DNA replication in $ E.coli $

DNA replication in $ E.coli $

We are searching data for your request:

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

$ E.coli $ has circular DNA which I guess implies one strand forms the outer circle and the other the inner one. So, is there a way to know if the replicated DNA forms the outer or inner circle? In the image attached it is seen that the replicated DNA forms the inner circle. Also mentioned is the fact that it is not possible to make out individual strands by TEM.

14.4 DNA Replication in Prokaryotes

By the end of this section, you will be able to do the following:

  • Explain the process of DNA replication in prokaryotes
  • Discuss the role of different enzymes and proteins in supporting this process

DNA replication has been well studied in prokaryotes primarily because of the small size of the genome and because of the large variety of mutants that are available. E. coli has 4.6 million base pairs in a single circular chromosome and all of it gets replicated in approximately 42 minutes, starting from a single site along the chromosome and proceeding around the circle in both directions. This means that approximately 1000 nucleotides are added per second. Thus, the process is quite rapid and occurs without many mistakes.

DNA replication employs a large number of structural proteins and enzymes, each of which plays a critical role during the process. One of the key players is the enzyme DNA polymerase, also known as DNA pol, which adds nucleotides one-by-one to the growing DNA chain that is complementary to the template strand. The addition of nucleotides requires energy this energy is obtained from the nucleoside triphosphates ATP, GTP, TTP and CTP. Like ATP, the other NTPs (nucleoside triphosphates) are high-energy molecules that can serve both as the source of DNA nucleotides and the source of energy to drive the polymerization. When the bond between the phosphates is “broken,” the energy released is used to form the phosphodiester bond between the incoming nucleotide and the growing chain. In prokaryotes, three main types of polymerases are known: DNA pol I, DNA pol II, and DNA pol III. It is now known that DNA pol III is the enzyme required for DNA synthesis DNA pol I is an important accessory enzyme in DNA replication, and along with DNA pol II, is primarily required for repair.

How does the replication machinery know where to begin? It turns out that there are specific nucleotide sequences called origins of replication where replication begins. In E. coli, which has a single origin of replication on its one chromosome (as do most prokaryotes), this origin of replication is approximately 245 base pairs long and is rich in AT sequences. The origin of replication is recognized by certain proteins that bind to this site. An enzyme called helicase unwinds the DNA by breaking the hydrogen bonds between the nitrogenous base pairs. ATP hydrolysis is required for this process. As the DNA opens up, Y-shaped structures called replication forks are formed. Two replication forks are formed at the origin of replication and these get extended bi-directionally as replication proceeds. Single-strand binding proteins coat the single strands of DNA near the replication fork to prevent the single-stranded DNA from winding back into a double helix.

DNA polymerase has two important restrictions: it is able to add nucleotides only in the 5' to 3' direction (a new DNA strand can be only extended in this direction). It also requires a free 3'-OH group to which it can add nucleotides by forming a phosphodiester bond between the 3'-OH end and the 5' phosphate of the next nucleotide. This essentially means that it cannot add nucleotides if a free 3'-OH group is not available. Then how does it add the first nucleotide? The problem is solved with the help of a primer that provides the free 3'-OH end. Another enzyme, RNA primase , synthesizes an RNA segment that is about five to ten nucleotides long and complementary to the template DNA. Because this sequence primes the DNA synthesis, it is appropriately called the primer . DNA polymerase can now extend this RNA primer, adding nucleotides one-by-one that are complementary to the template strand (Figure 14.14).

Visual Connection

Question: You isolate a cell strain in which the joining of Okazaki fragments is impaired and suspect that a mutation has occurred in an enzyme found at the replication fork. Which enzyme is most likely to be mutated?

The replication fork moves at the rate of 1000 nucleotides per second. Topoisomerase prevents the over-winding of the DNA double helix ahead of the replication fork as the DNA is opening up it does so by causing temporary nicks in the DNA helix and then resealing it. Because DNA polymerase can only extend in the 5' to 3' direction, and because the DNA double helix is antiparallel, there is a slight problem at the replication fork. The two template DNA strands have opposing orientations: one strand is in the 5' to 3' direction and the other is oriented in the 3' to 5' direction. Only one new DNA strand, the one that is complementary to the 3' to 5' parental DNA strand, can be synthesized continuously towards the replication fork. This continuously synthesized strand is known as the leading strand . The other strand, complementary to the 5' to 3' parental DNA, is extended away from the replication fork, in small fragments known as Okazaki fragments , each requiring a primer to start the synthesis. New primer segments are laid down in the direction of the replication fork, but each pointing away from it. (Okazaki fragments are named after the Japanese scientist who first discovered them. The strand with the Okazaki fragments is known as the lagging strand .)

The leading strand can be extended from a single primer, whereas the lagging strand needs a new primer for each of the short Okazaki fragments. The overall direction of the lagging strand will be 3' to 5', and that of the leading strand 5' to 3'. A protein called the sliding clamp holds the DNA polymerase in place as it continues to add nucleotides. The sliding clamp is a ring-shaped protein that binds to the DNA and holds the polymerase in place. As synthesis proceeds, the RNA primers are replaced by DNA. The primers are removed by the exonuclease activity of DNA pol I, which uses DNA behind the RNA as its own primer and fills in the gaps left by removal of the RNA nucleotides by the addition of DNA nucleotides. The nicks that remain between the newly synthesized DNA (that replaced the RNA primer) and the previously synthesized DNA are sealed by the enzyme DNA ligase , which catalyzes the formation of phosphodiester linkages between the 3'-OH end of one nucleotide and the 5' phosphate end of the other fragment.

Once the chromosome has been completely replicated, the two DNA copies move into two different cells during cell division.

