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As we’ve seen, DNA strands have directionality, with a 5’ nucleotide-phosphate and a 3’ deoxyribose hydroxyl end. This is even true for circular bacterial chromosomes…, if the circle is broken! Because the strands of the double helix are antiparallel, the 5’ end of one strand aligns with the 3’end of the other at both ends of the double helix. The complementary pairing of bases in DNA means that the base sequence of one strand can be used as a template to make a new complementary strand. As we’ll see, this structure of DNA created some interesting dilemmas for understanding the biochemistry of replication. The puzzlement surrounding how replication proceeds begins with experiments that visualize replicating DNA.
A. Visualizing Replication and Replication Forks
Recall the phenomenon of bacterial conjugation allowed a demonstration bacterial chromosomes were circular. In 1963, John Cairns confirmed this fact by direct visualization of bacterial DNA. He cultured E. coli cells for long periods on 3Hthymidine (3H-T) to make all of their cellular DNA radioactive. He then disrupted the cells gently to minimize damage to the DNA. The DNA released was allowed to settle and adhere to membranes. A sensitive film was placed over the membrane and time was allowed for the radiation to expose the film. After Cairns developed the autoradiographs, he examined the results in the electron microscope. He saw tracks of silver grains in the autoradiographs (the same kind of silver grains that create an image on film in old-fashioned photography). Look at the two drawings of his autoradiographs on the next page.
Cairns measured the length of the “silver” tracks, which usually consisted of three possible closed loops, or circles. The circumferences of two of these circles were always equal, their length closely predicted by the DNA content of a single, nondividing cell. Cairns therefore interpreted these images to be bacterial DNA in the process of replication. Cairns’ autoradiographs and the measurements that led him to conclude that he was looking at images of bacterial circular chromosomes are illustrated below.
He arranged his autoradiograph images in a sequence (below) to make his point.
Because the replicating chromosomes looked (vaguely!) like the Greek letter ( heta ), Cairns called them theta images. He inferred that replication starts at a single origin of replication on the bacterial chromosome, proceeding around the circle to completion.
175 Seeing E.Coli Chromosomes
Subsequent experiments by David Prescott demonstrated bidirectional replication…, that replication did indeed begin at an origin of replication, after which the double helix was unwound and replicated in both directions, away from the origins, forming two replication forks (illustrated below).
Bacterial cells can divide every hour (or even less); the rate of bacterial DNA synthesis is about 2 X 106 base pairs per hour. A typical eukaryotic cell nucleus contains thousands of times as much DNA as a bacterium, and typical eukaryotic cells double every 15-20 hours. Even a small chromosome can contain hundreds or thousands of times as much DNA as a bacterium. It appeared that eukaryotic cells could not afford to double their DNA at a bacterial rate of replication! Eukaryotes solved this problem not by evolving a faster biochemistry of replication, but by using multiple origins of replication from which DNA synthesis proceeds in both directions. This results in the creation of multiple replicons.
Each replicon enlarges, eventually meeting other growing replicons on either side to replicate most of each linear chromosome, suggested in the illustration below.
Before we consider the biochemical events at replication forks in detail, let's look at the role of DNA polymerase enzymes in the process.
B. DNA Polymerases Catalyze Replication
The first of these enzymes was discovered in E. coli by Arthur Kornberg, for which he received the 1959 Nobel Prize in Chemistry. Thomas Kornberg, one of Arthur’s sons later found two more of DNA polymerases! All DNA polymerases require a template strand against which to synthesize a new complementary strand. They all grow new DNA by adding to the 3’ end of the growing DNA chain in successive condensation reactions. And finally, all DNA polymerases also have the odd property that they can only add to a pre-existing strand of nucleic acid, raising the question of where the ‘preexisting’ strand comes from! DNA polymerases catalyze the formation of a phosphodiester linkage between the end of a growing strand and the incoming nucleotide complementary to the template strand. The energy for the formation of the phosphodiester linkage comes in part from the hydrolysis of two phosphates (pyrophosphate) from the incoming nucleotide during the reaction. While replication requires the participation of many nuclear proteins in both prokaryotes and eukaryotes, DNA polymerases perform the basic steps of replication, as shown in the illustration below.
Although DNA polymerases replicate DNA with high fidelity with as few as one error per 107 nucleotides, mistakes do occur. The proofreading ability of some DNA polymerases corrects many of these mistakes. The polymerase can sense a mismatched base pair, slow down and then catalyze repeated hydrolyses of nucleotides until it reaches the mismatched base pair. This basic proofreading by DNA polymerase is shown below.
After mismatch repair, DNA polymerase resumes forward movement. Of course, not all mistakes are caught by this or other repair mechanisms (see DNA Repair, below). Mutations in the eukaryotic germ line cells that elude correction can cause genetic diseases. However, most are the mutations that fuel evolution. Without mutations in germ line cells (egg and sperm), there would be no mutations and no evolution, and without evolution, life itself would have reached a quick dead end! Other replication mistakes can generate mutations somatic cells. If these somatic mutations escape correction, they can have serious consequences, including the generation of tumors and cancers.
C. The Process of Replication
DNA replication is a sequence of repeated condensation (dehydration synthesis) reactions linking nucleotide monomers into a DNA polymer. Like all biological polymerizations, replication proceeds in three enzymatically catalyzed and coordinated steps: initiation, elongation and termination.
