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Relative quantification in proteomics: new approaches for biochemistry
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TRENDS in Biochemical Sciences
Vol.31 No.8
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Relative quantification in proteomics: new approaches for biochemistry Richard D. Unwin, Caroline A. Evans and Anthony D. Whetton Stem Cell and Leukaemia Proteomics Laboratory, University of Manchester, Kinnaird House, Kinnaird Road, Withington, Manchester, M20 4QL, UK
Recent developments in mass spectrometry and protein arrays provide opportunities to derive systematically proteomic information from small samples of cellular material. Relative quantification among samples can be achieved with either gel-based or gel-free approaches. Furthermore, the adaptation of specific techniques facilitates absolute quantification. Here, relative quantification in two-dimensional gel electrophoresis is contrasted with that in non-gel-based approaches such as isobaric tagging of peptides, pre-labelling of living cells with isotopomeric forms of essential amino acids and protein array platforms. In addition, novel flowcytometry-based approaches are considered. These technologies can all be used to determine accurately the levels of proteins or biomarkers in a wide range of samples. Introduction The opportunities offered by genome sequencing have had an impact on biochemistry in that there is increasing expectation that systematic analysis of the protein content of the cell – namely, the proteome – will form an objective approach to studies of biological systems. Initially, it was thought that the conversion of genome data into rapid and effective proteome analyses would be immediately feasible. This possibility has been confounded by issues based around methodologies for protein identification and for determining relative protein levels in biological samples. These obstacles have been addressed by developing analyses, including quantification, for multiple proteins using various platforms. It is becoming increasingly clear that studying relative mRNA expression cannot provide details on all of the changes in components of cell systems. Specifically, mRNA changes do not relate directly to changes in protein levels, owing to different rates of translation, posttranslational modification, subcellular localization and degradation [1,2], which underlines the need for proteomic relative quantification techniques. In addition, the processes of posttranslational regulation of protein levels and activity, including events such as phosphorylation, glycosylation, acetylation and ubiquitination, all require quantification. Here we describe recent developments that have enabled detailed investigations of the proteome and posttranslational regulation. Corresponding author: Whetton, A.D. ([email protected]). Available online 11 July 2006 www.sciencedirect.com
Protein microarray technologies Protein microarrays facilitate the parallel analysis of multiple proteins in a high-throughput, miniaturized format for global protein screening and are available in two forms: one that determines protein function, and one that provides relative quantification. Although such analyses present considerable technical challenges, important advances have been made since the first proteome array was described [3]. Microarrays are currently generated by immobilizing multiple biomolecules onto solid supports. A high specificity of interaction between the immobilized binding molecule and the analyte is required to minimize cross-reactivity with other analytes and to decrease background signal. Functional microarrays Function-based microarrays can be used to analyse biochemical properties such as enzyme activity and substrate specificity [4] (Figure 1). For example, in vitro substrates for 87 different yeast protein kinases have been identified with chips spotted with >4000 proteins and the data used to generate a map of protein phosphorylation [5]. Protein–protein interactions can also be quantified with these arrays. An example of this approach has been the use of protein microarrays containing immobilized phosphotyrosine-binding domains of the ErbB receptor family: the interaction of each domain with 61 peptides containing the physiological sites of tyrosine phosphorylation from four ErbB receptors facilitated identification of complex protein interaction networks [6]. Applications can be performed in parallel – for example, protein microarrays comprising allelic variants of the p53 oncoprotein have been assayed for their ability to bind DNA and the MDM2 regulatory protein and to act as potential substrates for casein kinase II [7]. Chemical probes for activity-based protein profiling (ABPP) that report on active-site availability have been developed for the quantification of enzyme activity. ABPP complements functional arrays that assign enzyme function and substrate specificity (reviewed by Cravatt and Sorenson [8]). Probes for ABPP contain a reactive group directed to the active site, and one or more chemical tags, such as biotin or a fluorophore, for isolation and detection after one-dimensional or two-dimensional (2D) gel electrophoresis. Low-abundance proteins can be detected because ABPP is dependent on activity rather than expression. Applications include the identification of novel cancerassociated enzymes [9] and screening of enzyme inhibitors.
0968-0004/$ – see front matter ß 2006 Elsevier Ltd. All rights reserved. doi:10.1016/j.tibs.2006.06.003
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Figure 1. Relative quantification using protein microarrays: functional arrays. These microarrays can be used for high-throughput analysis of protein function. (a) Enzyme– substrate interactions can be determined by arraying potential substrates, incubating with an enzyme, and then quantifying each spot for enzyme activity, for example, by the addition of a (radioactive) phosphate group to a bound peptide or protein to determine kinase activity. (b) To determine protein–protein interactions, peptides, protein domains or proteins are immobilized as discrete spots. Potential binding partners (comprising single or multiple molecules) are then incubated with the array. Here, binding partners are fluorescently tagged (indicated by a star) to facilitate the detection of specific interactions. This type of approach has several applications, including the identification of binding partners for motifs such as SH2 domains or basic-region leucine zipper domains. Such applications can be performed in parallel.
In terms of inhibitor screening, ABPP is particularly useful for enzymes lacking known substrates. ABPP technology has been combined with liquid chromatography MS to achieve greater coverage of the proteome [10] and could be combined with isotope-coded mass tagging for relative quantification. Protein profiling microarrays There are two main types of array for relative quantification by protein profiling: forward-phase (capture) arrays and reverse-phase arrays. These microarrays have found considerable application in cancer research [11,12], where they are used to identify biomarkers for diagnostics and to discover potential therapeutic targets. Forward-phase arrays. In forward-phase arrays, proteins for analysis are captured by an array of molecules immobilized either on a slide format (Figure 2) or on beads at an optimal concentration. Many different capture molecules can be present in the array, facilitating relative quantification of multiple proteins. Arrays are typically probed with cell lysate or serum, and protein binding is assessed by direct labelling via protein tagging or indirect detection (i.e. a ‘sandwich’ type approach in which captured proteins are detected by matched antibodies), enabling signal amplification [13]. Differential protein expression profiling has been used to compare serum proteins from normal individuals and cancer patients (reviewed by Hamelinck [14]) and to analyse discrete protein subsets such as secreted cytokines [15]. Relative quantification is achieved by comparing control or test samples with a reference sample (usually a pool of all samples in the analysis) using two-colour fluorescence labelling with Cy dyes. An adaptation of the www.sciencedirect.com
forward-phase approach facilitates analysis of both protein levels and phosphorylation status by using two different antibodies that bind a protein independent of and dependent on its phosphorylation status to determine the total amount of protein and the amount phosphorylated at a specific site, respectively [16,17]. Antibodies remain the best-characterized capture molecules. Not all antibodies function in this format, however, because activity is not always retained after immobilization. An alternative approach is the use of aptamers – that is, oligonucleotide sequences with molecular recognition properties selected from combinatorial oligonucleotide libraries [18]. Aptamers bind protein ligands with high affinity and specificity and, because they are chemically synthesized, they can be easily labelled with reporter groups. They also have advantages over antibodies of being reversibly denatured and usually more stable. Reverse-phase arrays. In reverse-phase arrays, multiple samples, which are often complex mixtures such cell lysates, are immobilized by deposition as microspots and probed with a specific detection reagent. Reverse-phase protein blotting facilitates simultaneous analysis of multiple samples for a single analyte in a western blot type approach (Figure 3). Antibodies that have been used include those against housekeeping proteins, signalling proteins, cell-cycle proteins, apoptosis-related proteins and phosphorylation motifs present in signalling molecules. A chief application of reverse-phase arrays has been in analyses of the phosphoprotein profiles of discrete protein subsets, whereby arrays are probed with two different antibodies to infer protein posttranslational status (and therefore often activity). One antibody is specific for a
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easily observed, however, lower abundance signalling molecules might not be detectable. Prefractionation of crude extracts could alleviate this problem. Protein array approaches are based on determining quantitative changes by using specific predetermined assays in which defined protein targets are selected in a subjective manner. Novel proteins or sites of posttranslational modification are not considered in this type of approach. Discovery-led proteomics offers an objective means of determining differences in protein expression without knowledge of the identities of the proteins to be assayed. This approach facilitates the identification of proteins in a non-hypothesis-driven manner and can be taken forward by several different methodologies.
