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What Does It Take to Analyze Membrane Proteins? Ask Whitelegge

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At A Glance

Name: Julian Whitelegge

Age: 39

Position: Adjunct Associate Professor, The Pasarow Mass Spectrometry Laboratory, Departments of Psychiatry and Biobehavioral Sciences, Chemistry & Biochemistry and the Neuropsychiatric Institute, University of California, Los Angeles

Prior Experience: Postdoc in John Bowyer's lab at the University of London, visiting researcher in Fred McLafferty's lab at Cornell University

How did you get involved in proteomics?

In ‘86 there was a paper from Don Hunt’s lab, where they defined the N-termini of three different proteins that were key players in the plant membrane system I was particularly interested in. They used tandem mass spectrometry to define these structures. It was clear to me that mass spectrometry was going to be central to studying protein structure and its physiological adaptive responses.

I wasn’t in the position to be doing mass spec back then, [instead] I was using large amounts of 32P to do in vivo labeling, [but] knowing that there had to be a better way to proceed. The first postdoc that I did [at the University of London with John Bowyer] was in protein chemistry. We were using Edman sequencing to map around a herbicide binding site using photoaffinity labels. It was clear that we wanted to be mass spec compatible, so I developed some HPLC techniques that were compatible with very hydrophobic peptides derived from integral membrane proteins. One of these was a size exclusion method, and the other was a reverse phase method. These were eventually published in the European Journal of Biochemistry in 1991. In the meantime I did some molecular biology – basically some site-directed mutagenesis and some chloroplast transformation experiments. But around about ‘95-’96 it was clear to me that there were far too many people doing molecular biology, and [since] I already had protein chemistry experience, I wandered into the mass spec facility at UCLA and hooked up with Kym Faull. He’s the director of what’s now called the Pasarow Mass Spectrometry Laboratory.

It very soon became apparent that I could use the two HPLC techniques that I developed way back in the late ‘80s [as an] interface with electrospray ionization mass spectrometry to record peptide and protein mass spectra of integral membrane proteins and the peptides derived from them. There were really very few other people doing this. I went into mass spectrometry to be able to look at membrane proteins and their modifications such as phosphorylation and protein oxidation. We had to work quite a long time perfecting these HPLC techniques. We’ve been perfecting these techniques, and in the meantime we’ve gotten up to proteins with 12 transmembrane helices. That was the lactose permease from E. coli which we published in PNAS in 1999. What I’ve been doing since then is going into proteomics.

How are you applying protoemics?

We’ve proceeded in a couple different fashions. One way is to start looking at protein complexes. The nice thing about a protein complex is that it defines a number of different gene products that come together to perform a specific function. So you’ve got protein-protein interaction information that ties in with a number of different gene products. Very often you can assign function to a gene product based upon this type of association. Recently I’ve been looking at cytochrome b6f complex in collaboration with Bill Cramer of Purdue University. [That work is described in] a paper currently being considered [for publication in] Molecular and Cellular Proteomics. Basically what we have are two techniques – size exclusion chromatography and reverse phase chromatography – that allow us to achieve full subunit coverage of integral membrane proteins. That’s really a significant achievement because many chromatographic techniques do not fully represent all the integral membrane proteins. The larger ones tend to be rather sticky; they don’t necessarily elute from columns. In the case of 2D gel electrophoresis, the larger ones really don’t get past the first step – the isoelectric focusing step.

What allows your particular brand of chromatography to achieve this kind of coverage?

It’s a combination of both the mobile phases that are used and the stationary phases. I’ve found for the size exclusion chromatography the only suitable matrix is the SW series from Tosoh Biosep. [The matrix consists of] deactivated silica size exclusion beads, but the Tosoh Biosep deactivation process is different from everybody else’s, such that very few proteins, and certainly not the membrane proteins, stick to this matrix. [But] to achieve that type of chromatography, I actually go a little bit outside the specifications of the matrix provided by the manufacturer. I use chloroform methanol and one percent formic acid in water in the proportions 4:4:1. The manufacturer doesn’t recommend more than 50 percent organic solvent, but I’ve found that as long as I keep a reasonable proportion of water in the solvent – we’ve got over ten percent water – there really is no problem with these columns at all. In fact we’ve found that the lifetime is superb; they’re really quite inert under these conditions.