The process of DNA replication can be summarized as follows:

  1. DNA unwinds at the origin of replication.
  2. Helicase opens up the DNA-forming replication forks these are extended bidirectionally.
  3. Single-strand binding proteins coat the DNA around the replication fork to prevent rewinding of the DNA.
  4. Topoisomerase binds at the region ahead of the replication fork to prevent supercoiling.
  5. Primase synthesizes RNA primers complementary to the DNA strand.
  6. DNA polymerase III starts adding nucleotides to the 3'-OH end of the primer.
  7. Elongation of both the lagging and the leading strand continues.
  8. RNA primers are removed by exonuclease activity.
  9. Gaps are filled by DNA pol I by adding dNTPs.
  10. The gap between the two DNA fragments is sealed by DNA ligase, which helps in the formation of phosphodiester bonds.

Table 14.1 summarizes the enzymes involved in prokaryotic DNA replication and the functions of each.

Density-Gradient Centrifugation

A small amount of DNA in a concentrated solution of cesium chloride is centrifuged until equilibrium is closely approached. The opposing processes of sedimentation and diffusion have then produced a stable concentration gradient of the cesium chloride. The concentration and pressure gradients result in a continuous increase of density along the direction of centrifugal force. The macromolecules of DNA present in this density gradient are driven by the centrifugal field into the region where the solution density is equal to their own buoyant density. 11 This concentrating tendency is opposed by diffusion, with the result that at equilibrium a single species of DNA is distributed over a band whose width is inversely related to the molecular weight of that species (Fig. 1).

Ultraviolet absorption photographs showing successive stages in the banding of DNA from E. coli. An aliquot of bacterial lysate containing approximately 10 8 lysed cells was centrifuged at 31,410 rpm in a CsCl solution as described in the text. Distance from the axis of rotation increases toward the right. The number beside each photograph gives the time elapsed after reaching 31,410 rpm.

If several different density species of DNA are present, each will form a band at the position where the density of the CsCl solution is equal to the buoyant density of that species. In this way DNA labeled with heavy nitrogen (N 15 ) may be resolved from unlabeled DNA. Figure 2 shows the two bands formed as a result of centrifuging a mixture of approximately equal amounts of N 14 and N 15 Escherichia coli DNA.

a: The resolution of N 14 DNA from N 15 DNA by density-gradient centrifugation. A mixture of N 14 and N 15 bacterial lysates, each containing about 10 8 lysed cells, was centrifuged in CsCl solution as described in the text. The photograph was taken after 24 hours of centrifugation at 44,770 rpm. b: A microdensitometer tracing showing the DNA distribution in the region of the two bands of Fig. 2a. The separation between the peaks corresponds to a difference in buoyant density of 0.014 gm. cm. −3

In this paper reference will be made to the apparent molecular weight of DNA samples determined by means of density-gradient centrifugation. A discussion has been given 10 of the considerations upon which such determinations are based, as well as of several possible sources of error. 12



The production and activity of E. coli DnaA protein is regulated in several ways: by transcription, intracellular localization, and conformation (Katayama et al. 2010 Leonard and Grimwade 2010 Kaguni 2011). DnaA protein is stable, but replication cycle-specific dnaA gene transcription is important for sustaining well-timed initiation of replication (Bogan and Helmstetter 1997 Riber and Løbner-Olesen 2005). This can be explained by the role of ATP-DnaA in activating initiation (Kurokawa et al. 1999 Nishida et al. 2002) and the idea that newly synthesized DnaA preferentially binds ATP, the cellular level of which is 10-fold higher than that of ADP (Fig. 3).

Also, timely initiation of replication during the cell cycle requires specific chromosomal regions termed DARS (DnaA-reactivating sequence) 1 and DARS2. These regions bind ADP-DnaA molecules and promote the regeneration of ATP-DnaA by nucleotide exchange (Figs. 2 and 3) (Fujimitsu et al. 2009).

It has been estimated that there are 500–2000 DnaA molecules per cell, depending on strain backgrounds and growth rates (Sekimizu et al. 1988 Chiaramello and Zyskind 1989 Hansen et al. 1991a). oriC can bind 10–20 DnaA molecules (Messer 2002). A considerable number of DnaA molecules can be titrated at a specific chromosomal locus termed datA that is required for repressing rifamcipin-resistant unregulated initiation events (Figs. 2 and 3) (Kitagawa et al. 1998 Morigen et al. 2005).

The ATP-DnaA level fluctuates during the replication cycle, peaking around the time of initiation (Kurokawa et al. 1999). ATP-DnaA hydrolysis is required to reduce ATP-DnaA levels after initiation (Fig. 3). This regulation, termed RIDA (regulatory inactivation of DnaA), is coupled with the action of the DNA polymerase III holoenzyme. RIDA is crucial for regulating initiation so that it occurs only once per generation (Katayama et al.1998 Kato and Katayama 2001 Su’etsugu et al. 2004 Camara et al. 2005).

Regulation of DnaA Gene Transcription

The cellular DnaA concentration was found to be constant irrespective of growth medium and the cell cycle (Hansen et al. 1991a). However, the transcription of the dnaA gene varies in a replication cycle-dependent manner (Bogan and Helmstetter 1997). The main reason for the fluctuation seems to be that the dnaA gene promoter is sequestered by SeqA for almost the same duration as the origin (Fig. 3 Table 1) (Campbell and Kleckner 1990 Lu et al. 1994 Riber and Løbner-Olesen 2005). During sequestration, the promoter is unavailable to the transcription machinery. Because the dnaA gene is situated near oriC, this contributes to reducing the production of DnaA and thus the initiation potential (i.e., the ability to initiate replication) soon after new replication forks have been launched (Riber and Løbner-Olesen 2005). The dnaA promoter area also contains DnaA boxes and the promoter was found to be capable of autoregulation (Fig. 3 Table 1) (Messer and Weigel 1997 Hansen et al. 2007). Translation of dnaA mRNA has not been well characterized, but it is known that the start codon, GTG, functions inefficiently in E. coli.

