryotic Cdc45 homologous protein. The GINS-MCM-Cdc45 combination then reconstitutes a GINS-Cdc45 complex that is capable of strongly stimulating MCM helicase activity, forming a GMC complex similar to that of eukaryotes (Greci and Bell 2020) (Figure 1.1(d)).
We have seen that a single strand of DNA is present in the central channel of the MCM hexamer, enabling it to move from 3′ to 5′ on the single-stranded DNA serving as a template for continuous replication. To enable this advance and the opening of the DNA, the helicase hydrolyzes ATP. The transition that allows complete opening of the original region and correct positioning of MCM helicases on single-chain DNA has yet to be determined. To form a bidirectional replication bubble, it has been proposed that the two helicases cross over one another, passing one above the other, each on its own strand. Single-stranded DNA-binding proteins would then act, essentially on the discontinuous strand, with the primase on the continuous strand. Archaea possess two types of single-stranded DNA-binding proteins, one very similar to the RP-A protein of eukaryotes, the other very similar to the SSB protein of bacteria. Primase (PriSLX) is then likely to be positioned simultaneously, as primase interacts with the GMC complex via the GINS complex (Figure 1.1(d)). Presumably, this interaction is what ensures the coupling between the progression of the replication fork, created by the helicase and primer synthesis (Greci and Bell 2020) (Figure 1.1(e)). At this stage, activation of the GMC complex leads to the formation of a replication bubble, on which other replisome components are recruited to establish two replication forks and initiate bidirectional DNA synthesis.
1.1.4. Primer synthesis
DNA primases play an essential role in initiating DNA replication by synthesizing RNA primers on both the leading and lagging strands. In fact, DNA polymerases are unable to synthesize DNA de novo, and require a primer with a 3′-OH end to elongate DNA chains. In contrast, DNA primases are able to synthesize de novo short RNA fragments on template DNA, which serve as primers and are extended by the DNA polymerase (Figure 1.1(e)). Archaeal DNA primases consist of two subunits, PriS and PriL, which are conserved throughout the archaeal domain. However, a third component of the primase complex, PriX, is also present in many representatives of the Crenarchaeota phylum. PriS contains the enzyme’s active site, and PriL or PriLX regulates primase activity and contains an iron–sulfur cluster. In addition, PriS and PriL have orthologs in eukaryotes that form the primase module of the DNA polymerase α-primase complex. The PriSL primase complex is thus sometimes referred to as archaeal eukaryotic primase. Both subunits are essential for cell viability. However, the archaeal primase has the unique feature of being able to synthesize both DNA and RNA in vitro. Indeed, the PriS subunit of Pyrococcus furiosus synthesizes long DNA fragments, but the addition of the PriL subunit regulates the process by reducing DNA polymerase activity and increasing RNA polymerase activity, while decreasing the length of the fragments being generated (Liu et al. 2015). Furthermore, studies on the Pyrococcus abyssi enzyme suggest that the primase may also be involved in DNA repair, due to its DNA polymerase nucleotide gap filling and strand displacement activities, also observed in vitro (Le Breton et al. 2007). Although archaeal DNA primases exhibit both DNA and RNA synthesis activities in vitro, it has been shown that the primase is predominantly an RNA synthesis enzyme when the ratio of ribonucleotides (NTPs) to deoxyribonucleotides (dNTPs) is close to physiological values (Yan et al. 2018). Thus, the most likely hypothesis is that archaeal DNA primase is able to synthesize a short RNA fragment in the replication bubble on the forward strand, then steadily on the lagging strand, and that a primase conformational change mechanism governs the length of the synthesized fragments of 12–14 nucleotides (Greci and Bell 2020) (Figure 1.1(e)).