As we have seen, DNA synthesis starts at one or more origins or replication. These are DNA sequences targeted by initiator proteins in E. coli (below).
After breaking hydrogen bonds at the origin of replication, the DNA double helix is progressively unzipped in both directions (i.e., by bidirectional replication). The separated DNA strands serve as templates for new DNA synthesis. Sequences at replication origins that bind to initiation proteins tend to be rich in adenine and thymine bases. This is because A-T base pairs have two hydrogen (H-) bonds that require less energy to break than the three H-bonds holding G-C pairs together. Once initiation proteins loosen H-bonds at a replication origin, DNA helicase uses the energy of ATP hydrolysis to unwind the double helix. DNA polymerase III is the main enzyme that then elongates new DNA. Once initiated, a replication bubble (replicon) forms as repeated cycles of elongation proceed at opposite replication forks.
179 Replication Initiation in E. coli
Recalling that new nucleotides can only be added to the free 3' hydroxyl group of a pre-existing nucleic acid strand. Since no known DNA polymerase can start synthesizing new DNA strands from scratch, this is a problem! The action of DNA polymerases therefore requires a primer, a nucleic acid strand to which to add nucleotides. The questions were…, what is the primer and where does it come from? Since RNA polymerases (enzymes that catalyze RNA synthesis) are the only nucleotide polymerase that can grow a new nucleic acid strand against a DNA template from scratch (i.e., from the first base), it was suggested that RNA might be the primer, After synthesis of a short RNA primer, new deoxynucleotides would be added to its 3’ end by DNA polymerase. The discovery of short stretches of RNA nucleotides at the 5’ end of Okazaki fragments confirmed the notion of RNA primers. We now know that cells use primase, a special RNA polymerase active during replication, to make those RNA primers against DNA templates before a DNA polymerase can grow the DNA strands at replication forks. As we will see now, the requirement for RNA primers is nowhere more in evidence in events at a replication fork.
Looking at elongation at one replication fork (below), we see another problem:
One of the two new DNA strands can grow continuously towards the replication fork as the double helix unwinds. But what about the other strand? Either this other strand must grow in pieces in the opposite direction, or it must wait to begin synthesis until the double helix is fully unwound. If one strand of DNA must be replicated in fragments, then those fragments would have to be stitched (i.e., ligated) together. The problem is illustrated below.
According to this hypothesis, a new leading strand of DNA is lengthened continuously by sequential addition of nucleotides to its 3’ end against its leading strand template. The other strand however, would be made in pieces that would be joined in phosphodiester linkages in a subsequent reaction. Because of the extra step and presumably extra time it takes to make and join these new DNA fragments, this new DNA is called the lagging strand, making its template the lagging strand template.
Reiji Okazaki and his colleagues were studying mutants of T4 phage that grew slowly in their E. coli host cells. They graphed the growth rates of wild-type and mutant T4 phage and demonstrated that slow growth was due to a deficient DNA ligase enzyme, already known to catalyze the circularization of linear phage DNA molecules being replicated in infected host cells. The graph below summarizes their results.
Okazaki’s hypothesis was that the deficient DNA ligase in the mutant phage not only slowed down circularization of replicating T4 phage DNA, but would also be slow at joining phage DNA fragments replicated against at least one of the two template DNA strands. When the hypothesis was tested, the Okazakis found that short DNA fragments did indeed accumulate in E. coli cells infected with ligasedeficient mutants, but not in cells infected with wild type phage. The lagging strand fragments are now called Okazaki fragments.
You can check out Okazaki’s original research at this link.
Each Okazaki fragment would have to begin with a 5’ RNA primer, creating yet another dilemma! The RNA primer must be replaced with deoxynucleotides before stitching the fragments together. This in fact happens, and the process illustrated below
Removal of RNA primer nucleotides from Okazali fragments requires the action of DNA polymerase I, an enzyme that can also catalyze hydrolysis of the phosphodiester bonds between the RNA (or DNA) nucleotides from the 5’-end of a nucleic acid strand. Flap Endonuclease 1 (FEN 1) also plays a role in removing ‘flaps’ of nucleic acid from the 5’ ends of the fragments often displaced by polymerase as it replaces the replication primer. At the same time as the RNA nucleotides are removed, DNA polymerase I catalyzes their replacement by the appropriate deoxynucleotides. Finally, when a fragment is entirely DNA, DNA ligase links it to the rest of the already assembled lagging strand DNA. Because of its 5’ exonuclease activity (not found in other DNA polymerases), DNA polymerase 1 also plays unique roles in DNA repair (discussed further below). As Cairn’s suggested and others demonstrated, replication proceeds in two directions from the origin to form a replicon with its two replication forks (RFs). Each RF has a primase associated with replication of Okazaki fragments along lagging strand templates.
As Cairn’s suggested and others demonstrated, replication proceeds in two directions from the origin to form a replicon with its two replication forks (RFs). Each RF has a primase associated with replication of Okazaki fragments along lagging strand templates.
The requirement for primases at replication forks is shown below.
Now we can ask what happens when replicons reach the ends of linear chromosomes in eukaryotes.
182 Replication Elongation in E.coli
In prokaryotes, replication is complete when two replication forks meet after replicating their portion of the circular DNA molecule. In eukaryotes, many replicons fuse to become larger replicons, eventually reaching the ends of the chromosomes…, where there is yet another problem (below)!
