4. Quantitative Mass
4.1. Relative Quantification
shift the separation as well as quantification problem from protein to peptide level as peptides are much easier to handle than pro-teins due to their physic-chemical characteristics. Today, several MS-based quantification methods, including chemical, metabolic, enzymatic labeling, and label-free approaches ranging from the quantification of single peptides up to the quantification of pro-teins from whole cell lysates, exist that can be used as an alterna-tive or complementary setup to 2D-PAGE for analyzing complex protein and/or peptide mixtures. They include methods for rela-tive and absolute quantification such as label-free approaches (see Subheading 4.1.1); isotope labeling, e.g., isotope-coded affinity tags (ICAT) (92), isotope-coded protein labeling (ICPL) (93), isobaric tags for relative and absolute quantification (iTRAQ, TMT) (94), enzymatic labeling during protein hydrolysis in the presence of heavy (18O-containing) water (95, 96), and stable iso-tope labeling with amino acids in cell culture (SILAC) (97); and absolute quantification of proteins (AQUA) (98, 99). For a gen-eral overview, see (28, 29, 100, 101). All the listed methods hold their advantages and disadvantages. Global internal standard (GIST) approaches where proteins are digested to peptides prior to labeling hold two major limitations: the high sample complexity results in the detection and quantification of only a limited num-ber of peptides (undersampling of the mass spectrometer), and by protein digestion prior to labeling, all information about the origi-nal belonging to the resulting peptide is lost. For protein-based chemical labeling, the main limitation is the incomplete labeling of the proteins resulting in falsified results. Today, the most accurate results are obtained with SILAC; this method is indeed mainly restricted to cells grown in culture and simple organisms.
In the next two chapters, most frequently used methods for MS-based relative protein/peptide quantification are described shortly.
Labeling of proteins or peptides with isotopes or other kinds of reagents distinguishable by MS is the most common strategy for gel-free protein quantification in proteomics. It is a universal approach as labeling is done after protein extraction. Over the years, several strategies have been developed which each suit dif-ferent needs. Usually, they are used for “shotgun” experiments starting directly on peptide level using LC–MS for separation, quantification, and sometimes even identification in one step. It is to be noted that these parameters depend much on the capabili-ties of the mass spectrometer used. Disadvantages of isotope labeling include cost expensiveness and the possibility of incom-plete labeling. Most of the state-of-the-art labeling chemistries are summarized by Julka and Regnier (100).
4.1. Relative Quantification 4.1.1. Isotope Labeling
17 Instruments and Methods in Proteomics
As the first method using isotopic labels for quantitative MS, the ICAT or cleavable ICAT (cICAT) was invented by Aebersold and co-workers in 1999 (92). The reagent with specificity toward side chains of cysteinyl residues consists of three elements: first, a reac-tive group toward thiol groups (cysteines); second, a linker con-taining either 12C (light ICAT) or 13C(heavy ICAT) atoms; and third, a biotin group that can be used for affinity purification before MS analysis. To quantify protein expression levels, e.g., of two different cell states, the protein mixture of the first cell state is labeled with light ICAT and the protein mixture of the second is labeled with the heavy ICAT. After pooling of both samples, they are enzymatically digested to peptides, separated with HPLC, and analyzed via MS. The light or heavy ICAT-modified peptides co-elute in HPLC and can be easily distinguished from each other by a 9-Da mass shift. The relative quantification is determined by the ratio of the peptide pairs (102). The main drawback is that ICAT cannot be used to quantify all proteins due to the fact that the number of proteins containing cysteines is restricted and only limited sequence coverage of the protein can be reached (28). As a result, information about protein isoforms, degradation prod-ucts, or posttranslational modifications, which are not located in the cysteine-containing peptide, are lost.
The techniques isobaric tags for relative and absolute quantifi-cation (iTRAQ) and tandem mass tagging (TMT) were first introduced by Ross and Thompson, respectively (94, 103). Either protein or peptide labeling can be performed on lysine residues and/or the N-terminus. To date, eight different iTRAQ with eight different isobaric (same mass) mass tags, and six TMT reagents are available, allowing for multiplexing of samples.
Isobaric peptides hold the advantage of identical migration prop-erties in the HPLC before MS analysis. Quantification is done after peptide fragmentation by the generation of label-specific low molecular weight reporter ion and signal integration. The different tags can be distinguished after peptide fragmentation as they result in different mass spectra. Therefore, this method allows the simultaneous determination of both identity and rela-tive abundance of the peptide species (104, 105). iTRAQ and TMT can also be used for absolute quantification. Indeed, both methods hold the described limitations of GIST approaches.