Figure 2. Relative quantification using protein microarrays: forward-phase (capture) arrays. (a) Specific antibodies raised against proteins of interest are immobilized with a single capture antibody present in each spot. Many different capture molecules can be arrayed in this format for the simultaneous detection of multiple analytes. Recombinant antibody fragments, ‘affibodies’ (non-immunoglobulin-based affinity capture proteins) or aptamers (oligonucleotide sequences that bind epitopes with high affinity and selectivity) can be also used for forward capture arrays. Antibody arrays are used for protein quantification, protein binding studies and protein phosphorylation assays. (b) For relative quantification, comparison can be made to a reference sample (comprising a pool of all samples in the analysis) by using two-colour fluorescence labelling with Cy3 and Cy5 fluors, as shown here. Increased or decreased protein expression is identified from the ratio of the two dyes, analogous to DNA microarray analyses.
phosphorylated epitope and the second assesses total protein abundance. Reverse-phase protein arrays have been used to analyse signalling pathways with antibodies for specific phosphorylation sites in phosphoproteins, including MEK, MAPK, Raf and AKT kinases and the STAT family of cytoplasmic transcription factors [19]. A disadvantage of reverse-phase protein arrays is that only one analyte can be measured on a single array, although this can be addressed by printing arrays in a multisector format, thereby enabling parallel analysis of multiple analytes in the same experiment. Generally speaking, reverse-phase blots have the advantage that proteins do not require labelling. Although high-abundance proteins present in cell lysates can be www.sciencedirect.com
Two-dimensional PAGE Two-dimensional PAGE (2-DE) has been used for proteomic analysis for >30 years (reviewed by Gorg et al. [20]). Proteins are resolved on the basis of two independent parameters: overall electric charge and size (Figure 4a). In the first dimension, isoelectric focusing (IEF) separates proteins in a pH gradient gel on the basis of isoelectric point (pI). Proteins are then resolved orthogonally by molecular size using SDS–PAGE. Protein spots can be detected and quantified with chemical stains (reviewed by Patton [21]), which are selected on the basis of sensitivity (0.05–500 ng per spot), reproducibility of staining, and compatibility with downstream methods of protein characterization (Figure 4b). Coomassie blue stain and silver stain have been traditionally used, but fluorescent protein stains offer greater quantitative accuracy owing to their wider linear dynamic range (1000-fold) and greater compatibility with downstream protein characterization methods. Fluorescent stains specific for posttranslational modifications have been also developed [22]. One advantage of 2-DE over alternative gel-free peptide-based approaches is that, because protein modification can induce alterations in both mass and pI, different isoforms and posttranslationally modified protein species can be resolved. In addition, parallel profiles of phosphorylated and total proteins can be obtained by a multiplex proteomic approach [22] using sequential gel staining with Pro-QTM Diamond phosphoprotein stain, followed by a total protein stain such as Sypro Ruby (Figure 4c). The inclusion of Pro-QTM Emerald glycoprotein stain in the procedure facilitates analysis of glycosylated proteins in the same sample [23]. Relative quantification can be achieved by a computerbased comparison of gel images, which matches and compares protein spot intensities between gels. Inter-gel comparisons, however, rely on high levels of reproducibility in both separation and staining. Intra-gel comparisons based on fluorescent dyes with distinct spectral properties overcome this problem. In ‘difference in-gel electrophoresis, cyanine dyes are used to label protein samples fluorescently; these samples are then mixed and resolved on the same 2-DE gel. Proteins can be labelled on lysine (minimal dyes) [24] or, for higher sensitivity of detection, cysteine residues (saturation dyes) with Cy3 and Cy5 [25]. Fluorescently labelled proteins are imaged with different excitation and emission filters to detect the protein profiles for
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Figure 3. Relative quantification using protein microarrays: reverse-phase arrays. Mixtures of proteins from different samples, for example cell lysates, are spotted onto the microarrays and probed with antibodies targeted against specific proteins. A single target is analysed for each sample spotted onto the array. To assay protein phosphorylation, parallel arrays are probed with an antibody that is phosphorylation independent (left) and one that is phosphorylation dependent (right). The first antibody detects the protein of interest in both its nonphosphorylated and phosphorylated (P) states, thereby assessing its concentration in the sample; the second assays only the phosphorylated form. The antibodies can be fluorescently labelled for visualization, as shown here, and the amount of each protein can be relatively quantified for a set of samples. This type of array has been applied to parallel analyses of the site-specific phosphorylation of a range of phosphoproteins, including MEK, MAPK, Raf and AKT kinases and the STAT family of cytoplasmic transcription factors, and has been used to profile the activation status of cell signalling pathways in samples from individuals with cancer.
each sample (Figure 4d). The introduction of a third minimal dye (Cy2) has refined the method to include an internal standard for each gel, which is typically a mixture of the two samples being compared [26]. The key limitations of 2-DE are dynamic range and protein solubility. Abundant proteins, present at >10 000 copies per cell, can obscure proteins expressed at a low level (10–1000 copies per cell). Prefractionation strategies alleviate this difficulty (see later). For 2-DE approaches, using narrow-range pH gradients (‘zoom’ gels) increases resolution and protein loading [27]. IEF remains problematic owing to its incompatibility with poorly soluble proteins such as membrane proteins (hydrophobic) and very basic proteins. This problem can be partially overcome by using non-ionic and zwitterionic detergents with different solubilization properties [28] and non-equilibrium pH gradients. Although 2-DE remains a useful means of globally comparing proteins from several samples, the issues of sensitivity, protein exclusion and reproducibility abound. Furthermore, when a difference in staining intensity is shown (Figure 4), it remains necessary to identify the protein by mass spectrometry (MS). Role of MS in relative quantification proteomics The development of more sensitive and efficient mass spectrometers has brought MS to the forefront of proteomics research. Most mass spectrometers used for relative quantification work operate on broadly similar principles. First, the peptide sample is ionized, either in solution phase by spraying from a charged needle (electrospray ionization) or from solid phase by using a laser to transfer a proton from an energy-absorbing matrix to the peptide (matrix-assisted laser desorption–ionization). Once inside the instrument, peptide masses are determined by ion time of flight. Although this is a common type of mass analyser, www.sciencedirect.com
other types such as Fourier-transform ion cyclotron resonance [29], ion traps [29] and Orbitraps [30] can be used. The masses determined for the peptides can be used to identify proteins (Box 1). For complex mixtures of peptides, a liquid chromatography step before MS enables the samples to be fractionated and peptides to be analysed sequentially. Combining two types of chromatographic separation enables further fractionation of complex mixtures, thereby increasing both the number of peptides identified and coverage of the proteome [31]. Relative quantification by MS Although MS provides sensitive detection and identification of proteins and peptides, it is not directly quantitative. Two peptides from the same protein will not necessarily generate the same signal because differences in amino acid sequence will affect the efficiency with which they ionize. Indeed, often the same peptide will not produce exactly the same signal when analysed a second time, perhaps from a different sample, owing to slight differences such as coeluting peptides, solvent composition, and the age, condition and position of the electrospray needle. To overcome this problem, several methods have been developed. Chemically identical but isotopically distinct tags can be used to modify the peptide populations being compared. These peptides are then pooled, and analysed in a single MS run. Because the tags are chemically identical, the same peptides from each sample will behave identically; that is, they will elute from a chromatography system at the same time, and be ionized at the same time and therefore with the same efficiency. Relative quantification is obtained by comparing the signal intensities of peptides with a defined, isotope-induced mass difference (Figure 5a). Several methods for incorporating ‘stable isotope’ have been described for relative quantification (Box 2).
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Box 1. Protein identification by MS For identification of a single protein, such as a spot from a 2-DE gel, digestion of a protein with a specific enzyme produces a predictable, unique profile of peptide fragments, termed a ‘peptide mass fingerprint’. Measurement of these masses facilitates data comparison with theoretical peptide mass fingerprints derived in silico for all proteins in a selected database. This method is usually faster for the identification of large numbers of samples, because few spectra have to be generated per sample. However, the confidence associated with protein identification by peptide mass fingerprinting is lower than that associated with identification based on peptide sequence information.
Higher confidence peptide sequence information can be derived by MS/MS. In this technique, a specific peptide ion is selected by filtering out all other ions using a quadrupole. This peptide is induced to fragment by collision with an inert gas, such as nitrogen or argon. Because fragmentation occurs along the peptide backbone, the masses of the fragment ions indicate the peptide sequence. Figure I shows fragmentation of a peptide from bovine serum albumin, AEFVEVTK, together with its major fragment ions and the amino acid sequence to which these ions correspond.
Figure I. Peptide sequencing by MS. A tandem mass spectrometer consisting of a quadrupole and a time-of-flight analyser is shown. Peptide ions are generated by electrospray ionization from a fine needle. On entering the mass spectrometer, these ions are filtered on the basis of their mass-to-charge (m/z) ratio in the quadrupole. A peptide ion selected on the basis of its m/z ratio undergoes collisional induced decomposition after colliding with an inert gas. The fragment ions are then ‘pushed’ into the time-of-flight mass analyser in a synchronous fashion. There, peptide mass is inferred by measuring the time taken for a peptide ion fragment to travel from the source to the detector. The signal (y-axis of each spectrum) represents the number of ions that have hit the detector at a specific m/z ratio (x-axis). The pattern of ion masses is then used to infer peptide sequence information; in this example, the peptide is identified as one from bovine serum albumin. All labelled ions are singly charged. For clarity, not all ions have been labelled; these unlabelled ions might be multiply charged, might be fragments from the N-terminal half of the peptide, or might be derived from multiple fragmentation events on the same peptide.
