24.1: DNA Replication

Search Fundamentals of Biochemistry

Introduction

The elucidation of the structure of the double helix by James Watson and Francis Crick in 1953 provided a hint as to how DNA is copied during the process of DNA replication. Separating the strands of the double helix would provide two templates for the synthesis of new complementary strands, but exactly how new DNA molecules were constructed was still unclear. In one model, semiconservative replication, the two strands of the double helix separate during DNA replication, and each strand serves as a template from which the new complementary strand is copied. After replication in this model, each double-stranded DNA includes one parental or “old” strand and one daughter or “new” strand. There were two competing models also suggested: conservative and dispersive, which are shown in Figure \(\PageIndex{1}\).

Figure \(\PageIndex{1}\): Three Models of DNA replication. In the conservative model, parental DNA strands (blue) remained associated in one DNA molecule while new daughter strands (red) remained associated in newly formed DNA molecules. In the semiconservative model, parental strands separated and directed the synthesis of a daughter strand, with each resulting DNA molecule being a hybrid of a parental strand and a daughter strand. In the dispersive model, all resulting DNA strands have regions of double-stranded parental DNA and regions of double-stranded daughter DNA. Figure by Parker, N., et.al. (2019) Openstax

Matthew Meselson and Franklin Stahl devised an experiment in 1958 to test which of these models correctly represents DNA replication, as shown in Figure \(\PageIndex{2}\). They grew the bacterium, Escherichia coli for several generations in a medium containing a “heavy” isotope of nitrogen (¹⁵N) that was incorporated into nitrogenous bases and, eventually, into the DNA. This labeled the parental DNA. The E. coli culture was then shifted into a medium containing ¹⁴N and allowed to grow for one generation. The cells were harvested and the DNA was isolated. The DNA was separated by ultracentrifugation, during which the DNA formed bands according to its density. DNA grown in ¹⁵N would be expected to form a band at a higher density position than that grown in ¹⁴N. Meselson and Stahl noted that after one generation of growth in ¹⁴N, the single band observed was intermediate in position in between DNA of cells grown exclusively in ¹⁵N or ¹⁴N. This suggested either a semiconservative or dispersive mode of replication. Some cells were allowed to grow for one more generation in ¹⁴N and spun again. The DNA harvested from cells grown for two generations in ¹⁴N formed two bands: one DNA band was at the intermediate position between ¹⁵N and ¹⁴N, and the other corresponded to the band of ¹⁴N DNA.

Figure \(\PageIndex{2}\): Meselson and Stahl experimented with E. coli grown first in heavy nitrogen (¹⁵N) then in ¹⁴N. DNA grown in ¹⁵N (blue band) was heavier than DNA grown in ¹⁴N (red band), and sedimented to a lower level on ultracentrifugation. After one round of replication, the DNA sedimented halfway between the ¹⁵N and ¹⁴N levels (purple band), ruling out the conservative model of replication. After a second round of replication, the dispersive model of replication was ruled out. These data supported the semiconservative replication model. Figure by Parker, N., et.al. (2019) Openstax

These results could only be explained if DNA replicates in a semiconservative manner. Therefore, the other two models were ruled out. As a result of this experiment, we now know that during DNA replication, each of the two strands that make up the double helix serves as a template from which new strands are copied. The new strand will be complementary to the parental or “old” strand. The resulting DNA molecules have the same sequence and are divided equally into the two daughter cells.

Think about It: What would have been the conclusion of the Meselson-Stahl experiment if, after the first generation, they had found two bands of DNA?

To synthesize double-stranded DNA, the parental strands must separate so DNA polymerases can copy both strands. As all DNA polymerases synthesize new DNA in a 5' to 3' direction from a 3' to 5' template, different mechanisms are used to faithfully synthesize both parental strands. The general mechanism is shown in Figure \(\PageIndex{3}\).

Figure \(\PageIndex{3}\): The replication fork. Leading-strand synthesis proceeds continuously in the 5' to 3' direction. Lagging-strand synthesis also occurs in the 5' to 3' direction, but in a discontinuous manner. An RNA/DNA primer (labeled in green) initiates leading-strand synthesis and every Okazaki fragment on the lagging strand.

Small RNA primers are needed for the new strands. Short (1000-2000 NT) DNA (Okazaki) fragments are made on the 3'-5' parental strand. Ultimately the RNA primers are degraded and filled, and the Okazaki fragments ligated. We will discuss replication in detail for E. Coli, a model prokaryote, followed by replication in eukaryotes.

Figure \(\PageIndex{4}\) shows a general overview of a DNA "replication fork" from where DNA strand synthesis proceeds..

Figure \(\PageIndex{4}\): General Overview of a DNA Replication Fork. At the origin of replication, topoisomerase II relaxes the supercoiled chromosome. Two replication forks are formed by the opening of the double-stranded DNA at the origin, and helicase separates the DNA strands, which are coated by single-stranded binding proteins to keep the strands separated. DNA replication occurs in both directions. An RNA primer complementary to the parental strand is synthesized by RNA primase and is elongated by DNA polymerase III through the addition of nucleotides to the 3′-OH end. On the leading strand, DNA is synthesized continuously, whereas on the lagging strand, DNA is synthesized in short stretches called Okazaki fragments. RNA primers within the lagging strand are removed by the exonuclease activity of DNA polymerase I, and the Okazaki fragments are joined by DNA ligase. Figure by Parker, N., et.al. (2019) Openstax

DNA Replication in E. Coli

DNA replication has been well studied in bacteria primarily because of the small size of the genome and the mutants that are available. E. coli has 4.6 million base pairs (Mbp) in a single circular chromosome and all of it is replicated in approximately 42 minutes, starting from a single origin of replication and proceeding around the circle bidirectionally (i.e., in both directions), as shown in Figure \(\PageIndex{5}\). This means that approximately 1000 nucleotides are added per second. The process is quite rapid and occurs with few errors. E. coli has a single origin of replication, called oriC, on its one chromosome. The origin of replication is approximately 245 base pairs long and is rich in adenine-thymine (AT) sequences.

Figure \(\PageIndex{5}\): Prokaryotic DNA Replication. Replication of DNA in prokaryotes begins at a single origin of replication, shown in the figure to the left, and proceeds in a bidirectional manner around the circular chromosome until replication is complete. The bidirectional nature of replication creates two replication forks that are actively mediating the replication process. The right-hand figure shows a dynamic model of this process. The red and blue dots represent the incorporation of daughter strand nucleotides during the process of replication. Figures from: Daniel Yuen at David Tribe Derivatives and Catherinea228

Replication Overview - E. Coli

The open regions of DNA that are actively undergoing replication are called replication forks. All the proteins involved in DNA replication aggregate at the replication forks to form a replication complex called a replisome. The initial assembly of the complex that initiates primer synthesis is called the primosome. Table \(\PageIndex{1}\) below show the components that assemble at the replication fork to form the E. Coli replisome.

Table \(\PageIndex{1}\): Enzymes involved in DNA Replication in the prokaryote, E. coli

In E. coli, DNA replication is initiated at the single origin of replication, oriC. Binding of the initiator protein, DnaA, locally unfolds the DNA to form two template ssDNA, which bind DnaB helicase. A DnaB hexamer adds to each strand in a process promoted by DnaC, a helicase loader. The single-stranded DNA binding protein B (SSPB)binds to and protects the rest of the ssDNA, preventing further binding by DnaB. The primase, DnaG, is recruited to the site by the DnaB hexamer and synthesizes the RNA primers. DnaB also recruits DNA polymer III holoenzyme (PolIII HE) which binds through a β clamp. All of the bound proteins collectively form the replisome. An overview of E. Coli replisome is shown in Figure \(\PageIndex{6}\).

Figure \(\PageIndex{6}\): The bacterial replisome. Ilic, S.; Cohen, S.; Singh, M.; Tam, B.; Dayan, A.; Akabayov, B. DnaG Primase—A Target for the Development of Novel Antibacterial Agents. Antibiotics 2018, 7, 72. https://doi.org/10.3390/antibiotics7030072 Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/)

Once assembled, replisomes move in opposite directions from the single oriC in the E. Coli chromosome. They meet at the opposite ends at a termination site (ter) to which Tus proteins are bound that create ‘replication fork traps'. After completion of DNA replication, the newly synthesized genomes are separated and segregated to daughter cells.

An alternative term, the primosomes, is used to describe a subcomplex of the replisome which starts replication of the E. Coli chromosome, as well as some phages and plasmids. It contains 6 proteins including helicases and primases, and catalyzes the movement of the replication form by unwinding and primer synthesis. The motor protein helicases, use ATP to move along the ds-DNA backbone, unraveling it as it proceeds. The human genome has genes for 64 RNA and 31 DNA helicases (about 1% of eukaryotic genes).

Primase and Polymerase activities

The synthesis of both RNA strands by the DnaG primase, and DNA strands by DNA polymerase III holoenzyme (pol III) occurs at each start site for an Okazaki fragment. Both enzymes bind to the conserved carboxy-terminal tail of the single-stranded DNA-binding protein (SSB). It turns out that they can both be bound simultaneously.

The primase (DnaG) has three domains:

• N-terminus that binds the template
• RNA polymerase domain
• C-terminus that binds helicase and the C-terminus of SSB.

Primase is displaced by polIII after about 10 nucleotides have been added to the RNA primer so DNA synthesis can now occur at the 3' end of the primer.

Figure \(\PageIndex{7}\) shows how the primase to polymerase switch is made.

Figure \(\PageIndex{7}\): Schematic representation of the primase-to-polymerase switch during DNA replication in E. coli. Bogutzki, A., Naue, N., Litz, L. et al. Sci Rep 9, 14460 (2019). https://doi.org/10.1038/s41598-019-51031-0. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/

Panel (a): Two primase molecules cooperate in the synthesis of the RNA primer (red).

Panel (b): For elongation of the primer, pol III enters the complex, whereupon primase and pol III are concurrently bound to the primed site, possibly via interactions with the C-termini of an adjacent SSB tetramer.

Panel (c): During pol III-mediated elongation of the primer by several nucleotides, both enzymes stay bound to the template. Only after the primer has been elongated by more than 10 nucleotides, one of the primases is released in the G4ori system. It is presumably the displacement of SSB by pol III that causes the consequent dissociation of primase. Whereas in the G4ori system, the two primases are positioned by hairpin structures that prevent SSB from binding to this part of the origin.  At the E. coli replication fork, primases are brought into contact via their interaction with the replicative helicase DnaB.

What happens if the replication fork does not move to the termination site? If the DNA is damaged or if the replisome falls off of the chromosome, it can rebind and restart using Pri proteins. PriA is a DNA helicase that can bind to replication forks through DNA motifs and through interactions with SSBs. Other proteins involved include PriB, PriC, DnaT, DnaC, DnaB helicase, and DnaG primase as illustrated in Figure \(\PageIndex{8}\).

