Current Project – DNA traffic jam

November 20, 2012

Since November 1st I am doing research on the mechanisms of polymerase exchange under DNA damage conditions in the laboratory of Dr. Joseph Loparo at Harvard Medical School in Boston, USA.

In case you do not belong to the small group of people who intuitively know what my projects entails, this page intends to make it more clear.

Below you see a photograph taken out of my window at night. It symbolically stands for the need to regulate complex processes which are required to achieve a certain task. As with traffic a constant “flow” is also important for DNA replication. Every cell contains the same DNA so replication events are constantly occurring during life. However, there are certain “stop-lights” involved which come into play under certain conditions. During rush-hour traffic lights are controlled in a different manner then at 3 o’clock at night (at least this is desired).

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In a living cell it’s almost always rush-hour. Still, many checks lead to a surprisingly perfect flow with very few mistakes occurring. Despite this, external “stress factors” can distort the flow. Imagine the driver on the right would not care about his or her red sign. The following accident would require the set-up of a detour around the site of the accident so that a minimum of traffic flow can be guaranteed. However, this detour will make the overall traffic situation less efficient (especially during rush-hour!!) and everything will take way more time than needed. But at least a total breakdown can be prevented.

The sun’s UV-light leads to sort like accidents during DNA replication flow, because it changes the regular DNA structure which causes the formation of a so-called “roadblocks”. Regular enzymes called Polymerase III that catalyze the DNA replication can not replicate across these roadblocks. Polymerase III normally guarantees a high fidelity DNA replication meaning that there is an extremely low copy error rate. A problem related to this high accuracy is that Polymerase III is not very tolerant to DNA damage. So in order to prevent a total DNA replication breakdown a “detour” enzyme comes in (for example Polymerase II or IV). These enzymes are build up differently and are more tolerant to DNA roadblocks. However, they slow down the whole replication process and can lead to even more errors when used for too long.

This is why the use of Polymerases II and IV needs to be tightly regulated. Despite the importance of this process currently not a lot is known about it. And this is what my work will focus on.

The following explanations might become a bit more technical, but I will try to keep it to a minimum. To recap you knowledge about the general DNA replication process I want to introduce Fig.1. It depicts the protein complex that is required and sufficient for DNA replication in Escherichia coli bacteria. Together these proteins constitute the replisome.

Fig. 1: Schematic overview of the replisome. DNA helicase unwinds the double stranded DNA. Two single DNA strands arise; one in 5’-3’ and the other one in 3’-5’ direction. Because the DNA polymerase complex is only able to synthesize into the 5’-3’ direction in a continuous fashion, the inverse direction needs to be synthesized in small bits called Okazaki fragments. The strands are therefore termed “leading” and “lagging” strand, respectively. Primase attaches primers to the DNA so that replication can be repeatedly initialized on the lagging strand. In E. coli the two DNA polymerases are tethered to the helicase by the γ clamp-loader complex which is especially required to load the β clamps through which the αεθ DNA polymerase subunit attaches to the single DNA strand. Single strand binding proteins (SSBs) ensure that the single strand does not coil up and remains accessible for the lagging strand polymerase (based on: van Oijen, Loparo, Annual Reviews Biochemistry, 2010).

Back to the switching between the polymerases under DNA damage conditions: This process is termed translesion synthesis response (TLS) because the DNA roadblock (lesion) is bridged. A few aspects of TLS are already known, but in order to understand what they entail it is important that you have good grasp of Fig. 1.

It has been elucidated by O’Donnell and coworkers (Molecular Cell, 2005) and Sutton and colleagues (PNAS, 2009) that the β clamps which tethers the αεθ polymerase III subunits to the DNA plays an essential role during TLS. Fig. 2 explains why this is the case.

Fig. 2: Structure of the DNA β clamp with its rim and cleft contacts sites for Polymerase IV. Within the structure of the β clamp two hydrophobic clefts have been identified. These seem to be able to accommodate certain amino acid residues of the so-called “little finger” domain of Polymerase IV. This essential key-lock mechanism is supported by additional rim contacts (Loparo, based on Bunting et al., EMBO, 2003).

Therefore the β clamp seems to serve as the basis for polymerase exchange during TLS.

The next question would be: How can we study the dynamical changes between Polymerase III and the other polymerases such as Polymerase II or IV?

In Dr. Loparos’s laboratory several different methods have been developed how these changes can be observed on a single-molecule basis. The focus of my project will especially  lie on the TLS polymerase II (Pol II) and how it is able to replace the replicative polymerase III (Pol III). Pol II is special in regard to its high-fidelity and proofreading capability by its 3’-5’ exonuclease. These features are not present in the other existing translesion polymerase and make Pol II an interesting object to study. Also there are indications existing that the Pol III – Pol II exchange might work differently than the Pol III – Pol IV exchange. Single-molecule biology only works when methodology originating from physics is combined with biologically relevant questions, and classical biochemical techniques.
Traditional biochemical studies have obtained almost all of the current DNA replication knowledge and are continuing to do so. However, bulk effects seem to average out the dynamic states which are so essential for protein functioning. Single-molecule and fluorescence methods are a powerful means to study the functional trajectories of proteins while they are functioning. Studying the formation of (replisome) complexes is a very interesting application and has been demonstrated to be successful. The development of procedures to very locally and quantitatively study the mechanisms and stoichiometry of polymerase exchange will be central to this project.
Fluorescent organic molecules (dyes) and inorganic molecules (quantum dots) will be used to label proteins and DNA. Innovative laser microscopic techniques such as Total Internal Reflexion
Fluorescence (TIRF) and Förster Resonance Energy Transfer (FRET) microscopy can then be applied to image and quantify the associations of the labelled single molecules.

Central to these approaches is a single-molecule primer extension assay. It works by the application of a microfluidic flow-cell in which a DNA molecule is attached to a glass surface and is consequently stretched by a laminar buffer flow (Fig. 3, top). Extending this linear molecular and observing the change via fluorescence microscopic techniques allows to make conclusions about polymerase behaviour and dynamics (Fig. 3, bottom).

Fig. 3: The principle of a laminar flow cell which can be used to determine the single-molecule dynamics of polymerases (top) and an example of the fluorescence patterns that can consequently be observed by dark field microscopy (bottom). See the text for more details (based on: Tanner & van Oijen, Methods in Enzymology, 2010).

Based on these data (DNA length extension and time) it now becomes possible to actually observe polymerase switching. Previous studies have shown that Pol III is much faster in synthesizing long DNA molecules than other polymerases. By determining the difference between polymerase extension speeds it therefore becomes possible to determine which polymerase is active. Furthermore, it has been determined that a single Pol III synthesizes about 900 base pairs before it leaves the replication fork and a new Pol III comes in. Between these events always a small pause occurs. The lengths of the pause depends on the polymerase concentration. This is logical because at a higher polymerase concentration a switch can occur faster because there simply more molecules available in a certain location. By measuring a number of single-molecule trajectories under different conditions and with different polymerase types it becomes feasible characterize polymerase dynamics. Fig. 4 shows the result of a plot where two different polymerases were tested. Because the pause length and the reaction speed are known it is possible to distinguish between Pol III (fast) and Pol IV (slow).

Fig. 4: Time and DNA length info from a flow-stretching assay helps to identify switching dynamics of polymerases if synthesis speed pausing behaviour is known (see also Fig. 3).

Most of the above described approaches and the derived knowledge is only valid for Pol III/Pol IV switching. The situation might be different for Pol III/Pol II switching. Elucidating Pol II switching behaviour with the above describes assay will be a main aspect of my project.

Please stay tuned for updates!

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