While browsing recently published papers in other journals, I came across Supramolecular fishing for plasma membrane proteins using an ultrastable synthetic host-guest binding pair (published in Nature Chemistry). The idea here is that proteins on the surface of the cell are chemically modified by reaction with 1-trimethylammoniomethylferrocene (AFc). After cell lysis, the modified proteins bind to beads coated with cucurbit-7-uril. Cucurbit-7-uril and AFc interact to form an ultrastable host-guest binding pair. The captured proteins are then recovered either by treatment with a strong competitor or by heating to 95 C in buffer conditions.
I thought that this was rather pleasing! It reminded me of a protocol we had published on the metabolic labeling of glycans with azido sugars, and got me to thinking: “What sorts of protocols do we have for looking at membrane proteins?”.
Using our Browse functionality plus a PubMed search I came up with 17 that I thought would be relevant (please forgive me (and let me know!!) if I have missed something vitally important). These protocols can be roughly divided into those working with a specific protein that you already know (the majority), and those where you are working with membranes from whole cells. When looking at the structure and function of purified membrane proteins, one of the main challenges seems to be the design of a matrix such that you can get meaningful data from proteins that are in conditions as similar as possible to the cell membrane itself.
These are some notes that I made while going through the protocols:
Expression and purification
– This protocol includes a method for transforming a gene-vector construct into S. cerevisiae.
– Purification is via a His-tag
Looking mostly at structure
– His-tagged proteins are expressed in E.coli. The membranes are isolated and solubilised. Purification on a nickel column is followed by removal of the his-tag and size exclusion chromatography. Crystallisation is by hanging-drop vapour diffusion.
– Isotope labelled proteins are analysed in bicelles. The lipid bilayers of these are aligned perpendicular to the magnetic field forming a liquid crystalline phase.
– The protein is synthesised via FMOC chemistry. When analysed by electron paramagnetic resonance, the TOAC residue accurately reports the position, orientation and dynamics of the peptide backbone near the labelled site.
– The authors use purple membrane from Halobacterium salinarum as an example. The stylus of the AFM is used to obtain the topography of the membrane as well as to mechanically manipulate the protein. Measurements taken in the process of unfolding the protein provide information regarding the molecular interactions within it.
– This protocol uses GFP-fusion proteins. The link will take you to a cartoon showing a single cell before and after exposure to digitonin and trypsin.
– Each residue in the protein is systematically mutated to cysteine. The proteins are reacted with [N-ethyl-1-^14^C]ethyl-maleimide, separated by SDS–PAGE and analysed using a PhosphoImager.
Looking mostly at function using specific proteins
Gel chromatography and analytical ultracentrifugation to determine the extent of detergent binding and aggregation, and Stokes radius of membrane proteins using sarcoplasmic reticulum Ca2+-ATPase as an example
– This method uses the radioactive detergent ^14^C-n-dodecyl-β-D-maltoside. The Stokes radius is determined either by size-exclusion chromatography or by ultracentrifugation sedimentation velocity analysis.
– This protocol is at the border of a few sections as it uses a functional assay (uptake of a radiotracer appropriate to the type of membrane protein that you are interested in) to either find new proteins or work what mutations will result in changes in transport activity.
– cRNA is injected into the oocytes.
Two protocols that I wasn’t certain how to classify
– The membrane proteins are isolated by centrifugation (usually from a transfected cell line containing a target protein) and attached to a stationary phase.
– Frontal affinity chromatography using labelled ligands is used to measure molecular interactions.
– This protocol is for the MYTH system shown in the figure below.
– The method used is called LOPIT (localisation of organelle proteins by isotope tagging).
– Cell membranes are separated into fractions by equilibrium density gradient centrifugation. Each fraction is labelled using a different iTRAQ reagent, and the samples are pooled.
– The key features of this protocol are (1) the use of 60 % methanol to dissolve the membrane-protein sample, and (2) that the strong cation exchange chromatography is done off-line to allow for optimisation of the fractionation to improve the capacity of the LC-MS/MS analysis. This is significant, because it allows the identification of 1500-2500 unique proteins.