Other relative quantification approaches have been subsequently developed, including ‘isobaric tags for relative and absolute quantification’ (iTRAQ) [32]. This method can incorporate up to four mass tags that label the N terminus of every peptide (along with free amines in lysine side chains). Each tag is isobaric (i.e. has the same mass) such that, when the four labelled samples are pooled, the same peptides from each sample appear at the same mass in the MS spectrum. Because the distribution of isotopes in each tag is different, however, on fragmentation each tag generates a specific distinct reporter ion at mass:charge ratio 114, 115, 116 or 117. The relative abundances of these reporter ions provide relative quantification (Figure 5b). iTRAQ offers some advantages. First, because all peptides are labelled it is possible to obtain data on more peptides as compared with other methods, thereby increasing confidence in the quantification of each protein [33]. Second, because the tags are isobaric, the signal from up to four samples is summed in MS mode, providing an increase www.sciencedirect.com
in sensitivity. For iTRAQ, tandem mass spectrometry (MS/MS) must be carried out on all peptides to obtain quantification, whereas for ICATs or SILAC (Box 2) MS/ MS is carried out only on proteins that are differentially expressed. Although this approach could be viewed as ‘wasting analysis time’, the definition of a large population of unchanging proteins enables the use of normalization procedures found in microarray analyses and comparison to such data sets [2]. When samples are labelled at the peptide level as in iTRAQ, it is important that the prefractionation methods (e.g. immunoprecipitation) applied are highly robust and reproducible to maintain the reliability of relative quantification. With ICATs or SILAC (Box 2), by contrast, proteins can be pooled and then prefractionated and digested together, thereby reducing potential sources of experimental variation. A key aspect of all stable isotope labelling procedures is that the efficiency must approach 100%. Underlabelling leads to the generation of background ‘noise’ because the
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Figure 4. Two-dimensional gel-based analysis of the proteome. (a) Principle of 2-DE. Proteins are resolved according to charge and mass. In the first dimension, proteins are separated on the basis of their net charge, which is the sum of the negative and positive charges conferred by the amino acid side chains and the N and C termini, and is pH dependent. The isoelectric point (pI) is the pH at which the net charge of the protein is zero. During IEF, the proteins migrate in a pH gradient to the specific pH corresponding to their pI. Proteins are solubilized and denatured before IEF to expose internal ionizing groups. In the second dimension, proteins are separated by molecular size using SDS–PAGE. For detection, there are several staining protocols that detect either all proteins or specific posttranslational modifications and that can be used for relative quantification. (b) Representative 2-DE data. Replicate 2-DE gels of samples for comparison (‘Control’ and ‘Test’) are run in parallel. Relative quantification is performed by gel image analysis using specialist software that performs spot detection and spot matching across data sets. Comparison is often based on normalized spot volume, which is defined as the volume of a particular spot relative to the total volume of all spots present in the gel. A representative set of histograms is shown for the normalized spot volumes of a selected spot difference across a gel set. (c) Multiplexed proteomic analysis using fluorescent stains. This method facilitates parallel profiling of protein expression and posttranslational modification in a single 2-DE gel. Protein gels are run and stained with a posttranslational-modification-specific stain and the gels are imaged. Gels are then stained for total protein expression with Sypro Ruby and imaged again. The resulting images are then matched by overlaying. Shown here is a representative sequence of protein spots that are a mixture of phosphorylated and nonphosphorylated proteins. Protein spots containing phosphate groups are detected by Pro-Q Diamond staining, whereas Sypro Ruby detects all protein spots. The phosphorylated proteins migrate to a more acidic position in the gel relative to the parent protein, owing to the addition of negative charge to the protein. Multiple samples can be analysed by inter-gel comparison of the staining profiles. (d) Difference in gel electrophoresis. Proteins from two different samples are each labelled with a single fluorophore that has distinct spectral properties (here, Cy3 and Cy5). Samples are then combined and resolved by 2-DE gel electrophoresis on a single gel for intra-gel comparison. The labelled proteins are detected by using different excitation and emission filters, and two separate images are generated for comparison. An additional sample, typically a mixture of the samples to be compared, can also be included (labelled with Cy2) to provide an internal standard. Differences in protein expression are determined by gel image comparison using specialist software that assigns protein spot differences on a ratiometric basis.
same protein might be labelled to various extents in different experimental samples. This labelling efficiency requires several population doublings for labelling in culture, or the development of efficient chemical reactions. Several groups have recently described a label-free relative quantification approach, termed ‘differential MS’, which is based on normalization strategies that enable comparison of the signal strength of the same peptide in two distinct sample runs [34] (Figure 5c). The advantages of this approach are that labelling is not required and that, provided both MS and chromatography remain relatively constant over time, a large number of samples can be analysed [34,35]. www.sciencedirect.com
Improving sensitivity: subproteome analysis A chief difficulty in defining and quantifying a proteome is sample complexity. Fractionating organelles or isolating a protein set with a specific property (e.g. tyrosinephosphorylated proteins) provides increased penetration of the proteome. The most common approaches to organelle fractionation are based on centrifugation and differential solubility, or on antibody-directed methods (reviewed by Yates et al. [36]). Although these approaches have existed for several years, the impact that they have on state-of-the-art proteomics is substantial. Good fractionation strategies enhance penetration: for example, analysis of nucleoli
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Box 2. Common methods of stable isotope labeling Isotope coded affinity tags Isotope coded affinity tags (ICATs) are chemical modifiers that covalently bind cysteine residues. Thus, ICATs can be labelled with either a heavy or light isotope and contain a biotin tag for affinity purification. Initially, the mass difference between the tags was generated with deuterium [67], but this method has since been improved and the ‘heavy’ tag now contains nine 13C residues. Each tag contains a cleavable linker attached to a biotin moiety so that labelled peptides can be purified; the biotin tag is then removed to generate a mass addition of 227 Da or 236 Da for the light or heavy tag, respectively [68]. At present, however, only two samples can be compared by this method and only cysteine-containing peptides are enriched. Stable isotope labelling in culture In stable isotope labelling in culture (SILAC), cells are cultured in medium containing a ‘heavy’ essential amino acid [69] e.g. [d3]leucine, [13C6]arginine or [13C6]lysine for one sample of a pair to be compared. Combining these isotopic amino acids enables up to three samples to be compared [70]. This approach is suitable only for material obtained
has yielded relative quantification data on 489 proteins [37], and analysis of Saccharomyces cerevisiae mitochondria has identified 750 proteins [38]. Other organelles from higher eukaryotes that have been specifically enriched for proteomic analysis include the Golgi apparatus (421 proteins) [39], phagosomes (up to 520 proteins) [40], lysosomes (215 proteins) [41], lipid rafts (241 proteins) [42] and metaphase chromosomes (102 proteins) [43]. In principle, it is relatively easy to adapt these protocols to include relative quantification by using the strategies described in Box 2 and Figure 5. Cell-surface proteins can be analysed specifically after protein tagging with biotin reagents, which are hydrophilic and cannot cross the hydrophobic plasma membrane. Biotin-tagged proteins can be enriched with streptavidin beads, which bind the biotin moiety. The recovery of integral membrane proteins is improved by removing plasmamembrane-associated cytoskeletal proteins with high salt and alkaline washes. On the basis of this approach, MS analysis has identified 781 mammalian plasma membrane proteins [44]. These data show that it is increasingly feasible to identify and to quantify relatively many proteins from a given cell system if sufficient fractionation is performed. In most types of cell, proteomics technologies make the relative quantification and localization of thousands of proteins an achievable aim. Improving sensitivity: posttranslationally modified proteins As an alternative to subcellular fractionation, other protein subsets such as glycosylated [45], ubiquitinated [46] and sumoylated [47] proteins (the latter two representing the ‘degradome’) can be isolated. So far, however, the best-studied modification is phosphorylation [48]. Isolation of phosphoproteins can be achieved by several methods. At present, antibody-based isolation is normally targeted at phosphotyrosine. Using this approach coupled with SILAC (Box 2), Kratchmarova et al. [49] achieved relative quantification of 282 tyrosine phosphorylated proteins. iTRAQ relative quantification has also been www.sciencedirect.com
by cell culture (at least five population doublings are required for isotopic equilibrium to be reached) and cannot be applied to primary material. 18 O incorporation Tryptic hydrolysis of peptides requires H2O. By performing the digestion in ‘heavy’ water (H218O), all tryptic peptides gain C-terminal 18 O, generating an apparent mass shift of 4 Da owing to incorporation of the isotope during peptide cleavage [71]. 13
C/15N guanidination Modification of the N-terminal lysine with methylisourea to form homoarginine was initially reported as a strategy to improve the ionization and/or transmission of lysine-containing peptides. By using a 15N-containing version of the reagent, a mass difference of 3 Da can be introduced [72]. This method is a cheap, efficient and simple way in which to introduce a stable isotope, but data are hard to interpret because often the natural isotopic ‘envelope’ or mass range of the differentially labelled peptides will overlap – a factor that must be corrected for before relative abundance can be calculated.