When a replicon nears the end of a chromosome (i.e. a double-stranded DNA molecule), the strand synthesized continuously stops when it reaches the 5’ end of its template DNA. In theory, synthesis of a last Okazaki fragment can be primed from the 3’ end of the lagging template strand. The illustration above implies removal of a primer from the penultimate Okazaki fragment and DNA polymerase catalyzed replacement with DNA nucleotides. But what about the last Okazaki fragment? Would its primer be hydrolyzed? Moreover, without a free 3’ end to add to, how are those RNA nucleotides replaced with DNA nucleotides? The problem here is that every time a cell replicates, one strand of new DNA (likely both) would get shorter and shorter. Of course, this would not do…, and does not happen! Eukaryotic replication undergoes a termination process involving extending the length of one of the two strands by the enzyme telomerase. The action of telomerase is summarized in the illustration below
Telomerase consists of several proteins and an RNA. From the drawing,, the RNA component serves as a template for 5’-> 3’ extension of the problematic DNA strand. The protein with the requisite reverse transcriptase activity is called Telomerase Reverse Transcriptase, or TERT. The Telomerase RNA Component is called TERC. Carol Greider, Jack Szostak and Elizabeth Blackburn shared the 2009 Nobel Prize in Physiology or Medicine for discovering telomerase.
183 Telomerase Replication Prevents Chromosome Shorteninghttps://youtu.be/M4dmfrxGKKU
One of the more interesting recent observations was that differentiated, nondividing cells no longer produce the telomerase enzyme. On the other hand, the telomerase genes are active in normal dividing cells (e.g., stem cells) and cancer cells, which contain abundant telomerase.
4. Is Replication Processive?
Drawings of replicons and replication forks suggest separate events on each DNA strand. Yet events at replication forks seem to be coordinated. Replication may be processive, meaning both new DNA strands are replicated in the same direction at the same time, smoothing out the process. How might this be possible? The drawing below shows lagging strand template DNA bending, so that it faces in the same direction as the leading strand at the replication fork.
The replisome structure cartooned at the replication fork consists of clamp proteins, primase, helicase, DNA polymerase and single-stranded binding proteins among others.
Newer techniques of visualizing replication by real-time fluorescence videography call the processive model into question, suggesting that the replication process is anything but smooth! Are lagging and leading strand replication not in fact coordinated? Alternatively, is the jerky movement of DNA elongation in the video an artifact, so that the model of smooth, coordinated replication integrated at a replisome still valid? Or is coordination defined and achieved in some other way? Check out the video yourself in the article here.
5. One More Problem with Replication
Cairns recorded many images of E.coli of the sort shown below.
The coiled, twisted appearance of the replicating circles were interpreted to be a natural consequence of trying to pull apart helically intertwined strands of DNA… or intertwined strands of any material! As the strands continued to unwind, the DNA should twist into a supercoil of DNA. Increased DNA unwinding would cause the phosphodiester bonds in the DNA to rupture, fragmenting the DNA. Obviously, this does not happen. Experiments were devised to demonstrate supercoiling, and to test hypotheses explaining how cells relax the supercoils during replication. Testing these hypotheses revealed the topoisomerase enzymes. These enzymes bind and hold on to DNA, catalyze hydrolysis of phosphodiester bonds, control unwinding of the double helix, and finally catalyze the re-formation of the phosphodiester linkages. It is important to note that the topoisomerases are not part of a replisome, but can act far from a replication fork, probably responding to the tensions in overwound DNA. Recall that topoisomerases comprise much of the protein lying along eukaryotic chromatin.
We have considered most of the molecular players in replication. Below is a list of the key replication proteins and their functions (from here).
|Enzyme||Function in DNA Replication|
|DNA Helicase||Also known as helix destabilizing enzyme. Unwinds the DNA double helix at the Replication Fork.|
Builds a new duplex DNA strand by adding nucleotides in the 5' to 3' direction. Also performs proof-reading and error correction.
|DNA clamp||A protein which prevents DNA polymerase III from dissociating from the DNA parent strand.|
|Single-Strand Binding (SSB) Proteins||Bind to ssDNA and preven the DNA double helix from re-annealing after DNA helicase unwinds it thus maintaining the strand separation.|
|Topoisomerase||Relaxes the DNA from its super-coiled nature.|
|DNA Gyrase||Relieves strain of unwinding by DNA helicase; this is a specific type of topoisomerase|
|DNA Ligase||Re-anneals the semi-conservative strands and joins Okazaki Fragments of the lagging strand|
|Primase||Provides a starting point of RNA (or DNA) for DNA polymerase to begin synthesis of the new DNA strand.|
|Telomerase||Lengthens telomeric DNA by adding repetitive nucleotide sequences to the ends of eukaryotic chromosomes.|
Clamp loader ATPases and the evolution of DNA replication machinery
Clamp loaders are pentameric ATPases of the AAA+ family that operate to ensure processive DNA replication. They do so by loading onto DNA the ring-shaped sliding clamps that tether the polymerase to the DNA. Structural and biochemical analysis of clamp loaders has shown how, despite differences in composition across different branches of life, all clamp loaders undergo the same concerted conformational transformations, which generate a binding surface for the open clamp and an internal spiral chamber into which the DNA at the replication fork can slide, triggering ATP hydrolysis, release of the clamp loader, and closure of the clamp round the DNA. We review here the current understanding of the clamp loader mechanism and discuss the implications of the differences between clamp loaders from the different branches of life.
