Professor Sheena Radford

Professor Sheena Radford

Profile

I joined the University of Leeds in 1995 as a Lecturer in the School of Biochemistry and Molecular Biology, progressing to Reader in 1998 and Professor in 2000. In 2009 I became the Deputy Director of the Astbury Centre for Structural Molecular Biology, and its Director in 2012. I also became Astbury Professor of Biophysics in 2014.

Before coming to Leeds, I graduated with a BSc in Biochemistry at the University of Birmingham, completed my PhD in Biochemistry at the University of Cambridge with Professor R.N. Perham, FRS, and held various postdoctoral posts and a Royal Society University Research Fellowship at the Oxford Centre for Molecular Sciences.

I currently supervise about 30 PhD students and postdoctoral researchers in my laboratory. In total, more than 60 PhD students have been, or are being, supervised: about half of them are employed in academic research, 10% have other academic posts, about a third have industrial posts, and many of the rest have positions such as technical editing and school teaching.  More than 50 post-doctoral research assistants have been or are being supervised; of those who have moved on, about 15% have academic posts, and the majority of others are post-doctoral research in the UK and overseas.

I have published more than 280 peer-reviewed papers and spoken at over 360 invited lectures at national and international conferences, including more than 200 invited lectures at international conferences in countries including the UK, Germany, Denmark, USA, Australia, Japan, Sweden, Ireland, Belgium, Switzerland, Greece, Spain, Italy, France, The Netherlands, Portugal, Croatia, Israel, Austria, Canada, and Thailand.

In the last five years I have served, or am serving on, 6 major research funding panels, 12 Scientific Advisory Boards at prestigious institutions and 8 editorial boards for leading journals.

My prizes and awards include the 1996 Biochemical Society Colworth Medal, the 2005 Royal Society of Chemistry Astra-Zeneca prize in Proteins and Peptides, the 2009 Hites Award from the American Society for Mass Spectrometry (joint with Professor Alison Ashcroft), and the Protein Society Carl Branden award in 2013, and the 2015 Rita and John Cornforth Award of the Royal Society of Chemistry (also jointly with Professor Alison Ashcroft). I became a Fellow of the Academy of Medical Sciences in 2010, a Fellow of the Royal Society and an honorary member of the British Biophysical Society in 2014, and a Fellow of the Biophysical Society in 2018.

My research is focused on fundamental structural molecular biology, specifically in the measurement of the conformational dynamics of proteins and the elucidation of the role that these motions play in protein folding and misfolding of both water-soluble and membrane proteins. Using a wide range of biophysical methods and combining these with protein chemistry and molecular biology, my research focus over the last 30 years has been the delineation of the mechanisms by which proteins fold or misfold; how dynamic excursions enable proteins to self-associate into amyloid fibrils - the complex macromolecular assemblies associated with some of the deadliest human diseases; and how proteins fold into the bacterial outer membranes of Gram-negative organisms.

My major research achievements to date have included the use of native mass spectrometry, NMR and single molecule methods to characterise the intermediates in protein folding and in amyloid formation; to identify and delineate the mechanisms of action of small molecules able to interrupt protein aggregation, and the discovery of how and why different protein-protein interactions propagate amyloid formation whilst others inhibit assembly. In parallel, we have been using our biophysical toolkit of methods to understand how Gram-negative outer membrane proteins (OMPs) fold, how folding is supported by ATP-independent chaperones in the periplasm and how the β-barrel assembly machinery (BAM) catalyses OMP folding and assembly into the bacterial cell envelope.

Responsibilities

  • Astbury Professor of Biophysics
  • Director of the Astbury Centre
  • Group Leader of the Radford Laboratory

Research interests

The molecular details of how proteins fold from the linear amino acid sequence to their unique three-dimensional structures is one of the major challenges in biochemistry today. Although it has been known for more than forty years that proteins can fold to their native structures spontaneously in vitro, how this is achieved is still not understood in any great detail. The problem has become even more fascinating recently because of the discovery that folding in the cell is assisted by chaperone proteins and that misfolding events in vivo are responsible for several diseases. These represent major challenges to the pharmaceutical industry. These issues are the major focus of my research and are being tackled using a broad range of techniques including protein chemistry, structural molecular biology and sophisticated biophysical methods.