Figure \(\PageIndex{8}\): Primosome restart assembly. The proposed assembly mechanism is as follows. (i) PriA recognizes and binds to a replication fork, (ii) PriB joins PriA to form a PriA-PriB-DNA ternary complex, (iii) DnaT participates in this nucleocomplex to form a triprotein complex, in which PriB is released from ssDNA due to recruitment of DnaT, (iv) the PriA-PriB-DnaT-DNA quaternary complex loads the DnaB/C complex, and (v) DnaB is loaded on the lagging strand template. Yen-Hua Huang and Cheng-Yang Huang. BioMed Research International (2014). https://doi.org/10.1155/2014/195162. Huang and Cheng-Yang Huang. This is an open access article distributed under the Creative Commons Attribution License,

The primosome and replisome are complicated in structure and in their functional activity. Words go only so far in painting an image of how it works. To help we show a few different images of the replisome of E. Coli below.

The first is shown in Figure \(\PageIndex{9}\).

Figure \(\PageIndex{9}\): Replisome architecture in bacteria. (A) Architecture of the E. coli replisome, derived from in vitro studies and direct observation in vivo. Beattie TR, Reyes-Lamothe R. A. Front Microbiol. 2015 doi: 10.3389/fmicb.2015.00562. Creative Commons Attribution License (CC BY)

In this diagram, the leading strand is shown in the upper right end of the diagram. The central bottom loop shows the lagging strand. The τ₃δδ′ψχ is the clamp loader and the DnaB (red) is hexameric. The ssDNA in the lagging strand loop is bound by ssDNA binding proteins (SSB).

Figure \(\PageIndex{10}\) shows the rebind primosome which is mostly similar to the regular one.

Figure \(\PageIndex{10}\): Mechanisms of helicase loading leading to replisome assembly in E. coli. (A)Recognition and melting of the oriC locus during initiation by DnaA. (B)Recognition of abandoned fork structures during replisome reloading by PriA and PriC. All pathways converge on the loading of the replicative helicase DnaB, which acts as an assembly platform for the remaining replisome components.

Finally, Figure \(\PageIndex{11}\) shows models of DNA polymerase for lagging strand synthesis.

 

 

Figure \(\PageIndex{11}\): Usage of DNA polymerase during lagging strand synthesis. (A)S chematic of the E. coli replisome during the elongation step of an Okazaki fragment. (B )Lagging strand polymerase meets the RNA primer of the previous Okazaki fragment and stops synthesis. (C) Current model of events following completion of an Okazaki fragment. DNA polymerase is released from the β clamp (step 1) and the same molecule rebinds to a new β clamp to start the next Okazaki fragment (step 2). (D)An alternative model based on evidence from T4 and T7 replisomes. After completing the Okazaki fragment, the DNA polymerase detaches from the rest of the replisome (step 1). A new molecule of DNA polymerase is recruited to the replisome (step 2) and engages in the synthesis of a new Okazaki fragment. In this tentative model, a local pool of “spare” polymerases may facilitate their exchange and additional components may exchange along with the polymerase (not depicted)

E. Coli DNA Polymerases

E. Coli has 5 DNA polymerases. DNA polymerase I aids in lagging strand synthesis as it removes the RNA primers and incorporates DNA in its place. DNA polymerase II, may play an editing role following lagging strand synthesis by DNA polymerase I. DNA polymerases I and II also play a role in DNA repair, as do DNA polymerases IV and V.

DNA polymerases are shaped like a right hand in overall shape with three domains named palm, fingers, and thumb. The bottom of the cleft formed by the three domains forms the polymerase active site in the Palm domain, The monomeric nucleotides to be added bind through the finger domain, while the thumb domains facilitates the dissociation of the newly synthesized DNA. These features are illustrated for a polymerase that requires host thioredoxin for a bacteriophage T7 DNA polymerase in Figure \(\PageIndex{12}\).

Figure \(\PageIndex{12}\): Structure of T7 DNA replication complex. Melum 103 - Own work, CC BY-SA 4.0, https://commons.wikimedia.org/w/inde...curid=38408627

DNA Polymerase III

Pol III is a fascinating enzyme. It consists of an αεθ core with both 5'-3' polymerase and 3′−5′ proofreading activities, a β₂ ring-shaped "sliding clamp" that keeps the enzyme on the DNA track (processive) without iteratively jumping off and rebinding (distributive), and a (τ/γ)₃δδ′ψχ clamp loader. The SSB protein has a conserved amphiphilic C-terminus that binds both DnaG (primase) and the χ subunit of the clamp loader. A

After primer addition by DnaG, the β₂clamp of polIII is brought to the end of the primer terminus by the clamp loader, after which α and ε subunits bind the clamp. The holoenzyme can add ∼1000 Nt/s and over 150 kb without falling off. Hence it is a very processive enzyme.

Figure \(\PageIndex{13}\) shows an interactive iCn3D model of the the E. coli replicative DNA polymerase III (alpha, beta₂, epsilon, tau complex) bound to DNA (5FKV)

Figure \(\PageIndex{13}\): E. coli replicative DNA polymerase III (alpha, beta₂, epsilon, tau complex) bound to DNA (5FKV). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...vf4pCugQms7tw8

• N-terminal domains of α (αNTD, residues 1–963, are colored in salmon
• OB (964–1072) on the C-term domain of α (αCTD) colored brown,
• τ-binding domains (TBD, 1173–1160) on the C-term domain of α (αCTD) colored dark salmon,
• ε in yellow
• θ in orange (?)
• β₂ in aquamarine
• τ_C in slate gray
• DNA in spacefill, backbone magenta and purple, bases in CPK colors

Overall, there are significant conformational changes in the DNA Polymerase III complex upon binding to the DNA that cause the tail of the polymerase to move from interacting with the clamp in the DNA-bound state to a position 35 Å away from the clamp in the DNA-free state. It has been hypothesized that this large conformational change may help the polymerase act as a switch to facilitate the lagging strand synthesis. On the lagging strand, the polymerase repositions to a newly primed site every ∼1000 bp. To do so, the polymerase needs to release both clamp and DNA. The switch-like movement of the polymerase tail may play a part in the release and consequent repositioning of the polymerase at the end of the Okazaki fragment.

Video 25.1.1: DNA Binding Induces Large Conformational Changes in the DNA Polymerase III Complex(click link to view). The video shows the linear morphing of the DNA-free to the DNA-bound state showing the large conformation change between the two states. The green subunit is the β-clamp, The α-subunit is shown in orange with the active-site residues in magenta, the α-C-terminal domain (α-CTD shown in brown, the ε-subunit in yellow, and the τ-tail shown in blue. Video from: Fernandez-Liero, R., et al. (2015) eLife 4:e11134

The complex can also proofread the newly synthesized DNA. This requires some conformational changes in the polIII complex, including a rotation/tilt of the dsDNA against the β₂ ring-shaped "sliding clamp". The thumb domain moves between the two DNA strands containing a mismatch and produced a distorted DNA. The epsilon subunit, a nuclease, can reach the mismatched nucleotide and clip it off.

Figure \(\PageIndex{14}\) shows an interactive iCn3D model of the E. coli replicative DNA polymerase III-clamp-exonuclease-theta complex bound to DNA in the editing mode (5M1S)

Figure \(\PageIndex{14}\): E. coli replicative DNA polymerase III-clamp-exonuclease-theta complex bound to DNA in the editing mode (5M1S). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...gedpgkHdqG1PGA

• PolIII α brown,
• PolIII ε in yellow
• PolIII θ in orange
• PolIII β₂ in cyan
• DNA in spacefill, backbone primer in magenta, the template in purple, and bases in CPK colors

DNA Polymerase I

DNA polymerase I, as does polIII, has a 5' to 3' polymerase activity. Also, both have a 3' to 5' exonuclease activity for proofreading as well as a 5'-3' exonuclease to remove RNA primers. It contains three domains, a 5'-3' exonuclease followed by a 3'-5' exonuclease, then the polymerase domain. Selective proteolysis between the first two domains produces the Klenow fragment. In contrast, the 5'-3' exonuclease of polIII is in the separate epsilon subunit.

Figure \(\PageIndex{15}\) shows an interactive iCn3D model of the predicted AlphaFold structure of E. Coli DNA Polymerase I (P00582)

Figure \(\PageIndex{15}\): Predicted AlphaFold structure of E. Coli DNA Polymerase I (P00582). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...YhbSsUtpnkTfz8

https://structure.ncbi.nlm.nih.gov/i...oNSC2GJAsMEUk6

• 5' to 3' exonuclease, 1-323, magenta
• 3' to 5' exonuclease, 324-517, orange
• 5' to 3' polymerase, 521-928, cyan
• Val700-Arg713, Motif A, yellow
• Klenow fragment: 324-928

Key aspartate and glutamates involved in the polymerase active site are shown in sicks and labeled. Motif A is conserved in prokaryotic DNA polymerases. Essential roles of motif A in catalysis include interaction with the incoming dNTP and coordination with two divalent metal ions that are required for the polymerization reaction. Note the distance between the 3' to 5' exonuclease and the 5'-3' polymerase.

Other enzyme activities

DNA Ligase

DNA Ligase enzymes seal the breaks in the backbone of DNA that are caused during DNA replication, DNA damage, or during the DNA repair process. The biochemical activity of DNA ligases results in the sealing of breaks between 5′-phosphate and 3′-hydroxyl termini within a strand of DNA. DNA ligases have been differentiated as being ATP-dependent or NAD⁺-dependent depending on the co-factor (or co-substrate) that is used during their reaction. Typically, more than one type of DNA ligase is found within an organism.

Figure \(\PageIndex{16}\) shows the structure of E. coli LigA in complex with nicked adenylated DNA from PDB 2OWO, visualized by UCSF Chimera. The various domains are indicated by different colors and relate to Pfam domains indicated.

Figure \(\PageIndex{16}\): Structure of DNA ligase. Pergolizzi, G., Wagner, G.K, and Bowater, R.P. (2016) Biosci Rep 36(5) e00391

DNA ligase enzyme is covalently modified by the addition of the AMP moiety to a Lysine residue on the enzyme. The AMP derives from the ATP or NADH cofactor. The downstream 5'-phosphate at the site of the DNA nick is able to mediate a nucleophilic attack on the AMP-enzyme complex, causing the AMP to transfer to the 5'-phosphate position of the DNA. The AMP serves as a good leaving group for the nucleophilic attack of the upstream 3'-OH with the 5'-phosphate to seal the DNA backbone, and release the AMP. DNA ligase can use either adenosine triphosphate (ATP) or nicotinamide adenine dinucleotide (NAD⁺) as a cofactor. Figure \(\PageIndex{17}\) shows a mechanism of the ligation reaction, which is powered by ATP hydrolysis.