High-speed replication of chromosomal DNA requires the DNA polymerase to be attached to a sliding clamp (known as proliferating cell nuclear antigen, or PCNA, in eukaryotes) that prevents the polymerase from falling off DNA [1, 2]. In all cells and in some viruses, the clamp is a ring-shaped protein complex that encircles DNA, forming a sliding platform on which DNA polymerases and other proteins that move along DNA are assembled. Sliding clamps play a part in DNA replication, DNA repair, cell cycle control and modification of chromatin structure [3, 4], and defects in several clamp-associated factors are associated with cancer and other disorders caused by abnormalities in DNA replication and repair .
Because sliding clamps encircle DNA but do not interact tightly with it, they can slide along the double helix by diffusion [6–9]. Sliding clamps from different branches of life have different subunit stoichiometry (they are dimers in bacteria  and trimers in eukarya, archaea and bacteriophage [11–15]) and their sequences have diverged beyond recognition. Nevertheless, their structures are remarkably similar. The conserved structure is an elegant symmetrical elaboration of a simple β-α-β motif, repeated 12 times around a circle [10, 14]. The circular geometry is broken when the clamp is opened for loading onto DNA, but the elegance is retained during the loading step as the clamp assumes a helical symmetry that reflects the helical symmetry of DNA (see below).
The increase in the processivity and speed of DNA synthesis when DNA polymerases are engaged to sliding clamps is very considerable. For example, in the absence of the clamp, the polymerase subunit of the bacterial replicase synthesizes DNA at a rate of about 10 base pairs per second  and is hardly processive. In contrast, the same polymerase subunit synthesizes 500 to 1,000 nucleotides per second when bound to the sliding clamp [17–19]. To consider a startling analogy based on scaling linear dimensions, if the bacterial replicase were a car, it would travel only about 5 to 10 miles per hour without the clamp and faster than the speed of sound with the clamp. The bacterial replicase has a processivity of about 10 base pairs in the absence of the clamp , but has an average processivity of approximately 80 kilobases when bound to the sliding clamp in the replisome . To invoke another analogy based on scaling dimensions, if the polymerase were a tightrope walker, without the aid of the clamp only about 20 feet of the tightrope would be traversed before the polymerase 'walker' fell off. The clamp allows the polymerase to hold on to the DNA 'rope' without letting go, and now it would 'walk' almost 30 miles before falling off.
The enhancement in speed and processivity that the clamp confers on the polymerase would not be possible without the clamp loader, the less glamorous but much more hardworking handmaiden of the sliding clamp, which diligently loads the clamps onto primed DNA throughout the process of DNA replication. Together, the clamp and the clamp loader lie at the heart of the replisome - the DNA replication machinery, which, with the polymerases (leading and lagging strand), includes the helicases that unwind the double-stranded DNA ahead of the polymerase at the replication fork, the primase that synthesizes the RNA primer required for the initiation of DNA synthesis, and the single-strand DNA-binding proteins that prevent the DNA from re-annealing in the wake of the helicases (Figure 1). The clamp loader opens sliding clamps and places them on the DNA at the site of the primer-template junction in the correct orientation for polymerase to bind, both at the initiation of DNA synthesis on the leading strand and continually at the start of each Okazaki fragment on the lagging strand. Thus, the clamp loader is critical for the tight coupling of leading and lagging strand synthesis. Indeed, in bacteria the clamp loader acts physically to hold the leading and lagging strand polymerases together [22–25] so that the two polymerases progress in tandem, with the lagging strand wrapped around the replisome in trombone fashion . How leading and lagging strand polymerases are coupled in eukaryotic DNA synthesis is not known, and this is one of the major open questions about the operation of the eukaryotic replisome.
Architecture of the bacterial replication fork. The helicase is a homohexamer that encircles the lagging strand and binds directly to the primase synthesizing the primer RNA. The clamp loader acts to hold the replisome together by binding directly to the helicase as well as three polymerase subunits for simultaneous synthesis of the leading and lagging strands. The leading strand polymerase synthesizes DNA continuously, while the other two polymerases presumably cycle on and off the lagging strand, which is coated in single-strand binding protein (SSB). The polymerase subunits are attached to circular clamps that encircle duplex DNA for enhanced processivity and speed.
Despite the uncertainty in the precise architectural role of the clamp loader in the eukaryotic replisome, it is clear that sliding clamps are centrally important. The sliding clamp recruits the polymerase as well as other factors to the replication fork, including the chromatin-modifying proteins required to reassemble chromatin on the newly synthesized DNA [27, 28].
Along with many other vital functions, mitochondria generate most of the cell’s energy through aerobic respiration. In humans, as in most eukaryotes, mitochondria possess their own indispensable genome (mtDNA) which replicates separately from the nuclear genome. Human mtDNA is a 16.6-kb closed circular molecule encoding 37 genes required to make 13 essential subunits of the oxidative phosphorylation (OXPHOS) complexes, as well as critical elements of the mitochondrial translational machinery (Fig. 1a). OXPHOS drives adenosine triphosphate (ATP) production, generating energy for cellular consumption. Each human cell contains hundreds to thousands of mtDNA copies existing in a mixture of wild type and mutated molecular species that is collectively called heteroplasmy [1,2,3,4]. Low levels of heteroplasmy have been detected in most healthy individuals , whereas high levels of heteroplasmy have been implicated in a broad range of neurometabolic disorders under the umbrella term of “mitochondrial diseases” [4,5,6].
