Imperial College London

Professor David S. Rueda

Faculty of MedicineDepartment of Infectious Disease

Chair of Molecular and Cellular Medicine
 
 
 
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Contact

 

+44 (0)20 3313 1604david.rueda Website

 
 
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Location

 

10N12ACommonwealth BuildingHammersmith Campus

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Summary

 

CRISPR/Cas9 off-target mechanism

Over the past decade, the application of CRISPR/Cas9 technology has revolutionised genome editing in biological research. Cas9 is a programmable endonuclease routinely used to generate sequence deletions, insertions, and even to regulate gene expression from bacteria to mammals. However, spurious off-target edits have represented a critical barrier to therapeutic applications. The molecular mechanism by which Cas9 binds and cleaves off-targets remains largely unknown, which is a significant problem that hinders the development of new and improved CRISPR/Cas9 systems with high accuracy and efficiency.
Using a combination of single-molecule approaches, traditional biochemistry, structural biology and cutting-edge genomics, we are elucidating the molecular mechanism by which Cas9 discriminates between on- and off-targets. Our goal is to help develop new and improved CRISPR/Cas9 technology to generate accurate and efficient genome editing tools for therapeutic applications.

Force stretching λ-DNA from 5 to 50 pN induces reversible off-target binding at multiple sites. Confocal time-lapse movie initially shows a single dCas9 complex (green dot) bound on-target to λ-DNA held between two beads by optical tweezers at low force (5 pN). As force is increased to 50 pN, multiple off-target binding events appear. Off-targets dissociate while decreasing the force back to 5 pN, and only the on-target complex remains bound.

DNA repair and chromatin remodelling dynamics

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Each human cell contains the equivalent of two meters of DNA packed in a small, micrometre-sized nucleus in the form of chromatin. The DNA in each of those cells experiences tens of thousands of lesions per day resulting from a variety of chemical, mechanical and radiation sources. Unless this damage is repaired, mutations arise that can lead to numerous diseases such as cancer and neurodegenerative disorders (including Alzheimer’s, Huntington’s and Parkinson’s diseases) that are linked with accumulated DNA damage and defective DNA repair. Elegant systems have evolved to act on chromatin to facilitate the repair process. DNA damage repair requires nucleosome remodelling to allow access to the DNA repair machinery. A group of ATP-dependent enzymes that modify chromatin structure are involved in these processes. These complex multi-subunit machines carry out multiple tasks on nucleosomes including chemical modifications, histone exchange and sliding them on DNA in what appears to be a highly coordinated process. SWR1 complex (ySWR1 in yeast, and hSRCAP in humans) is a 1.1 MDa multi-subunit complex that utilizes ATP to replace canonical H2A histones with the Htz1 variant (H2A.Z in mammalian cells). This chromatin remodelling activity is associated with regulation of gene expression in heterochromatin regions of plant and mammal chromosomes and with the cellular response to DNA damage. Despite a large number of genetic, biochemical and structural studies on ySWR1, its detailed exchange mechanism is still unknown. In collaboration with Prof Dale Wigley (Imperial College London), we are investigating the molecular mechanism of ATP-dependent replacement of the canonical two H2A histones with the Htz1 variant by the ySWR1 complex. To this aim, we are developing various single-molecule FRET assays to characterize the multi-step DNA unwrapping process required for remodelling.

Nucleosome unwrapping model by SWR1

Dynamics of SMC complexes

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Throughout the life of a eukaryotic cell, chromosomes undergo drastic conformational rearrangements that play essential roles in almost all nuclear processes, including gene expression, DNA repair and cell division. The super-structure of chromatin is regulated by ring-shaped, ATP-dependent molecular motors belonging to the SMC family of protein complexes. In eukaryotic cells, the three types of SMC complexes are cohesin, condensin and SMC5/6. While some of their biological functions have been well described, the molecular mechanism by which these complexes function remains poorly understood. In collaboration with Prof. Luis Aragon (Cell Cycle Group, MRC-LMS), we are developing single-molecule approaches to investigate these mechanisms.

Two DNA molecules held by a quad-optical trap (Lumicks) are held together by a Cohesin complex, which can slide on the DNA without letting go of the DNA strands.

Single-molecule dynamics in live cells

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Imaging individual RNA molecules in live cells is key to understanding fundamental cellular processes such as transcription, translation, splicing, transport and decay. To this aim, we are developing bright and photostable Mango RNA aptamers in collaboration with Prof Peter Unrau (Simon Fraser University). More specifically, we have developed stably folding Mango aptamer arrays and demonstrated their ability to image both coding and non-coding RNAs at single-molecule resolution without affecting their known localisation patterns. Thanks to the rapid exchange of photobleached dyes, Mango II arrays enable extended imaging times, which in turn benefits super-resolution techniques such as Structured Illumination Microscopy (SIM). In addition, our Mango II arrays are readily compatible with immunostaining, RNA-FISH, and orthogonal labelling by MS2-arrays. Our Mango arrays enable accurate determination of RNA transcription, nuclear export and subcellular localisation. We are currently exploiting this new technology to investigate several aspects of RNA metabolism, gene expression and chromatin structure in mammalian cells.

Imaging single RNA molecules in live mammalian cells with fluorogenic RNA-Mango aptamers.

Guest Lectures

Harden Conference, Harden Conference, Helicases, Cambridge, UK, Helicases, Cambridge, UK, 2013

Spanish Biophysical Society Meeting, Spanish Biophysical Society Meeting, Valencia, ESP, 2013

Annual Meeting of the RNA Society (session chair),, Annual Meeting of the RNA Society (session chair), Davos, SUI, 2013

Pittcon Conference, Pittcon Conference, Philadelphia, PA, 2013

Physics of the Genome, Physics of the Genome,, Amsterdam, NL, 2013

Gordon Research Conference, Single Molecule, Gordon Research Conference, Single Molecule, Mount Snow, VT (July 2012), 2012

Telluride Conference, Telluride Conference, Telluride, Colorado, 2012

FASEB Nucleic Acid Enzymes, FASEB Nucleic Acid Enzymes, Snowmass, CO, 2012

Gregorio Weber Symposium, University of Buenos Aires, Buenos Aires, Argentina, 2011

Riboclub, Sherbrooke University, Quebec, Canada, 2011

A. Paul Schaap Symposium, Wayne State University, Detroit, MI, 2011

Telluride Conference, Telluride Conference, Telluride, Colorado, 2011

2nd Workshop on Frontiers of Biophysics, Seoul National University, Korea, 2011

Future of Biophysics Burroughs Welcome Fund Symposium, Future of Biophysics Burroughs Welcome Fund Symposium, Baltimore, MD, 2011

Zing Nucleic Acids Conference, Zing Nucleic Acids Conference, Cancun, Mexico, 2010

EURASNET Interdisciplinary Focus Meeting, EURASNET Interdisciplinary Focus Meeting, Poznan, Poland (August 2010), 2010

European Biophysical Society Meeting, European Biophysical Society Meeting, Genoa, Italy, 2009

Zing Nucleic Acids Conference, Zing Nucleic Acids Conference, Cancun, Mexico, 2009

Annual meeting of the Anachem, Annual meeting of the Anachem, Detroit, MI, 2008

Emerging Nanoscience Applications in Technology and Biomedicine, WSU, Detroit, MI, 2007

Annual Meeting of the American Academy of Nanomedicine, Annual Meeting of the American Academy of Nanomedicine, San Diego, CA, 2007

Annual meeting of the Anachem, Annual meeting of the Anachem, Detroit, MI, 2006