1.2. The elongation phase of replication
Following primer synthesis, the RNA primer synthesized by the primase is extended by a so-called “replicative” DNA polymerase. DNA polymerases have been classified into seven families on the basis of amino acid sequence similarity (A, B, C, D, E, X and Y). Bacteria and eukaryotes have five and 15 DNA polymerases, respectively. Replicative polymerases belong to the C family in bacteria and to the B family in eukaryotes. In archaea, the diversity of DNA polymerases is less rich, given that depending on the species, only two to three polymerase families are represented, by two to four different polymerases. These are the B, D and Y families. The DNA polymerases of the Y family are involved in DNA repair mechanisms, while the B and D families contain replicative DNA polymerases, which have a different phyletic distribution within the archaea domain. Archaeal DNA polymerase D, PolD, was the last DNA polymerase to be discovered, in the late 1990s (Cann et al. 1998). It is specific to archaea and is present in all phyla, except the Crenarchaeota. PolD is an enzyme complex composed of two subunits: DP1, which carries the proofreading activity, and DP2, which carries the DNA polymerization activity. The recent acquisition of the crystallographic structure of PolD has shown that the structure of DP2 resembles the catalytic center of RNA polymerases, thus opening a window into the evolutionary transition between the RNA and DNA worlds (Sauguet 2016). In terms of function, PolD is essential for viability, suggesting that it is the replicative DNA polymerase in species that possess it. Indeed, these species also generally possess another B-family polymerase, PolB, which is not essential for viability and is more likely to be involved in DNA repair activities (Kushida et al. 2019). Crenarchaeota, which do not possess PolD, have at least two B-family DNA polymerases that ensure genome replication (PolB I and PolB II, sometimes accompanied by PolB III), one of which is essential, namely PolB I.
Figure 1.2.Diagram of the composition and organization of the replisome.
As with all replicative DNA polymerases, PolD or PolB possess both DNA synthesis and error correction activities. These capabilities enable them to accurately replicate the genome, generating few errors. However, while this characteristic is essential for chromosome replication, it is not sufficient to ensure the completion of this cellular process. In particular, replicative DNA polymerases are always associated with ancillary factors that give them qualities of processivity (average number of nucleotides added to the DNA strand being synthesized, without the polymerase detaching itself from the template strand; this can exceed 100,000 nucleotides for replicative polymerases) and speed. This is particularly true of one of the essential factors in the architecture of the replisome for all three branches of life, the so-called “sliding clamp” protein (Figure 1.2). This sliding clamp surrounds the DNA and binds to the DNA polymerases, enabling the enzyme to remain in contact with the template DNA strand, thus improving the enzyme’s processivity. This factor (PCNA in eukaryotes and archaea, or β-clamp in bacteria) has a ring-like structure, formed by the assembly of two (β-clamp) or three (PCNA) subunits, and functions as a molecular platform that is capable of binding to DNA polymerases, as well as many other DNA-modifying proteins. To load the clamp onto the double-stranded DNA, a second factor is required: replication factor C (RFC). Its role is to attach the sliding clamp and load it onto the DNA at the 3′-OH end of the primer (Kelman and Kelman 2014). The organization of these factors, such as replicative DNA polymerase, PCNA, CMG complex and DNA primase, that make up the replisome allows the coupling of the replication fork progression and DNA synthesis. As the replication fork advances, it causes a separation of the two upstream DNA strands, generating two single-stranded DNAs that serve as templates for DNA synthesis (Figure 1.2). These single-stranded portions of DNA are highly unstable and prone to degradation. In order to prevent such damage, the single-stranded portions of DNA at the fork are colonized by a single-stranded DNA-binding protein (RPA for Euryarchaeota and SSB for Crenarchaeota), which prevents their degradation by nucleases (Taib et al. 2021). The spatial and time-based organization of these different components (DNA polymerases, primase, CMG complex, PCNA, RFC and RPA) at the replication fork level is not yet well defined in archaea (Figure 1.2). However, this replication fork will enable a coordinated DNA synthesis on the two strands, even though they have opposite polarities. Indeed, the anti-parallel nature of the DNA double helix and the 5′→3′ directionality of replicative DNA polymerases dictate that the leading strand, which has been newly synthesized at the replication fork, is polymerized continuously, while the other is replicated discontinuously. In other words, the primase regularly synthesizes RNA primers on the lagging strand, which are then elongated by the DNA polymerase, generating a succession of multiple segments known as Okazaki fragments (Okazaki et al. 1968) (Figure 1.2), whose maturation will generate a continuous strand of neosynthesized DNA.