Figure \(\PageIndex{17}\): DNA Ligase Reaction. After Showalter, A. (2002). .

 

Figure \(\PageIndex{18}\) shows an interactive iCn3D model of Human DNA Ligase I bound to 5'-adenylated, nicked DNA (1X9N)

Figure \(\PageIndex{18}\): Human DNA Ligase I bound to 5'-adenylated, nicked DNA (1X9N). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...VNgobsRuLTsXZA

The protein has three domains:

• DNA binding domain (DBD), shown in magenta. In contrast to most DNA binding proteins, Ligase I binds to the minor grove around the area of DNA damage.
• Adenylation domain (AdD), shown in cyan, has covalently attached cofactor AMP and key catalytic residues shown as sticks and labeled. It ligates the broken DNA and forms a phosphodiester bond.
• The OB-fold domain (OBD), shown in yellow, facilitates catalysis as it binds and unwinds over the short region.

The DNA strands are as follows:

• ss DNA terminated with dideoxy is shown in green
• ss template DNA is shown in brown
• ss 5'-phosphorylated DNA is shown in gray

The AdD and OBD domains are similar in structure to other covalent nucleotidyltransferases involved in DNA and RNA ligation and capping of messenger RNA. Glu 566, Glu 621, and Arg 573 interact with AMP and probably help determine the specificity of the AMP cofactor (over GTP). Divalent cations are required for catalysis but are not present in the above structure. A E566K mutation leads to severe immunodeficiency. Lys 568 forms the covalent AMP adduct. The dideoxynucleoside in the structure is not optimally positioned for reaction with 5'P of AppDNA. Glu 720 and Glu 621 are highly conserved and presumably involved in metal ion binding.

Zoom into the structure to observe the changes at the 3' end of the dideoxy DNA and 5' end of the phosphorylated DNA. These nucleotides are given the abbreviation X in iCn3D as they are modified.

Topoisomerases

We just studied these in a previous section but here is a review for this section. The unwinding of the double-stranded helix at the replication fork generates winding tension in the DNA in the form of positive supercoils upstream of the replication fork. Enzymes called topoisomerases counteract this by introducing negative supercoils into the DNA in order to relieve this stress in the helical molecule during replication. There are four known topoisomerase enzymes found in E. coli that fall into two major classes, Type I Topoisomerases and Type II Topoisomerases, as shown in Figure \(\PageIndex{19}\). Topoisomerase I and III are Type I topoisomerases, whereas DNA gyrase and Topoisomerase IV are Type II topoisomerases.

Figure \(\PageIndex{19}\): Structures of Type I and Type II Topoisomerase. (A) Crystal structure of a Type I Topoisomerase, shown in blue, bound to DNA, shown in orange and yellow. (B) Crystal structure of a Type II Topoisomerase. Topoisomerase II forms a tetrameric structure, shown in green and blue. ATP cofactors (pink) are shown bound to the enzyme.

Goodsell, D.S. (2015) RCSD PDB-101 Molecule of the Month

Type I Topoisimerase

Type I Topoisomerases relieve tension caused during the winding and unwinding of DNA. One way that they can do this is by making a cut or nick in one strand of the DNA double helix as shown in Figure \(\PageIndex{19}\). The 5'-phosphoryl side of the nicked DNA strand remains covalently bound to the enzyme at a tyrosine residue, while the 3'-end is held noncovalently by the enzyme. The Type I topoisomerases rotate or spin the 3'-end of the DNA around the intact DNA strand. This releases the overwinding in the DNA and effectively releases tension. The enzyme completes the reaction by resealing the phosphodiester backbone or ligating the broken strand back together. Overall, only one strand of the DNA is broken during the reaction mechanism and there is NO requirement of ATP during the reaction. The E. coli Topo I enzyme can only remove negative DNA supercoils, but not positive ones. Thus, this enzyme is not involved in relieving the positive supercoiling caused by the DNA helicase during replication. This is in contrast to eukaryotic Topo I that can relieve both positive and negative supercoiling. Although E. coli Topoisomerase I is not directly involved in relieving the tension caused by DNA replication, it is essential for E. coli viability. It is thought to help balance the actions of the Type II topoisomerases and help maintain optimal supercoiling density within the chromosomal DNA. Thus, Topo I is thought to help maintain the homeostatic balance of chromosome supercoiling within E. coli. Topo III, which is also a Type I Topoisomerase, appears to play a role in the decatenation of the daughter chromosomes during DNA replication, but does not play a role in the relaxation of supercoiling.

Figure \(\PageIndex{19}\): Reaction of Type I Topoisomerases. During the reaction, Type I Topoisomerases nick one strand of the DNA. One end of the nicked DNA is covalently attached to the enzyme, shown in light green in the lower diagrams. The other end is held non-covalently and rotated around the double helix to unwind the supercoiling and relax the DNA. Once supercoiling tension is released, the backbone is resealed and the Topoisomerase I enzyme is released. emixed from: Notahelix and JoKalliauer, and the National Human Genome Research Institute

Type II Topoisomerases have multiple functions within the cell. They can increase or decrease winding tension within the DNA or they can unknot or decatanate DNA that has become tangled with another strand as shown in Figure \(\PageIndex{20}\). It does so by a more dangerous method than their Type I counterparts, by breaking both strands of the DNA during their reaction mechanism. The enzyme is covalently attached to both broken sides while the other DNA helix is passed through the break. The double-stranded break is then resealed.

Figure \(\PageIndex{20}\): Reaction of Type II Topoisomerases. Figure modified fromLaponogov, I., et al. (2018) Nature Communications 9:2579. .

The proposed type II topoisomerase reaction cycle is exemplified by topoisomerase IV. Topoisomerase IV subunits are denoted in grey, cyan, and yellow. The gate or G-DNA is in green and the transported or T-DNA is in mauve. ATP bound to the ATPase domains is denoted by a red dot. In step 1, the G-DNA binds with the enzyme. ATP and the T-DNA segment associated with the enzyme in step 2. In step 3, the G-DNA is cleaved and the T-DNA is passed through the break. Drug-targetable domains within the type II topoisomerase complex are highlighted in subsections A, B, and C with examples on the right-hand side of the figure.

Type II Topoisomerase - D

DNA gyrase is the type II topoisomerase enzyme that is primarily involved in relieving positive supercoiling tension that results due to the helicase unwinding at the replication fork. Type II Topoisomerases, especially Topo IV, also address a key mechanistic challenge that faces the bacterial replisome during the termination of DNA replication. The circular nature of the bacterial chromosome dictates that a pair of replisomes that initiate from a single origin of replication will eventually converge on each other in a head-to-head orientation. Positive supercoiling accumulates between the the two replisomes as they converge, but the activity of DNA gyrase, which normally removes positive supercoils, becomes limited by the decreasing amount of template DNA available. Instead, supercoils may diffuse behind the replisomes, forming precatenanes between newly replicated DNA; in E.coli these must be resolved by Topo IV for chromosome segregation to occur.

Termination of Replication

If starting replication is critically important and obviously highly controlled as illustrated above, then termination of replication must be equally critical, otherwise genome instability would arise. There is one discrete origin of replication in E. Coli, oriC, with a defined sequence. In contrast, there are 10, 23 base-pair, nonpalindromic termination sites (Ter)of slightly different sequences. These bind the termination protein, Tus. The affinity of Tus for the Ter site depends on the Ter sequence and, in general,is tight with a K_D in the picomolar range. There are two types of Ter-Tus complexes, one an open"permissive" conformation that allows replication to continue, and a locked, "nonpermissive" form that stops it. In the nonpermissive conformation, a key and conserved cytosine on the leading strand at a conserved GC base pair is flipped out into a cytosine binding pocket, which you can think of as a "stop sign" for replication.

If you think of the E. Coli circular chromosome as a clock with the oriC at 12 o'clock, there are 5 Ter sequences as the replication fork move counter-clockwise at about 7 o'clock and another 5 as the fork move clockwise at around 5 o'clock. The sequences run in opposite polarity to prevent the left-side replication fork from entering the right-hand side as it moves around the chromosome and vice versa. The replisome displaces the Tus proteins at the permissive Ter sites but stops at the nonpermissive site, where DnaB helicase unwinds the DNA, flipping out the cytosine as the locked conformation forms. These processes are illustrated in Figure \(\PageIndex{21}\).

Figure \(\PageIndex{21}\): DNA replication and Tus-ter termination trap in E. coli. Katie H. Jameson et al., JBC, 297, (2021). DOI:h ttps://doi.org/10.1016/j.jbc.2021.101409. Creative Commons Attribution (CC BY 4.0)

Panel A, E. coli contains a single circular chromosome, which replicates bidirectionally from a single origin (small oval). The direction of replisome travel from the origin is depicted by arrows. Chromosomal midpoint indicated by a straight line. The location of the ter sites on the E. coli chromosome is shown relative to oriC. Permissive orientation is displayed in light blue, nonpermissive orientation is displayed in dark blue.

Panel B, the structure of Tus-ter (PDB ID: 2I06) illustrating the nonpermissive and permissive faces (left) and the “locked” conformation formed by DNA unwinding at the nonpermissive face (right). The cytosine base at position 6 of ter (C6), which flips into a specific binding site on the nonpermissive face of Tus to form the “lock,” is indicated.

Replication can proceed at the Ter site if the replisome is moving from the light blue to dark blue sequences of the TER site. The TER site hence exhibits polarity.

Panel A, E. coli contains a single circular chromosome, which replicates bidirectionally from a single origin (small oval). The direction of replisome travel from the origin is depicted by arrows. Chromosomal midpoint indicated by a straight line. The location of ter sites on the E. coli chromosome is shown relative to oriC. Permissive orientation is displayed in light blue, nonpermissive orientation is displayed in dark blue.

Panel B, structure of Tus-ter (PDB ID: 2I06) illustrating the nonpermissive and permissive faces (left) and the “locked” conformation formed by DNA unwinding at the nonpermissive face (right). The cytosine base at position 6 of ter (C6), which flips into a specific binding site on the nonpermissive face of Tus to form the “lock,” is indicated.

Figure \(\PageIndex{22}\) Crystal structure (PDB code: 2I06) of the locked Tus–Ter complex shows the flipped C(6) base at the non-permissive face (5)

Figure \(\PageIndex{22}\): Crystal structure (PDB code: 2I06) of the locked Tus–Ter complex shows the flipped C(6) base at the non-permissive face (5). Pandey et al. Nucleic Acids Res. 2015 Jul 13; 43(12): 5924–5935. doi: 10.1093/nar/gkv527. Creative Commons Attribution License (http://creativecommons.org/licenses/by-nc/4.0/),

Figure \(\PageIndex{23}\) shows interactive iCn3D models of the Escherichia Coli Replication Terminator Protein (Tus) Complexed With TerA DNA in open (left) or locked form (right).