Binding of DnaA to Sites Other than oriC

As mentioned, the DnaA protein functions not only as the initiator but also as a gene regulatory protein. There are about 300 high-affinity DnaA binding sites and a very large number of low-affinity sites around the chromosome (Kitagawa et al. 1996 Roth and Messer 1998 Hansen et al. 2007). As the chromosome is replicated, the DnaA binding sites are duplicated and contribute to titration of DnaA away from oriC. The main titration site, datA, is situated near oriC and is duplicated soon after initiation of replication (Kitagawa et al. 1996, 1998 Ogawa et al. 2002 Morigen et al. 2003, 2005). In vivo studies indicate that the datA site binds on average 60 DnaA molecules (Hansen et al. 2007). The datA site should therefore contribute to reducing the initiation potential at oriC when oriC is still in sequestration. The datA site is about 1 kb in size and contains five high-affinity DnaA binding sites and about 25 low-affinity sites (Fig. 2 Table 1) (Kitagawa et al. 1996, 1998 Hansen et al. 2007). High-affinity DnaA boxes 2 and 3 are crucial for efficient binding of DnaA to datA. It is possible that the DnaA bound to these sites function as a core for further cooperative DnaA binding (Ogawa et al. 2002). The production of DnaA protein as cells grow and the generation of binding sites as the chromosome is replicated, as well as sequestration and RIDA, has been simulated in silico (Hansen et al. 1991 Browning et al. 2004 Atlas et al. 2008) and may indicate that initiation occurs as soon as enough ATP-DnaA has accumulated at oriC. However, cells with similar numbers of origins and chromosomal DnaA binding sites but different growth conditions have been reported to contain different amounts of DnaA per origin at initiation (Torheim et al. 2000 Flåtten et al. 2009). So, although origin firing as soon as enough ATP-DnaA has accumulated is an attractive model of cell cycle regulation, a complete understanding of regulation encompassing different growth conditions is still lacking even in the well-characterized E. coli bacterium.

Regulation of the DnaA Nucleotide Form by DARS and Acidic Phospholipids

E. coli cells can convert ADP-DnaA to ATP-DnaA by nucleotide exchange (Fig. 3) (Kurokawa et al. 1999 Fujimitsu et al. 2009). The DARS1 and DARS2 sequences promote this reaction and are located halfway within the intergenic region between oriC and terC, to the right and left of oriC, respectively (Fig. 2A) (Fujimitsu et al. 2009). A common feature of DARS1 and DARS2 is the presence of a DnaA box cluster, in which three DnaA boxes are similarly oriented and are located at a similar distance (Fig. 2B). Multiple ADP-DnaA molecules can form complexes with DARS, which facilitates the release of ADP from DnaA. The resultant apo-DnaA molecules are likely released from DARS because of reduced complex formation activity, which allows the binding of ATP and DnaA reactivation. Increasing the cellular copy number of DARS1 or DARS2 increases the ATP-DnaA level, inducing extra initiation events, whereas deleting both DARS1 and DARS2 causes a delay in initiation (Fujimitsu et al. 2009). Newly translated DnaA protein binds ATP, providing cells with a basal level of ATP-DnaA. However, this alone is not enough to initiate replication in a timely manner. Thus, the function of DARSs is crucial for timely initiation (Fig. 3 Table 1). Functional regulation of DARS remains to be clarified, except for the fact that each DARS contains a regulatory region in addition to the common sequence bearing the DnaA box cluster (Fujimitsu et al. 2009 Leonard and Grimwade 2009). In addition to DARS, acidic phospholipids such as cardiolipin and phosphatidylglycerol play an important role in the nucleotide release of ADP-DnaA (Sekimizu and Kornberg 1988), and DnaA is reactivated by exchange of the bound nucleotide in vitro in the presence of oriC and ATP (Crooke et al. 1992). Reduction of the acidic phospholipids in vivo can inhibit initiation at oriC (Xia and Dowhan 1995), but exactly how acidic phospholipids affect DnaA activity during the cell cycle remains to be elucidated.

Regulation of DnaA Multimer Formation by DiaA

DiaA forms homotetramers and each protomer contains a specific site for binding to DnaA domain I (Fig. 3) (Ishida et al. 2004 Keyamura et al. 2007, 2009). These features allow a single DiaA tetramer to bind multiple molecules of DnaA, which can stimulate cooperative binding of DnaA to oriC and the unwinding reaction (Fig. 3 Table 1). The binding of ATP-DnaA to low-affinity binding sites within oriC is enhanced by DiaA (Fig. 1) (see Bell and Kaguni 2013 Leonard and Méchali 2013). In diaA-disrupted mutant cells, replication initiation is delayed and initiation at sister oriC copies occurs asynchronously in rapidly growing cells (Ishida et al. 2004 Keyamura et al. 2007, 2009). These data are consistent with the observation that replication is initiated asynchronously in mutants bearing DnaA box R4-deleted oriC (Bates et al. 1995), because the binding of DnaA to high-affinity DnaA box R4 enhances cooperative DnaA binding to low-affinity sites.

After DUE unwinding in oriC complexes, DiaA must be released from DnaA (Keyamura et al. 2009). The DiaA-binding site of DnaA domain I is also used to bind DnaB helicase. However, the mechanism of DiaA-DnaA dissociation has not yet been elucidated.

DiaA orthologs are evolutionarily conserved in bacterial species (Keyamura et al. 2007). In addition, the HobA protein of Helicobacter pylori (Hp), a member of ε-Proteobacteria, displays functional and structural similarity to DiaA, although there is no significant sequence similarity (Table 1) (Natrajan et al. 2007, 2009 Zakrzewsak-Czerwinska et al. 2007 Zawilak-Pawlik et al. 2007, 2011 Terradot et al. 2010).

