Extract from IMMS: The Next Five Years - Prof. Helen J. Cooper

Published on 17 Jul 17, under FAIMS, Science & Research

Here's an extract from the 2nd edition of our free ebook Ion Mobility Mass Spectrometry: The Next 5 Years. Download the complete ebook here.

Prof. Helen J. Cooper

University of Birmingham, UK



Professor Helen J. Cooper is EPSRC Fellow and Professor of Mass Spectrometry in the School of Biosciences at the University of Birmingham. Her research focuses on the development and application of advanced mass spectrometry for the analysis of biomolecules, particularly peptides and proteins. Current areas of research include liquid extraction surface analysis of biological substrates such as dried blood spots, thin tissue sections and bacterial colonies; high field asymmetric waveform ion mobility spectrometry, and electron mediated dissociation. She obtained her BSc and PhD in Chemistry from the University of Warwick. She started her postdoctoral research at the University of Warwick before moving to the National High Magnetic Field Laboratory at Florida State University. She took a position at the University of Birmingham in 2003, and in 2004 was awarded a Wellcome Trust University Technology Fellowship. She became Professor of Mass Spectrometry in 2013 and was awarded her EPSRC Fellowship in 2014.

Q1: What are your main research activities in ion mobility - mass spectrometry (past or present)?

We became interested in ion mobility in 2008 as a result of some work we had undertaken with

phosphopeptides. We had noticed that isomeric phosphopeptides fragmented differently by electron capture dissociation and concluded that this was the result of intramolecular interactions between the phosphate group and basic amino acid side chains1. We hypothesised that, as a result of these interactions, phosphopeptides with identical sequences but differing sites of modification would have differing conformations and therefore should be separable by ion mobility. We were able to show that was the case using FAIMS2, and it has since been demonstrated using drift tube ion mobility spectrometry and travelling wave ion mobility spectrometry. We subsequently moved on to consider the use of FAIMS, and other ion mobility approaches, in the analysis of other protein post-translational modifications including nitration3 and glycosylation4.

Our current research activities have two strands. Firstly, we are interested in developing and applying FAIMS methods for proteomics. We demonstrated the complementarity of FAIMS and strong cation exchange chromatography for improved proteome coverage5. We also demonstrated the benefits of FAIMS for the large scale analysis of peptide sequence variants6. More recently, we applied FAIMS for the comprehensive mapping of O-glycosylation in flagellin from Campylobacter jejuni7 and the analysis of phosphorylation in fibroblast growth factor signalling8.

The second strand focuses on the coupling of FAIMS with liquid extraction surface analysis (LESA) mass spectrometry. We have shown that high resolution LESA mass spectrometry may be coupled with FAIMS for the analysis of intact proteins from a range of biological substrates, including thin tissue sections, dried blood spots and bacterial colonies growing on agar9,10. The use of FAIMS results in improved S/N and separation of different molecular classes. The improvements in S/N due to FAIMS result in shorter analysis times, thereby allowing LESA FAIMS mass spectrometry imaging of intact proteins. We have also developed software for visualisation of FAIMS data. Figure 1 shows the total ion transmission map obtained following a 1-dimensional FAIMS analysis (DF = 270 Td, CF = -1 – 4 Td) after LESA sampling of a mouse liver section.

1D FAIMS analysis after LESA sampling of mouse liver
Figure 1: Total ion transmission map obtained following 1D FAIMS analysis after LESA sampling of a mouse liver section. DF = 270 Td, CF = -1 – 4 Td. Mean mass spectrum is projected in red and total ion chromatogram is projected in blue. Data collected on an Orbitrap Elite (Thermo Fisher) mass spectrometer coupled with a Triversa Nanomate (Advion) and ultraFAIMS (Owlstone).

Q2: What have been the most significant instrumentation or applications developments in ion mobility - mass spectrometry?

A truly significant development was the application of FAIMS to proteomics by Pierre Thibault’s group11. More recent work by the same group12, and by Robert Moritz and co-workers13, further served to confirm the potential of FAIMS in the proteomics field. We are now in a position where FAIMS-based proteomics has moved beyond proof-of-principle and is being applied to real-world biological problems.

For our work on integrating FAIMS with LESA mass spectrometry, the most significant advance was the development of the miniaturised FAIMS device by Owlstone. We are now able to couple the three key pieces of hardware – the sampling robot, FAIMS and the mass spectrometer – and are able to undertake imaging experiments.

Q3: Where do you see ion mobility - mass spectrometry making the most impact in the next 5 years? Any predictions for where the field will go?

The use of FAIMS in proteomics is growing and I believe this will continue providing that user-friendly integration of nano liquid chromatography and FAIMS is made readily available. Improvements in this aspect are ongoing in the field.

Since the last edition of this ebook, our group have coupled FAIMS with LESA mass spectrometry for intact protein analysis from a range of biological substrates. This development has potential applications in a range of areas, including mass spectrometry imaging, and we are very excited to pursue this research!

To find out how other experts in the field of ion mobility - mass spectrometry are using the technique, download our free ebook Ion Mobility - Mass Spectrometry: the Next Five Years. 

IMMS ebook CTA


  1. Creese, A.J. and H.J. Cooper, J. Am. Soc. Mass Spectrom., 2008. 19: p. 1263-1274.
  2. Xuan, Y., et al., Rapid Commun. Mass Spectrom., 2009. 23: p. 1963-1969.
  3. Shvartsburg, A.A., et al., Anal. Chem., 2011. 83: p. 6918-6923.
  4. Creese, A.J. and H.J. Cooper, Anal. Chem., 2012. 84 : p. 2597-2601.
  5. Creese, A.J., et al., J. Am. Soc. Mass Spectrom., 2013. 24: p. 431-443.
  6. Creese, A.J., J. Smart, and H.J. Cooper, Anal. Chem., 2013. 85: p. 4836-4843.
  7. Ulasi, G.N., et al., Proteomics, 2015. 15(16): p. 2733-2745.
  8.  Zhao, H., et al., J. Proteome Res., 2015. 14(12): p. 5077-5087.
  9. Griffiths, R.L., et al., Analyst, 2015. 140: p. 6879-6885.
  10. Sarsby, J., et al., Anal. Chem., 2015. 87(13): p. 6794-6800.
  11. Saba, J., et al., J. Proteome Res., 2009. 8: p. 3355-3366.
  12. Bridon, G., et al., J. Proteome Res., 2012. 11(2): p. 927-940.
  13. Swearingen, K.E., et al., Mol. Cell Proteomics, 2011. 11(4): p. DOI: 10.1074/mcp.M111.014985

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