1.3. Okazaki fragment maturation
The initiator RNA at the 5′ end of the Okazaki fragments must be removed and replaced by DNA. These newly synthesized DNA fragments are then joined together, creating a fully replicated, continuous DNA. This process is called Okazaki fragment maturation. It involves the coordinated action of various proteins and enzymatic activities. In particular, RPA (replication protein A), PCNA (proliferating cell nuclear antigen), a DNA polymerase, two nucleases FEN1 (Flap endonuclease 1) and RNase H (ribonuclease H), and a DNA ligase are involved in this mechanism. The essential steps in Okazaki fragment maturation are based on (i) ribonucleotide removal by ribonuclease H type 2 (RNase HII); (ii) oligonucleotide removal by FEN1; and (iii) DNA fragment assembly by DNA ligase. This process mainly takes place on the discontinuous strand, but also acts at the origin level, in the region of divergences between the continuous and discontinuous strands.
1.3.1. Removal of ribonucleotides by RNase HII
RNase HII is an enzyme capable of recognizing and hydrolyzing ribonucleotides incorporated into double-stranded DNA. It specifically cuts the phosphodiester bonds of the RNA moiety, generating a DNA fragment containing a 5′-mono- ribonucleotide (Figure 1.3(a) II). Unlike ribonuclease H type 1 or RNase HI, which cleaves RNA-DNA hybrids containing at least four RNA residues, RNase HII can catalyze the 5′ cleavage of a single ribonucleotide. These various cutting specificities suggest the involvement of ribonucleases H in mechanisms that maintain genomic stability (Tadokoro and Kanaya 2009). With the exception of a few archaea that have both types of ribonuclease H (RNases HI and HII), such as Sulfurispaera tokodaii, H. volcanii, Halobacterium sp. NRC-1 or Pyrobaculum aerophilum, the majority of archaeal genomes contain a single ribonuclease H, that being RNase HII. Thus, the ubiquity of this enzyme suggests its functional multidisciplinarity in DNA repair or replication pathways in archaea, even though it does not seem essential for cell survival (Meslet-Cladiére et al. 2007; Sarmiento et al. 2013; Birien 2018). Moreover, PCNA, an essential protein factor in genomic maintenance processes, is able to modulate the enzymatic properties of RNase HII (Meslet-Cladiére et al. 2007).
Solved three-dimensional (3D) structure of RNases HII is available in various archaea (Lai et al. 2000). It contains a highly conserved catalytic core, a characteristic of all ribonuclease 3D structures (Clouet-d’Orval et al. 2018). Like most other replication proteins, the structure–function relationship of archaeal RNase HII shares more homologies with eukaryotes than with bacteria (Malfatti et al. 2019). The mechanism for excising ribonucleotides from DNA is the most representative cellular function of RNase HII in the three domains of life, with the enzyme cutting 5′ to the ribonucleotide, initiating the repair system (Heider et al. 2017). Although the enzyme is not essential in archaea, as it is in yeast (Jeong et al. 2004), it cannot be ruled out that RNase HII may be involved as a backup or in cooperation in the Okazaki fragment maturation process (Henneke 2012) (Figure 1.3(a)). In this case, the RNase HII cleavage product would initiate the Okazaki fragment maturation pathway, preceding the strand synthesis-displacement step by DNA polymerase, followed by FEN1 cleavage of the branched doublestranded DNA (Figure 1.3(a) II).