Regulation of the DnaA Nucleotide Form by RIDA

After ATP-DnaA promotes replication initiation, it is hydrolyzed in a manner dependent on a complex consisting of ADP-Hda protein and the DNA-loaded clamp (Fig. 4 Table 1) (Katayama et al. 1998 Kato and Katayama 2001 Su’etsugu et al. 2008). The resultant ADP-DnaA is inactive in initiation. This system is termed RIDA (regulatory inactivation of DnaA). RIDA is crucial for DnaA inactivation and thereby effectively supports once-per-generation initiation (Kurokawa et al. 1999 Camara et al. 2005 Riber et al. 2009). The hda gene is required for promoting cell proliferation, decreasing cellular ATP-DnaA levels and repressing overinitiation (Kato and Katayama 2001 Fujimitsu et al. 2008 Charbon et al. 2011). Incubation of temperature-sensitive hda mutant cells at the restrictive temperature leads to overinitiation of replication and induces inhibition of cell division, producing filamentous cells (Fujimitsu et al. 2008). Inhibition of cell division is thought to be a consequence of checkpoint regulation, but the exact mechanism by which this occurs remains unknown. DnaA AAA + sensor II motif Arg-334 is specifically required for ATP-DnaA hydrolysis and expression of a DnaA R334A mutant protein causes overinitiation and inhibition of cell growth in an oriC-dependent manner (Table 1) (Nishida et al. 2002).

The basic mechanism of RIDA. When DNA polymerase III holoenzyme completes Okazaki fragment synthesis on the lagging strand, the clamp subunit is released from the DNA polymerase III core and remains on the synthesized DNA. ADP-Hda binds to the hydrophobic pocket of the DNA-loaded form of the clamp via the clamp-binding motifs in the amino terminus of Hda. The resultant ADP-Hda-clamp-DNA complex interacts with and promotes the DnaA-bound ATP hydrolysis, releasing ADP-DnaA back into the DnaA cycle. The interaction between the AAA + domains of Hda and DnaA is crucial, and is assisted by the interaction between Hda and DnaA domain IV (DNA-binding domain). N, Hda-amino terminus I-II, DnaA domain I-II III, DnaA domain III (AAA + domain) IV, DnaA domain IV (DNA-binding domain).

Hda protein consists of a short amino-terminal region containing the clamp-binding motif and an AAA + domain that is homologous to DnaA domain III (Fig. 4) (Dalrymple et al. 2001 Kato and Katayama 2001 Kurz et al. 2004 Su’etsugu et al. 2005 Xu et al. 2009). The Hda clamp-binding motif is commonly present in clamp-binding proteins such as DNA polymerase III core subunit α (Dalrymple et al. 2001). It binds to the hydrophobic pocket of the clamp, which is the same site to which DNA polymerase III subunit α binds (Dalrymple et al. 2001 Kurz et al. 2004 Su’etsugu et al. 2005). The Hda AAA + domain specifically binds ADP, but not ATP (Su’etsugu et al. 2008). ADP-Hda is monomeric and active in RIDA, whereas apo-Hda is multimeric and inactive in RIDA (Su’etsugu et al. 2008). The Hda AAA + domain bears a specific Arg residue (i.e., Arg finger) that is crucial for promoting DnaA-ATP hydrolysis (Su’etsugu et al. 2005). This residue may participate in forming the ATP hydrolysis catalytic center by direct DnaA-Hda interaction. This is a common characteristic of many AAA + proteins (Neuwald et al. 1999 Ogura et al. 2004 Indiani and O’Donnell 2006). In addition to the Arg finger, specific residues within the AAA + domains of DnaA and Hda are required for DnaA-Hda interaction and DnaA-ATP hydrolysis (Nakamura et al. 2010). DnaA domain IV (DNA binding domain) promotes this interaction by binding to Hda (Fig. 4) (Keyamura and Katayama 2011).

During DNA elongation, the clamps remain on the lagging strand after Okazaki fragments are synthesized and the DNA polymerase III core is released (Yuzhakov et al. 1996 Balakrishnan and Bambara 2013 Goodman and Woodgate 2013 Hedglin et al. 2013 MacAlpine and Almouzni 2013). DNA-loaded, DNA polymerase-free clamps bind ADP-Hda, resulting in the activation of RIDA (Su’etsugu et al. 2004, 2008). In addition, it is possible that because a clamp is a homodimer, Hda and DNA polymerase III subunit α bind to each protomer of the same clamp to allow Hda to hydrolyze DnaA-ATP as soon as replication forks are under way (Johnsen et al. 2011). DNA-free clamps are inactive in RIDA, although they can bind Hda, which ensures the timely and replication-coupled activation of RIDA. The dsDNA region flanking the clamp is required for RIDA, and may be recognized by DnaA (Fig. 4) (Su’etsugu et al. 2004).

The ADP-Hda-clamp-DNA complex is stable, whereas the affinity of this complex for DnaA is weak (Su’etsugu et al. 2008). This is consistent with the fact that the complex is reused for cyclically interacting with multiple ATP-DnaA molecules, and that there are only about 100 Hda molecules per cell (Katayama et al. 2010).

The main principle of RIDA, which is the use of DNA-loaded clamps for replication-coupled negative feedback to the initiator protein, is evolutionarily conserved from bacteria to eukaryotes, including yeast, Xenopus, and human cells (Table 1) (Katayama et al. 2010 Zielke et al. 2012).

Southeast Student Exploring E. Coli DNA Replication in NSF Summer Research

Adrienne Brauer knows all about preparation, cadence, tempo and pace as a long-distance runner on the cross country and track teams at Southeast Missouri State University.

This summer, though, she’s repurposing those skills off the course in a National Science Foundation (NSF) Research Experiences for Undergraduates (REU) at Washington University in St. Louis where she’s working with Dr. Petra Levin, biology professor and co-director of the plant and microbial biosciences graduate program, putting in the hours, preparing for graduate school and earning the experience for a career centered in microbiology research.

Adrienne, of Oakford, Illinois, knows that success comes with training, laying the groundwork, conditioning, clearing the hurdles and striding to the finish line. A Southeast senior, Adrienne is a microbiology major with a minor in chemistry. She’s hoping her experience at Washington University helps prepare her for her next step in her educational journey.

Adrienne says she’s conducting her research this summer along with post-doctoral and other undergraduate students, examining E. coli’s ability to become more antibiotic resistant under stress. She says she’s studying how E. coli replicate their DNA during rapid growth. When the cell is growing too fast for DNA replication to keep up, it undergoes multifork replication this occurs when a cell’s DNA begins a second (or third or fourth) round of replication before the first round has been completed, she said.

