Successful DNA replication is essential to maintain genome integrity and prevent disease. In human cells, over 6 billion base pairs of DNA need to be replicated accurately and packaged into chromatin each cell division cycle. Thus, the replisome is a highly regulated, dynamic machine that must work with speed and precision to duplicate the chromosomes successfully. Furthermore, it must work in the context of defects in the DNA template such as DNA damage, which present challenges to both genetic and epigenetic inheritance. DNA damage response (DDR) proteins are recruited and activated in these circumstances to ensure the successful completion of chromosome replication. Failures can lead to mutation, epigenetic changes, and other chromosomal aberrations that ultimately cause disease such as cancer.
Maintaining genome integrity during DNA replication is critical to prevent disease. We are interested in how cells faithfully duplicate their genomes even when challenged by difficult to replicate sequences, stable protein-DNA complexes, and DNA lesions. In addition, epigenetic inheritance also is challenged during DNA replication. We are developing new technologies to allow us to understand the processes that ensure the proper chromatin modifications (such as histone post-translational modifications) are maintained through the cell division cycle.
One example of a project in this area is our studies of SMARCAL1 -- a gene we identified in an RNAi screen as a genome maintenance protein that functions at stalled replication forks. Mutations in SMARCAL1 cause a human disease called Schimke immunoosseous dysplasia (SIOD). Our data indicate that th epleiotorpic phenotypes of this disease are due to defects in genome maintenance during DNA replication. We are currently using a variety of approaches including biochemistry, genetics, and cell biology to understand how SMARCAL1 functions during DNA replication and why exactly why defects in SMARCAL1 activity cause genome instability and disease.
Mammalian cells lack highly efficient, sequence defined origins of replication. Hence, it has been difficult (if not impossible) to track proteins as they move with a replication fork, visualize proteins that are recruited to damaged forks at low copy number, or to follow the process of chromatin replication. We have largely been limited to the sensitivity and resolution of light microscopy and immunofluorescent imaging. To overcome this technical limitation we developed iPOND (Isolation of Proteins on Nascent DNA). iPOND provides a unique opportunity to use a biochemical purification procedure to track proteins at replication forks and study chromatin deposition and maturation in mammalian cells. The technique provides high resolution (on the order of 1000-3000 base pairs) and sensitivity (we can monitor proteins like polymerases which are likely only present at one or two copies per fork). Importantly, the technique is compatible with unbiased approaches like mass spectrometry.
We used iPOND to define the timing histone deposition and chromatin maturation during DNA replication. Class 1 histone deacetylases are enriched at replisomes and remove pre-deposition marks on histone H4. Chromatin maturation continues even when decoupled from replisome movement. Furthermore, fork stalling causes changes in the recruitment and phosphorylation of proteins at the damaged fork. Checkpoint kinases catalyze H2AX phosphorylation, which spreads from the stalled fork to include a large chromatin domain even prior to fork collapse and double-strand break formation. Finally, we demonstrated a switch in the DDR at persistently stalled forks that includes MRE11-dependent and independent RAD51 assembly. iPOND promises to be a powerful technology to study protein dynamics during DNA replication, the DNA damage response, and chromatin maturation.