The Green Lab - Replication Fork Dynamics
DNA replication is a fundamental cellular process. A cell must copy its genome accurately and completely in order for the two daughter cells not to contain changes in the DNA sequence, or mutations. Despite decades of work attempting to understand this process much remains to be learned about the mechanisms and control of replicative processes. We are attempting to fill some of the gaps in our knowledge by dissecting protein-protein interactions occurring at replication forks. Our intention is to clarify how transitions between the sequential steps required for accurate replication and its associated processes are regulated. We are also using biochemical approaches to identify novel proteins involved in replication and its control.
Figure 1. PCNA organises many steps at the replication fork.
Here it is shown interacting with the replicative polymerases on both the leading and lagging strands,
and on the lagging strand also coordinating Okazaki fragment processing by Fen-1 and Ligase1.
The homotrimeric sliding clamp, PCNA, plays a key role in DNA replication by coordinating many of the enzymes involved.
This is achieved by direct interactions with key proteins, including the replicative polymerases delta (Polδ) and epsilon (Polε),
and the Okazaki fragment processing proteins flap endonuclease 1 (Fen-1) and ligase 1 (Lig1) (Figure 1).
However, this model is dramatically over-simplistic. In addition to Okazaki fragment processing, other replication associated functions,
including mismatch correction, DNA methylation, and chromatin assembly must also be spatially and temporally coordinated for
accurate genome duplication in an unperturbed cell cycle. All these processes utilise PCNA-interacting proteins, many of which
are implicated in cancer development.
DNA replication has to be controlled within the cell cycle to ensure that the genome is copied precisely once.
PCNA is implicated in this control through interactions with cell cycle regulators and licensing factors.
Replication is further complicated when the DNA template is damaged. Damaged DNA is not a substrate for replicative polymerases,
but specialized polymerases (such as Polη, Polι and Polκ) can utilize such templates for translesion synthesis (TLS).
PCNA is an essential cofactor for these TLS polymerases. Damage-induced replication stalling results in ubiquitination of PCNA which enhances interaction between TLS polymerases and PCNA,
facilitating a switch from replicative to TLS polymerases. This is just one cellular example of a protein handover mediated by PCNA,
but the detailed molecular mechanism of even this well studied case is still unclear and its analysis is currently a key aim of the TLS field.
More than 70 proteins have now been shown to interact with PCNA, many of them implicated in the maintenance of genomic stability;
clearly not all of these can associate on a single PCNA trimer at one time.
Therefore key questions include:
• How are the appropriate PCNA partners chosen during the sequential processes of replication?
• How are inappropriate interactions, such as the association of a TLS polymerase if the template is not damaged,
or association of chromatin assembly factors before ligation is complete, prevented?
Figure 2. Replication foci (or factories) visualised in the nucleus of a human cell co-transfected with RFP-PCNA (red) and GFP-polη (green).
Overlaying these two images (lower picture) gives yellow colour where the two proteins co-localise.Within the nucleus DNA synthesis and post-replication processes occur at discrete sites, known as replication foci or factories, where multiple replication forks, PCNA and replication proteins co-localize (Figure 2). The size of these structures is approximately 200 nm, at the resolution limit of the light microscope. At this resolution, co-localisation does not necessarily represent a true in vivo interaction. Many of the interactions between PCNA and replication or replication-associated proteins have been identified by yeast 2-hybrid, bioinformatic, or proteomics approaches. Such studies, however, report on interactions as they occur in isolation, not necessarily within the context of the presence of other proteins, let alone within a replication factory.
The aim of our studies is to obtain greater detail concerning protein-protein interactions at replication forks, and in particular protein transitions on PCNA.To this end we utilise in vitro and in vivo techniques to analyse protein interactions.
We are a molecular and cell biology lab that also uses biochemical techniques to address questions concerning DNA replication. We routinely express and purify recombinant proteins (using E. coli) and analyse protein interactions by pull downs, immunoprecipitations and western blots. Our model system is mammallian tissue culture cells, which we stably and transiently transfect with constructs that express fluorescent replication proteins or siRNAs to deplete specific proteins from cells. These cells are analysed by confocal microscopy, live cell imaging (for some of our cool movies click here) and FACS and assessed for tolerance to replication inhibitors such as hydroxyurea.
More specialised techniques include Surface Plasmon Resonance (SPR) analysis, which we perform on a Biacore machine. This enables us to measure the kinetic parameters of interactions between replication proteins in vitro. A good introduction to the principles of this technique is available here, and a comprehensive survey of the recent SPR literature can be found here. The machine that we use is situated in the Department of Biochemistry. Information from these studies enables us to build models to explain the transitions between the steps of replication, which can then be tested by altering in vivo conditions and analysing replicative outcomes.
Because the size of replication factories approaches the resolution limit of the light microscope, even state-of-the-art confocal microscopy is not able to give us information about protein interactions in vivo. We have been using a super-resolution STED (stimulated emission depletion) microscope manufactured by Leica to look in finer detail at replication structures, our recent paper on this is available here. We also overcome microscope resolution limits by the use of Fluorescence Resonance Energy Transfer (FRET) techniques to probe events at the nanometer scale in vivo and in real time. A good introduction to FRET can be found here, with reviews here or here. We are assisted in the application of these technically demanding microscopy techniques by the excellent confocal facilities here in the Department of Zoology.
More recently we have been developing screens to attempt to identify novel proteins involved in DNA replication. These include some utilising biochemical purification strategies and others based on mammalian 2-hybrid systems. The identification of novel players in this dynamic pathway might help us to understand how these complex events can be properly controlled in vivo.