Additionally, iTRAQ/TMT quantification cannot be obtained on all kinds of mass spectrometers as low molecular mass reporter ion region is not accessible in all instruments.
Isotope-coded protein labeling is based on isotopic labeling of all free amino groups in proteins (93). Proteins from two differ-ent samples are extracted, alkylated, and labeled with either the isotope-free ICPL (light) or the isotope ICPL tag (heavy). After labeling, the protein mixtures are combined, optionally separated, e.g., by 1D-PAGE to reduce complexity, enzymatically digested, 4.1.1.1. Chemical Labeling
18 May et al.
and subsequently analyzed by MS (93). The heavy and light peptides differ in mass, and are visible as doublets in the mass spectra. Again, the peak intensities reflect relative quantitative information of the original proteins. The main advantage of this approach is the labeling already on protein level, circumventing all described limitations of the GIST approaches, although it holds the risk of incomplete protein labeling.
Enzymatic labeling with heavy water (16O/18O method) uses the fact that during protein digestion with trypsin, Glu-C or Lys-C up to two O atoms are incorporated into the peptide.
Thus, digestion in the presence of H218O results in a peptide mass shift of 4 Da compared to that in peptides generated during diges-tion in the presence of normal H216O. In a workflow using the
16O/18O method, the samples are independently digested in the presence of either H216O or H218O, and the samples are pooled and separated by HPLC, followed by peptide quantification and identification. This method is relatively simple; indeed, it holds the risk of back exchange of the O atoms and does not allow for multiplexing.
Stable isotope labeling by amino acids in cell culture (SILAC) is a metabolic labeling based on the in vivo incorporation of specific amino acids into mammalian proteins (106). For example, mam-malian cells are grown up in a medium with normal essential amino acids (light label) and concomitantly in a medium with isotopic modified forms of essential amino acids (heavy label).
After some proliferation cycles, the isotopic/normal amino acids incorporate completely into the cells. Protein extracts can be pooled, digested, and analyzed by MS. The heavy and light pep-tides elute as peak pairs separated by a defined mass difference.
The ratios of the resulting relative peak intensities reflect the abundances of each measured peptide (107). Mainly, the isotopes
13C, 15N, 2H, and 18O are used for stable isotope labeling. The incorporation of the isotopes in proteins can be performed in cell culture and even in vivo in simple organisms such as Drosophila melanogaster, Caenorhabditis elegans, or mice (107, 108). For higher organisms, especially humans, this kind of metabolic label-ing is technically not feasible or completely impossible due to ethical reasons.
To overcome the limitations of incomplete labeling, and also to spare costs and to reduce loss of proteins in the cause of sample prepa-ration, label-free MS approaches have been developed (101, 109).
One disadvantage of label-free quantification indeed is that this technique does not allow multiplexing, and has a slight lack of sensitivity compared to labeling assays. Nevertheless, label-free approaches offer the opportunity to analyze samples with a 4.1.1.2. Metabolic Labeling
4.1.2. Label-Free Quantification
19 Instruments and Methods in Proteomics
protein amount that would be too low for labeling or 2D-DIGE strategies, since they omit many preparation steps.
In spectral counting, the number of mass spectra repeatedly measured for one protein serves as a value for quantitation of this ion (109, 110). It could be shown that this number is propor-tional to the concentration of a peptide in a sample when ana-lyzed by nano-LC–MS (111). This is due to the fact that the higher the concentrations of a peptide, the longer it will take to elute from the HPLC system. Modern mass spectrometers can produce several MS² spectra in the time interval the peptides need to completely elute and be ionized by ESI. Disadvantages of spec-tral counting rise from the complexity of biological samples: Even with the best available LC system, co-eluting of peptides will still occur when analyzing complex mixtures such as cell lysates. Mass spectrometers will not be able to identify all co-eluting peptides at once. As a consequence, several replicated LC–MS runs will be needed to reach maximum identification results from one sample (111). This also leads to the second disadvantage of spectral counting that quantitative information can be obtained only from the peptides chosen as precursors, while information on less intensive or unselected peptides will be lost. Nevertheless, spec-tral counting is a cost-sparing alternative to labeling assays taking into account that this approach seems to be accurate, especially for high abundance proteins, but is highly sensitive to run-to-run variations (normalization is mandatory!).