The human mtDNA reference map and the mtDNA deletion mapping pipeline. a Human mtDNA reference map, color-coded by feature. Inset: respiratory chain complexes encoded by mtDNA (color) and nuclear genes (gray). MtDNA-encoded genes and feature abbreviations: tRNA genes, IUPAC single-letter amino acid codes rRNA genes, sedimentation coefficients (e.g., 16S) previously proposed heavy and light strand features, _H and _L promoters, P_ replication origins, O_ (e.g., PH1, OL) and MT- prefixes are omitted. The 7S-3′-terminus and oriL are indicated (red and green arrows). b LostArc method outline. c The mtDNA fractions (mean ± 95% CL) indicate enrichment during library preparation and a subsequent lack of selection in sequencing (qPCR for steps 1, 3, and 4b, fraction of reads mapped to mtDNA reference for step 7). See also Additional file 1: Fig. S1. d Estimated mtDNA frequencies. Species with frequencies over 5 × 10 −4 must reside in multiple truncated fibers (i.e., they predate adulthood or mark mutational hotspots). Deletions observed ≥ 2x must have ≥ 20 copies in the muscle sample. Deletions observed once will have frequencies inflated to roughly 1/depth. These collectively represent all less frequent species
Pathogenic variants of at least 24 nuclear genes whose protein products are responsible for mtDNA maintenance co-segregate with mitochondrial disease [7,8,9,10]. Two such genes (POLG and POLG2, respectively) encode the catalytic and accessory subunits of DNA polymerase γ (Pol γ). Pol γ is a family A DNA polymerase with 3′-5′ exonuclease and 5′-dRP lyase activities and the only known mitochondrial replicase [11, 12]. Pol γ works in concert with the Twinkle helicase, a mitochondrial ssDNA binding protein and other accessory factors to carry out efficient mtDNA replication [13, 14]. Pathogenic mutations in POLG are the most common cause of mitochondrial disease linked to improper maintenance of the mitochondrial genome . Over three hundred missense mutations in POLG have been reported to cause a wide spectrum of mitochondrial diseases with ages of onset ranging from early (childhood myocerebrohepatopathy or Alpers-Huttenlocher syndrome) to late (ataxia neuropathies or progressive external ophthalmoplegia) [16, 17]. Pathogenic POLG variants encode proteins with assorted biochemical deficiencies, including catalytic deficits, structural instability, and defects in binding DNA or accessory proteins . Early-onset POLG disorders are associated with a quantitative loss of mtDNA copies (mtDNA depletion), whereas later-onset POLG disorders are associated with accumulation of mtDNA deletions .
Point mutations and deletions in mtDNA accumulate with age and have been implicated in normal aging and age-related pathologies [18,19,20,21,22,23]. However, as assayed in human brain tissue, the accumulation of point mutations is orders of magnitude too low to explain the phenotypes of aging . Prior work by this group had shown that point mutations in mtDNA do not limit the natural lifespan of mice . These observations lead to the hypothesis that the greater genetic disruption caused by mtDNA deletions may be a significant contributor to mechanisms of aging. Disruption of Pol γ’s exonuclease function prevents proofreading of DNA replication errors , resulting in a 160-fold increase in mtDNA deletions in yeast . In homozygous POLG exonuclease deficient mice [28, 29], random mutation capture assays identified mtDNA deletions as a causative factor in the premature aging phenotype . The biochemical mechanisms underlying the formation of mtDNA deletions are not well defined. Several models have been proposed, including primer relocation between direct repeats during mtDNA replication [31, 32] and ectopic re-annealing of broken strands within short homologous DNA sequences .
To gain insight into the molecular mechanisms underlying formation of mtDNA deletions and to define the roles of deletions in aging and human mitochondrial disease, we sought to delineate the position, length, sequence context, and abundance of individual mtDNA deletions in human tissue samples. Because mtDNA represents only 0.93% of total DNA in human skeletal muscle, existing deletion detection techniques rely on PCR to selectively enrich for mtDNA sequences [34, 35]. In order to avoid PCR artifacts and target biasing due to primer selection, we developed LostArc, an ultra-sensitive high throughput deletion detection pipeline. Here, we used LostArc to analyze mtDNA sequences from skeletal muscle samples of mitochondrial disease patients bearing inherited pathogenic POLG variants and from age-matched individuals with wild type POLG. Hundreds of thousands of unique deletions were identified with frequencies sufficient to explain the musculoskeletal phenotypes of aging and disease. Patterns in the locations and sequence contexts suggest that DNA replication is the primary driver of deletion formation and strongly support asynchronous strand displacement models of mtDNA replication.
Dr. Meselson has made important contributions to the areas of DNA replication, repair and recombination as well as isolating the first restriction enzyme. Currently, he is Professor of Molecular and Cellular Biology at Harvard University, where his lab studies aging in the model organism bdelloid rotifers. Meselson is also a long-time advocate for the abolition… Continue Reading
More Talks in Genetics and Gene Regulation
Youreka Science Video about the Meselson and Stahl Paper
Research Paper Discussed in this Talk
Meselson, M. & F. W. Stahl. 1958. The replication of DNA in Escherichia coli. PNAS 44:671-682.