Escherichia Coli Replication Terminator Protein (Tus) Complexed With TerA DNA (2I05)	Escherichia Coli Replication Terminator Protein (Tus) Complexed With DNA- Locked form (2I06)
(Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?7mc7UTqLBbgRZP8p8	(Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?bScTvhNGAMD456vt9

In the figures, the C6 that is flipped out in the locked nonpermissive form (right) is shown in spacefill and labeled as C324. The Arg 198 alters its orientation to allow the C6 to flip out and form the locked conformational form.

A summary of the process of DNA replication is shown in Video 25.1.2

Click Here to View Video

Video 9.2 Overview of the DNA Replication Process

Video from: Yourgenome, animated by Polymime Animation Company, Ltd

DNA Replication of Extrachromosomal Elements: Plasmids and Viruses

To copy their nucleic acids, plasmids and viruses frequently use variations on the pattern of DNA replication described for prokaryote genomes. We will focus here on one style known as the rolling circle method.

Whereas many bacterial plasmids replicate by a process similar to that used to copy the bacterial chromosome, other plasmids, several bacteriophages, and some viruses of eukaryotes use rolling circle replication as shown in Figure \(\PageIndex{24}\).

Figure \(\PageIndex{24}\): Rolling Circle Replication. The process of rolling circle replication is initiated by a single stranded nick in the DNA. Within prokaryotes, DNA polymerase III is utilized to generate the daughter strand. DNA ligase rejoins nicks in the backbone and enables the initiation of DNA synthesis of the second daughter strand. Figure by Parker, N., et.al. (2019) Openstax.

The circular nature of plasmids and the circularization of some viral genomes on infection make this possible. Rolling circle replication begins with the enzymatic nicking of one strand of the double-stranded circular molecule at the double-stranded origin (dso) site. In bacteria, DNA polymerase III binds to the 3′-OH group of the nicked strand and begins to unidirectionally replicate the DNA using the un-nicked strand as a template, displacing the nicked strand as it does so. Completion of DNA replication at the site of the original nick results in the full displacement of the nicked strand, which may then recircularize into a single-stranded DNA molecule. RNA primase then synthesizes a primer to initiate DNA replication at the single-stranded origin (sso) site of the single-stranded DNA (ssDNA) molecule, resulting in a double-stranded DNA (dsDNA) molecule identical to the other circular DNA molecule.

DNA Replication in Eukaryotes

The Cell Cycle

The cell cycle is an ordered series of events involving cell growth and cell division that produces two new daughter cells. Cells on the path to cell division proceed through a series of precisely timed and carefully regulated stages of growth, DNA replication, and division that produce two genetically identical cells. The cell cycle has two major phases, interphase, and the mitotic phase, as shown in Figure \(\PageIndex{25}\). During interphase, the cell grows and DNA is replicated. During the mitotic phase, the replicated DNA and cytoplasmic contents are separated and the cell divides. Watch this video about the cell cycle: http://openstax.org/l/biocellcyc

Figure \(\PageIndex{25}\): Diagram of the Cell Cycle. Fowler, S., et.al. (2013) Openstax

A cell moves through a series of phases in an orderly manner. During interphase, G₁ involves cell growth and protein synthesis, the S phase involves DNA replication and the replication of the centrosome, and G₂ involves further growth and protein synthesis. The mitotic phase follows interphase. Mitosis is nuclear division during which duplicated chromosomes are segregated and distributed into daughter nuclei. Usually, the cell will divide after mitosis in a process called cytokinesis in which the cytoplasm is divided and two daughter cells are formed.

During interphase, the cell undergoes normal processes while also preparing for cell division. For a cell to move from interphase to the mitotic phase, many internal and external conditions must be met. The three stages of interphase are called G₁, S, and G₂. The first stage of interphase is called the G₁ phase, or first gap, because little change is visible. However, during the G₁ stage, the cell is quite active at the biochemical level. The cell is accumulating the building blocks of chromosomal DNA and the associated proteins, as well as accumulating enough energy reserves to complete the task of replicating each chromosome in the nucleus. Throughout interphase, nuclear DNA remains in a semi-condensed chromatin configuration. In the S phase (synthesis phase), DNA replication results in the formation of two identical copies of each chromosome—sister chromatids—that are firmly attached at the centromere region,as shown in Figure \(\PageIndex{26}\). At this stage, each chromosome is made of two sister chromatids and is a duplicated chromosome. The centrosome is duplicated during the S phase. The two centrosomes will give rise to the mitotic spindle, the apparatus that orchestrates the movement of chromosomes during mitosis. In mammals, the centrosome consists of a pair of rod-like centrioles at right angles to each other. Centrioles help organize cell division. Centrioles are not present in the centrosomes of many eukaryotic species, such as plants and most fungi.

Figure \(\PageIndex{26}\): Human Chromosome Structure. Figure A from: The National Human Genome Research Institute, and Figure B from: The School of Biomedical Sciences Wiki

(A) Shows a spectral karyogram of a normal human female. Humans have a total of 23 pairs of chromosomes for a total of 46. Each pair of chromosomes are referred to as homologous chromosomes as they contain copies of the same gene regions. Each of the homologous pairs of chromosomes is stained the same color. Chromosomes are shown in their condensed, unreplicated state. (B) Shows a schematic diagram of a single chromosome before (lower diagram) and after (upper diagram) replication. Upon replication, the identical copies of the chromosome are called sister chromatids and are linked together at the centromere structure.

Figure A from: The National Human Genome Research Institute, and Figure B from: The School of Biomedical Sciences Wiki

In the G₂ phase, or second gap, the cell replenishes its energy stores and synthesizes the proteins necessary for chromosome manipulation. Some cell organelles are duplicated, and the cytoskeleton is dismantled to provide resources for the mitotic spindle. There may be additional cell growth during G₂. The final preparations for the mitotic phase must be completed before the cell is able to enter the first stage of mitosis. To make two daughter cells, the contents of the nucleus and the cytoplasm must be divided. The mitotic phase is a multistep process during which the duplicated chromosomes are aligned, separated, and moved to opposite poles of the cell, and then the cell is divided into two new identical daughter cells. The first portion of the mitotic phase, mitosis, is composed of five stages, which accomplish nuclear division. The second portion of the mitotic phase, called cytokinesis, is the physical separation of the cytoplasmic components into two daughter cells.

If cells are not traversing through one of the phases of interphase or mitosis, they are said to be in G₀ or a resting state. If cells enter G₀ permanently, they are said to have entered a stage of replicative senescence and will no longer be maintained for long-term viability within the organism.

The progression of cells through the cell cycle requires the coordinated actions of specific protein kinases, known as cyclin-dependent kinases. Cyclin-dependent kinases are usually abbreviated as CDK or CDC proteins. CDK/CDC proteins require the binding of a regulatory cyclin protein to become activated, as shown in Figure \(\PageIndex{27}\). The major cyclin proteins that drive the cell cycle in the forward direction, are expressed only at discrete times during the cell cycle. When activated by a cyclin counterpart, CDK/CDC enzymes phosphorylate downstream targets involved with cell cycle progression. For example, the primary cyclin-CDK complex involved in the initiation of DNA replication during S-phase is the CyclinE-CDK2 complex. CDK2 is activated by the expression and binding of Cyclin E during late G1 phase. This causes CDK2 to phosphorylate downstream targets, including the retinoblastoma tumor suppressor protein, pRb. pRB normally binds and inhibits the activity of transcription factors from the E2F family. Following the release of E2F transcription factors from pRb, E2Fs activate the transcription of genes involved in DNA replication and the progression of cells into S-phase.

Figure \(\PageIndex{26}\): Cyclin Dependent Kinases (CDKs) and their Cyclin Regulatory Subunits. Figure A from: Aleem, E., and Arceci, R.J. (2015) Front. Cell and Dev 3 (16) and Figure B from: Cyclinexpression_waehrend-Zellzyklus

 

Panel (A) shows CDK-cyclin complexes with direct functions in regulating the cell cycle are shown. CDK3/cyclin C drives cell cycle entry from G₀. CDK4/6/cyclin D complexes initiate phosphorylation of the retinoblastoma protein (pRb) and promote the activation of CDK2/cyclin E complex. In late G1, CDK2/cyclin E complex completes phosphorylation and inactivation of pRb, which releases the E2F transcription factors and G1/S transition takes place. DNA replication takes place in S phase. CDK2/cyclin A complex regulates progression through S phase and CDK1/cyclin A complex through G2 phase in preparation for mitosis (M). Mitosis is initiated by CDK1/cyclin B complex.

Panel (B) Shows the cyclical nature of cyclin expression during cell cycle progression. Cyclin abundance is regulated by transcriptional expression and rapid protein degradation. Thus, their biological activity is targeted at very specific time points during the cell cycle progression.

Replication Initiation

Origin organization, specification, and activation in eukaryotes are more complex than in bacterial or archaeal kingdoms and significantly deviate from the paradigm established for prokaryotic replication initiation. The large genome sizes of eukaryotic cells, which range from 12 Mbp in S. cerevisiae to 3 Gbp in humans, necessitates that DNA replication starts at several hundred (in budding yeast) to tens of thousands (in humans) origins to complete DNA replication of all chromosomes during each cell cycle, as shown in Figure \(\PageIndex{27}\).

Figure \(\PageIndex{27}\): Eukaryotic chromosomes are typically linear, and each contains multiple origins of replication. The top figure is a graphic representation of the eukaryotic origins of replication, while the bottom image is a Cryo-electron micrograph image. The figure on the top is from Parker, N. et al. and the figure on the bottom is from Fritensky, B. and Brien, N

With the exception of S.cerevisiae and related Saccharomycotina species, eukaryotic origins do not contain consensus DNA sequence elements but their location is influenced by contextual cues such as local DNA topology, DNA structural features, and chromatin environment. Nonetheless, eukaryotic origin function still relies on a conserved initiator protein complex to load replicative helicases onto DNA during the late M and G1 phases of the cell cycle, a step known as origin licensing. In contrast to their bacterial counterparts, replicative helicases in eukaryotes are loaded onto origin duplex DNA in an inactive, double-hexameric form and only a subset of them (10–20% in mammalian cells) is activated during any given S phase, events that are referred to as origin firing. The location of active eukaryotic origins is therefore determined on at least two different levels, origin licensing to mark all potential origins, and origin firing to select a subset that permits assembly of the replication machinery and initiation of DNA synthesis. The extra licensed origins serve as backup and are activated only upon slowing or stalling of nearby replication forks, ensuring that DNA replication can be completed when cells encounter replication stress. Together, the excess of licensed origins and the tight cell cycle control of origin licensing and firing embody two important strategies to prevent under- and overreplication and to maintain the integrity of eukaryotic genomes.