1.3.2. Removal of oligonucleotides by FEN1
FEN1 is a structure-specific nuclease for DNA and/or RNA. It has endonuclease and 5′–3′ exonuclease activity (Friedrich-Heineken et al. 2003; Henneke et al. 2003). Like its eukaryotic counterpart, FEN1 plays a significant role in Okazaki fragment maturation and DNA repair (Henneke et al. 2003; Balakrishnan and Bambara 2013; Burkhart et al. 2017). It interacts directly and is stimulated by PCNA (Henneke 2012; Heider et al. 2017). In the presence of RNase HII (Figure 1.3(a)), the Okazaki fragment maturation process requires the removal of the mono-ribonucleotide from the preceding fragment in order for ligation by DNA ligase to proceed correctly. In this pathway, the replicating DNA polymerase (family B or D) during DNA synthesis encounters and displaces the fragment containing the mono-ribonucleotide, generating a branched double-stranded DNA substrate recognized and cut by FEN1 (Sato et al. 2003; Henneke 2012). In the absence of RNase HII (Figure 1.3(b)), complete removal of the initiator RNA from Okazaki fragments relies on the coordinated action of synthesis and strand displacement by the replicative DNA polymerase (B or D family), as well as FEN1.
The length of the displaced single strand is maintained at a minimal size, so as to avoid exposure to various rearrangements or degradation (Henneke 2012). Occasionally, strand-displacement activity by DNA polymerase can become ineffective. In this case, degradation of the DNA-hybridized initiator RNA would rely on activation of the 5′–3′ exonuclease function of FEN1. Although not reported in archaea, this maturation mechanism could be similar to the nick translation process encountered in eukaryotes and bacteria (Lundquist and Olivera 1982; Stodola and Burgers 2016).
The three-dimensional structure of FEN1, alone or in the presence of its substrate, has been obtained in various archaea (Hosfield et al. 1998; Chapados et al. 2004). The catalytic domain of FEN1 resembles that of nucleases and contains an active site with two metal ions. Catalysis relies on a conformational change in the FEN1 helical loop which, after binding to the intermediate structures of the Okazaki fragments, lead to the 5′ cleavage specificity of the fragment (Storici et al. 2002). It would appear that the archaeal FEN1 protein is capable of recognizing, binding and cutting a branched double-stranded fragment with a 1-nucleotide unpaired 3′ end (downstream strand) and a short flap containing the initiator 5′ RNA (upstream strand) (Chapados et al. 2004). Given the importance of FEN1 in the maturation process of Okazaki fragments, it is however surprising that its genetic deletion in archaea (Meslet-Cladiére et al. 2007) does not result in cell death, with the exception of Halobacterium sp. NRC-1 (Berquist et al. 2007). Like in yeast (Storici et al. 2002), it is highly probable that archaea use substitution nucleases. For example, the exonuclease activity of the GINS-associated nuclease (GAN) complex in T. kodakarensis has been suggested to replace the combined action of the two nucleases RNase HII and FEN1 (Burkhart et al. 2017).
1.3.3. DNA fragment assembly by DNA ligase
After complete removal of the initiator RNA from the Okazaki fragments, DNA ligase (LIG1) sutures the newly synthesized DNA strand, replacing the RNA and adjacent DNA fragment (Figure 1.3(a) V and (b) IV).
Catalysis involves the formation of a phosphodiester bond between the 3′-OH carbon of the upstream strand and the 5′-phosphate of the adjacent strand. It involves a cofactor as an energy source, namely, ATP and/or NAD+, the use of which by LIGI is dependent on the type of archaea (Rolland et al. 2004; Zhao et al. 2006). DNA ligase interacts physically with PCNA, which stabilizes it on the DNA duplex (Vijayakumar et al. 2007).
With the exception of archaea that possess heterotrimeric PCNA, such as Saccharolobus solfataricus, it has been suggested that DNA polymerase, FEN1 and DNA ligase compete for the same homotrimeric PCNA-binding sites (Henneke 2012), as described in eukaryotes (Ayyagari et al. 2003).