Both DNA replication and transcription involve binding complementary nucleic acids to DNA, yielding a new strand of either DNA or RNA. Both processes can lead to errors if an incorrect nucleotide is incorporated. An error in either DNA replication or transcription can cause a change in the gene, by either changing the DNA sequence in one of the daughter cells leading to transcription of the incorrect mRNA sequence, or by causing the mRNA to incorporate an incorrect base pair resulting in the wrong protein sequence being translated.
Another area of human biology tightly linked to Neanderthal variants in the genome is the immune system. Given that human ancestors were exposed to a menagerie of different pathogens—some of which came directly from the Neanderthals—as they migrated through Eurasia, the Neanderthal sequences introgressed into the human genome may have helped defend against these threats, to which Neanderthals had long been exposed.
“Viral challenges, bacterial challenges are among the strongest selective forces out there,” says Kelso. Unlike changes in other environmental conditions such as daylight patterns and UV exposure, “pathogens can kill you in one generation.”
Hints of archaic DNA’s role in immune function surfaced as early as 2011, as soon as the Neanderthal genome was available for cross-referencing with sequences from modern humans. A team led by researchers at Stanford University found that certain human leukocyte antigen (HLA) alleles, key players in pathogen recognition, held signs of archaic ancestry—from Neanderthals, but also from another hominin cousin, the Denisovans. 6 “It’s a cool paper and one that contributed to a lot of people thinking about the effects of introgression,” says Capra.
Several other studies since then have strengthened the link between archaic DNA and immune function, branching out from the HLA system to include numerous other pathways. 7 For example, multiple labs have tied Neanderthal variants to altered expression levels of genes encoding toll-like receptors (TLRs), key players in innate immune responses. In 2016, Kelso, Dannemann, and a colleague found that pathogen response and susceptibility to develop allergies were associated with Neanderthal sequences that affect TLR production. 8
Viruses, in particular, appear to be potent drivers of adaptation. Last year, University of Arizona population geneticist David Enard and colleagues found that one-third of Neanderthal variants under positive selection were linked to genes encoding proteins that interact with viruses. 9
Viral challenges, bacterial challenges are among the strongest selective forces out there. Pathogens can kill you in one generation.
Researchers have also identified several less-easily explainable phenotypic associations with Neanderthal introgression. In their 2017 analysis, for example, Kelso and Dannemann found that Neanderthal variants were associated with chronotype—whether people identify as early birds or night owls—as well as links with susceptibility to feelings of loneliness or isolation and low enthusiasm or interest. The associations with mood-related phenotypes jibe with what Capra’s group had found the year before in its dataset of medical information, which linked Neanderthal variants to risks for depression and addiction. “These were associations that were quite strong,” says Capra. “And when we looked at genes where these regions of Neanderthal ancestry fell, in many cases they made sense given what we already know about those genes.”
Why these associations exist is still a mystery. Kelso suspects that light might be a unifying factor, with both changes in day-length patterns and UV exposure reductions as they moved to more-northern latitudes. But that’s just a hunch, she emphasizes.
“It’s fun speculating about how [Neanderthal introgression] could have been advantageous, or how variants that make us depressed in the modern environment could have been beneficial,” says Capra. “I don’t really even know what depression meant 40,000 years ago. That’s both the challenge and the fun, provocative part about all this.”
The Watson-Crick Model
A molecular structure for DNA has been proposed by Watson and Crick. 15 It has undergone preliminary refinement 16 without alteration of its main features and is supported by physical and chemical studies. 17 The structure consists of two polynucleotide chains wound helically about a common axis. The nitrogen base (adenine, guanine, thymine, or cytosine) at each level on one chain is hydrogen-bonded to the base at the same level on the other chain. Structural requirements allow the occurrence of only the hydrogen-bonded base pairs adenine-thymine and guanine-cytosine, resulting in a detailed complementariness between the two chains. This suggested to Watson and Crick 18 a definite and structurally plausible hypothesis for the duplication of the DNA molecule. According to this idea, the two chains separate, exposing the hydrogen-bonding sites of the bases. Then, in accord with the base-pairing restrictions, each chain serves as a template for the synthesis of its complement. Accordingly, each daughter molecule contains one of the parental chains paired with a newly synthesized chain (Fig. 6).
Illustration of the mechanism of DNA duplication proposed by Watson and Crick. Each daughter molecule contains one of the parental chains (black) paired with one new chain (white). Upon continued duplication, the two original parent chains remain intact, so that there will always be found two molecules each with one parental chain.
The results of the present experiment are in exact accord with the expectations of the Watson-Crick model for DNA duplication. However, it must be emphasized that it has not been shown that the molecular subunits found in the present experiment are single polynucleotide chains or even that the DNA molecules studied here correspond to single DNA molecules possessing the structure proposed by Watson and Crick. However, some information has been obtained about the molecules and their subunits it is summarized below.