Human Primosome

In humans,, the primosome contains primase and DNA polymerase α (Polα), and makes RNA-DNA primers to which deoxynucleotides are added by polymerases δ and ϵ. Hence there are two catalytic sites for addition or ribo- and deoxyribonucleotides. The structure of the human primosome and the C-terminal domain of the primase large subunit (p58_C) with bound DNA/RNA duplex is presented below. p58_C coordinates the catalytic activities.

As with other polymerases, primase synthesis of RNA primers has the following steps:

• initiation (rate limiting): primase binds to DNA and makes a dinucleotide RNA;
• elongation, which is not as fast as DNA replication since it is less processive, adding only around 10 nucleotides. These short fragments are moved to Polα where deoxynucleotides are added with inactivation of the primase
• termination.

The structures of the enzymes are as follows:

Human Polα consists of a :

• large catalytic subunit (p180) with a C-terminal p180_C domain with two Zn²⁺ binding modules.
• smaller accessory subunit (p70) with an N-terminal (p70_N), a phosphodiesterase, and oligonucleotide/oligosaccharide-binding (OB) domains.

Human primase consist of

• catalytic (p49)
• regulatory (p58) subunits with two domains, the N-terminal (p58_N), which interacts with p49 and which connects primase and Polα, and a C-terminal (p58_C) which contains an iron-sulfur cluster involved in substrate binding and primase activity.

The structures are shown in Figure \(\PageIndex{28}\).

Figure \(\PageIndex{28}\): Structure of the human primosome hetero-tetramer complex. Baranovskiy et al. JBC, 291, 10006-10020 (2016). DOI: https://doi.org/10.1074/jbc.M116.717405. Creative Commons Attribution (CC BY 4.0)

Panel A shows a schematic representation of the domain organization. The flexibly tethered domains are shown as separate parts. p58_C coordinates the iron-sulfur cluster. Exo* is an exonuclease domain with no associated activity due to the evolutionary substitution of the catalytic amino acid residues; PDE, phosphodiesterase.

Pane B shows the crystal structure of the primosome. Subunits are shown as schematics and colored as in A. The α-carbons of catalytic aspartates are shown as purple spheres.

Figure \(\PageIndex{29}\) shows an interactive iCn3D model of the Human primosome without nucleic acids (5EXR)

Figure \(\PageIndex{29}\): Human primosome without nucleic acids (5EXR). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...12dKTjz7hkcD97

DNA primase small: catalytic (p49) - dark green

DNA primase large: regulatory (p58) subunits. p58 has two distinct domains, N-terminal (p58_N light blue) and C-terminal (p58_C gray/purple), connected with an 18-residue linker (253–270) . p58_N interacts with p49 and connects primase with Polα ), and an iron-sulfur cluster containing p58_C plays an important role in substrate binding and primase activity

DNA polymerase alpha catalytic subunit: large catalytic subunit (p180). has p180core (orange) and linker 1251-1265 then the C-terminal domain (p180_C - blue) connects to small subunit p70. (p180_C) contains Zn1 and Zn2 bind site

DNA polymerase alpha subunit B: smaller accessory subunit (p70) with 3 domains: p70_N (light green) then linker 79-156 BOTH NOT SHOWN IN STRUCTURE), the P70 phosphodiesterase, and oligonucleotide/oligosaccharide-binding (OB) domains (combined magenta).

Figure \(\PageIndex{30}\) shows an interactive iCn3D model of the C-terminal domain of the human DNA primase large subunit with bound DNA template/RNA primer (5F0Q)

Figure \(\PageIndex{30}\): C-terminal domain of the human DNA primase large subunit with bound DNA template/RNA primer (5F0Q). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...jqV6UxcgeYNbM7

The ss-DNA is shown with a pale green backbone while the RNA backbone is shown in magenta. The FeS cluster and a Mg²⁺ ion are shown in the catalytic subunit. The Mg²⁺ is shown interacting with a terminal GTP of the RNA.

Figure \(\PageIndex{31}\) shows an interactive iCn3D model of the catalytic core of human DNA polymerase alpha in a ternary complex with an RNA-primed DNA template and dCTP (4QCL)

Figure \(\PageIndex{31}\): Catalytic core of human DNA polymerase alpha in ternary complex with an RNA-primed DNA template and dCTP (4QCL). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...gYFMnVSVJZiYA6

The ss-DNA is shown with a cyan backbone while the RNA primer backbone is shown in magenta. dCTP is shown in spacefill.

Eukaryotic DNA polymerases

Similar to DNA replication in prokaryotes, DNA replication in eukaryotes occurs in opposite directions between the two new strands at the replication fork. Within eukaryotes, two replicative polymerases synthesize DNA in opposing orientations, as shown in Figure \(\PageIndex{32}\). Polymerase ε (epsilon) synthesizes DNA in a continuous fashion, as it is “pointed” in the same direction as DNA unwinding. Similar to bacterial replication, this strand is known as the leading strand. In contrast, polymerase δ (delta) synthesizes DNA on the opposite template strand in a fragmented, or discontinuous, manner and this strand is termed the lagging strand. The discontinuous stretches of DNA replication products on the lagging strand are known as Okazaki fragments and are about 100 to 200 bases in length at eukaryotic replication forks. Owing to the “lagging” nature, the lagging strand generally contains a longer stretch of ssDNA that is coated by single-stranded binding proteins, which stabilizes ssDNA templates by preventing secondary structure formation or other transactions at the exposed ssDNA. In eukaryotes, ssDNA stabilization is maintained by the heterotrimeric complex known as replication protein A (RPA) (Figure 9.19). Each Okazaki fragment is preceded by an RNA primer, which is displaced by the procession of the next Okazaki fragment during synthesis. In eukaryotic cells, a small amount of the DNA segment immediately upstream of the RNA primer is also displaced, creating a flap structure. This flap is then cleaved by endonucleases (such as Fen1, discussed later). At the replication fork, the gap in DNA after removal of the flap is sealed by DNA ligase I. Owing to the relatively short nature of the eukaryotic Okazaki fragment, DNA replication synthesis occurring discontinuously on the lagging strand is less efficient and more time consuming than leading-strand synthesis.

Figure \(\PageIndex{32}\): The Eukaryotic Replisome Complex Coordinates DNA Replication. Lemanm A.R. and Noguchi, E. (2013) Genes 4(1):1-32.

Replication on the leading and lagging strands is performed by Pol ε and Pol δ, respectively. Many replisome factors (including the FPC [fork protection complex], Claspin, And1, and RFC [the replication factor C clamp loader]) are charged with regulating polymerase functions and coordinating DNA synthesis with the unwinding of the template strand by Cdc45-MCM [mini-chromosome maintenance]-GINS [go-ichi-ni-san]. The replisome also associates with checkpoint proteins as DNA replication and genome integrity surveillance mechanisms.

Figure \(\PageIndex{33}\) shows an interactive iCn3D model of the Core human replisome (7PFO). (long load time)

Figure \(\PageIndex{33}\): Core human replisome (7PFO). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...XQQijVnottcwa7 (long load time)

The leading DNA strand backbone is shown in spacefill magenta while the lagging strand backbone is shown in cyan. The DNA bases are shown as CPK spheres. The ATP analog, phosphaminophosphonic acid-adenlate ester, is shown in spacefill with CPK colors and labeled. The C-alpha traces of the different protein subunits are all shown in different colored alpha-C traces, except the DNA polymerase epsilon catalytic subunit A which is shown in cartoon form and colored by secondary structure. (long load time)

At the eukaryotic replication fork, three distinct replicative polymerase complexes contribute to canonical DNA replication: α, δ, and ε. These three polymerases are essential for the viability of the cell. Because DNA polymerases require a primer on which to begin DNA synthesis, first, polymerase α (Pol α) acts as a replicative primase. Pol α is associated with an RNA primase and this complex accomplishes the priming task by synthesizing a primer that contains a short ~10-nucleotide RNA stretch followed by 10 to 20 DNA bases. Importantly, this priming action occurs at replication initiation at origins to begin leading-strand synthesis and also at the 5' end of each Okazaki fragment on the lagging strand.

However, Pol α is not able to continue DNA replication. From in vitro studies, it was observed that DNA replication must be “handed off” to another polymerase to continue synthesis. The polymerase switching requires clamp loaders. Initially, it was thought that Pol δ performed leading-strand replication and that Pol α completed each Okazaki fragment on the lagging strand. Using mutator polymerase variants and mapping nucleotide misincorporation events, Kunkel and colleagues found that Pol ε and Pol δ mutations lead to mismatched nucleotide incorporation only on the leading and lagging strands, respectively. Thus, normal DNA replication requires the coordinated actions of three DNA polymerases: Pol α for priming synthesis, Pol ε for leading-strand replication, and Pol δ for generating Okazaki fragments during lagging-strand synthesis.

In eukaryotes, DNA polymerases are grouped into seven families (A, B, C, D, X, Y, and RT). Crystal structures of the three nuclear replicative DNA polymerases demonstrate that they belong to the B family (Figure 25.1.17). All three replicative DNA polymerases are multi-subunit enzymes as shown in Table \(\PageIndex{2}\) below.

Table 25.1.2 Subunits of the Major Eukaryotic Replicative DNA Polymerases

Table \(\PageIndex{2}\): Subunits of the Major Eukaryotic Replicative DNA Polymerases. Doublié, S. and Zahn, K.E. (2014) Front. Microbiol 5:444

All B family polymerases are composed of five subdomains, the fingers, thumb, and palm which constitute the core of the enzyme, as well as an exonuclease domain and an N-terminal domain (NTD). The palm, a highly conserved fold composed of four antiparallel β strands and two helices, harbors two strictly conserved catalytic aspartates, located in motif A, DXXLYPS and motif C, DTDS , as shown in Figure \(\PageIndex{34}\).

Figure \(\PageIndex{34}\): Structure of polymerases. Doublié, S. and Zahn, K.E. (2014) Front. Microbiol 5:444

This fold is shared by a very large group of enzymes, including DNA and RNA polymerases, reverse transcriptases, CRISPR polymerase, and even reverse (3′–5′) transferases. In contrast, the thumb and the fingers subdomains exhibit substantially more structural diversity. The fingers undergo a conformational change upon binding DNA and the correct incoming nucleotide. This movement allows residues in the finger subdomain to come in contact with the nucleotide in the nascent base pair. The thumb holds the DNA duplex during replication and plays a part in processivity. The exonuclease domain carries a 3′–5′ proofreading activity, which removes misincorporated nucleotides. The NTD seems to be devoid of catalytic activity. In pol δ the NTD comprises three motifs, one has a topology resembling an OB fold, one a single-stranded DNA binding motif, and the another has a RNA-binding motif (RNA Recognition Motif or RRM). The NTD likely plays a role in polymerase stability and fidelity through its interactions with other domains.

