Figure 1. Quantification of protein expression levels based on 2D gel electrophoresis. Proteins can be visualized using a variety of protein stains; in this example, silver staining was used. The images are of 2D gels loaded with 150 μg protein extract from yeast cells, which were grown in aerobic chemostat cultures: (a) carbon-limited for ethanol or (b) carbon-limited for glucose, respectively. In this experiment, protein expression levels were determined by comparing spot intensities on triplicate 2D gels. The differences in spot density correlate with differences in protein abundance. Magnified regions of triplicate 2D gel images are shown in (c) and (d). In (c), examples of proteins that were detected on the 2D gels of only one of the nutrient-limited chemostat cultures and were undetectable on the 2D gels of the other nutrient-limited chemostat culture are shown and in (d) several other, statistically significant changes in protein expression levels between the ethanol-limited and glucose-limited yeast chemostat cultures are shown. In both (c) and (d), protein spots of interest are indicated with a circle, with the corresponding protein names on the left hand side. Histograms show the protein abundance, with protein intensity of the 2D gels indicated in black for the glucose-limited and white for the ethanol-limited cultures. Figure adapted, with permission, from Ref. [44].

Figure 2. Overview of recent technologies in quantitative proteomics. (a) Two-dimensional differential in gel electrophoresis (2D DiGE). 2D DiGE is used to analyze more than one sample on a single 2D gel. This method employs pre-electrophoretic labelling of proteins with different spectrally resolvable fluorescent cyanide dyes (e.g. Cy2, Cy3, Cy5). Proteins common to the three samples are subsequently labelled with three structurally similar tags, which are size and charge matched. Identical proteins have the same mobility regardless of the dye and will migrate to a single spot on a 2D gel. A typical experimental setup is designed in such a way so that two dyes (Cy3 and Cy5) are used to label different protein samples, while the third (Cy2), is used to label an internal standard. The internal standard can be used on every 2D gel within the experiment, which enables accurate comparison of protein abundance between samples and allows improved spot detection between different gels. (b) Stable isotope labelling. In this approach, all peptides and/or proteins that are present in a sample are labelled with a stable isotope. After mixing the unlabelled (‘light’) sample with the stable isotope labelled (‘heavy’) sample, the sample is processed and analyzed by mass spectrometry to determine relative protein expression levels. Figure adapted, with permission, from Ref. [63].

Figure 3. An overview of different strategies for quantification through stable-isotope labelling and mass spectrometry. These protein quantification methods make use of stable isotope labels (i.e. chemically identical but mass-differentiated tags) to label all peptides and/or proteins that are present in a sample. After mixing the unlabelled sample with the stable-isotope-labelled sample, the sample is processed and analyzed by mass spectrometry to determine relative protein expression levels. The mass spectrometric intensity response is independent of the isotopic composition of the peptides. The ratio of the integrated signal peak areas of the light and heavy isotopic form of the same peptide, therefore directly reflect the protein abundance ratio. The stable isotope label can be incorporated into proteins or peptides at different moments during sample preparation (labelling is indicated with black bold boxes). The methods are: (a) Metabolic labelling. Metabolic labelling is achieved by growing an organism in the presence of a stable- isotope-labelled nitrogen or carbon source, such as ¹⁵N ammonium sulphate, ¹³C-labelled glucose or labelled amino acids. In a typical labelling study, cells are grown under different conditions and labelled with different forms of the stable isotope. Subsequently, these cells are mixed and analyzed using mass spectrometry; (b) Protein labelling (e.g. the ICAT approach); and (c) Peptide labelling, either during digestion in ¹⁸O labelled water, via derivatization of the peptides or via methods such as iTRAQ.

Figure 4. General scheme for protein complex analyses. Protein complexes can be purified using different techniques, such as affinity purification, immune purification and tandem affinity (TAP) purification. In the affinity-based purification method, proteins are tagged (fused) with a bait, such as one or more proteins or peptides (epitopes), that enables affinity purification. The tagged protein and its interacting partners are co-purified, separated by 1D gel electrophoresis, proteolyzed and then identified by mass spectrometry. In this way, protein-interaction networks in the cell can be established. Several comprehensive studies have been reported to unravel the functional organization of the yeast proteome by systematic analysis of protein complexes 65 and 66. Comparison of yeast and human complexes showed that conservation across species extends from single proteins to the protein assemblies. It will be interesting to extend these studies to unravel how these complex dynamic networks of protein interactions adapt to biological processes, such as cell cycle and budding, and also to changes in environmental growth conditions.