Proteomics Moves From Expression to Turnover

Update and Future Perspective

Mary K Doherty; Phillip D Whitfield


Expert Rev Proteomics. 2011;8(3):325-334. 

In This Article

Protein Turnover in Cellular Systems

It is now possible to overcome these difficulties using proteomics, which has allowed the field of research to move forward to the point where we can rapidly profile the synthesis and degradation rates of individual proteins from complex samples on a proteome-wide scale. In particular, the latest generation of mass spectrometers, with their increased sensitivity, resolution and mass accuracy, have allowed proteomics to move from a purely qualitative discipline to a robust, quantitative tool. A key technique, used in conjunction with MS is stable isotope labeling of amino acids in cell culture (SILAC).[25] This is the primary method for incorporating the tracer label into proteins. Although used predominantly for relative quantification of proteins,[26–29] the technique can be adapted to determine protein synthesis and degradation rates. In essence, the cells are grown in a medium that has been depleted of the chosen amino acid and subsequently supplemented with either the unlabeled or labeled form of that amino acid (Figure 2). As the cells divide, the amino acid is incorporated into the proteins, which can be monitored using MS. A synthesis-based approach measures the incorporation of a label into a protein, whereas a degradation-based approach measures the loss of a label from previously labeled proteins (Figure 3). The major distinction between conventional SILAC and the adaptation of the technique for investigating protein dynamics is that instead of comparing one experimental state with another, it is the rate of incorporation (or loss) of a label that is determined.

Figure 2.

Stable isotope labeling of amino acids in cell culture. (A) A conventional stable isotope labeling of amino acids in cell culture experiment. Cells are grown in medium containing either unlabeled or labeled amino acids. The biochemical intervention of choice can then be applied to individual cultures at specific time points. The cell lysate is then combined and processed. For a simple global screen, proteins are generally separated by 1DGE, digested with trypsin and analyzed by mass spectrometry. Alternatively, the samples can be enriched for either a subcellular fraction of interest or for a particular post-translational modification prior to mass spectrometric analysis. (B) A representative spectrum of a peptide that is present in all three conditions, each offset by the mass of the labeled amino acid. Using the relative intensities of the ions, the change in concentration of the peptide and hence protein over the time course of the experiment can be determined.
1DGE: 1D gel electrophoresis.

Figure 3.

The dynamic stable isotope labeling of amino acids in cell culture approach. Cells were grown in media supplemented with the labeled amino acid and subsequently 'chased' by transfer to a medium containing an unlabeled amino acid. For a synthesis-based approach, samples are collected during the incorporation phase, the proteins separated by 1D gel electrophoresis, digested and analyzed by mass spectrometry. The relative incorporation of the heavy and light amino acids into multiple proteins is determined over the time course, from which the first-order rate constant for synthesis of the protein is determined by nonlinear curve fitting. In a degradation-based approach, samples are taken during the chase period and the rate of loss of labeled amino acid calculated.

The choice of label is important and has been discussed extensively. It is important that the tryptic peptide pair can be separated in the mass spectrometer. This has become relatively facile, but it is important to consider that each peptide-derived ion has an associated natural abundance isotope profile. If the mass offset between the unlabeled and labeled peptide ions is too small, deconvolution of the ion pairs becomes complicated and impedes accurate quantification. Other considerations include: the nature of the label and co-elution of the peptide pair; the ability of the cell to synthesize and process the precursor (ideally the cell will be auxotrophic for the amino acid); abundance of the amino acid in the proteome and in the proteolytically derived peptides; lability of the label; and metabolic activity of the precursor pool. This has led to a preference for 13C- and 15N-labeled amino acids, in particular arginine and lysine, for cells in culture. It should also be ensured that the proteins are homogeneously labeled with the amino acid of choice.[30]

The first organism to be probed using this new technology was the yeast Saccharomyces cerevisiae, which was supplemented with 2H10 leucine in the growth medium.[31] The yeast used was auxotrophic for leucine, minimizing any dilution of the label by de novo synthesis. Proteins were labeled for approximately 50 h (over seven doubling times) before switching the cells to medium containing unlabeled leucine and commencing the 'chase' period for an additional 50 h. Mass spectral profiles of protein-derived tryptic peptides allowed the replacement of the labeled protein by unlabeled protein to be tracked, providing a value for the rate of loss of label for each protein. Values for each protein were derived from multiple peptides, allowing accurate first-order rate constants to be calculated with confidence. The intracellular degradation rates were obtained by applying an appropriate correction factor to account for loss through dilution into daughter cells.

A similar approach was used in 3T3-L1 adipocytes, with L-(2,3,4,5,6[2H5]) phenylalanine as the tracer.[32] It was possible to discriminate between rapidly synthesized proteins (collagen) and slowly synthesized proteins (cytoskeletal), indicating an active metabolism of the extracellular matrix. A primary flaw in the approach was the selection of the tracer, which resulted in an under-representation of the proteome. In a more recent study, cultures of HeLa cells were metabolically labeled with either unlabeled arginine, 13C614N4 arginine or 13C615N4 arginine.[26] Each cellular population was treated with actinomycin D, a transcription inhibitor, at different times, permitting the kinetic profiles of nucleolar proteins to be determined.

In a related approach to SILAC, Cargile et al. used 13C glucose in their study of Escherichia coli protein turnover, in a method reliant on culture doping to measure the relative ratio of new protein synthesis to protein degradation.[33] This approach requires that the cells have enough time to take up and start incorporating the substrate into proteins. For prokaryotes, this problem is solved by waiting approximately one cell doubling time, which is possible as the measurement is relative, not absolute. However, the choice of glucose as a carbon source introduces additional complexity, as it labels different amino acids to different extents. This variation in labeling dynamics between amino acid pools means that it is difficult to define absolute rates of protein labeling and, thus, absolute rates of synthesis, degradation and protein turnover.


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