DNA polymerases require additional factors to support DNA replication in vivo. DNA polymerases have a semi-closed hand structure, which allows them to load onto DNA and translocate. This structure permits DNA polymerase to hold the single-stranded template, incorporate dNTPs at the active site, and release the newly formed double strand. However, the conformation of DNA polymerases does not allow for their stable interaction with the template DNA. To strengthen the interaction between template and polymerase, DNA sliding clamps have evolved, promoting the processivity of replicative polymerases. In eukaryotes, this sliding clamp is a homotrimer known as proliferating cell nuclear antigen (PCNA), which forms a ring structure. The PCNA ring has polarity with a surface that interacts with DNA polymerases and tethers them securely to DNA. PCNA-dependent stabilization of DNA polymerases has a significant effect on DNA replication because it enhances polymerase processivity up to 1,000-fold (Figure 25.1.19).

The DNA helicases (MCM proteins) and polymerases must also remain in close contact at the replication fork (Figure 25.1.19). If unwinding occurs too far in advance of synthesis, large tracts of ssDNA are exposed. This can activate DNA damage signaling or induce aberrant DNA repair processes. To thwart these problems, the eukaryotic replisome contains specialized proteins that are designed to regulate the helicase activity ahead of the replication fork. These proteins also provide docking sites for physical interaction between helicases and polymerases, thereby ensuring that duplex unwinding is coupled with DNA synthesis.

Control of Origin Firing

Origin usage in eukaryotes can be dynamic, with origin firing at different sites depending on cell type and developmental stage. Nevertheless, the mechanism of replisome assembly and origin firing is highly conserved. During late mitosis and G₁ phase, cell cycle proteins, such as Cdc6, associate with Ori sites throughout the genome and recruit the helicase enzymes, MCMs 2-7 as shown in Figure \(\PageIndex{35}\). At this time, double hexamers of the MCM2-7 complex are loaded at replication origins. This generates a pre-replication complex (pre-RC). Origins with an associated pre-RC are considered licensed for replication. Licensed replication origins can then be “fired,” when replication actually initiates at the Ori. Origin firing is brought about by multiple phosphorylation events carried out by the cyclin E-CDK2 complex at the onset of S phase and by other cyclin-dependent kinases (CDKs) prior to individual origin firing (Figure \(\PageIndex{35}\)). Cyclin-dependent kinases (CDKs) are the families of protein kinases first discovered for their role in regulating the cell cycle. They are also involved in regulating transcription, mRNA processing, and the differentiation of nerve cells. CDKs are activated through the binding of an associated cyclin regulatory protein. Without a cyclin, CDKs exhibit little kinase activity. Following the phosphorylation of the pre-RC, origin melting occurs and DNA unwinding by the helicase generates ssDNA, exposing a template for replication (Figure \(\PageIndex{35}\)). The replisome then begins to form with the localization of replisome factors such as Cdc45. DNA synthesis begins on the melted template, and the replication machinery translocates away from the origin in a bidirectional manner.

Figure \(\PageIndex{35}\): MCM2-7 loads onto DNA at replication origins during G1 and unwinds DNA ahead of replicative polymerases. Figure from: Lemanm A.R. and Noguchi, E. (2013) Genes 4(1):1-32

Pane (A) shows the combined activities of Cdc6 and Cdt1 bring MCM complexes (shown as blue circles of varying shades) to replication origins.

Panel (B) shows CDK/DDK-dependent phosphorylation of pre-RC components leads to replisome assembly and origin firing. Cdc6 and Cdt1 are no longer required and are removed from the nucleus or degraded

Panel (C) shows MCMs and associated proteins (GINS and Cdc45 are shown) unwinding DNA to expose template DNA. At this point, replisome assembly can be completed and replication initiated. “P” indicates phosphorylation.

Replication through Nucleosomes

Eukaryotic genomes are substantially more complicated than the smaller and unadorned prokaryotic genomes. Eukaryotic cells have multiple noncontiguous chromosomes, each of which must be compacted to allow packaging within the confined space of a nucleus. As seen in chapter 4, chromosomes are packaged by wrapping ~147 nucleotides (at intervals averaging 200 nucleotides) around an octamer of histone proteins, forming the nucleosome. The histone octamer includes two copies each of histone H2A, H2B, H3, and H4. In chapter 8, it was highlighted that histone proteins are subject to a variety of post-translational modifications, including phosphorylation, acetylation, methylation, and ubiquitination that represent vital epigenetic marks. The tight association of histone proteins with DNA in nucleosomes suggests that eukaryotic cells possess proteins that are designed to remodel histones ahead of the replication fork, in order to allow smooth progression of the replisome. It is also essential to reassemble histones behind the fork to reestablish the nucleosome conformation. Furthermore, it is important to transmit the epigenetic information found on the parental nucleosomes to the daughter nucleosomes, in order to preserve the same chromatin state. In other words, the same histone modifications should be present on the daughter nucleosomes as on the parental nucleosomes. This must all be done while doubling the amount of chromatin, which requires the incorporation of newly synthesized histone proteins. This process is accomplished by histone chaperones and histone remodelers, which are discussed below and shown in Figure \(\PageIndex{36}\).

Figure \(\PageIndex{36}\): Nucleosome displacement and deposition during DNA replication. Lemanm A.R. and Noguchi, E. (2013) Genes 4(1):1-32

Histones are removed from chromatin ahead of the replication fork. FACT may facilitate this process. Asf1 recruits histone H3-H4 dimers to the replication fork. CAF-1 and Rtt106 load newly synthesized (light purple) histones to establish chromatin behind the fork. Previously loaded histones (dark purple) are also deposited on both daughter DNA strands. The histone chaperones involved in these processes are associated with replisome proteins: CAF-1/Rtt106 with PCNA and FACT/Asf1 with MCMs.

Several histone chaperones are known to be involved in replication-coupled nucleosome assembly, including the FACT complex. The FACT complex components were originally identified as proteins that greatly stimulate transcription by RNA polymerase II. In budding yeast, FACT was found to interact with DNA Pol α-primase complex, and the FACT subunits were found to interact genetically with replication factors. More recently, studies showed that FACT facilitates DNA replication in vivo and is associated with the replisome in budding yeast and human cells. The FACT complex is a heterodimer that does not hydrolyze ATP, but facilitates the “loosening” of histones in nucleosomes

Replication Fork Barriers and the Termination of Replication

In prokaryotes, such as the E. coli, bidirectional replication initiates at a single replication origin on the circular chromosome and terminates at a site approximately opposed from the origin. This replication terminator region contains DNA sequences known as Ter sites, polar replication terminators that are bound by the Tus protein. The Ter-Tus complex counteracts helicase activity, resulting in replication termination. In this way, prokaryotic replication forks pause and terminate in a predictable manner during each round of DNA replication.

In eukaryotes, the situation differs. Replication termination typically occurs by the collision of two replication forks anywhere between two active replication origins. The location of the collision can vary based on the replication rate of each of the forks and the timing of origin firing. Often, if a replication fork is stalled or collapsed at a specific site, replication of the site can be rescued when a replisome traveling in the opposite direction completes copying the region. However, there are numerous programmed replication fork barriers (RFBs) and replication “challenges” throughout the genome. To efficiently terminate or pause replication forks, some fork barriers are bound by RFB proteins in a manner analogous to E. coli Tus. In these circumstances, the replisome and the RFB proteins must specifically interact to stop replication fork progression.

Telomeres and Replicative Senescence

The End Replication Problem

We have discussed the structure of telomers in the previous section. Let's look now at their activity/function. In humans, telomeres consist of hundreds to thousands of repetitive sequences of TTAGGG at chromosomal ends for maintaining genomic integrity. Because the DNA replication is asymmetric along double strands, RNA primer sequence at the 3′-hydroxyl end cannot be replaced by DNA polymerase I, as there is no 3'-OH primer group present for the polymerase to extend the DNA chain. This causes the loss of 30–200 nucleotides with each DNA replication and cell division and is known as the end replication problem. Telomeres provide a repetitive noncoding sequence of DNA at their 3′ en, to prevent the loss of critical genetically encoded information during replication. Moreover, telomeres are coated with a complex of six capping proteins, also known as shelterin proteins, which are packed into a compact T-loop structure that hides the ends of the chromosomes. This prevents the DNA repair machinery from mistaking chromosomal ends for double-stranded DNA breaks, as shown in Figure \(\PageIndex{37}\). Therefore, telomeres have been proposed as a mitotic clock that measures how many times a cell has divided and in essence, gives a cell a defined lifetime.

Figure \(\PageIndex{37}\): Telomere Structure. Image (A) by: MBInfo and Image (B) by:Thomas Splettstoesser

Pane (A): shows telomeres located at the end of chromosomes, where they help protect against the loss of DNA during replication.

Panel (B) shows DNA quadruplex formed by telomere repeats. The looped conformation of the DNA backbone is very different from the typical DNA helix, this is known as T-loop formation. The green spheres in the center represent potassium ions.

The human telomerase enzyme is responsible for maintaining and elongating telomeres and consists of an RNA component (TERC) and a reverse transcriptase (TERT), that serves as the catalytic component, as shown in Figure \(\PageIndex{38}\). The TERT uses the TERC as a template to synthesize new telomeric DNA repeats at a single-stranded overhang to maintain telomere length (Figure 25.1.26). Some cells such as germ cells, stem cells, hematopoietic progenitor cells, activated lymphocytes, and most cancer cells constitutively express telomerase and maintain telomerase activity to overcome telomere shortening and cellular senescence. However, most other somatic cells generally have a low or undetectable level of telomerase activity and concomitantly limited longevity. Interestingly, overall telomerase activity decreases with age, but increases markedly in response to injury, suggesting a role for telomerase in cellular regeneration during wound healing. The telomere length and integrity are regulated through the interplay between the telomerase and shelterin proteins.

Figure \(\PageIndex{37}\): Conceptual Model of Telomerase Activity. Abbexa Ltd.

The active site of the telomerase enzyme contains the RNA template, TERC (shown in red) and aligns with the last few telomeric bases at the end of the chromosome (shown in blue). This creates a single-stranded overhang that can be used as a template by the TERT reverse transcriptase to extend the telomere sequence.