Current major projects include:

  • Mechanism(s) of protein misfolding and assembly into amyloid
  • Membrane protein folding mechanisms and role of chaperones and the BAM complex
  • Stabilising proteins of therapeutic and industrial interest against aggregation
  • Method development (MS, NMR, single molecule methods)

Watch a lecture by Professor Radford discussing her work and celebrating her FRS award. Go to https://www.youtube.com/watch?v=r1eK3DLCMcM

Detailed research programme:
1. Mechanism(s) of protein mis-folding and assembly into amyloid

A major project in the group focuses on using our knowledge of protein folding mechanisms to develop new understandings of how proteins misfold and cause disease. Specifically, we are exploring the mechanism of onset of several human amyloid diseases, including Alzheimer’s, Parkinson’s disease, type II diabetes and dialysis-related amyloidosis, caused by Aβ, α-synuclein, amylin and β2-microglobulin, respectively. Our approach combines structural analysis of the species formed during aggregation obtained using fluorescence, single molecule methods (FRET and dynamic force microscopy), mass spectrometry and NMR, with detailed analysis of the kinetics of aggregation. In collaboration with Drs Eric Hewitt and Patricija van Oosten Hawle (Astbury Centre for Structural Molecular Biology and Faculty of Biological Sciences), analysis of the effects of the different species identified on cellular function and in C elegans are being investigated. With Professor Neil Ranson of the Astbury Centre for Structural Molecular Biology and Faculty of Biological Sciences, we are using cryo-electron microscopy to investigate the structure of amyloid fibrils and other aggregated species. Overall, our aim is to derive a detailed molecular understanding of the aggregation process from monomer to amyloid and to use the power of combinatorial chemistry combined with cell biological assays and structural analysis to find new therapies for amyloid diseases.

Highlights over recent years have included using NMR and other biophysical methods to map the energy landscape for the formation of amyloid fibrils of β2m (Fig. 1) of different morphological types, and analysis of the structure of amyloid fibrils using cryo-electron microscopy (with Professor Neil Ranson, Astbury Centre for Structural Molecular Biology and Faculty of Biological Sciences), and solid state NMR (with Robert Griffin (Massachusetts Institute of Technology, USA)). In addition, we have used solution NMR methods to determine the structure of the amyloidogenic precursor of β2m and have shown that this species is not only highly amyloidogenic in itself, but it is also able to convert the non-amyloidogenic wild-type protein into a conformation able to self-assemble into amyloid at neutral pH. Reported in Molecular Cell in 2011 and 2014, this work revealed that conformational conversion is not restricted to prions but, instead, many proteins may possess the ability to convert a benign conformer to an amyloidogenic form by bimolecular collision. We are now continuing this work, extending the ideas found to other protein systems and using NMR to obtain more direct structural insights into the mechanism by which conformational conversion occurs, as well as the conformational properties of higher order oligomeric states on-pathway to fibril formation.

In parallel with this work, in a long-standing collaboration with Professor Alison Ashcroft (Astbury Centre for Structural Molecular Biology and Faculty of Biological Sciences), and now with Professor Frank Sobott (Astbury Centre for Structural Molecular Biology and Faculty of Biological Sciences), we are developing ion mobility mass spectrometry (IMS) coupled with mass spectrometry (MS) to identify and individually characterise the structural properties, population and stability of different oligomeric species of aggregation-prone sequences that are co-populated in the early stages of amyloid assembly. In addition, we are developing this approach to search for small molecules able to inhibit amyloid assembly and to determine their mechanism of action in detail. (Fig. 2). Published in Nature Chemistry in 2015 and Nature Chemical Biology in 2016 in collaboration with Professor Andrew Wilson (School of Chemistry, University of Leeds, and Astbury Centre for Structural Molecular Biology) and Dr Richard Foster (School of Chemistry, Astbury Centre for Structural Molecular Biology), we have identified novel small molecules and fragments via screening experiments, and are currently further optimising these. This work is being funded by an ERC Advanced Award, Wellcome Trust Investigator Award and the EPSRC.