The DNA molecules derived from E. coli by detergent-induced lysis have a buoyant density in CsCl of 1.71 gm. cm. −3 , in the region of densities found for T2 and T4 bacteriophage DNA, and for purified calf-thymus and salmon-sperm DNA. A highly viscous and elastic solution of N 14 DNA was prepared from a dodecyl sulfate lysate of E. coli by the method of Simmons 19 followed by deproteinization with chloroform. Further purification was accomplished by two cycles of preparative density-gradient centrifugation in CsCl solution. This purified bacterial DNA was found to have the same buoyant density and apparent molecular weight, 7 × 10 6 , as the DNA of the whole bacterial lysates (Figs. 7, 8).
Microdensitometer tracing of an ultraviolet absorption photograph showing the optical density in the region of a band of N 14 E. coli DNA at equilibrium. About 2 μg. of DNA purified as described in the text was centrifuged at 31,410 rpm at 25° in 7.75 molal CsCl at pH 8.4. The density gradient is essentially constant over the region of the band and is 0.057 gm./cm. 4 . The position of the maximum indicates a buoyant density of 1.71 gm. cm. −3 In this tracing the optical density above the base line is directly proportional to the concentration of DNA in the rotating centrifuge cell. The concentration of DNA at the maximum is about 50 μg./ml.
The square of the width of the band of Fig. 7 plotted against the logarithm of the relative concentration of DNA. The divisions along the abscissa set off intervals of 1 mm. 2 . In the absence of density heterogeneity, the slope at any point of such a plot is directly proportional to the weight average molecular weight of the DNA located at the corresponding position in the band. Linearity of this plot indicates monodispersity of the banded DNA. The value of the the slope corresponds to an apparent molecular weight for the Cs·DNA salt of 9.4 × 10 6 , corresponding to a molecular weight of 7.1 × 10 6 for the sodium salt.
A selectable marker is usually a gene that confers resistance to an antibiotic that would otherwise kill the cells.
Identify the purpose of selection in genetic engineering
- Recombinant DNA is introduced into the organism from which the replication sequences were obtained, then the foreign DNA will be replicated along with the host cell’s DNA in the transgenic organism.
- Artificial genetic selection is the process in which cells that have not taken up DNA are selectively killed, and only those cells that can actively replicate DNA containing the selectable marker gene encoded by the vector are able to survive.
- When bacterial cells are used as host organisms, the selectable marker is usually a gene that confers resistance to an antibiotic that would otherwise kill the cells, typically ampicillin.
- molecular cloning: a set of experimental methods in molecular biology that are used to assemble recombinant DNA molecules and to direct their replication within host organisms.
- PCR: polymerase chain reaction
Scientists who do experimental genetics employ artificial selection experiments that permit the survival of organisms with user-defined phenotypes. Artificial selection is widely used in the field of microbial genetics, especially molecular cloning.
DNA recombination has been used to create gene replacements, deletions, insertions, inversions. Gene cloning and gene/protein tagging is also common. For gene replacements or deletions, usually a cassette encoding a drug-resistance gene is made by PCR.
Molecular cloning is a set of experimental methods in molecular biology that are used to assemble recombinant DNA molecules and to direct their replication within host organisms. The use of the word cloning refers to the fact that the method involves the replication of a single DNA molecule starting from a single living cell to generate a large population of cells containing identical DNA molecules. Molecular cloning generally uses DNA sequences from two different organisms: the species that is the source of the DNA to be cloned, and the species that will serve as the living host for replication of the recombinant DNA. Molecular cloning methods are central to many contemporary areas of modern biology and medicine.
In a conventional molecular cloning experiment, the DNA to be cloned is obtained from an organism of interest. It is then treated with enzymes in the test tube to generate smaller DNA fragments. Subsequently, these fragments are then combined with vector DNA to generate recombinant DNA molecules. The recombinant DNA is then introduced into a host organism (typically an easy-to-grow, benign, laboratory strain of E. coli bacteria ). This will generate a population of organisms in which recombinant DNA molecules are replicated along with the host DNA. Because they contain foreign DNA fragments, these are transgenic or genetically-modified microorganisms (GMO). This process takes advantage of the fact that a single bacterial cell can be induced to take up and replicate a single recombinant DNA molecule. This single cell can then be expanded exponentially to generate a large amount of bacteria, each of which contain copies of the original recombinant molecule. Thus, both the resulting bacterial population, and the recombinant DNA molecule, are commonly referred to as “clones”. Strictly speaking, recombinant DNA refers to DNA molecules, while molecular cloning refers to the experimental methods used to assemble them.
Molecular cloning takes advantage of the fact that the chemical structure of DNA is fundamentally the same in all living organisms. Therefore, if any segment of DNA from any organism is inserted into a DNA segment containing the molecular sequences required for DNA replication, and the resulting recombinant DNA is introduced into the organism from which the replication sequences were obtained, then the foreign DNA will be replicated along with the host cell’s DNA in the transgenic organism.
Molecular cloning is similar to polymerase chain reaction (PCR) in that it permits the replication of a specific DNA sequence. The fundamental difference between the two methods is that molecular cloning involves replication of the DNA in a living microorganism, while PCR replicates DNA in an in vitro solution, free of living cells. Whichever method is used, the introduction of recombinant DNA into the chosen host organism is usually a low efficiency process that is, only a small fraction of the cells will actually take up DNA. Experimental scientists deal with this issue through a step of artificial genetic selection, in which cells that have not taken up DNA are selectively killed, and only those cells that can actively replicate DNA containing the selectable marker gene encoded by the vector are able to survive. When bacterial cells are used as host organisms, the selectable marker is usually a gene that confers resistance to an antibiotic that would otherwise kill the cells, typically ampicillin. Cells harboring the vector will survive when exposed to the antibiotic, while those that fail to take up vector sequences die. When mammalian cells (e.g. human or mouse cells) are used, a similar strategy is used, except that the marker gene (in this case typically encoded as part of the kanMX cassette) confers resistance to the antibiotic Geneticin.