In vivo, shortened telomeres and damaged telomeres generally caused by reactive oxygen species (ROS) are usually assumed to be the main markers of cellular aging and are thought to be the main cause of replicative senescence. In vitro, telomeres lose approximately 50–200 bp at each division due to the end-replication problem. Approximately 100 mitoses are thought to be sufficient to reach the Hayflick limit, or the maximum number of mitotic events allowed prior to entering replicative senescence. Cells in continual renewal, such as blood cells, compensate for telomere erosion by expressing telomerase, the only enzyme able to polymerize telomeric sequences de novo at the extremity of telomeres. Knocking out telomerase components, such as the catalytic subunit (TERT) or the RNA template (TERC), induces several features of aging in mice. In humans, germline mutations in telomerase subunits are responsible for progeroïd syndromes, such as Dyskeratosis congenita, a rare genetic form of bone marrow failure. Furthermore, healthy lifespan in humans is positively correlated with longer telomere length and patients suffering from age-related diseases and premature aging have shorter telomeres compared with healthy individuals. An accumulation of unrepaired damage within telomeric regions has also been shown to accumulate in aging mice and non-human primates, suggesting that damage of telomeres with age may also be contributing to age-driven disease states and reduced health span.

Thus, one could argue that the activation and expression of telomerase may be a way of reducing age-related diseases and increasing overall longevity. However, the constitutive expression of telomerase, unfortunately, is a characteristic of almost all cancer cells. It is therefore, no surprise that transgenic animals over-expressing the catalytic subunit of telomerase (mTERT), develop cancers earlier in life. However, over-expression of telomerase in mice that are highly resistant to cancers has shown large increases in median lifespan and significantly reduced age-associated disorders. Since humans are not highly resistant to cancer, this is not a feasible option for humans. However, additional studies in mice, where constitutive expression of telomerase is only introduced into a small percentage of host cells using adenovirus gene therapy techniques has yielded more promising results. Adenoviruses are a group of viruses that form an icosahedral protein capsid that houses a linear double-stranded DNA genome. Infections in humans typically cause symptoms of the common cold and are usually mild in nature. These are a good target for gene therapy, as the DNA that they carry can be mutated, so that they are deficient in their ability to replicate once they have infected the host. They can also be transformed to carry a gene of interest into the host, where that gene can then integrate into the host genome. Experiments in mice that were infected with an adenovirus carrying the mTERT gene showed that mTERT was delivered to a wide range of tissues within the body, and increased telomere length within those tissues. Furthermore, the mTERT expressing mice were healthier than their litter mates and displayed a reduction in disabling conditions associated with physiological aging such as osteoporosis and insulin resistance, as shown in Figure \(\PageIndex{38}\). Cognitive skills and metabolic functions were also improved. Noticeably, mice treated with gene therapy did not have increased incidence in cancer rates, suggesting that in at least for the short-lived mouse species, gene therapy approach to increased telomerase activity is safe. Within these animals, the median lifespan was increased by 24% when animals were treated at 1 year of age, and by 13% if treated at 2 years of age.

Figure \(\PageIndex{38}\): Promoting healthspan in Mice using a Telomerase Gene Therapy. Delivery of the catalytic subunit of telomerase (TERT) using a modified adenovirus vector (rAAV) suppresses aging-associated telomere erosion and extends short telomeres in a variety of tissues. Consequently, animals display improved healthspans and extended lifespans. Boccardi, V. and Herbig, U. (2012) EMBO Mol Med 4:685-687.

 

Replication and Repair of Telomere Sequences

In addition to the end replication problem, telomeric DNA (telDNA) replication and repair is a real challenge due to the different structural features of telomeres. First, the nucleotide sequence itself consists of a hexanucleotide motif (TTAGGG) repeated over kilobases, with the 5′-3′ strand named the “G-strand” due to its high content in guanine. During the progression of the replication fork, the lagging strand, corresponding to the G-strand, forms G-quadruplex (G4) structures, which have to be resolved to allow fork progression and to complete replication, as shown in Figure \(\PageIndex{39}\). Secondly, R-loops corresponding to highly stable RNA:DNA hybrids, involving the long non-coding telomeric transcript TERRA (telomeric repeat-containing RNA) also have to be dissociated. Thirdly, the extremity of telomeres adopts a specific loop structure, the T-loop, which has to be unraveled. This is the loop that hides the double-stranded end from the DNA damage sensors, and is locked by the hybridization of the 3′ single strand overhang extremity with the above 3′-5′ strand, thereby displacing the corresponding 5′-3′ strand to form a D-loop (displacement loop) structure (Figure \(\PageIndex{38}\)). Lastly, replication also has to deal with barriers encountered elsewhere in the genome, such as torsions and a condensed heterochromatic environment.

Figure \(\PageIndex{39}\): Obstacles and solutions to replicate telomeres. Billiard, P. and Poncet, D.A. (2019) Int J. Mol. Sci. 20(19) 4959

Panel (a) shows the Telomeric sequence, with the G-strand in a solid red line and the C-strand in a solid green line, is depicted. The terminal D-loop structuring the much larger T-loop is stabilized by the shelterin complex. The replisome (PCNA, pol ε, etc) polymerizes a new G-strand (depicted in a dotted red line) and frees the parental G-strand, enabling the formation of G4 secondary structure. R-loops corresponding to TERRA hybridization (in dotted black lines) with the 3'-5' strand and torsions due to the fork progression are also shown.

Panel (b) shows replication helpers, such as helicases, either helping in G4 unwinding or in D-loop unlocking are depicted. The DNAses (Top2a, DNA2) and RNAses (RNAse H1 and FEN1) help in resolving torsions and RNA:DNA heteroduplexes, while Timeless stimulates the replisome and POT1 competes with RPA1 for binding of the single-strand and helps in G4 dissolution. The shelterin components, POT1, TRF1, and TRF2 help in loading the helper proteins (fine green arrows)

Since telomeres face a host of obstacles to completing the replication process, as discussed in Figure 25.1.28, the cell possesses a set of specialized machinery to fully achieve their replication, such as the RTEL1, TRF1, and TRF2 proteins, DNAses, RNAsses, and Timeless. The recruitment of these factors is orchestrated by the shelterin complex.

At the molecular level, the GGG telomeric repeats are particularly sensitive to ROS, which produce stretches of 8-oxoguanine that are especially difficult to repair. Coupled with inefficient telomere repair, these ROS-induced lesions produce single and double-strand breaks, and/or generate replicative stress, ultimately resulting in telomere shortening. The presence of unrepaired single or tandem 8-oxoguanine drastically inhibits the binding of TRF1 and TRF2, and impairs the recruitment of telomerase, especially when ROS damage is localized in the 3′ overhang. This type of damage contributes to telomere deprotection and shortening. Strikingly, ROS (and other metabolic stresses) also induce the relocation of TERT to mitochondria, as observed (i) in primary neurons after oxidative stress; (ii) in neurons exposed to the tau protein; (iii) in Purkinje neurons subjected to excitotoxicity; and (iv) in cancer cell lines treated with a G4 ligand. Mitochondrial TERT increases the inner membrane potential, as well as the mtDNA copy number, and decreases ROS production with a protective effect on mtDNA. Mitochondria are also critical sensors of cellular damage and contribute to the processes of autophagy and apoptosis (programmed cell death). The relocalization of TERT following chromosomal damage in the nucleus, may indicate one mechanism the mitochondria utilizes to monitor cellular stress and damage.

Replication of Mitochondrial DNA

Mammalian mitochondria contain multiple copies of a circular, double-stranded DNA genome approximately 16.6 kb in length, as shown in Figure \(\PageIndex{40}\). The two strands of mtDNA differ in their base composition, with one being rich in guanines, making it possible to separate a heavy (H) and a light (L) strand by density centrifugation. The mtDNA contains one longer noncoding region (NCR) also referred to as the control region. In the NCR, there are promoters for polycistronic transcription, one for each mtDNA strand; the light strand promoter (LSP) and the heavy strand promoter (HSP). The NCR also harbors the origin for H-strand DNA replication (O_H). A second origin for L-strand DNA replication (O_L) is located outside the NCR, within a tRNA cluster.

Figure \(\PageIndex{40}\): Map of human mtDNA. Falkenberg, M. (2018) Essays Biochem 62(3):287-296

Falkenberg, M. (2018) Essays Biochem 62(3):287-296

As shown in Figure \(\PageIndex{40}\), the genome encodes for 13 mRNA (green), 22 tRNA (violet), and 2 rRNA (pale blue) molecules. There is also a major noncoding region (NCR), which is shown enlarged at the top in blue. The major NCR contains the heavy strand promoter (HSP), the light strand promoter (LSP), three conserved sequence boxes (CSB1-3, orange), the H-strand origin of replication (O_H), and the termination-associated sequence (TAS, yellow). The triple-stranded displacement-loop (D-loop) structure is formed by a premature termination of nascent H-strand DNA synthesis at TAS. The short H-strand replication product formed in this manner is termed 7S DNA. A minor NCR, located approximately 11,000 bp downstream of O_H, contains the L-strand origin of replication (O_L).

A dedicated DNA replication machinery is required for its maintenance. Mammalian mtDNA is replicated by proteins distinct from those used for nuclear DNA replication and many are related to replication factors identified in bacteriophages. DNA polymerase γ (POLγ) is the replicative polymerase in mitochondria. In human cells, POLγ is a heterotrimer with one catalytic subunit (POLγA) and two accessory subunits (POLγB). POLγA belongs to the A family of DNA polymerases and contains a 3′–5′ exonuclease domain that acts to proofread the newly synthesized DNA strand. POLγ is a highly accurate DNA polymerase with a frequency of misincorporation lower than 1 × 10⁻⁶. The accessory POLγB subunit enhances interactions with the DNA template and increases both the catalytic activity and the processivity of POLγA. The DNA helicase TWINKLE travels in front of POLγ, unwinding the double-stranded DNA template. TWINKLE forms a hexamer and requires a fork structure (a single-stranded 5′-DNA loading site and a short 3′-tail) to load and initiate unwinding. Mitochondrial single-stranded DNA-binding protein (mtSSB) binds to the formed ssDNA, protects it against nucleases, and prevents secondary structure formation

The most accepted model of DNA replication in the mitochondria is the strand displacement model, as shown in Figure \(\PageIndex{41}\). Within this model, DNA synthesis is continuous on both the H- and L-strand. There is a dedicated origin for each strand, O_H and O_L. First, replication is initiated at O_H and DNA synthesis then proceeds to produce a new H-strand. During the initial phase, there is no simultaneous L-strand synthesis and mtSSB covers the displaced, parental H-strand. By binding to single-stranded DNA, mtSSB prevents the mitochondrial RNA polymerase (POLRMT) from initiating random RNA synthesis on the displaced strand. When the replication fork has progressed about two-thirds of the genome, it passes the second origin of replication, O_L. When exposed in its single-stranded conformation, the parental H-strand at O_L folds into a stem–loop structure. The stem efficiently blocks mtSSB from binding and a short stretch of single-stranded DNA in the loop region, therefore remains accessible, allowing POLRMT to initiate RNA synthesis. POLRMT is not processive on single-stranded DNA templates. After adding approximately 25 nucleotides, it is replaced by POLγ and L-strand DNA synthesis is initiated. From this point, H- and L-strand synthesis proceeds continuously until the two strands have reached full circle. Replication of the two strands is linked, since H-strand synthesis is required for the initiation of L-strand synthesis. DNA Ligase III is used to complete the ligation of the newly formed DNA strands.