“We use fluorescent proteins to label certain sites on the chromosome, and when you view these cells under the microscope, you see the origin and terminus glow bright red and green,” Adrienne said. “Using fluorescent imaging has been such a neat experience for me here. The microscope we use is very high tech and even allows us to make ‘movies’ of cells growing and moving over a course of hours.”

Later in the summer, she plans to clone a handful of genes into the strains she’s been working with and observing the effect on DNA replication.

“My time in the lab has been particularly rewarding and productive thanks to the two years of research I’ve spent in Dr. (Jeremy) Ellermeier’s lab at Southeast,” she said. “He taught me so much that has come into play in Dr. Levin’s lab!”

Adrienne also is attending seminars and research classes at Washington University to learn helpful skills for graduate school, such as the “ins and outs of good researchers,” ethical considerations, communication with audiences, writing a mock grant proposal, conducting a literature review and participating in a poster presentation of her research at the conclusion of the program.

“Everyone is helpful and as a team they’ve already taught me so much,” she said. “I’ve had a taste of what my future will be like as a grad student. I’m in the lab and attending workshops and classes up to nine hours a day, learning about careers in biological sciences, new ways to analyze bacterial cells and how to become a more independent researcher creating hypotheses of my own.”

Outside of the lab, Adrienne and the other undergraduate interns have been attending classes and workshops to improve their research skills and prepare them for graduate school.

Adrienne Brauer, center, is a member of the Southeast women’s track and field team that placed second at the recent Ohio Valley Conference Track and Field Championships held at Southeast in May.

“We meet and talk with professionals from biotech companies like Pfizer and Bayer and have one-on-one advising meetings with faculty who help guide us in the right direction, whether that be pursuing a Ph.D., medical school or MTSP (Medical Scientist Training) program,” she said. “By the end of the summer, I’ll feel confident applying to some of the best microbiology programs in the country.”

Adrienne and her fellow interns also have explored St. Louis and have spent down time in the Danforth residence hall, where they’ve hosted unofficial journal clubs in the evening to discuss best ways to observe substrate localization in bacteria, she said. They’ve also had game nights, and, because she’s living near Forest Park, she’s also running in the park almost every day.

She’s says she’s appreciated the opportunity to work with both talented students and faculty members in the NSF REU this summer.

“I’m surrounded by great professionals in the field I want a career in,” she said. “Not only am I feeding my curiosity in microbiology but I’m gaining skills and knowledge necessary for me getting into a great grad school. I’m planning on getting my Ph.D. in microbiology, and the schools with the best programs are very competitive. This REU will help me immensely.”

She’s not exactly sure what her future career will look like, but it will definitely involve research.

“As much as I’ve enjoyed working with Salmonella at SEMO, I can’t wait to branch out and work with other bacteria. I would love to work for the National Institutes of Health or the CDC, but I would also enjoy teaching at a university with a research lab myself,” she said.

Adrienne says her high school science teachers sparked her interest in the sciences early on and set her on a path in science, technology, engineering and mathematics (STEM).

“They challenged me in class and made me love the questions in science that really make you think, which were more exciting to me than simply memorizing the definitions of vocabulary words,” she said. “The ‘aha moment’ I get studying biology is incredibly rewarding and makes me keep coming back for more,” she said. “Understanding the intricate mechanisms by which cells operate is really exciting and discovering previously unknown mechanisms is the goal we’re after.”

Replication in Prokaryotes

The replication begins at a specific initiation point called OriC point or replicon. (Replicon: It is a unit of the genome in which DNA is replicated it contains an origin for initiation of replication) It is the point of DNA open up and form open complex leading to the formation of prepriming complex to initiate replication process.

The OriC site is situated at 74″ minute near the ilv gene. The OriC site consists of 245 basepairs, of which three of 13 basepair sequence are highly conserved in many bacteria and forms the consensus sequences (GATCTNTTNTTTT). Close to OriC site, there are four of 9 basepair sequences each (TTATCCACA).

The sequence of reactions in the initiation process is as follows:

a) Dna A protein recognizes and binds up to four 9bp repeats in OriC to form a complex of negatively supercoiled OriC DNA wrapped around a central core of Dna A protein monomers. This process requires the presence of the histone like HU or 1 HC proteins to facility DNA bending.

b) Dna A protein subunits then successively melt three tandemly repeated 13bp segments in the presence of ATP at >=22*C (open complex).

c) The Dna A protein then guides a Dna B – Dna C complex into the melted region to form a so called prepriming complex. The Dna C is subsequently released. Dna B further unwinds open complex to form prepriming complex.

d) DNA gyrase, single stranded binding protein (SSB), Rep protein and Helicase – II are bound to prepriming complex and now complex is called as priming complex.

e) In the presence of gyrase and SSB, helicases further unwinds the DNA in both directions so as to permit entry of primase and RNA polymerase. Then RNA polymerase forms primer for leading strand synthesis while primase in the form of primosome synthesis primer for lagging strand synthesis.

f) To the above complex, DNA polymerase – III will bind and forms replisome.

REPLISOME: It is the multiprotein structure that assembles at the bacterial replicating fork to undertake synthesis of DNA. It contains DNA polymerase and other enzymes.