Sheena Radford

Fig. 1: Mechanisms of amyloid assembly. The schematic represents some of the possible routes of amyloid formation through primary (black arrows) or secondary pathways (green arrows). Taken from Mechanisms of amyloid formation revealed by solution NMR, Karamanos, T., Kalverda, A.P., Thompson, G.S. and Radford S.E. (2015) Prog. NMR Spectros., 88 - 89, 86 -104

Sheena Radford

Fig. 2: Schematic of the ESI-IMS-MS experimental procedure for small molecule screening to an amyloid precursor protein. The protein of interest is mixed individually with small molecules from a compound library in 96-well plate format. Via a Triversa NanoMate automated nano-ESI interface, the samples are infused into the mass spectrometer, wherein separation occurs based on the mass to charge ratio (m/z) and collisional cross-sectional area (CCS). Taken from ESI-IMS-MS: a method for rapid analysis of protein aggregation and its inhibition by small molecules, Young, L.M., Saunders, J.C., Mahood, R.A., Revill, C.H., Foster, R.J., Ashcroft, A.E. & Radford, S.E. (2016) Methods, 95, 62-69

2. Membrane protein folding mechanisms and role of chaperones and the BAM complex

Although we have learned much about protein folding mechanisms in recent years, principally through the development of new methods and studies of simple and experimentally tractable systems, our understanding of how proteins fold rapidly and efficiently to their unique native conformation both in vitro and in vivo remains an exciting challenge. In order to develop new and more detailed models of protein folding, we have studied the folding of the small helical bacterial immunity proteins (principally Im7 and Im9) for the last decade.  By combining stopped flow methods, ultra-rapid mixing experiments, single molecule fluorescence (FRET) and NMR analysis we have shown that Im7 folds through an intermediate that is on-pathway to the native state and has a distorted three helical structure stabilised in a large part by non-native inter-helical interactions (Nature SMB, 2009).

Our most recent research on protein folding has moved to the challenging field of membrane protein folding, focusing on bacterial outer membrane (OMPs). In collaboration with Dr David Brockwell, Dr Roman Tuma and Professor Neil Ranson (all of the Astbury Centre for Structural Molecular Biology and Faculty of Biological Sciences), we are investigating how OMPs are able to cross the inner membrane via the SecYEG translocon, traverse the periplasm (aided by chaperones) and assemble into bacterial outer membrane. Our first inroads into this field (published and highlighted in PNAS, 2010) exploited phi-value analysis to reveal a structural model of the transition state for folding of the E.coli outer membrane protein, PagP. The results revealed a complex, tilted insertion mechanism, previously predicted for membrane insertion of this class of proteins. Current work is building on these insights by exploring the role of the molecular chaperones Skp and SurA and the BAM complex in assisting the folding of OMPs (Figure 3).

Most recently, we have elucidated the first solution structure of the BAM complex using cryo-EM, shown that Skp is a multi-valent chaperone, and have started to reveal how the entire pathway of OMP folding is choreographed from passage across the inner membrane via SecYEG, chaperoned across the periplasm by Skp and SurA and finally folded into the OM by BAM. This work is funded by the Wellcome Trust, BBSRC and MRC, published in Nature Structural and Molecular Biology (2016 and 2017) and Nature Communications (2016) and Elife (2016, 2018).

Sheena Radford

Fig. 3: OMP assembly pathway across the periplasm. Taken from Outer membrane protein folding from an energy landscape perspective, Schiffrin, B., Brockwell, D.J. & Radford, S.E. (2017) BMC Biology, 15, 123

3. Stabilising proteins of therapeutic and industrial interest against aggregation

In the third area of our research interests, we are exploiting our fundamental knowledge of folding for practical benefits. Combining our skills with the microbiological expertise of Professor Jim Bardwell (Michigan) we developed a β-lactamase host-guest system to select for protein sequences with enhanced stability. The results (published in Molecular Cell in 2009) showed that the vast majority of mutations that enhance stability for Im7 occur in residues that are required for function. The results support the view that protein sequences are highly frustrated (i.e. function compromises stability and folding capability). They also demonstrate the utility of the β-lactamase system developed to generate proteins that retain function, but are optimised for stability. Further developing this approach has enabled us to use the split β-lactamase system to screen for protein sequences that are less aggregation-prone and to screen for small molecules able to protect proteins from aggregation (Nature Chem. Biol (2016)) (Figure 4).

Our current work is focused on developing the split β-lactamase system, and other approaches (including fragment-based and other design strategies), to screen for sequence hot spots that cause aggregation of proteins, particularly those of interest and relevance to the biopharmaceutical industry, as well as to screen for small molecules able to arrest protein aggregation. The aim is to use evolutionary pressure to reveal new insights into protein stability, dynamics, allostery and binding, and to harness these fundamental insights into practical benefits both to academia and to the bio-pharmaceutical sector.