successfully used alongside phosphotyrosine immunoprecipitation [50]. An alternative method is to modify phosphorylation sites chemically to enable purification of phosphoserine- and phosphothreonine-containing proteins; quantification can be achieved by including a stable isotope tag [51]. Phosphoprotein enrichment can also be achieved with immobilized metal affinity chromatography. Although in most cases these enrichments have been combined with 2-DE for protein analysis [52], it is possible to use gel-free MS methods to gain even greater penetration into the phosphoproteome. Another common approach to assess phosphorylation is the enrichment of phosphopeptides. This approach has several advantages, including simplification of the analyte mixture and the ability to obtain both information about the site of the modification and relative quantification. Often, however, it does not reveal whether any difference is due to changing phosphorylation stoichiometry or overexpression of a constitutively phosphorylated protein. Nevertheless, by using immobilized metal affinity chromatography for enrichment of phosphopeptides [53] and sufficient starting material, it has been possible to identify >700 phosphopeptides in yeast [54] and 2000 phosphorylation sites from >900 proteins in mammalian cells [55]. Other methods for phosphopeptide enrichment such as titanium oxide [56], anti-phosphotyrosine antibodies [57] and diagonal chromatography [58] have been also used. All offer opportunities for increased proteomic penetration to identify posttranslational modifications crucial to various cellular processes. Absolute quantification Relative quantification is sometimes insufficient for the biochemical process or issue to be addressed fully. Several strategies now enable absolute protein quantification by MS. The first such method, called AQUA (absolute quantification) [59], uses a stable isotope-labelled synthetic peptide in a protocol similar to the stable isotope methods used for relative quantification (Box 2; Figure 5). This peptide is spiked into the analyte mixture at a known concentration, and the ratio of the test peptide to the
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Figure 6. The multiple reaction monitoring (MRM) technique. (a) Specific peptide detection by MRM. Triple-quadrupole mass spectrometers can be configured to enable only peptides of a specified m/z value into the collision cell, and a second mass filter to enable only a specific diagnostic ion to reach the detector. Several of these ‘transitions’, each performed in a short (millisecond) timescale, can be analysed in an experiment – a method known as ‘MRM’. The filtering vastly reduces background and therefore dramatically increases the signal to noise. Here, the first quadrupole of a mass spectrometer is set to detect a peptide ion at m/z 575.7 from a mixture of peptides delivered to the mass spectrometer. Ions of this mass are fragmented by collisional induced dissociation. If a diagnostic fragment ion with an m/z of 529.7 is generated, this ion passes to the detector. Peptides of m/z 575.7, which do not generate this product ion, do not produce a signal, demonstrating the selectivity of this analysis. (b) Absolute quantification by MRM. The inclusion of a standard peptide labelled with a stable isotope enables MRM transitions to be set up for both the standard and test peptides. Because the sample and standard peptides have different masses, they can be monitored in parallel by using different MRM transitions; because they are chemically identical, however, their response in the mass spectrometer should be the same. Therefore, comparison of the total signal generated by each peptide facilitates absolute quantification of the peptide and thus the protein of interest. Here, the signal of the sample peptide is 65% of that of the standard.
Figure 5. Strategies for relative quantification of peptides by MS. (a) Stable isotope labelling can be used for relative quantification. Various methods exist for incorporating the ‘heavy’ and ‘light’ isotopes (Box 2). Although being chemically identical, isotopes enable peptides from each sample to be discriminated and relatively quantified in the mass spectrometer by a comparison of the intensities of peaks that elute from a liquid chromatography system at the same time. Isotopomeric peptide ions from two samples have a defined mass difference but coelute. Here, the peptide at m/z 686.8 and its heavy counterpart at m/z 691 have equivalent intensity, whereas the ion at m/z 708.8 has higher intensity than its heavy counterpart at m/z 712.8. The ion at m/z 708.8 is thus selected for fragmentation to obtain sequence information and to show that this protein, radixin, is differentially expressed in samples A and B. (b) iTRAQ tags have identical overall mass and can react with all peptides via free amine groups. Here, each chemically identical peptide from the four samples appears at the same mass in an MS experiment. Colour coding in the stacked bars represents the fact that the isobarically tagged peptide has come from four different samples. In MS mode, there is no means of distinguishing from which of the four samples the peptides were derived. On fragmentation, however, specific reporter ions are released that differ in mass owing to differential isotope usage (this mass difference is offset in the intact tag by isotopic differences in a ‘balance’ group moiety). The ratios between these ions are thus representative of the relative expression of that peptide or protein. Here, fragment ions (bottom spectrum) clearly identify the peptide as one from a-internexin, an intermediate filament protein, whereas the reporter ions (top spectrum) show that this peptide is present at similar levels in sample A, B and C, but is more abundant in sample D. (c) Label-free relative quantification involves analysing samples in separate experimental runs in series. Peptides are separated by liquid chromatography and analysed by MS. A peptide-specific elution profile is then generated, which shows how much of the peptide (m/z 632.2) was detected throughout the experimental run. The area under this profile is then compared between two (or more) experimental runs to determine the relative quantity of that peptide. There is an assumption (which requires formal proof for each experimental system used in this way) that the ion current associated with a particular peptide is proportional to the molar quantity of the peptide present in each run in the sample series. This assumption is partly dealt with by complex data normalization to correct for inter-run variation owing to changes in temperature, sample loading and ionization efficiency. MS/MS is performed on every peptide in each sample to obtain data for inferred peptide sequence; here, it shows that sample B (blue) contains more glycogen phosphorylase than sample A (red). www.sciencedirect.com
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Figure 7. Analysis of cell-surface markers by inductively coupled plasma MS (ICP-MS). Cells express antigens on their surface that define their phenotype and that can be recognized specifically by antibodies. Tagging these antibodies with elements facilitates their detection by MS. Examples of elemental tags include the rare earth metals, europium (Eu), terbium (Tb) and samarium (Sm), as shown here. Cells are atomized and ionized on introduction into the inductively coupled plasma, and multiple elemental tags can be quantified simultaneously. The presence and intensity of the mass signals correlate to the relative expression of the specific cell markers. ICP-MS offers advantages over flow cytometry for multiplex analysis because, unlike fluorophores, the elemental tags produce signals that are essentially non-overlapping and a greater number of signals can be quantified in parallel [66].
‘heavy’ labelled standard is used to calculate the concentration of the protein. By using a synthetic ‘posttranslationally modified’ peptide, the absolute amounts of modified (phosphorylated) peptide can also be calculated, along with the total amount of that protein, giving information regarding modification stoichiometry. The QCAT method [60] uses a synthetic gene encoding a hybrid of several tryptic peptides from proteins of interest that can be isotopically labelled. Because all standard peptides are at identical concentrations, the simultaneous absolute quantification of many proteins becomes a realistic prospect. For complex mixtures, Anderson et al. [61] designed the ‘stable isotope standards and capture by anti-peptide antibodies (SISCAPA) workflow, in which standard labelled peptides are spiked into the analyte, and peptides of interest are enriched using antibodies. This approach can be improved by multiple reaction monitoring – a highly selective and exquisitely sensitive method for detecting a specific ion (peptide) in a sample (Figure 6a). Multiple reaction monitoring can be used to quantify the peptide and its labelled standard (Figure 6b) and is both more sensitive and linear over a greater range than is seen in time-of-flight MS [62].
can be used to discriminate selectively and to purify cell populations on the basis of their expression of characteristic cell-surface antigens. Cells can be fixed, permeabilized and labelled with antibodies directed against intracellular proteins of interest, such as phosphoproteins and apoptosis-related proteins [63,64]. In this way, defined cell populations, and even single cells, can be analysed in the context of complex cell populations for the presence of proteins of interest in a multiplex approach [65]. A flow-cytometry-based approach and MS have now been coupled. Specific cell marker proteins can be affinity labelled by using antibodies (or potentially aptamers) containing metal-containing labels that provide elemental tags [66]. Cells are analysed by inductively coupled plasma MS (ICP-MS) to determine elemental composition. This approach has been developed for the simultaneous identification of multiple cellular antigens using four diagnostic elemental tags (Au, Sm, Eu and Tb) coupled to secondary antibodies. It has the potential for higher throughput than can be achieved with current flow cytometry approaches. Potential applications include identification and analysis of low-abundance cancer stem cells in tumour samples (Figure 7).
Flow-cytometry-based relative quantification Flow cytometry measures physical and chemical characteristics of cells or particles in suspension. It is used routinely to analyse cell populations labelled with fluorescent antibodies directed against antigens of interest. When combined with fluorescence-activated cell sorting, it
Concluding remarks Many novel technologies are currently being applied to the multiplex-based assay of relative protein quantities. These technologies afford new opportunities to the biochemist and can be applied subjectively (antibody-based arrays) or objectively (MS-based approaches) to offer improved
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sensitivity and scope for the identification of large numbers of alterations in the proteome in response to a specific state or stimulus. The advent of more approaches as new techniques are developed is undoubted. In addition, improvements in MS instrumentation in terms of sensitivity and speed, together with better sample preparation and purification procedures will increase penetration into the proteome. The development of more complex and wide-ranging protein array technologies will soon make the simultaneous quantification of thousands of proteins, protein modifications and protein activities a realistic goal. The opportunities for systematic investigation based on these approaches abound and offer new horizons for quantification in biological systems. Acknowledgements Work in the authors’ laboratory is supported by the Leukaemia Research Fund, UK, and the Biotechnology and Biological Sciences Research Council.