Now the stage is set for the initiation of synthesis and the elongation to proceed. But this occurs in two mechanistically different pathways in the 5′–>3′ template strand and 3′–>5′ template strand.

a) Initiation of synthesis and Elongation on the 5′–>3′ template (synthesis of leading strand) (If replication fork moves in 3′–>5′ direction)

The DNA daughter strand that is synthesized continuously on 5′–>3′ template is called leading strand. DNA pol-III synthesizes DNA by adding 5′-P of deoxynucleotide to 3′-OH group of the already presenting fragment. Thus chain grows in 5′–>3′ direction. The reaction catalyzed by DNA pol-III is very fast. The enzyme is much more active than DNA pol – I and can add 9000 nucleotides per minute at 37*C. The RNA primer that was initially added by RNA polymerase is degraded by RNase.

b) Initiation of synthesis and Elongation on 3′–>5′ template when fork moves in 3′–>5′ direction (Synthesis of lagging strand)

The daughter DNA strand which is synthesized in discontinuous complex fashion on the 3′–>5′ template is called lagging strand. It occurs in the following steps:

i) Synthesis of Okazaki fragment:

To the RNA primer synthesized by primosome, 1000-2000 nucleotides are added by DNA pol-III to synthesis Okazaki fragments.

ii) Excision of RNA primer:

When the Okazaki fragment synthesis was completed up to RNA primer, then RNA primer was removed by DNA pol – I using its 5′–>3′ exonuclease activity.

iii) Filling the gap (Nick translation)

The gap created by the removal of primer, is filled up by DNA pol – I using the 3′-OH of nearby Okazaki fragment by its polymerizing activity.

iv) Joining of Okazaki fragment: (Nick sealing)

Finally, the nick existing between the fragments are sealed by DNA ligase which catalyze the formation of phosphodiester bond between a 3′-OH at the end of one strand and a 5′ – phosphate at the other end of another fragment. The enzyme requires NAD for during this reaction.


Termination occurs when the two replicating forks meet each other on the opposite side of circular E.Coli DNA. Termination sites like A, B, C, D, E and F are found to present in DNA. Of these sites, Ter A terminates the counter clockwise moving fork while ter C terminates the clockwise moving forks. The other sites are backup sites. Termination at these sites are possible because, at these sites tus protein (Termination utilizing substance) will bound to Dna B protein and inhibits its helicase activity. And Dna B protein released and termination result.

After the complete synthesis, two duplex DNA are found to be catenated (knotted). This catenation removed by the action of topoisomerase. Finally, from single parental duplex DNA, two progeny duplex DNA synthesized.


Especially initiation of replication is regulated. Dna A protein when available in high concentration then ratio of DNA to cell mass is quiet high but at low Dna A concentration, the ratio found to be low. This shows that Dna A protein regulates the initiation of replication.

DNA Replication, Repair, and Mutagenesis

Polymerase III

Polymerase I plays an essential role in the replication process in E. coli, but it is not responsible for the overall polymerization of the replicating strands. The enzyme that accomplishes this is a less abundant enzyme, polymerase III (pol III). (A DNA polymerase II has also been isolated from E. coli, but it probably plays no role in DNA synthesis.) Pol III catalyzes the same polymerization reaction as pot I but has certain distinguishing features. It is a very complex enzyme and is associated with eight other proteins to form the pol III holoenzyme. (The term holoenzyme refers to an enzyme that contains several different subunits and retains some activity even when one or more subunits is missing.) Pol III is similar to pol I in that it has a requirement for a template and a primer but its substrate specificity is much more limited. For a template pol III cannot act at a nick nor can it unwind a helix and carry out strand displacement. The latter deficiency means that an auxiliary system is needed to unwind the helix ahead of a replication fork. Pol III, like pol I, possesses a 3′ → 5′ exonuclease activity, which performs the major editing function in DNA replication. Polymerase III also has a 3′exonuclease activity, but this activity does not seem to play a role in replication.

Pol I and pol III holoenzyme are both essential for E. coli replication. The need for two polymerases seems to be characteristic of all cellular organisms but not all viruses, e.g., E. coli. Phage T4 synthesizes its own DNA polymerase, which is capable of carrying out all functions necessary for synthesizing phage DNA.

In the usual polymerization reaction, the activation energy for phosphodiester bond formation comes from cleaving of the triphosphate. Since DNA ligase can use a monophosphate, another source of energy is needed. This energy is obtained by hydrolyzing either ATP or NAD the energy source depends on the organism from which the DNA ligase is obtained.

Ligases have two major functions: the sealing of single-strand breaks produced randomly in DNA molecules by nucleases and the joining of fragments during a particular stage of replication. DNA ligases are enzymes that can form a phosphodiester bond at a single-strand break in DNA, a reaction between a 3′-OH group and a 5′-monophosphate. These groups must be termini of adjacent base-paired deoxynucleotides ( Figure 24-4 ).

Bacteria usually contain a single species of ligase. Mammalian cells possess two DNA ligases (I and II) present in very small amounts compared with bacteria. Both eukaryotic ligases are located in the nucleus. Ligase I is predominant in proliferating cells and presumably plays a role in DNA replication ligase II predominates in resting cells.


I've started reading microcosm by my favorite science writer, Carl Zimmer [Buy This Book!]. Watch for a review, coming soon.

I was mildly disappointed to see Carl repeat a common myth about DNA replication in E. coli on page 29. Since we often use this myth to teach critical thinking in our undergraduate classes, I thought it would be worthwhile to discuss it here.

Today I'm going to present the problem and let everyone think about a possible solution. On Sunday, I'll publish the answer. (If you know the solution, you are not allowed to post it in the comments&mdashI'll delete those comments. You can ask for clarification or speculate.)

Here's what Carl says at the top of page 29.

Today, we're not concerned about the 20 minute generation time but I note, for the record, that the average generation time of E. coli, in vivo, is about one day. I also want to mention that the 20 minute generation time is an extreme example that's achieved only under the most extraordinary circumstances. Typical generation times in the lab are about 30 minutes.

However, that's not the problem. Let's assume a generation time of 20 minutes.

In the next paragraph Carl says .

What Carl is referring to the the assembly of replication complexes (replisomes) at the origin of replication. Once those complexes are assembled, replication fires off and proceeds in opposite directions (bidirectionally) until the two fork meet at the opposite side of the chromosome.

Carl is correct when he says that the forks move at 1000 nucleotides per second. Later on in his book he mentions that the size of the E. coli chromosome is 4,600,000 base pairs or 4,600 kb (p. 116). At 1000 nucs per second it would take 4600 second to replicate this DNA if there was only one replication fork. Since there are two, it will take 2,300 seconds.