The three-dimensional structure of LIG1 has been solved in various archaea (Pascal et al. 2006), and the architectural organization of its three domains (the nucleotidyl transferase domain (NT), the oligonucleotide/oligosaccharide folding domain (OB-fold) and the DNA-binding domain (DBD)) strongly resembles that of human LIG1 in complex with DNA (Pascal et al. 2004). Although the conformational dynamics of these three domains differ between archaeal and eukaryotic LIG1s, they are involved in the implementation of catalysis.
In the case of ATP-dependent LIG1, the OB domain transfers AMP to a lysine in the active site of the NT domain during the first step of the reaction (Sriskanda and Shuman 1998). AMP is then transferred from LIGI to the DNA 5′-phosphate group, and LIG1 catalyzes the final ligation step, releasing the AMP.
The DBD domain, which is largely responsible for the physical interaction of DNA ligase with its substrate, enables the three domains (DBD-NT-OB) to completely envelop and accurately position the NT and OB domains involved in catalysis at the 3′-OH and 5′-phosphate ends of the DNA duplex (Pascal et al. 2004).
Figure 1.3.Maturation mechanism of archaeal Okazaki fragments.
Comment on Figure 1.3.– (a) Reaction pathway involving RNase HII. The replicating DNA polymerase family B or D) of the lagging strand replicates DNA in the presence of PCNA, and RNase HII specifically cuts into the initiating RNA (step I and step II). Removal of the mono-ribonucleotide involves the coordinated activities of DNA synthesis and strand displacement by the replicative DNA polymerase (family B or D), as well as cutting of the displaced strand by FEN1 (step III and step IV). DNA ligase forms the phosphodiester bond between the two adjacent DNA strands (step V); (b) RNase HII-independent reactive pathway. DNA polymerase replicates the lagging strand in the presence of PCNA (step I). Complete removal of the initiator RNA involves the coordinated activities of DNA synthesis and strand displacement by the replicative DNA polymerase (family B or D), and cutting of the displaced strand by FEN1 (step II and step III). DNA ligase forms the phosphodiester bond between the two adjacent DNA strands (step IV).
The ligation of DNA ends is an essential step during the maturation process of Okazaki fragments. DNA ligase appears to be an essential protein for archaeal survival (Sarmiento et al. 2013). When multiple copies of genes exist in genomes, only their simultaneous genetic deletion is lethal, indicating that functional redundancy is essential for cell survival (Zhao et al. 2006). Biochemical experiments have shown in vitro that DNA ligase plays a significant role in the Okazaki fragment maturation process, beyond its function of ligating DNA strands. Indeed, LIG1 interacts functionally with DNA polymerases and FEN1, stimulating strand-displacement activity and cleavage activity, respectively (Henneke 2012), with ligation taking place after complete removal of primer ribonucleotides. Unlike the Okazaki fragment maturation process in eukaryotes, ligation in archaea can take place close to the RNA–DNA junction, since the DNA fragment downstream of the RNA is synthesized by a high-fidelity DNA polymerase (Henneke et al. 2005).
1.4. Restarting replication forks
Once the replisome components are in place at the replication fork, DNA synthesis and Okazaki fragment maturation take place as described above. However, this process is never as straightforward. In the course of a cell’s life, DNA can be subjected to numerous natural stresses, both endogenous and exogenous. In particular, cellular metabolism produces reactive oxygen species, which can cause DNA damages. Similarly, exposure of cells to exogenous genotoxic agents, such as ultraviolet radiation or ionizing radiation, is a source of damage. Numerous repair mechanisms exist, enabling an adapted response to each type of lesion. However, some DNA damages are not repaired prior to replication, which can cause a replication fork to stop at the DNA lesions. Other events, such as collision between the replication fork and the transcription machinery, or the presence of repeated regions of DNA capable of forming secondary structures, can also hinder replication. These various events can lead the replication fork to slowdown or even stop. These events are known as replication stress. Replication stress leads to DNA damage and genetic instability, contributing to the development of cancer in higher eukaryotes. Understanding the molecular basis of the cellular response to replication stress is therefore a major challenge. The decade-long study of the response to replication stress in bacteria, then in unicellular eukaryotes and mammalian cells (Carr and Lambert 2013; Michel and Sandler 2017), has revealed a mechanistic link between replication and recombination. Indeed, the restart of replication forks that have stopped involves proteins of the homologous recombination pathway, which can lead to chromosomal rearrangements at replication arrest sites. These studies have enabled us to understand the molecular basis for the stability of arrested replication forks and their restart, depending on the nature of the replication arrest.