Figure 25.1.24 Replication of the human mitochondrial genome. Mitochondrial DNA replication is initiated at O_H and proceeds unidirectionally to produce the full-length nascent H-strand. mtSSB binds and protects the exposed, parental H-strand. When the replisome passes O_L, a stem–loop structure is formed that blocks mtSSB binding, presenting a single-stranded loop-region from which POLRMT can initiate primer synthesis. The transition to L-strand DNA synthesis takes place after about 25 nt, when POLγ replaces POLRMT at the 3′-end of the primer. Synthesis of the two strands proceeds in a continuous manner until two full, double-stranded DNA molecules have been formed.Figure from: Falkenberg, M. (2018) Essays Biochem 62(3):287-296

During DNA replication, the parental molecule remains intact, which poses a steric problem for the moving replication machinery. Topoisomerases belonging to the type 1 family can relieve torsional strain formed in this way, by allowing one of the strands to pass through the other. In mammalian mitochondria, TOP1MT a type IB enzyme can act as a DNA “swivel”, working together with the mitochondrial replisome. Furthermore, replication of circular DNA often causes the formation of catenanes, or interlocked circles that need to be separated from one another. The type 1A topoisomerase, topoisomerase 3α (Top3α), is required to resolve the hemicatenane structure that can form during mtDNA replication.

Curiously, not all replication events initiated at O_H continue to full circle. Instead, 95% are terminated after about the first 650 nucleotides at a sequence known as the termination associated sequences (TAS) (Figure 25.1.23). This creates a short DNA fragment known as the 7S DNA, that remains bound to the parental L-strand, while the parental H-strand is displaced (Figure 25.1.23). As a result, a triple-stranded displacement loop structure, a D-loop, is formed. The functional importance of the D-loop structure is unclear and how replication is terminated at TAS is also not known.

 

Mastering the Content

Which of the following is the enzyme that replaces the RNA nucleotides in a primer with DNA nucleotides?

1. DNA polymerase III
2. DNA polymerase I
3. primase
4. helicase

[reveal-answer q=”628075″]Show Answer[/reveal-answer]
[hidden-answer a=”628075″]Answer b. DNA polymerase I is the enzyme that replaces the RNA nucleotides in a primer with DNA nucleotides.[/hidden-answer]

Which of the following is not involved in the initiation of replication?

1. ligase
2. DNA gyrase
3. single-stranded binding protein
4. primase

[reveal-answer q=”820951″]Show Answer[/reveal-answer]
[hidden-answer a=”820951″]Answer a. Ligase is not involved in the initiation of replication.[/hidden-answer]

Which of the following enzymes involved in DNA replication is unique to eukaryotes?

1. helicase
2. DNA polymerase
3. ligase
4. telomerase

[reveal-answer q=”650146″]Show Answer[/reveal-answer]
[hidden-answer a=”650146″]Answer d. Telomerase is unique to eukaryotes.[/hidden-answer]

Which of the following would be synthesized using 5′-CAGTTCGGA-3′ as a template?

1. 3′-AGGCTTGAC-4′
2. 3′-TCCGAACTG-5′
3. 3′-GTCAAGCCT-5′
4. 3′-CAGTTCGGA-5′

[reveal-answer q=”429167″]Show Answer[/reveal-answer]
[hidden-answer a=”429167″]Answer c. 3′-GTCAAGCCT-5′[/hidden-answer]

The enzyme responsible for relaxing supercoiled DNA to allow for the initiation of replication is called ________.
[reveal-answer q=”855893″]Show Answer[/reveal-answer]
[hidden-answer a=”855893″]The enzyme responsible for relaxing supercoiled DNA to allow for the initiation of replication is called DNA gyrase or topoisomerase II.[/hidden-answer]

Unidirectional replication of a circular DNA molecule like a plasmid that involves nicking one DNA strand and displacing it while synthesizing a new strand is called ________.
[reveal-answer q=”378861″]Show Answer[/reveal-answer]
[hidden-answer a=”378861″]Unidirectional replication of a circular DNA molecule like a plasmid that involves nicking one DNA strand and displacing it while synthesizing a new strand is calledrolling circle replication.[/hidden-answer]

More primers are used in lagging strand synthesis than in leading strand synthesis.
[reveal-answer q=”25479″]Show Answer[/reveal-answer]
[hidden-answer a=”25479″]True[/hidden-answer]

1. Why is primase required for DNA replication?
2. What is the role of single-stranded binding protein in DNA replication?
3. Below is a DNA sequence. Envision that this is a section of a DNA molecule that has separated in preparation for replication, so you are only seeing one DNA strand. Construct the complementary DNA sequence (indicating 5′ and 3′ ends).DNA sequence: 3′-T A C T G A C T G A C G A T C-5′
4. Review Figure 1 and Figure 2. Why was it important that Meselson and Stahl continue their experiment to at least two rounds of replication after isotopic labeling of the starting DNA with¹⁵N, instead of stopping the experiment after only one round of replication?
5. If deoxyribonucleotides that lack the 3′-OH groups are added during the replication process, what do you expect will occur?

25.1.7 References

1. Parker, N., Schneegurt, M., Thi Tu, A-H., Lister, P., Forster, B.M. (2019) Microbiology. Openstax. Available at: https://opentextbc.ca/microbiologyopenstax/
2. Principles of Biochemistry/Cell Metabolism I: DNA replication. (2017, August 6). Wikibooks, The Free Textbook Project. Retrieved 19:07, October 31, 2019 from en.wikibooks.org/w/index.php?title=Principles_of_Biochemistry/Cell_Metabolism_I:_DNA_replication&oldid=3259729.
3. Kaiser, G.E. (2015) Prokaryotic Cell Anatomy. Community College of Baltimore County. Available at: http://faculty.ccbcmd.edu/~gkaiser/SoftChalk%20BIOL%20230/Prokaryotic%20Cell%20Anatomy/nucleoid/nucleoid/nucleoid3.html
4. The RCSB PDB "Molecule of the Month": Inspiring a Molecular View of Biology D.S. Goodsell, S. Dutta, C. Zardecki, M. Voigt, H.M. Berman, S.K. Burley (2015) PLoS Biol 13(5): e1002140. doi: 10.1371/journal.pbio.1002140
5. Wikipedia contributors. (2020, May 7). Helicase. In Wikipedia, The Free Encyclopedia. Retrieved 13:38, June 9, 2020, from en.Wikipedia.org/w/index.php?title=Helicase&oldid=955303097
6. Windgassen, T.A., Wessel, S.R., Bhattacharyya, B., and Keck, J.L. (2017) Mechanisms of bacterial DNA replication restart. Nuc Acids Res 46(2):504-519. Available at: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5778457/
7. Xu, Z-Q., Dixon, N.E. (2018) Bacterial Replisomes. Curr Opin Struct Biol 53:159-168. Available at: https://www.sciencedirect.com/science/article/pii/S0959440X18300952
8. Liu, B., Eliason, W.K., and Steitz, T.A. (2013) Structure of a helicase-helicase loader complex reveals insights into the mechanism of bacterial primosome assembly. Nature Comm 4:2495. Available at: https://www.researchgate.net/publication/256764134_Structure_of_a_helicase-helicase_loader_complex_reveals_insights_into_the_mechanism_of_bacterial_primosome_assembly
9. Xu, Z-Q., and Dixon, N.E. (2018) Bacterial Replisomes. Curr Op Struc Biol 53:159-168. Available at: https://www.sciencedirect.com/science/article/pii/S0959440X18300952
10. Fernandez-Leiro, R., Conrad, J., Scheres, S.HW., and Lamers, M.H. (2015) cryo-EM structures of the E. coli replicative DNA polymerase reveal its dynamic interactions with the DNA sliding clamp, exonuclease and τ. eLife 4:e11134. Available at: https://elifesciences.org/articles/11134
11. Ekundayo, B. and Bleichert, F. (2019) Origins of DNA replication. PLOS 15(12): e1008556. Available at: https://journals.plos.org/plosgenetics/article?id=10.1371/journal.pgen.1008320
12. Leman, A.R., and Noguchi, E. (2013) The replication fork: understanding the eukaryotic replication machinery and the challenges to genome duplication. Genes 4(1): 1-32. Available at: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3627427/
13. Doublié, S. and Zahn, K.E. (2014) Structural insights into eukaryotic DNA replication. Front. Microbiol. 5:444. Available at: https://www.frontiersin.org/articles/10.3389/fmicb.2014.00444/full
14. Billard, P., and Poncet D.A. (2019) Replication stress at telomeric and mitochondrial DNA: Common origins and consequences on ageing. Int J. Mol Sci 20(19):4959. Available at: https://www.mdpi.com/1422-0067/20/19/4959/htm
15. Yeh, J-K., and Wang, C-Y. (2016) Telomeres and telomerase in cardiovascular diseases. Genes 7(9)58. Available at: https://www.mdpi.com/2073-4425/7/9/58/htm
16. Boccardi, V. and Herbig, U. (2012) Telomerse gene therapy: a novel approach to combat aging. EMBO Mol Med 4:685-687. Available at: https://www.embopress.org/doi/epdf/10.1002/emmm.201200246
17. Wikipedia contributors. (2020, April 26). Cyclin-dependent kinase. In Wikipedia, The Free Encyclopedia. Retrieved 18:52, June 30, 2020, from https://en.Wikipedia.org/w/index.php?title=Cyclin-dependent_kinase&oldid=953307433
18. Falkenberg, M. (2018) Mitochondrial DNA replication in mammalian cells: overview of the pathway. Essays Biochem 62(3):287-296. Available at: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC6056714/
19. Folwer, S., et. al. (2013) Concepts of Biology. Openstax. Available at: https://openstax.org/details/books/concepts-biology?Book%20details
20. Aleem, E. and Arceci, R.J. (2015) Targeting cell cycle regulators in hematologic malignancies. Frontiers in Cell and Developmental Biology 3(16). Available at: https://www.researchgate.net/publication/275354547_Targeting_cell_cycle_regulators_in_hematologic_malignancies