You can do the math. This is 38 minutes. It is a correct number&mdashit takes at least 38 minutes to replicate the E. coli chromosome, not 20 minutes as stated earlier. It is true that the generation time of E. coli can be as short as 20 minutes under extraordinary circumstances.

Here's the problem. How can E. coli divide faster than it can replicate it's chromosome?

Because Science!

Good morning, everyone! It hasn’t been very long since you heard from me last, but I hope you all had a good night! Although I don’t normally get up at this hour on Thursdays, I have a test to study for, so I’m here to bring you the wonders of DNA replication. Today’s subject: prokaryotes!

We know that, when one cell splits into two, it has to copy its DNA so that each of the daughter cells will have a copy. We also know that the mechanism for this is a pretty straightforward one: the strands of the double helix are pulled apart, and each strand is used as a template to make its complementary strand. (This is called semiconservative, since each daughter copy has one old strand and one new one.)

High school biology students are also probably aware that DNA replication occurs at origins of replication, and replication forks move away from origins at both directions. In E. coli, the origin is OriC.

As mentioned in my post on transcription, unwinding the double helix creates tension (positive supercoiling) that can stop replication if it’s not alleviated. Thus, an enzyme called DNA gyrase (a type II topoisomerase) uses ATP to introduce negative supercoiling.

Other proteins are also important to the process of replication: helicase, an ATP-dependent enzyme, binds to a single-stranded region of DNA and then unwinds the rest of it by disrupting the hydrogen bonds between base pairs. SSBs, or single-stranded binding proteins, keep the single strands from reannealing before our polymerase can do its magic.

And, speaking of the polymerase, let’s introduce our main player! The main part of replication is carried out by an enzyme (or, I guess, a class of enzymes) called DNA polymerase. These use the single-stranded template of DNA to synthesize a complementary strand by assembling dNTPs in the proper order.

However, there’s a problem with this enzyme: it can only add nucleotides in a 5′ to 3′ direction. That’s no problem for the 3′-5′ strand, whose complementary strand (the “leading” strand) runs in that direction, but a little bit of a problem arises when dealing with the opposite template. In fact, the best our DNA polymerases can do is bend the 5′-3′ strand around and replicate it in small (1000-2000 bp) backwards. This leads to the creation of fragments called Okazaki fragments, which are then sealed together in the final product to form a whole “lagging” strand.

Another limitation of DNA polymerases is that they can’t start synthesizing a complementary strand from scratch. This is easily alleviated, though, by an enzyme called primase, an enzyme that synthesizes short stretches of RNA called “primers.” Primers give the DNA polymerase a starting point, and they’re replaced with DNA in the final product.

Now, you’re noticing that I’m referring to polymerases in the plural, implying that we don’t have just one. That’s true for both prokaryotes and eukaryotes (unlike RNA polymerase, which is the only prokaryotic RNA polymerase). To keep things a little ([coughs up a lung] A LOT) simpler, we’re going to start by looking at just prokaryotic replication, and there’s no better place to start than the polymerases.

Prokaryotes have five DNA polymerases that are designated with numerals I through V. I, II and V mostly just deal with DNA repair, so we won’t talk about them too much. Wikipedia describes IV as “an error-prone polymerase involved in non-targeted mutagenesis.” III is the one that carries out most DNA replication it’s not very abundant in cells, but it’s extremely processive (good at hanging on to the DNA template) in comparison to the others.

DNA polymerase III holoenzyme is composed of seventeen different subunits, each of which confers it some sort of functionality. The most noteworthy of these is the gamma-complex, which acts as a “clamp loader” by helping clamp the beta-dimer rings of the enzyme (which help keep it on the DNA) to the strand using ATP.

Now, once we actually get our polymerase III going and making DNA, DNA polymerase I gets its time to shine. This enzyme carries out the very important job of replacing the RNA primers with DNA. It’s also special because, in addition to being able to do this, is can remove nucleotides from a DNA strand in both the 3′ to 5′ and 5′ to 3′ direction.

The 3′ to 5′ exonuclease activity is useful because it means that this enzyme can quite literally check its work as it goes. If it discovers that it’s made a mistake, all it has to do is back up and chew the incorrect base off of the growing strand. Pretty nifty, if you think about it.

After polymerase I does its thing, an enzyme called ligase comes in an seals the nicks in the backbone left behind by all of this priming and Okizaki fragment-forming nonsense.

Finally, we need to terminate our DNA replication. In prokaryotes, this isn’t too difficult: it just requires the Ter sequence (GTGTGTTGT). Tus protein, a contrahelicase, binds here and basically just goes, “Nope nope nope, no helicase is passing through here!” The replication forks end, and replication is terminated.

See, not too bad, huh? Now we get to the really fun stuff—those stupid, show-off eukaryotes!

See a lot of errors? Feel the need to point them out? Please have mercy—I’m posting this without proofing it, because I’m kinda in a hurry here.

70 DNA Replication in Eukaryotes

By the end of this section, you will be able to do the following:

  • Discuss the similarities and differences between DNA replication in eukaryotes and prokaryotes
  • State the role of telomerase in DNA replication

Eukaryotic genomes are much more complex and larger in size than prokaryotic genomes. Eukaryotes also have a number of different linear chromosomes. The human genome has 3 billion base pairs per haploid set of chromosomes, and 6 billion base pairs are replicated during the S phase of the cell cycle. There are multiple origins of replication on each eukaryotic chromosome humans can have up to 100,000 origins of replication across the genome. The rate of replication is approximately 100 nucleotides per second, much slower than prokaryotic replication. In yeast, which is a eukaryote, special sequences known as autonomously replicating sequences (ARS) are found on the chromosomes. These are equivalent to the origin of replication in E. coli.

The number of DNA polymerases in eukaryotes is much more than in prokaryotes: 14 are known, of which five are known to have major roles during replication and have been well studied. They are known as pol α, pol β, pol γ, pol δ, and pol ε.

