Professor Andrew Hudson
Professor of Biophysical Chemistry
I am a biophysical chemist with a track record for interdisciplinary collaborative research. I graduated with a Bachelor’s degree in Chemistry from the University of Oxford and a PhD in Chemical Physics from the University of Toronto where I worked with Professor John Polanyi FRS. Between 2000 and 2005 I worked for a number of companies in the optical technologies industry EFOS Inc. (Mississauga Canada) EXFO Inc. (Mississauga Canada) and Novx Systems (Richmond Hill Canada). In 2005 I returned to academia and started an independent research group moving to the University of Leicester in 2008. I specialise in applying optical methods including optical tweezing molecular spectroscopy and microscopy (and combinations of these) to address problems at the life science interface.
My research has been funded by EPSRC the Wellcome Trust the Royal Society BBSRC and the Leverhulme Trust. I have been using methods in single molecule fluorescence microscopy to study the conformational dynamics of forked-DNA and the reaction mechanisms of RNA splicing complexes. I have also applied Raman microspectroscopy to monitor haem proteins in live cardiomyocytes and determine mechanisms for how small molecules (NO, CO) confer cardioprotection via their interactions with haem proteins. I have expertise with developing hybrid techniques: for example the integration of holographic optical tweezing and spectroscopy and microfluidics and spectroscopy; both of which are currently being used for single cell studies. I have also recently designed a genetically-encoded sensor to measure exchangeable haem in live cells using fluorescence lifetime imaging.
Biospectroscopy and imaging
My research group’s aim is to bring frontier technologies in spectroscopy, imaging, photonics and microfluidics to bear on problems at the life-science interface.
A particular area of interest is the application of specialised techniques in fluorescence imaging to monitor the dynamics of single molecules. Differences in chemical reactivity might arise from small variations in the conformational structure of proteins and nucleic acids, however, the dynamics of individual molecules are not revealed in traditional experiments. Instead, the results reflect the ensemble average across the entire population of molecules. It is the objective of single-molecule research to reveal sub-population heterogeneity. With Professor Ian Eperon, we are applying single molecule methods to address otherwise inaccessible problems in RNA splicing. These have included validation of the spice-site selection model and the pathways for the early stages of spliceosome assembly and 3′ splice-site selection. We have also invented a technique to encapsulate single biological molecules in aqueous microdroplets within a water-in-oil emulsion. The approach enables single molecule fluorescence measurements to be made on freely-diffusing molecules, and eliminates the normal requirement to tether molecules to a solid support (the technology has been patented by the University).
On a different theme within the research group, we apply a number of imaging modalities to quantifying the distribution of hemoproteins in living cells, and how this distribution evolves in response to different stimulants. We are able to distinguish between the identity of the proteins, and the oxidation and coordination state of the metal. Recently, using Raman imaging, we have been able to provide mechanistic evidence for how small molecules (NO, CO) might confer protection on cardiomyocytes (heart cells) via their interactions with hemoproteins. We are also working with Professor Emma Raven to develop optical sensor technology for in vivo quantification of labile heme.
A number of years ago, whilst working on lab-based atmospheric measurements, the group invented a technique to monitor the binary coalescence of liquid-aerosol microdroplets using holographic optical tweezing combined with elastic and inelastic scattering measurements. The method was able to report on the binary coalescence time, and surface tension & viscosity of the composite microdroplet. We now employ the same technique to reveal chemical and physical changes in cells and synthetic lipid vesicles. With Professor Russell Wallis, we are looking at the real-time process of pore formation by the toxin, pneumolysin, in liposomal membranes, and with Dr Natalie Garton (Department of Respiratory Sciences), we are looking at how an aerosolised bacterium responds to osmostic stress.
The set-up mixing aquous and oil phases into a reverse emulsion.
Schematic showing mixing of aquous and oil phases into a reverse emulsion.
Leung, G. C.; Fung, S. S.; Gallio, A. E.; Blore, R.; Alibhai, D.; Raven, E. L.; Hudson, A. J., Unravelling the mechanisms controlling heme supply and demand. Proc Natl Acad Sci U S A 2021, 118 (22).
Faraj, B. H. A.; Collard, L.; Cliffe, R.; Blount, L. A.; Lonnen, R.; Wallis, R.; Andrew, P. W.; Hudson, A. J., Formation of pre-pore complexes of pneumolysin is accompanied by a decrease in short-range order of lipid molecules throughout vesicle bilayers. Sci Rep 2020, 10 (1), 4585.
Leung, G. C.; Fung, S. S.; Dovey, N. R. B.; Raven, E. L.; Hudson, A. J., Precise determination of heme binding affinity in proteins. Anal Biochem 2019, 572, 45-51.
Fernandez, M. O.; Thomas, R. J.; Garton, N. J.; Hudson, A.; Haddrell, A.; Reid, J. P., Assessing the airborne survival of bacteria in populations of aerosol droplets with a novel technology. J R Soc Interface 2019, 16 (150), 20180779.
Jobbins, A. M.; Reichenbach, L. F.; Lucas, C. M.; Hudson, A. J.; Burley, G. A.; Eperon, I. C., The mechanisms of a mammalian splicing enhancer. Nucleic Acids Res 2018, 46 (5), 2145-2158.
Chen, L.; Weinmeister, R.; Kralovicova, J.; Eperon, L. P.; Vorechovsky, I.; Hudson, A. J.; Eperon, I. C., Stoichiometries of U2AF35, U2AF65 and U2 snRNP reveal new early spliceosome assembly pathways. Nucleic Acids Res 2017, 45 (4), 2051-2067.
Collard, L.; Perez-Guaita, D.; Faraj, B. H. A.; Wood, B. R.; Wallis, R.; Andrew, P. W.; Hudson, A. J., Light Scattering By Optically-Trapped Vesicles Affords Unprecedented Temporal Resolution Of Lipid-Raft Dynamics. Sci Rep 2017, 7 (1), 8589.
Wright, A. J.; Richens, J. L.; Bramble, J. P.; Cathcart, N.; Kitaev, V.; O'Shea, P.; Hudson, A. J., Surface-enhanced Raman scattering measurement from a lipid bilayer encapsulating a single decahedral nanoparticle mediated by an optical trap. Nanoscale 2016, 8 (36), 16395-16404.
Weinmeister, R.; Freeman, E.; Eperon, I. C.; Stuart, A. M.; Hudson, A. J., Single-Fluorophore Detection in Femtoliter Droplets Generated by Flow Focusing. ACS Nano 2015, 9 (10), 9718-30.
Almohammedi, A.; Kapetanaki, S. M.; Hudson, A. J.; Storey, N. M., Monitoring Changes in the Redox State of Myoglobin in Cardiomyocytes by Raman Spectroscopy Enables the Protective Effect of NO Donors to Be Evaluated. Anal Chem 2015, 87 (20), 10605-12..
- Applications of single molecule fluorescence microscopy
- Design of genetically-encoded sensors to detect small molecules in cells
- Optical tweezing and microspectroscopy of cells
- Mechanistic studies in haem biology
- Raman spectroscopic analysis of biopigments
Physical chemistry component of the BSc MChem and MSc degrees in Chemistry; including topics on thermodynamics kinetics mathematics for chemists molecular symmetry and molecular spectroscopy.
Press and media
Biophysical measurements; chemical and physical properties of protein and nucleic acids; microscopy and optics; spectroscopic identification of compounds; optical forces.