For the reverse phase, I gave up reverse phase columns [packed with silica] quite a long time ago. I actually use polystyrene divinyl benzene copolymer, or PLRPS, from Polymer Labs. The reason I prefer that particular matrix is that they seem to have the best quality control on their chromatography beads, so that when I have a new column I have the ultimate resolution with their matrix. But obviously there are many other manufacturers of polystyrene divinyl benzene copolymer beads, such as Hamilton and PerSeptive.

In the most recent paper I sent to MCP on the cytochrome b6f complex, I was able to elute all of the known major components of the complex using an isopropanol/acetonitrile solvent mix. Then if you go back and do an elution with 60 percent formic acid with isopropanol you see the proteins that are left behind, and a notable protein there is cytochrome b. It’s actually only 24 kDa; it has four transmembrane helices but it’s one of these very sticky integral membrane proteins.

What other projects in proteomics are you working on?

A more general proteomics project that I’m working on is funded by the Department of Energy’s microbial cell project. The goal is basically to do proteomics on the photosynthetic cyanobacterium Synechocystis 6803. We’re particularly interested in membrane composition changes associated with different mutants of this organism. Basically our approach to the proteome of this prokaryote, which has about 4,000 open reading frames from the genome, is to break open the cells, and immediately separate the soluble proteins from the membrane proteins. The soluble proteins we’re looking at by two different techniques: 2D electrophoresis and 2D chromatography.

The integral membrane proteome is really [my] area of expertise. What I’m doing with the integral membrane proteins is using a 2D chromatography approach. In [John Yates’] MudPIT approach they proteolyze the starting material, and then look at the peptides. The way that I’m working is to actually do 2D separations on the intact proteins, record the intact mass spectra of the proteins, and collect fractions at the same time to do downstream identification experiments. It probably doesn’t achieve the same throughput [as the MudPIT approach] but we have the added benefit of this intact molecular weight readout, and we see any heterogeneity that’s associated with the protein, [such as] posttranslational modifications.

How do you isolate particular complexes – wouldn’t you also get extraneous proteins not associated with the complex you’re interested in?

The separations that you get in size exclusion chromatography are by no means perfect. You do get overlapping elution of different complexes and that’s a disadvantage of this particular technique. But toward a general technique to look at whole membranes, we are finding that this is one way that we can proceed. It’s not perfect – I’d be the first to admit that, but it’s certainly giving us useful insights.

What kind of equipment do you have in the Pasarow Mass Spectrometry Lab at UCLA?

We’ve got a Fourier transform mass spectromter with a seven Tesla magnet combined with both the MALDI source and the electrospray source from Ion Spec. We’ve got a LCQ DECA with their Surveyor LC system, which is dedicated to proteomics-type experiments – LC/MS/MS – generating sequence tags for protein identification. We use SEQUEST software for identifications; we also use SONAR over the web, and MASCOT as well. We’ve got an Applied Biosystems Voyager DE-STR MALDI-TOF, which we use for our peptide mass fingerprinting, and we find that with an internal standard we can very often achieve 5 ppm on our MALDI, which is a great asset in protein identification. We’ve got a couple of old API III triple quads, which we still use for a lot of our LC/MS applications. I’ve got one running right now for this process that I call LC/MS+. We’re doing LC on intact proteins, running half the eluent into the mass spectrometer for intact protein measurements, and we’re collecting fractions with the rest of the eluent. This is a central theme to what I do.

At the moment we’re actually installing the Bio-Rad ProteomeWorks system, combined with a Micromass MALDI-TOF, and their MassPrep [sample preparation robot] as well. In association with that we’re currently buying a Q-TOF, but we haven’t decided between Q-TOF and QSTAR. We’ve just heard that Applied Biosystems has something coming out called the QSTAR XL, which supposedly has a hardware fix to remove singly-charged ions. So we’re going to be checking out the QSTAR XL to see if it can really get rid of the singly-charged ions but leave all the doubly-charged ions behind. Of course we’re a little bit skeptical about that, but that’s why we’re going to the factory.

 

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