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Relative quantification in proteomics: new approaches for biochemistry Richard D. Unwin, Caroline A. Evans and Anthony D. Whetton Stem Cell and Leukaemia Proteomics Laboratory, University of Manchester, Kinnaird House, Kinnaird Road, Withington, Manchester, M20 4QL, UK
Recent developments in mass spectrometry and protein arrays provide opportunities to derive systematically proteomic information from small samples of cellular material. Relative quantification among samples can be achieved with either gel-based or gel-free approaches. Furthermore, the adaptation of specific techniques facilitates absolute quantification. Here, relative quantification in two-dimensional gel electrophoresis is contrasted with that in non-gel-based approaches such as isobaric tagging of peptides, pre-labelling of living cells with isotopomeric forms of essential amino acids and protein array platforms. In addition, novel flowcytometry-based approaches are considered. These technologies can all be used to determine accurately the levels of proteins or biomarkers in a wide range of samples. Introduction The opportunities offered by genome sequencing have had an impact on biochemistry in that there is increasing expectation that systematic analysis of the protein content of the cell – namely, the proteome – will form an objective approach to studies of biological systems. Initially, it was thought that the conversion of genome data into rapid and effective proteome analyses would be immediately feasible. This possibility has been confounded by issues based around methodologies for protein identification and for determining relative protein levels in biological samples. These obstacles have been addressed by developing analyses, including quantification, for multiple proteins using various platforms. It is becoming increasingly clear that studying relative mRNA expression cannot provide details on all of the changes in components of cell systems. Specifically, mRNA changes do not relate directly to changes in protein levels, owing to different rates of translation, posttranslational modification, subcellular localization and degradation [1,2], which underlines the need for proteomic relative quantification techniques. In addition, the processes of posttranslational regulation of protein levels and activity, including events such as phosphorylation, glycosylation, acetylation and ubiquitination, all require quantification. Here we describe recent developments that have enabled detailed investigations of the proteome and posttranslational regulation. Corresponding author: Whetton, A.D. ([email protected]). Available online 11 July 2006 www.sciencedirect.com
Protein microarray technologies Protein microarrays facilitate the parallel analysis of multiple proteins in a high-throughput, miniaturized format for global protein screening and are available in two forms: one that determines protein function, and one that provides relative quantification. Although such analyses present considerable technical challenges, important advances have been made since the first proteome array was described [3]. Microarrays are currently generated by immobilizing multiple biomolecules onto solid supports. A high specificity of interaction between the immobilized binding molecule and the analyte is required to minimize cross-reactivity with other analytes and to decrease background signal. Functional microarrays Function-based microarrays can be used to analyse biochemical properties such as enzyme activity and substrate specificity [4] (Figure 1). For example, in vitro substrates for 87 different yeast protein kinases have been identified with chips spotted with >4000 proteins and the data used to generate a map of protein phosphorylation [5]. Protein–protein interactions can also be quantified with these arrays. An example of this approach has been the use of protein microarrays containing immobilized phosphotyrosine-binding domains of the ErbB receptor family: the interaction of each domain with 61 peptides containing the physiological sites of tyrosine phosphorylation from four ErbB receptors facilitated identification of complex protein interaction networks [6]. Applications can be performed in parallel – for example, protein microarrays comprising allelic variants of the p53 oncoprotein have been assayed for their ability to bind DNA and the MDM2 regulatory protein and to act as potential substrates for casein kinase II [7]. Chemical probes for activity-based protein profiling (ABPP) that report on active-site availability have been developed for the quantification of enzyme activity. ABPP complements functional arrays that assign enzyme function and substrate specificity (reviewed by Cravatt and Sorenson [8]). Probes for ABPP contain a reactive group directed to the active site, and one or more chemical tags, such as biotin or a fluorophore, for isolation and detection after one-dimensional or two-dimensional (2D) gel electrophoresis. Low-abundance proteins can be detected because ABPP is dependent on activity rather than expression. Applications include the identification of novel cancerassociated enzymes [9] and screening of enzyme inhibitors.
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Figure 1. Relative quantification using protein microarrays: functional arrays. These microarrays can be used for high-throughput analysis of protein function. (a) Enzyme– substrate interactions can be determined by arraying potential substrates, incubating with an enzyme, and then quantifying each spot for enzyme activity, for example, by the addition of a (radioactive) phosphate group to a bound peptide or protein to determine kinase activity. (b) To determine protein–protein interactions, peptides, protein domains or proteins are immobilized as discrete spots. Potential binding partners (comprising single or multiple molecules) are then incubated with the array. Here, binding partners are fluorescently tagged (indicated by a star) to facilitate the detection of specific interactions. This type of approach has several applications, including the identification of binding partners for motifs such as SH2 domains or basic-region leucine zipper domains. Such applications can be performed in parallel.
In terms of inhibitor screening, ABPP is particularly useful for enzymes lacking known substrates. ABPP technology has been combined with liquid chromatography MS to achieve greater coverage of the proteome [10] and could be combined with isotope-coded mass tagging for relative quantification. Protein profiling microarrays There are two main types of array for relative quantification by protein profiling: forward-phase (capture) arrays and reverse-phase arrays. These microarrays have found considerable application in cancer research [11,12], where they are used to identify biomarkers for diagnostics and to discover potential therapeutic targets. Forward-phase arrays. In forward-phase arrays, proteins for analysis are captured by an array of molecules immobilized either on a slide format (Figure 2) or on beads at an optimal concentration. Many different capture molecules can be present in the array, facilitating relative quantification of multiple proteins. Arrays are typically probed with cell lysate or serum, and protein binding is assessed by direct labelling via protein tagging or indirect detection (i.e. a ‘sandwich’ type approach in which captured proteins are detected by matched antibodies), enabling signal amplification [13]. Differential protein expression profiling has been used to compare serum proteins from normal individuals and cancer patients (reviewed by Hamelinck [14]) and to analyse discrete protein subsets such as secreted cytokines [15]. Relative quantification is achieved by comparing control or test samples with a reference sample (usually a pool of all samples in the analysis) using two-colour fluorescence labelling with Cy dyes. An adaptation of the www.sciencedirect.com
forward-phase approach facilitates analysis of both protein levels and phosphorylation status by using two different antibodies that bind a protein independent of and dependent on its phosphorylation status to determine the total amount of protein and the amount phosphorylated at a specific site, respectively [16,17]. Antibodies remain the best-characterized capture molecules. Not all antibodies function in this format, however, because activity is not always retained after immobilization. An alternative approach is the use of aptamers – that is, oligonucleotide sequences with molecular recognition properties selected from combinatorial oligonucleotide libraries [18]. Aptamers bind protein ligands with high affinity and specificity and, because they are chemically synthesized, they can be easily labelled with reporter groups. They also have advantages over antibodies of being reversibly denatured and usually more stable. Reverse-phase arrays. In reverse-phase arrays, multiple samples, which are often complex mixtures such cell lysates, are immobilized by deposition as microspots and probed with a specific detection reagent. Reverse-phase protein blotting facilitates simultaneous analysis of multiple samples for a single analyte in a western blot type approach (Figure 3). Antibodies that have been used include those against housekeeping proteins, signalling proteins, cell-cycle proteins, apoptosis-related proteins and phosphorylation motifs present in signalling molecules. A chief application of reverse-phase arrays has been in analyses of the phosphoprotein profiles of discrete protein subsets, whereby arrays are probed with two different antibodies to infer protein posttranslational status (and therefore often activity). One antibody is specific for a
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easily observed, however, lower abundance signalling molecules might not be detectable. Prefractionation of crude extracts could alleviate this problem. Protein array approaches are based on determining quantitative changes by using specific predetermined assays in which defined protein targets are selected in a subjective manner. Novel proteins or sites of posttranslational modification are not considered in this type of approach. Discovery-led proteomics offers an objective means of determining differences in protein expression without knowledge of the identities of the proteins to be assayed. This approach facilitates the identification of proteins in a non-hypothesis-driven manner and can be taken forward by several different methodologies.