The response to replication stress in archaea is still largely unknown. However, a close link between replication and recombination has also been found in archaea. In hyperthermophilic archaea, key homologous recombination proteins, notably the RadA recombinase responsible for strand invasion, are essential for cell survival (Fujikane et al. 2010). In these organisms, this close link between replication and recombination ensures replication fidelity and stability. Indeed, the mismatch repair pathway present in bacteria and eukaryotes does not appear to be present in hyperthermophilic archaea. It could therefore be that mismatches cause replication fork arrests, whose restart would be ensured by the homologous recombination pathway, thus coupling mismatch repair and fork restart (Grogan 2015) (Figure 1.4, steps c and d). As mentioned at the beginning of this chapter, it has also been suggested, in the hyperthermophilic archaea T. kodakarensis, that replication initiation occurs exclusively from homologous recombination intermediates, independently of the presence of an origin of replication associated with a replication initiation protein (Gehring et al. 2017). These intermediates, known as D-loops, correspond to three-branched DNA structures generated after strand invasion by the recombination machinery, onto which the replication machinery can be loaded to restart replication in the event of arrest, or even to initiate replication in the absence of canonical replication initiation at replication origins (Figure 1.4).
Biochemical studies aimed at identifying proteins with activity on branched DNA fork substrates in the hyperthermophilic archaea P. furiosus have identified the Hef protein, a helicase/nuclease of the XPF/MUS81/FANCM family. This protein is present in eukaryotes and archaea, but is absent in bacteria. Most eukaryotes possess at least four members, which are involved in DNA repair, meiotic recombination and/or replication restart (Ciccia et al. 2008). The biochemical characterisation of the P. furiosus hef protein has proposed a role for Hef in the remodeling of arrested replication forks (Komori et al. 2004) (Figure 1.4, steps b and c). Based on these results, in vivo studies in the mesophilic archaea H. volcanii have demonstrated a role for Hef in restarting replication, as well as a role for homologous recombination proteins (Lestini et al. 2010, 2013).
Figure 1.4.Restarting arrested replication forks .
Comment on Figure 1.4.– (a) After replication fork arrest, fork regression leads to pairing of the neosynthesized strands and the formation of a 4-branched DNA structure. A role for the Hef helicase in this remodeling has been proposed (Komori et al. 2004); (b) the action of nucleases (endonuclease or exonuclease), which can be coupled to helicases, restores a three-branch structure on which a replisome can, again, be assembled to enable replication to restart; (c) degradation of the 5′ end by a 5′-3′ exonuclease results in the formation of a single-stranded region with a 3′ end, on which the RadA recombination protein polymerizes to form a filament, while migration of the Holliday junction shifts this four-branch structure; (d) the RadAfilament catalyzes the strand invasion reaction, leading to the formation of a second Holliday junction; (e) resolution of the Holliday junctions restores a three-branch structure, known as the D-loop, on which a replisome can again be assembled to enable replication to restart. The neo-synthesized DNA strands are in red, while the template strands are in blue. The arrow represents the 3′ end. Dotted lines represent strand continuity. The blue rectangles on the template strands represent the region where the arrest occurred.
In archaea, we find a close link between replication and recombination, which seems to be involved in both the initiation of replication and the response to replication stress. A better understanding of these mechanisms in archaea will undoubtedly deepen our understanding of these fundamental processes, which are common to bacteria and eukaryotes alike.