Finally, in collaboration with Dr David Brockwell (Astbury Centre for Structural Molecular Biology and Faculty of Biological Sciences) and Dr Nikil Kapur (School of Mechanical Engineering, University of Leeds), we are examining how flow fields enhance, or cause, protein aggregation by flow-induced protein deformations (PNAS, 2017).

This work is funded by the BBSRC and EPSRC, and involves strong collaborations with Medimmune Limited and UCB.

Sheena Radford

Fig. 4: Split β-lactamase assay for protein aggregation identifies aggregation-prone sequences. Left: Inactive enzyme: if the test protein aggregates, the activity of β-lactamase is reduced and the bacteria become more sensitive to the β-lactam antibodies. Right: Small-molecule inhibitors (yellow) of protein aggregation diffuse into the periplasm via porins (violet trimers) and prevent aggregation of the β-lactamase tripartite fusion protein, restoring bacterial resistance to β-lactam antibiotics. Adapted from An in vivo platform for identifying inhibitors of protein aggregation, Saunders, J.C., Young, L.M., Mahood, R.A., Revill, C.H., Foster, R.J., Jackson, M.P., Smith, D.A.M., Ashcroft, A.E., Brockwell, D.J. & Radford, S.E. (2016) Nature Chem. Biol., 12, 94-101

4. Method development (MS, NMR, single molecule methods)

Major developments in instrumentation have played a key role in increasing in our understanding of folding and aggregation mechanisms to date. Future developments in these fields will require further innovative approaches that cross the boundaries between disciplines. We have been involved in many exciting collaborations to fulfil this aim. Together with Dr Roman Tuma (Astbury Centre for Structural Molecular Biology and Faculty of Biological Sciences), we have built and developed instruments capable of single molecule measurements using both FRET and FCS (Elife 2016 and 2018). Developing MS methods continues to be an aim of our laboratory (in collaboration with Professors Alison Ashcroft and Frank Sobott (both of the Astbury Centre for Structural Molecular Biology and Faculty of Biological Sciences)). In addition, developments in NMR methods remain a mainstay of our laboratory activities, whilst, in collaboration with Dr David Brockwell (Astbury Centre for Structural Molecular Biology and Faculty of Biological Sciences), we are involved in some exciting developments in the use of the AFM for force measurements of protein unfolding and protein binding. Finally, with Professor Andrew Wilson (School of Chemistry, University of Leeds, and Astbury Centre for Structural Molecular Biology) and Professor Nikil Kapur (School of Mechanical Engineering, University of Leeds) we are developing new cross-linking methodologies to track protein-protein interactions in vitro and in vivo. More information about these projects can be found on the websites of our collaborators on their Astbury web pages.

For further details about the Radford laboratory, people involved, molecular images and available opportunities please see:  

<h4>Research projects</h4> <p>Any research projects I'm currently working on will be listed below. Our list of all <a href="https://biologicalsciences.leeds.ac.uk/dir/research-projects">research projects</a> allows you to view and search the full list of projects in the faculty.</p>

Qualifications

  • BSc, Birmingham, 1984
  • PhD, Cambridge, 1987

Professional memberships

  • Fellow of the Royal Society, 1 May 2014
  • Fellow of Academy of Medical Sciences, 2010
  • Fellow of the Biophysical Society for leadership in protein biophysics, 2018
  • Fellow of the European Molecular Biology Organization (EMBO), 2007
  • Honorary Member of the British Biophysical Society, May 2014
  • Faculty of 1000, 2014
  • Fellow of the Royal Society of Chemistry, 2003
  • Fellow of the Biophysical Society for leadership in protein biophysics 2018
  • ASBMB and American Chemical Society
  • Biochemical Society and Protein Society

Student education

I currently teach Biochemistry at both undergraduate and postgraduate levels. Specifically, I teach basic mechanisms and concepts in protein folding, and contribute to Advanced Topic Units to final year students. Our laboratory also hosts students for their final year research projects (Biochemistry and Biological Sciences) for both BSc and MBiol schemes. Finally, I teach on a module in Protein Folding and Assembly in the Astbury Centre Wellcome Trust funded 4-year PhD scheme, the Molecular Basis of Biological Mechanisms.

Current postgraduate research students

<h4>Postgraduate research opportunities</h4> <p>We welcome enquiries from motivated and qualified applicants from all around the world who are interested in PhD study. Our <a href="https://biologicalsciences.leeds.ac.uk/research-opportunities">research opportunities</a> allow you to search for projects and scholarships.</p>