Figure 2. Relative quantification using protein microarrays: forward-phase (capture) arrays. (a) Specific antibodies raised against proteins of interest are immobilized with a single capture antibody present in each spot. Many different capture molecules can be arrayed in this format for the simultaneous detection of multiple analytes. Recombinant antibody fragments, ‘affibodies’ (non-immunoglobulin-based affinity capture proteins) or aptamers (oligonucleotide sequences that bind epitopes with high affinity and selectivity) can be also used for forward capture arrays. Antibody arrays are used for protein quantification, protein binding studies and protein phosphorylation assays. (b) For relative quantification, comparison can be made to a reference sample (comprising a pool of all samples in the analysis) by using two-colour fluorescence labelling with Cy3 and Cy5 fluors, as shown here. Increased or decreased protein expression is identified from the ratio of the two dyes, analogous to DNA microarray analyses.
phosphorylated epitope and the second assesses total protein abundance. Reverse-phase protein arrays have been used to analyse signalling pathways with antibodies for specific phosphorylation sites in phosphoproteins, including MEK, MAPK, Raf and AKT kinases and the STAT family of cytoplasmic transcription factors [19]. A disadvantage of reverse-phase protein arrays is that only one analyte can be measured on a single array, although this can be addressed by printing arrays in a multisector format, thereby enabling parallel analysis of multiple analytes in the same experiment. Generally speaking, reverse-phase blots have the advantage that proteins do not require labelling. Although high-abundance proteins present in cell lysates can be www.sciencedirect.com
Two-dimensional PAGE Two-dimensional PAGE (2-DE) has been used for proteomic analysis for >30 years (reviewed by Gorg et al. [20]). Proteins are resolved on the basis of two independent parameters: overall electric charge and size (Figure 4a). In the first dimension, isoelectric focusing (IEF) separates proteins in a pH gradient gel on the basis of isoelectric point (pI). Proteins are then resolved orthogonally by molecular size using SDS–PAGE. Protein spots can be detected and quantified with chemical stains (reviewed by Patton [21]), which are selected on the basis of sensitivity (0.05–500 ng per spot), reproducibility of staining, and compatibility with downstream methods of protein characterization (Figure 4b). Coomassie blue stain and silver stain have been traditionally used, but fluorescent protein stains offer greater quantitative accuracy owing to their wider linear dynamic range (1000-fold) and greater compatibility with downstream protein characterization methods. Fluorescent stains specific for posttranslational modifications have been also developed [22]. One advantage of 2-DE over alternative gel-free peptide-based approaches is that, because protein modification can induce alterations in both mass and pI, different isoforms and posttranslationally modified protein species can be resolved. In addition, parallel profiles of phosphorylated and total proteins can be obtained by a multiplex proteomic approach [22] using sequential gel staining with Pro-QTM Diamond phosphoprotein stain, followed by a total protein stain such as Sypro Ruby (Figure 4c). The inclusion of Pro-QTM Emerald glycoprotein stain in the procedure facilitates analysis of glycosylated proteins in the same sample [23]. Relative quantification can be achieved by a computerbased comparison of gel images, which matches and compares protein spot intensities between gels. Inter-gel comparisons, however, rely on high levels of reproducibility in both separation and staining. Intra-gel comparisons based on fluorescent dyes with distinct spectral properties overcome this problem. In ‘difference in-gel electrophoresis, cyanine dyes are used to label protein samples fluorescently; these samples are then mixed and resolved on the same 2-DE gel. Proteins can be labelled on lysine (minimal dyes) [24] or, for higher sensitivity of detection, cysteine residues (saturation dyes) with Cy3 and Cy5 [25]. Fluorescently labelled proteins are imaged with different excitation and emission filters to detect the protein profiles for
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Figure 3. Relative quantification using protein microarrays: reverse-phase arrays. Mixtures of proteins from different samples, for example cell lysates, are spotted onto the microarrays and probed with antibodies targeted against specific proteins. A single target is analysed for each sample spotted onto the array. To assay protein phosphorylation, parallel arrays are probed with an antibody that is phosphorylation independent (left) and one that is phosphorylation dependent (right). The first antibody detects the protein of interest in both its nonphosphorylated and phosphorylated (P) states, thereby assessing its concentration in the sample; the second assays only the phosphorylated form. The antibodies can be fluorescently labelled for visualization, as shown here, and the amount of each protein can be relatively quantified for a set of samples. This type of array has been applied to parallel analyses of the site-specific phosphorylation of a range of phosphoproteins, including MEK, MAPK, Raf and AKT kinases and the STAT family of cytoplasmic transcription factors, and has been used to profile the activation status of cell signalling pathways in samples from individuals with cancer.
each sample (Figure 4d). The introduction of a third minimal dye (Cy2) has refined the method to include an internal standard for each gel, which is typically a mixture of the two samples being compared [26]. The key limitations of 2-DE are dynamic range and protein solubility. Abundant proteins, present at >10 000 copies per cell, can obscure proteins expressed at a low level (10–1000 copies per cell). Prefractionation strategies alleviate this difficulty (see later). For 2-DE approaches, using narrow-range pH gradients (‘zoom’ gels) increases resolution and protein loading [27]. IEF remains problematic owing to its incompatibility with poorly soluble proteins such as membrane proteins (hydrophobic) and very basic proteins. This problem can be partially overcome by using non-ionic and zwitterionic detergents with different solubilization properties [28] and non-equilibrium pH gradients. Although 2-DE remains a useful means of globally comparing proteins from several samples, the issues of sensitivity, protein exclusion and reproducibility abound. Furthermore, when a difference in staining intensity is shown (Figure 4), it remains necessary to identify the protein by mass spectrometry (MS). Role of MS in relative quantification proteomics The development of more sensitive and efficient mass spectrometers has brought MS to the forefront of proteomics research. Most mass spectrometers used for relative quantification work operate on broadly similar principles. First, the peptide sample is ionized, either in solution phase by spraying from a charged needle (electrospray ionization) or from solid phase by using a laser to transfer a proton from an energy-absorbing matrix to the peptide (matrix-assisted laser desorption–ionization). Once inside the instrument, peptide masses are determined by ion time of flight. Although this is a common type of mass analyser, www.sciencedirect.com
other types such as Fourier-transform ion cyclotron resonance [29], ion traps [29] and Orbitraps [30] can be used. The masses determined for the peptides can be used to identify proteins (Box 1). For complex mixtures of peptides, a liquid chromatography step before MS enables the samples to be fractionated and peptides to be analysed sequentially. Combining two types of chromatographic separation enables further fractionation of complex mixtures, thereby increasing both the number of peptides identified and coverage of the proteome [31]. Relative quantification by MS Although MS provides sensitive detection and identification of proteins and peptides, it is not directly quantitative. Two peptides from the same protein will not necessarily generate the same signal because differences in amino acid sequence will affect the efficiency with which they ionize. Indeed, often the same peptide will not produce exactly the same signal when analysed a second time, perhaps from a different sample, owing to slight differences such as coeluting peptides, solvent composition, and the age, condition and position of the electrospray needle. To overcome this problem, several methods have been developed. Chemically identical but isotopically distinct tags can be used to modify the peptide populations being compared. These peptides are then pooled, and analysed in a single MS run. Because the tags are chemically identical, the same peptides from each sample will behave identically; that is, they will elute from a chromatography system at the same time, and be ionized at the same time and therefore with the same efficiency. Relative quantification is obtained by comparing the signal intensities of peptides with a defined, isotope-induced mass difference (Figure 5a). Several methods for incorporating ‘stable isotope’ have been described for relative quantification (Box 2).
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Box 1. Protein identification by MS For identification of a single protein, such as a spot from a 2-DE gel, digestion of a protein with a specific enzyme produces a predictable, unique profile of peptide fragments, termed a ‘peptide mass fingerprint’. Measurement of these masses facilitates data comparison with theoretical peptide mass fingerprints derived in silico for all proteins in a selected database. This method is usually faster for the identification of large numbers of samples, because few spectra have to be generated per sample. However, the confidence associated with protein identification by peptide mass fingerprinting is lower than that associated with identification based on peptide sequence information.
Higher confidence peptide sequence information can be derived by MS/MS. In this technique, a specific peptide ion is selected by filtering out all other ions using a quadrupole. This peptide is induced to fragment by collision with an inert gas, such as nitrogen or argon. Because fragmentation occurs along the peptide backbone, the masses of the fragment ions indicate the peptide sequence. Figure I shows fragmentation of a peptide from bovine serum albumin, AEFVEVTK, together with its major fragment ions and the amino acid sequence to which these ions correspond.
Figure I. Peptide sequencing by MS. A tandem mass spectrometer consisting of a quadrupole and a time-of-flight analyser is shown. Peptide ions are generated by electrospray ionization from a fine needle. On entering the mass spectrometer, these ions are filtered on the basis of their mass-to-charge (m/z) ratio in the quadrupole. A peptide ion selected on the basis of its m/z ratio undergoes collisional induced decomposition after colliding with an inert gas. The fragment ions are then ‘pushed’ into the time-of-flight mass analyser in a synchronous fashion. There, peptide mass is inferred by measuring the time taken for a peptide ion fragment to travel from the source to the detector. The signal (y-axis of each spectrum) represents the number of ions that have hit the detector at a specific m/z ratio (x-axis). The pattern of ion masses is then used to infer peptide sequence information; in this example, the peptide is identified as one from bovine serum albumin. All labelled ions are singly charged. For clarity, not all ions have been labelled; these unlabelled ions might be multiply charged, might be fragments from the N-terminal half of the peptide, or might be derived from multiple fragmentation events on the same peptide.
