Determination of the absolute amount of proteins in proteomic studies is often derived from a measurement of the absolute peptide concentration. This is achieved by either providing standard peptides or by label-free approaches. In addition, a procedure for absolute quantification of intact proteins using labeled standard proteins has recently been described (Waanders et al., 2007).
Absolute quantification using stable-isotope labeled standard peptides The use of synthetic standard peptides for absolute quantification was originally described in 1983, where enzymatically deuterium labeled standard peptides were used for absolute quantification of enkephalin in thalamus tissue (Desiderio and Kai, 1983). This approach has been refined by Gygi and co-workers, and is now commonly known as AQUA (absolute quantification; Gerber et al., 2003; Kirkpatrick et al., 2005). Heavy isotope labeled standard peptides (also termed AQUA peptides) are synthesized by incorporation of one 13C- and/or
15N-labeled amino acid. As a result, the standard and the endogenous peptide share the same physicochemical properties, including chromatographic co-elution, ionization efficiency, and fragmentation pattern during MS/MS experiments. Importantly, the endogenous and the standard peptide can be distinguished by a distinct mass shift in the MS spectrum caused by the incorporated heavy labeled amino acid. The endogenous and the corresponding standard peptide thus present a peak pair consisting of a light peak (the non-labeled endogenous peptide) and a heavy peak (belonging to the standard peptide harboring incorporated stable isotopes; see Figure 2.3 for an example). The signal intensities of the light (endogenous) and the heavy (standard) peptides reflect the relative amounts. Because the concentration of the standard peptide is known, the absolute amount of the endogenous peptide and finally the protein can thus be determined. Absolute quantification using standard peptides is often applied to measure the level of particular peptide modification (e.g.
phosphorylation (Gerber et al., 2003), ubiquitinylation (Kirkpatrick et al., 2005)) or to analyze and validate biomarkers in clinical studies (Pan et al., 2005). The selection of standard peptides is often empirically (Bantscheff et al., 2007), i.e. the choice of synthetic standard peptides results from the analysis of endogenous peptides generated from the proteins under investigation. There are several aspects that have to be taken into account when selecting standard peptides, e.g. possible modification of amino acid residues (e.g. oxidation of methionine), chromatographic elution, ionization efficiency etc. A study about prediction of frequently detected tryptic peptides in a given proteomic platform (so-called proteotypic peptides) might help when selecting standard peptides for absolute quantification (Mallick et al., 2007).
Figure 2.3: Example MS spectrum of an endogenous and the corresponding standard peptide for quantification. MS spectrum of ILLGGYQSR (CDC5L protein) and the corresponding synthetic standard peptide.
The endogenous and the standard peptide show according to the incorporated stable isotope labeled amino acid (arginine) a distinct mass shift (10 Da). The spectrum was acquired by nanoLC-offline MALDI-ToF/ToF-MS.
There are however limitations for the absolute quantification using standard peptides. First, the addition of standard peptides to a proteome digest provides quantification of only single or few proteins of the sample. For reliable absolute quantification of a protein in a mixture, several standard peptides per protein have to be first selected and then provided during quantification to achieve more than one reference value per protein. Furthermore, incomplete protein hydrolysis is one of the most critical issue in the event of absolute quantification as it dramatically affects the final result.
To increase confidence in quantification experiments, addition of multiple peptides for each protein to be quantified is recommended. This can be simplified by using labeled standard proteins, which provide multiple peptides for absolute quantification after their hydrolysis. To this end, heavily labeled amino acids are incorporated into entire proteins resulting in heavily labeled standard proteins, which are then added to the sample under investigation. After protein hydrolysis of the endogenous and the standard proteins, standard peptides for all generated endogenous peptides are available. Several approaches using labeled standard peptides have recently been introduced (PSAQ, Protein Standard Absolute Quantification (Brun et al., 2007); Absolute SILAC (Hanke et al., 2008); FLEXIQuant, Full-length expressed
stable isotope-labelled proteins for quantification (Singh et al., 2009)). Very similar is the use of artificial QconCAT proteins, which are assembled from different standard peptide sequences (concatenated signature peptides encoded by QconCAT genes; Pratt et al., 2006). During hydrolysis of QconCAT proteins several standard peptides belonging to diverse proteins are generated allowing the quantification of different proteins in a sample.