The essential steps of replication are the same as in prokaryotes. Before replication can start, the DNA has to be made available as a template. Eukaryotic DNA is bound to basic proteins known as histones to form structures called nucleosomes. Histones must be removed and then replaced during the replication process, which helps to account for the lower replication rate in eukaryotes. The chromatin (the complex between DNA and proteins) may undergo some chemical modifications, so that the DNA may be able to slide off the proteins or be accessible to the enzymes of the DNA replication machinery. At the origin of replication, a pre-replication complex is made with other initiator proteins. Helicase and other proteins are then recruited to start the replication process ((Figure)).

Difference between Prokaryotic and Eukaryotic Replication
Property Prokaryotes Eukaryotes
Origin of replication Single Multiple
Rate of replication 1000 nucleotides/s 50 to 100 nucleotides/s
DNA polymerase types 5 14
Telomerase Not present Present
RNA primer removal DNA pol I RNase H
Strand elongation DNA pol III Pol α, pol δ, pol ε
Sliding clamp Sliding clamp PCNA

A helicase using the energy from ATP hydrolysis opens up the DNA helix. Replication forks are formed at each replication origin as the DNA unwinds. The opening of the double helix causes over-winding, or supercoiling, in the DNA ahead of the replication fork. These are resolved with the action of topoisomerases. Primers are formed by the enzyme primase, and using the primer, DNA pol can start synthesis. Three major DNA polymerases are then involved: α, δ and ε. DNA pol α adds a short (20 to 30 nucleotides) DNA fragment to the RNA primer on both strands, and then hands off to a second polymerase. While the leading strand is continuously synthesized by the enzyme pol δ, the lagging strand is synthesized by pol ε. A sliding clamp protein known as PCNA (proliferating cell nuclear antigen) holds the DNA pol in place so that it does not slide off the DNA. As pol δ runs into the primer RNA on the lagging strand, it displaces it from the DNA template. The displaced primer RNA is then removed by RNase H (AKA flap endonuclease) and replaced with DNA nucleotides. The Okazaki fragments in the lagging strand are joined after the replacement of the RNA primers with DNA. The gaps that remain are sealed by DNA ligase, which forms the phosphodiester bond.

Telomere replication

Unlike prokaryotic chromosomes, eukaryotic chromosomes are linear. As you’ve learned, the enzyme DNA pol can add nucleotides only in the 5′ to 3′ direction. In the leading strand, synthesis continues until the end of the chromosome is reached. On the lagging strand, DNA is synthesized in short stretches, each of which is initiated by a separate primer. When the replication fork reaches the end of the linear chromosome, there is no way to replace the primer on the 5’ end of the lagging strand. The DNA at the ends of the chromosome thus remains unpaired, and over time these ends, called telomeres, may get progressively shorter as cells continue to divide.

Telomeres comprise repetitive sequences that code for no particular gene. In humans, a six-base-pair sequence, TTAGGG, is repeated 100 to 1000 times in the telomere regions. In a way, these telomeres protect the genes from getting deleted as cells continue to divide. The telomeres are added to the ends of chromosomes by a separate enzyme, telomerase ((Figure)), whose discovery helped in the understanding of how these repetitive chromosome ends are maintained. The telomerase enzyme contains a catalytic part and a built-in RNA template. It attaches to the end of the chromosome, and DNA nucleotides complementary to the RNA template are added on the 3′ end of the DNA strand. Once the 3′ end of the lagging strand template is sufficiently elongated, DNA polymerase can add the nucleotides complementary to the ends of the chromosomes. Thus, the ends of the chromosomes are replicated.

Telomerase is typically active in germ cells and adult stem cells. It is not active in adult somatic cells. For their discovery of telomerase and its action, Elizabeth Blackburn, Carol W. Greider, and Jack W. Szostak ((Figure)) received the Nobel Prize for Medicine and Physiology in 2009.

Telomerase and Aging

Cells that undergo cell division continue to have their telomeres shortened because most somatic cells do not make telomerase. This essentially means that telomere shortening is associated with aging. With the advent of modern medicine, preventative health care, and healthier lifestyles, the human life span has increased, and there is an increasing demand for people to look younger and have a better quality of life as they grow older.

In 2010, scientists found that telomerase can reverse some age-related conditions in mice. This may have potential in regenerative medicine. 1 Telomerase-deficient mice were used in these studies these mice have tissue atrophy, stem cell depletion, organ system failure, and impaired tissue injury responses. Telomerase reactivation in these mice caused extension of telomeres, reduced DNA damage, reversed neurodegeneration, and improved the function of the testes, spleen, and intestines. Thus, telomere reactivation may have potential for treating age-related diseases in humans.

Cancer is characterized by uncontrolled cell division of abnormal cells. The cells accumulate mutations, proliferate uncontrollably, and can migrate to different parts of the body through a process called metastasis. Scientists have observed that cancerous cells have considerably shortened telomeres and that telomerase is active in these cells. Interestingly, only after the telomeres were shortened in the cancer cells did the telomerase become active. If the action of telomerase in these cells can be inhibited by drugs during cancer therapy, then the cancerous cells could potentially be stopped from further division.

Section Summary

Replication in eukaryotes starts at multiple origins of replication. The mechanism is quite similar to that in prokaryotes. A primer is required to initiate synthesis, which is then extended by DNA polymerase as it adds nucleotides one by one to the growing chain. The leading strand is synthesized continuously, whereas the lagging strand is synthesized in short stretches called Okazaki fragments. The RNA primers are replaced with DNA nucleotides the DNA Okazaki fragments are linked into one continuous strand by DNA ligase. The ends of the chromosomes pose a problem as the primer RNA at the 5’ ends of the DNA cannot be replaced with DNA, and the chromosome is progressively shortened. Telomerase, an enzyme with an inbuilt RNA template, extends the ends by copying the RNA template and extending one strand of the chromosome. DNA polymerase can then fill in the complementary DNA strand using the regular replication enzymes. In this way, the ends of the chromosomes are protected.