Other relative quantification approaches have been subsequently developed, including ‘isobaric tags for relative and absolute quantification’ (iTRAQ) [32]. This method can incorporate up to four mass tags that label the N terminus of every peptide (along with free amines in lysine side chains). Each tag is isobaric (i.e. has the same mass) such that, when the four labelled samples are pooled, the same peptides from each sample appear at the same mass in the MS spectrum. Because the distribution of isotopes in each tag is different, however, on fragmentation each tag generates a specific distinct reporter ion at mass:charge ratio 114, 115, 116 or 117. The relative abundances of these reporter ions provide relative quantification (Figure 5b). iTRAQ offers some advantages. First, because all peptides are labelled it is possible to obtain data on more peptides as compared with other methods, thereby increasing confidence in the quantification of each protein [33]. Second, because the tags are isobaric, the signal from up to four samples is summed in MS mode, providing an increase www.sciencedirect.com
in sensitivity. For iTRAQ, tandem mass spectrometry (MS/MS) must be carried out on all peptides to obtain quantification, whereas for ICATs or SILAC (Box 2) MS/ MS is carried out only on proteins that are differentially expressed. Although this approach could be viewed as ‘wasting analysis time’, the definition of a large population of unchanging proteins enables the use of normalization procedures found in microarray analyses and comparison to such data sets [2]. When samples are labelled at the peptide level as in iTRAQ, it is important that the prefractionation methods (e.g. immunoprecipitation) applied are highly robust and reproducible to maintain the reliability of relative quantification. With ICATs or SILAC (Box 2), by contrast, proteins can be pooled and then prefractionated and digested together, thereby reducing potential sources of experimental variation. A key aspect of all stable isotope labelling procedures is that the efficiency must approach 100%. Underlabelling leads to the generation of background ‘noise’ because the
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Figure 4. Two-dimensional gel-based analysis of the proteome. (a) Principle of 2-DE. Proteins are resolved according to charge and mass. In the first dimension, proteins are separated on the basis of their net charge, which is the sum of the negative and positive charges conferred by the amino acid side chains and the N and C termini, and is pH dependent. The isoelectric point (pI) is the pH at which the net charge of the protein is zero. During IEF, the proteins migrate in a pH gradient to the specific pH corresponding to their pI. Proteins are solubilized and denatured before IEF to expose internal ionizing groups. In the second dimension, proteins are separated by molecular size using SDS–PAGE. For detection, there are several staining protocols that detect either all proteins or specific posttranslational modifications and that can be used for relative quantification. (b) Representative 2-DE data. Replicate 2-DE gels of samples for comparison (‘Control’ and ‘Test’) are run in parallel. Relative quantification is performed by gel image analysis using specialist software that performs spot detection and spot matching across data sets. Comparison is often based on normalized spot volume, which is defined as the volume of a particular spot relative to the total volume of all spots present in the gel. A representative set of histograms is shown for the normalized spot volumes of a selected spot difference across a gel set. (c) Multiplexed proteomic analysis using fluorescent stains. This method facilitates parallel profiling of protein expression and posttranslational modification in a single 2-DE gel. Protein gels are run and stained with a posttranslational-modification-specific stain and the gels are imaged. Gels are then stained for total protein expression with Sypro Ruby and imaged again. The resulting images are then matched by overlaying. Shown here is a representative sequence of protein spots that are a mixture of phosphorylated and nonphosphorylated proteins. Protein spots containing phosphate groups are detected by Pro-Q Diamond staining, whereas Sypro Ruby detects all protein spots. The phosphorylated proteins migrate to a more acidic position in the gel relative to the parent protein, owing to the addition of negative charge to the protein. Multiple samples can be analysed by inter-gel comparison of the staining profiles. (d) Difference in gel electrophoresis. Proteins from two different samples are each labelled with a single fluorophore that has distinct spectral properties (here, Cy3 and Cy5). Samples are then combined and resolved by 2-DE gel electrophoresis on a single gel for intra-gel comparison. The labelled proteins are detected by using different excitation and emission filters, and two separate images are generated for comparison. An additional sample, typically a mixture of the samples to be compared, can also be included (labelled with Cy2) to provide an internal standard. Differences in protein expression are determined by gel image comparison using specialist software that assigns protein spot differences on a ratiometric basis.
same protein might be labelled to various extents in different experimental samples. This labelling efficiency requires several population doublings for labelling in culture, or the development of efficient chemical reactions. Several groups have recently described a label-free relative quantification approach, termed ‘differential MS’, which is based on normalization strategies that enable comparison of the signal strength of the same peptide in two distinct sample runs [34] (Figure 5c). The advantages of this approach are that labelling is not required and that, provided both MS and chromatography remain relatively constant over time, a large number of samples can be analysed [34,35]. www.sciencedirect.com
Improving sensitivity: subproteome analysis A chief difficulty in defining and quantifying a proteome is sample complexity. Fractionating organelles or isolating a protein set with a specific property (e.g. tyrosinephosphorylated proteins) provides increased penetration of the proteome. The most common approaches to organelle fractionation are based on centrifugation and differential solubility, or on antibody-directed methods (reviewed by Yates et al. [36]). Although these approaches have existed for several years, the impact that they have on state-of-the-art proteomics is substantial. Good fractionation strategies enhance penetration: for example, analysis of nucleoli
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Box 2. Common methods of stable isotope labeling Isotope coded affinity tags Isotope coded affinity tags (ICATs) are chemical modifiers that covalently bind cysteine residues. Thus, ICATs can be labelled with either a heavy or light isotope and contain a biotin tag for affinity purification. Initially, the mass difference between the tags was generated with deuterium [67], but this method has since been improved and the ‘heavy’ tag now contains nine 13C residues. Each tag contains a cleavable linker attached to a biotin moiety so that labelled peptides can be purified; the biotin tag is then removed to generate a mass addition of 227 Da or 236 Da for the light or heavy tag, respectively [68]. At present, however, only two samples can be compared by this method and only cysteine-containing peptides are enriched. Stable isotope labelling in culture In stable isotope labelling in culture (SILAC), cells are cultured in medium containing a ‘heavy’ essential amino acid [69] e.g. [d3]leucine, [13C6]arginine or [13C6]lysine for one sample of a pair to be compared. Combining these isotopic amino acids enables up to three samples to be compared [70]. This approach is suitable only for material obtained
has yielded relative quantification data on 489 proteins [37], and analysis of Saccharomyces cerevisiae mitochondria has identified 750 proteins [38]. Other organelles from higher eukaryotes that have been specifically enriched for proteomic analysis include the Golgi apparatus (421 proteins) [39], phagosomes (up to 520 proteins) [40], lysosomes (215 proteins) [41], lipid rafts (241 proteins) [42] and metaphase chromosomes (102 proteins) [43]. In principle, it is relatively easy to adapt these protocols to include relative quantification by using the strategies described in Box 2 and Figure 5. Cell-surface proteins can be analysed specifically after protein tagging with biotin reagents, which are hydrophilic and cannot cross the hydrophobic plasma membrane. Biotin-tagged proteins can be enriched with streptavidin beads, which bind the biotin moiety. The recovery of integral membrane proteins is improved by removing plasmamembrane-associated cytoskeletal proteins with high salt and alkaline washes. On the basis of this approach, MS analysis has identified 781 mammalian plasma membrane proteins [44]. These data show that it is increasingly feasible to identify and to quantify relatively many proteins from a given cell system if sufficient fractionation is performed. In most types of cell, proteomics technologies make the relative quantification and localization of thousands of proteins an achievable aim. Improving sensitivity: posttranslationally modified proteins As an alternative to subcellular fractionation, other protein subsets such as glycosylated [45], ubiquitinated [46] and sumoylated [47] proteins (the latter two representing the ‘degradome’) can be isolated. So far, however, the best-studied modification is phosphorylation [48]. Isolation of phosphoproteins can be achieved by several methods. At present, antibody-based isolation is normally targeted at phosphotyrosine. Using this approach coupled with SILAC (Box 2), Kratchmarova et al. [49] achieved relative quantification of 282 tyrosine phosphorylated proteins. iTRAQ relative quantification has also been www.sciencedirect.com
by cell culture (at least five population doublings are required for isotopic equilibrium to be reached) and cannot be applied to primary material. 18 O incorporation Tryptic hydrolysis of peptides requires H2O. By performing the digestion in ‘heavy’ water (H218O), all tryptic peptides gain C-terminal 18 O, generating an apparent mass shift of 4 Da owing to incorporation of the isotope during peptide cleavage [71]. 13
C/15N guanidination Modification of the N-terminal lysine with methylisourea to form homoarginine was initially reported as a strategy to improve the ionization and/or transmission of lysine-containing peptides. By using a 15N-containing version of the reagent, a mass difference of 3 Da can be introduced [72]. This method is a cheap, efficient and simple way in which to introduce a stable isotope, but data are hard to interpret because often the natural isotopic ‘envelope’ or mass range of the differentially labelled peptides will overlap – a factor that must be corrected for before relative abundance can be calculated.