Plasmid vector map: This vector confers amplicillin resistance.
Department of Biology
In the late 1970s Carl Woese discovered that living organisms on Earth could be classified into one of three distinct domains, Eukarya, Bacteria and Archaea. In the time following that pivotal discovery, it has become apparent that, although morphologically resembling bacteria, the archaea in fact possess a number of molecular features more reminiscent of eukarya than bacteria. More specifically, the information processing pathways in archaea form a simplified version of the eukaryotic apparatus. Our work has primarily focussed on members of the genus Sulfolobus. In particular we have studied S. solfataricus and S. acidocaldarius. These species are hyperthermophilic acidophilic aerobes (they grow at 80°C and at pHs between 2 and 4). The hyperthermophilicity of the organisms is mirrored by an innate thermostability of the proteins that they encode. This greatly facilitates purification of native and recombinant proteins. There are also a growing number of genetic tools available for these species, further adding to their utility as model organisms.
DNA replication is a complex multi-step process involving the coordinated interplay of many proteins. During evolution, two distinct sets of cellular DNA replication proteins have evolved, one used by bacteria and a core machinery common to archaea and eukaryotes. In general the archaeal apparatus is a simplified version of that in eukaryotes, making Archaea a useful model system. We have mapped 3 origins of replication in the single chromosome of Sulfolobus and have characterized the interplay of initiator proteins with the origins. In addition, we are investigating the architecture of the replication fork assembly. In particular we are interested in the interplay between architectural and enzymatic components of the Okazaki fragment maturation machinery. Additionally, we have studied how replication termination is effected in Sulfolobus. Beyond the core replication architectures, we are interested in the twin processes of sister chromatid cohesion and chromosome dimer resolution in Sulfolobus.
As with DNA replication, the archaeal transcription apparatus is a slimmed down version of the eukaryotic machinery. The archaeal RNA polymerase is able to initiate transcription in vitro with the aid of just two general transcription factors, TBP and TFB. In collaboration with Nicola Abrescia (Bilbao) we are investigating the archaeal transcription apparatus using a combination of structural analyses and next generation sequencing technologies.
Cell division in Eukaryotes and most Bacteria and Archaea is dependent on proteins in the near ubiquitous FtsZ/Tubulin and MreB/Actin superfamilies. However, the genomes of Sulfolobus and a number of other crenarchaea notably lack genes for these proteins. We have identified Sulfolobushomologs of the eukaryotic ESCRT system as key players in Sulfolobus cell division. The ESCRT proteins in eukaryotes play a diverse variety of roles including endosome sorting, membrane abscission during cytokinesis and is a system that is highjacked by viruses, including HIV and Ebola, to exit from cells. The Sulfolobus ESCRT system contains a limited subset of the eukaryal machinery. In particular, the early components of the eukaryal machinery, that define positioning of the apparatus lack clear orthologs in Sulfolobus. In our recent work, we have identified the factor responsible for recruiting Sulfolobus ESCRT-III to membranes at mid-cell.
Ligation of DNA is a critical step in many modern molecular biology workflows. The sealing of nicks between adjacent residues of a single-strand break on a double-strand substrate and the joining of double-strand breaks are enzymatically catalyzed by DNA ligases. The formation of a phosphodiester bond between the 3' hydroxyl and 5' phosphate of adjacent DNA residues proceeds in three steps: Initially, the ligase is self-adenylated by reaction with free ATP. Next, the adenyl group is transferred to the 5'-phosphorylated end of the "donor" strand. Lastly, the formation of the phosphodiester bond proceeds after reaction of the adenylated donor end with the adjacent 3' hydroxyl acceptor and the release of AMP. In living organisms, DNA ligases are essential enzymes with critical roles in DNA replication and repair. In the lab, DNA ligation is performed for both cloning and non-cloning applications.
DNA Ligase Fidelity: When does it matter?
High fidelity polymerases are everywhere&mdashbut why would you need a high fidelity ligase? And what do we even mean by &ldquofidelity&rdquo when we&rsquore talking about ligation? In this webinar, NEB Scientist and ligase expert Greg Lohman discusses mismatch ligation by DNA ligases and the molecular diagnostics applications that depend on the use of high-fidelity DNA ligases like NEB&rsquos HiFi Taq DNA Ligase to detect single base differences in DNA.
Ligation, the process of joining DNA fragments with a DNA ligase, proceeds in three steps. Learn more about the function of ligation with our quick tutorial animation.
What molar ratios should I use for DNA Ligation?
The optimal reactant ratio is contingent upon the downstream application.
Why do I need to add PEG to my DNA ligation?
Polyethylene glycol (PEG) is an important reagent in ligation reactions, find out why.
What are the best conditions for DNA ligation?
Find out how the downstream application dictates the best reaction conditions for ligation.
Are some ligations more difficult than others?
Ligation of blunt ends and single-base overhangs require optimized reaction conditions.