One of the latest quantitative MS methods that is still under development is comparative or differential LC–MS (112). This method utilizes the ability of mass spectrometers to record not only m/z and the intensity of the MS signal, but also RT. Softwares use these data to build contour plots in the form of heat maps, in which RT and m/z span up a plane, and MS intensity will be displayed in a color code (101). Quantitative information is obtained by integration of the volume of the m/z–RT intensity peaks. Software calculates the features which are the sum of all peaks generated by one peptide as quantitative factors. Special algorithms are used for normalization between the LC–MS runs.
The advantage of this method is that it does not need any MS² spectra for quantitation, with the result that all signals recorded in one LC–MS run will be quantified. This could become the main advantage of comparative LC–MS, as the quantitative informa-tion should be more extensive than in spectral counting. Indeed, spectral counting still has advantages in sensitivity and reproduc-ibility (109). A major disadvantage of comparative LC–MS is that it allows no multiplexing and thus is more sensitive for run-to-run variations than labeling methods. Nevertheless, some studies report successful use of comparative LC–MS methods (example given by Johansson (113)).
20 May et al.
Intensive effort is spent currently to improve label-free quantification approaches, especially with respect to reproducibility, data analysis, and statistics.
Over the last years, proteome research is more and more focused on the Absolute quantification of proteins (AQUA). AQUA per-mits the direct quantification of differences in proteins and post-translational modified protein expression levels (98). Therefore, chemically synthesized isotope peptides, which are unique for the proteins of interest, are used as internal standards by adding a known quantity to the analytical sample (114, 115). The ratio of synthetic to endogenous peptide is measured and the absolute level of the endogenous peptide can be precisely and quantita-tively calculated and consequently the absolute levels of proteins and posttranslational modified proteins are known (98).
Although there are efforts to use MALDI, factors such as variable crystallization and laser ablation may lead to poor repro-ducibility, and thus generally ESI is the method of choice for AQUA (114). Before starting the AQUA approach, one has to adjust the peptide retention by RP chromatography, ionization efficiency, fragmentation via CID, and the amount of added stan-dards to fit with the dynamic detection range of the mass spec-trometer (see Gerber et al. for detailed information (98)). In a rather complex sample, the detection of the desired peptide likely competes with the detection of other isobaric peptides in the sample. This can be overcome by the combination of AQUA with MRM, allowing for a selective absolute quantification of the tar-get protein (115). This technique is of considerable benefit for, e.g., the absolute quantification of known biomarkers. Other available approaches for absolute quantification based on internal standards are QConCat (116) and protein standard for absolute quantification (PSAQ) (117).
In the past decade, major developments in instrumentation and methodology have been achieved in proteomics. Powerful tech-niques have been established to identify and differentially quan-tify protein species of complex biological samples. Many proteomic laboratories are investigating new techniques to overcome consis-tent obstacles. Beyond alterations of the genome, the increasing advances in proteomics hold great promise for a comprehensive description of protein isoforms or even posttranslational modifi-cations. With the ongoing improvement of sample preparation techniques and mass spectrometer sensitivities, the resolution of quantifiable compounds will be further improved in proteomics 4.2. Absolute
Quantification
5. Summary
21 Instruments and Methods in Proteomics
research allowing for the identification and especially reliable quantification of, e.g., physiologically relevant biomarkers indicating specific disease states.
1. For the electrophoretic separation of membrane proteins, conventional 2D-PAGE is not suitable. For this purpose, the application of specialized gel-based gel techniques such as CTAB- or BAC-SDS-PAGE, or MS-based methods is highly recommended (15, 118, 119).
2. Whenever a labeling approach is chosen for quantitative pro-teomics, labeling limitations have to be considered. For example, a saturation DIGE approach in 2D-DIGE will enhance the sensitivity but only cysteine residues will be labeled. Since cysteines are not found in all proteins, informa-tion about these proteins is lost. Moreover, peptide labeling might be more efficient than protein labeling.
3. In order to rule out labeling preferences, a dye swap should be included in 2D-DIGE experiments. This can be performed by switching the labeling dyes of samples A and B in two con-secutive experiments.
4. Protein differences between samples which have been found to be statistically valid in one technique need to be further validated by an independent method.
5. One has to consider that gel-based and MS-based techniques generally do not result in identical protein lists. Rather, both approaches complement each other. For a detailed and broad description of proteins within a sample, one may think about combining both approaches.
Acknowledgments
FB, PC, CS, BS, and KM are funded by the BMBF (grant 01 GS 08143). CM is supported by the Alma-Vogelsang Foundation.
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