successfully used alongside phosphotyrosine immunoprecipitation [50]. An alternative method is to modify phosphorylation sites chemically to enable purification of phosphoserine- and phosphothreonine-containing proteins; quantification can be achieved by including a stable isotope tag [51]. Phosphoprotein enrichment can also be achieved with immobilized metal affinity chromatography. Although in most cases these enrichments have been combined with 2-DE for protein analysis [52], it is possible to use gel-free MS methods to gain even greater penetration into the phosphoproteome. Another common approach to assess phosphorylation is the enrichment of phosphopeptides. This approach has several advantages, including simplification of the analyte mixture and the ability to obtain both information about the site of the modification and relative quantification. Often, however, it does not reveal whether any difference is due to changing phosphorylation stoichiometry or overexpression of a constitutively phosphorylated protein. Nevertheless, by using immobilized metal affinity chromatography for enrichment of phosphopeptides [53] and sufficient starting material, it has been possible to identify >700 phosphopeptides in yeast [54] and 2000 phosphorylation sites from >900 proteins in mammalian cells [55]. Other methods for phosphopeptide enrichment such as titanium oxide [56], anti-phosphotyrosine antibodies [57] and diagonal chromatography [58] have been also used. All offer opportunities for increased proteomic penetration to identify posttranslational modifications crucial to various cellular processes. Absolute quantification Relative quantification is sometimes insufficient for the biochemical process or issue to be addressed fully. Several strategies now enable absolute protein quantification by MS. The first such method, called AQUA (absolute quantification) [59], uses a stable isotope-labelled synthetic peptide in a protocol similar to the stable isotope methods used for relative quantification (Box 2; Figure 5). This peptide is spiked into the analyte mixture at a known concentration, and the ratio of the test peptide to the
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Figure 6. The multiple reaction monitoring (MRM) technique. (a) Specific peptide detection by MRM. Triple-quadrupole mass spectrometers can be configured to enable only peptides of a specified m/z value into the collision cell, and a second mass filter to enable only a specific diagnostic ion to reach the detector. Several of these ‘transitions’, each performed in a short (millisecond) timescale, can be analysed in an experiment – a method known as ‘MRM’. The filtering vastly reduces background and therefore dramatically increases the signal to noise. Here, the first quadrupole of a mass spectrometer is set to detect a peptide ion at m/z 575.7 from a mixture of peptides delivered to the mass spectrometer. Ions of this mass are fragmented by collisional induced dissociation. If a diagnostic fragment ion with an m/z of 529.7 is generated, this ion passes to the detector. Peptides of m/z 575.7, which do not generate this product ion, do not produce a signal, demonstrating the selectivity of this analysis. (b) Absolute quantification by MRM. The inclusion of a standard peptide labelled with a stable isotope enables MRM transitions to be set up for both the standard and test peptides. Because the sample and standard peptides have different masses, they can be monitored in parallel by using different MRM transitions; because they are chemically identical, however, their response in the mass spectrometer should be the same. Therefore, comparison of the total signal generated by each peptide facilitates absolute quantification of the peptide and thus the protein of interest. Here, the signal of the sample peptide is 65% of that of the standard.
Figure 5. Strategies for relative quantification of peptides by MS. (a) Stable isotope labelling can be used for relative quantification. Various methods exist for incorporating the ‘heavy’ and ‘light’ isotopes (Box 2). Although being chemically identical, isotopes enable peptides from each sample to be discriminated and relatively quantified in the mass spectrometer by a comparison of the intensities of peaks that elute from a liquid chromatography system at the same time. Isotopomeric peptide ions from two samples have a defined mass difference but coelute. Here, the peptide at m/z 686.8 and its heavy counterpart at m/z 691 have equivalent intensity, whereas the ion at m/z 708.8 has higher intensity than its heavy counterpart at m/z 712.8. The ion at m/z 708.8 is thus selected for fragmentation to obtain sequence information and to show that this protein, radixin, is differentially expressed in samples A and B. (b) iTRAQ tags have identical overall mass and can react with all peptides via free amine groups. Here, each chemically identical peptide from the four samples appears at the same mass in an MS experiment. Colour coding in the stacked bars represents the fact that the isobarically tagged peptide has come from four different samples. In MS mode, there is no means of distinguishing from which of the four samples the peptides were derived. On fragmentation, however, specific reporter ions are released that differ in mass owing to differential isotope usage (this mass difference is offset in the intact tag by isotopic differences in a ‘balance’ group moiety). The ratios between these ions are thus representative of the relative expression of that peptide or protein. Here, fragment ions (bottom spectrum) clearly identify the peptide as one from a-internexin, an intermediate filament protein, whereas the reporter ions (top spectrum) show that this peptide is present at similar levels in sample A, B and C, but is more abundant in sample D. (c) Label-free relative quantification involves analysing samples in separate experimental runs in series. Peptides are separated by liquid chromatography and analysed by MS. A peptide-specific elution profile is then generated, which shows how much of the peptide (m/z 632.2) was detected throughout the experimental run. The area under this profile is then compared between two (or more) experimental runs to determine the relative quantity of that peptide. There is an assumption (which requires formal proof for each experimental system used in this way) that the ion current associated with a particular peptide is proportional to the molar quantity of the peptide present in each run in the sample series. This assumption is partly dealt with by complex data normalization to correct for inter-run variation owing to changes in temperature, sample loading and ionization efficiency. MS/MS is performed on every peptide in each sample to obtain data for inferred peptide sequence; here, it shows that sample B (blue) contains more glycogen phosphorylase than sample A (red). www.sciencedirect.com
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Figure 7. Analysis of cell-surface markers by inductively coupled plasma MS (ICP-MS). Cells express antigens on their surface that define their phenotype and that can be recognized specifically by antibodies. Tagging these antibodies with elements facilitates their detection by MS. Examples of elemental tags include the rare earth metals, europium (Eu), terbium (Tb) and samarium (Sm), as shown here. Cells are atomized and ionized on introduction into the inductively coupled plasma, and multiple elemental tags can be quantified simultaneously. The presence and intensity of the mass signals correlate to the relative expression of the specific cell markers. ICP-MS offers advantages over flow cytometry for multiplex analysis because, unlike fluorophores, the elemental tags produce signals that are essentially non-overlapping and a greater number of signals can be quantified in parallel [66].
‘heavy’ labelled standard is used to calculate the concentration of the protein. By using a synthetic ‘posttranslationally modified’ peptide, the absolute amounts of modified (phosphorylated) peptide can also be calculated, along with the total amount of that protein, giving information regarding modification stoichiometry. The QCAT method [60] uses a synthetic gene encoding a hybrid of several tryptic peptides from proteins of interest that can be isotopically labelled. Because all standard peptides are at identical concentrations, the simultaneous absolute quantification of many proteins becomes a realistic prospect. For complex mixtures, Anderson et al. [61] designed the ‘stable isotope standards and capture by anti-peptide antibodies (SISCAPA) workflow, in which standard labelled peptides are spiked into the analyte, and peptides of interest are enriched using antibodies. This approach can be improved by multiple reaction monitoring – a highly selective and exquisitely sensitive method for detecting a specific ion (peptide) in a sample (Figure 6a). Multiple reaction monitoring can be used to quantify the peptide and its labelled standard (Figure 6b) and is both more sensitive and linear over a greater range than is seen in time-of-flight MS [62].
can be used to discriminate selectively and to purify cell populations on the basis of their expression of characteristic cell-surface antigens. Cells can be fixed, permeabilized and labelled with antibodies directed against intracellular proteins of interest, such as phosphoproteins and apoptosis-related proteins [63,64]. In this way, defined cell populations, and even single cells, can be analysed in the context of complex cell populations for the presence of proteins of interest in a multiplex approach [65]. A flow-cytometry-based approach and MS have now been coupled. Specific cell marker proteins can be affinity labelled by using antibodies (or potentially aptamers) containing metal-containing labels that provide elemental tags [66]. Cells are analysed by inductively coupled plasma MS (ICP-MS) to determine elemental composition. This approach has been developed for the simultaneous identification of multiple cellular antigens using four diagnostic elemental tags (Au, Sm, Eu and Tb) coupled to secondary antibodies. It has the potential for higher throughput than can be achieved with current flow cytometry approaches. Potential applications include identification and analysis of low-abundance cancer stem cells in tumour samples (Figure 7).
Flow-cytometry-based relative quantification Flow cytometry measures physical and chemical characteristics of cells or particles in suspension. It is used routinely to analyse cell populations labelled with fluorescent antibodies directed against antigens of interest. When combined with fluorescence-activated cell sorting, it
Concluding remarks Many novel technologies are currently being applied to the multiplex-based assay of relative protein quantities. These technologies afford new opportunities to the biochemist and can be applied subjectively (antibody-based arrays) or objectively (MS-based approaches) to offer improved
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sensitivity and scope for the identification of large numbers of alterations in the proteome in response to a specific state or stimulus. The advent of more approaches as new techniques are developed is undoubted. In addition, improvements in MS instrumentation in terms of sensitivity and speed, together with better sample preparation and purification procedures will increase penetration into the proteome. The development of more complex and wide-ranging protein array technologies will soon make the simultaneous quantification of thousands of proteins, protein modifications and protein activities a realistic goal. The opportunities for systematic investigation based on these approaches abound and offer new horizons for quantification in biological systems. Acknowledgements Work in the authors’ laboratory is supported by the Leukaemia Research Fund, UK, and the Biotechnology and Biological Sciences Research Council.
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