Label-free approaches for absolute quantification Label-free quantification is a method that determines relative or absolute protein amounts without using stable isotope containing compounds. The advantage of label-free quantification is that time-consuming steps of introducing labels to proteins or peptides are not required and that there are no costs for expensive labeling reagents. Furthermore, there is no limit as to the number of experiments to be compared and several peptides per protein are available for quantification.
Mass spectral complexity is not increased as in the case of differently labeled samples what provides a higher analytical depth. Unfortunately, label-free approaches are least accurate among the mass spectrometric quantification techniques and they require a high reproducibility at each step, i.e. all experiments need to be accurately reproduced to achieve reliable quantification (Bantscheff et al., 2007).
One possibility to determine absolute protein amounts without labeling of peptides or proteins is to make use of the number of observed and theoretically observable peptides. The protein abundance index (PAI) is then calculated by dividing the number of observed tryptic peptides by the number of theoretically observable peptides from a particular protein and gives an estimate for absolute protein amounts in a complex mixture (Rappsilber et al., 2002). For absolute quantification, the PAI was later converted to an exponentially modified form (emPAI), which is proportional to the protein content in a protein mixture. Ishihama et al.
have shown that the emPAI-abundances from the actual values are within 63 % on average.
Nonetheless, emPAI values are easily calculated but provide only a rough estimate of the absolute protein amounts (Ishihama et al., 2005).
Incomplete digestion is a critical issue for absolute quantification. One way to deal with this is to average the quantities of the three most abundant peptides of every protein. It is generally assumed that some parts of the protein are completely digested and thus the three most abundant peptides reflect the protein concentration. The protein mixture is spiked with an intact standard protein and after hydrolysis the average MS signal response of the standard protein is used to calculate an universal signal response factor (ion counts/mole of protein) for the particular experimental setup. This factor can then be used to determine the concentration of the analyzed proteins within the mixture (Silva et al., 2006).
Top-down quantification of SILAC-labeled proteins Top-down is the analysis of intact proteins instead of generated peptides. To date, only one study about top-down absolute quantification has been released. Waanders et al., 2007 introduced the quantification of intact SILAC-labeled proteins. During the SILAC (stable isotope labeling using amino acids in cell culture, see below) method, cells are grown in media containing different isotope labeled amino acids. When using heavy and light lysine and arginine for SILAC labeling, intact proteins do not interfere with peaks of different charge states between 10 and 200 kDa. The authors have shown that two SILAC proteins (light and heavily labeled) can be quantified with an average standard deviation of 6 % (Waanders et al., 2007).
Absolute quantification to determine the protein stoichiometry within protein complexes By absolutely quantifying proteins in a purified protein complex, the protein stoichiometry of the quantified proteins can be established. In recent years, only few studies addressed the protein stoichiometries within protein complexes using absolute quantification. Two studies, combining chemical labeling of endogenous and standard peptides, were recently introduced. Hochleitner et al., 2005 determined the protein stoichiometry of the spliceosomal U1 small nuclear ribonucleoprotein complex and Holzmann et al., 2009 determined the stoichiometry of the MP1-p14 complex. A different approach has been designed for affinity-tag purified protein complexes (Wepf et al., 2009). An amino acid sequence serving as standard peptide is embedded in the affinity tag and is released after tryptic digestion. The protein is then quantified by adding a stable isotope labeled reference peptide of the tag and the other proteins are quantified by correlational quantification to the tagged protein. This approach benefits from the fact that one stable isotope labeled standard peptide can be used for quantification of different proteins, but requires the protein of interest to be present as tagged protein in the sample.