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The role of mass spectrometry in proteome studies
Biomolecular Engineering 18 (2001) 221– 227 www.elsevier.com/locate/geneanabioeng
Review
The role of mass spectrometry in proteome studies Tuula A. Nyman * Turku Centre for Biotechnology, BioCity, P.O. Box 123, FIN-20521 Turku, Finland
Abstract Mass spectrometry (MS) is an important tool in modern protein chemistry. In proteome analyses the expression of hundreds or thousands of proteins can be monitored at the same time. First, complex protein mixtures are separated by two-dimensional gel electrophoresis (2-DE) and then individual proteins are identified by using MS followed by database searches. Recent developments in this field have made it possible to do automated, high-throughput protein identification that is needed in proteome analyses. MS can also be used to characterize post-translational modifications in proteins and to study protein complexes. This review will introduce the current MS methods used in proteome studies, and discuss their advantages and disadvantages. New instrumental MS developments are also presented that are useful in these analyses. © 2001 Elsevier Science B.V. All rights reserved. Keywords: Mass spectrometry (MS); Two-dimensional gel electrophoresis (2-DE); Proteome analyses; MALDI-TOF
1. Introduction Proteomics is the study of total protein complements, proteomes, e.g. from a given tissue or cell type. Proteome analyses are usually carried out by using two-dimensional gel electrophoresis (2-DE) followed by protein identification by mass spectrometry (MS) and database searches. 2-DE dates back to 1970s [1], but protein identification and characterization by mass spectrometry has become possible only during the last decade. The most important reasons for this are the new soft ionization methods, matrix-assisted laser-desorption ionization (MALDI) [2] and electrospray ionization (ESI) [3,4], together with developments in sample preparation techniques and MS instruments. Equally crucial for protein identification by MS is also the exponentially growing amount of sequence information available in databases. Compared to transcriptomics, where the DNA-microarray technology is a completely automated method, proteome analysis requires a lot of manual work and suffers from problems mainly associated with 2-DE * Tel.: +358-2-333-8027; fax: +358-2-333-8000. E-mail address: [email protected] (T.A. Nyman).
[5,6]. Despite the difficulties the importance of proteomics is well recognized. DNA-microarray technology is based on measuring steady-state mRNA levels whereas proteomics studies proteins, which are the active agents in cells. Recent studies have also shown that mRNA expression and protein levels do not always correlate [7,8]. Proteins can be post-translationally modified or present as different isoforms, and these modifications cannot be predicted from the corresponding DNA-sequences. Mass spectrometry has an important role in characterization of these modifications.
2. Protein identification by mass spectrometry Mass spectrometers consist of the ion source, mass analyzer, ion detector and data acquisition unit. First, molecules are ionized in the ion source, then they are separated according to their mass-to-charge ratio in the mass analyzer and the separated ions are detected. Mass spectrometry has become a widely used method in protein analysis after the invention of MALDI and electrospray ionization methods (Fig. 1). There are several options for mass analyzer, the most common combinations being time-of-flight (TOF) connected to
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MALDI and triple quadrupole, quadrupole-TOF or ion trap mass analyzer coupled to ESI. In proteome analysis electrophoretically separated proteins can be identified by mass spectrometry with two different approaches. The simplest way is a technique called peptide mass fingerprinting (PMF) [9– 13]. In this approach the protein spot of interest is in-gel digested with a specific enzyme, the resulting peptides are extracted from the gel and the molecular weights of these peptides are measured. Database search programs can create theoretical PMFs for all the proteins in the database, and compare them to the obtained one (Fig. 2). In the second approach peptides after in-gel digestion are fragmented in the mass spectrometer yielding partial amino acid sequences from the peptides (sequence tags), and database searches are performed using both molecular weight and sequence information [14,15]. PMF is usually carried out with MALDI-TOF, and sequence tags by nano-ESI tandem mass spectrometry (MS/MS) [16– 18]. The sensitivity of protein identification by MS is practically in the femtomole range, even though sensitivities at the attomole level have been reported [19].
3. Peptide mass fingerprinting by MALDI-TOF Protein identification by in-gel digestion and peptide mass fingerprinting is nowadays a routine method in most MS laboratories. This method requires optimized in-gel digestion [20– 22] and desalting protocols before MALDI analysis. Trypsin is the most often used enzyme for in-gel digestion, since it cleaves proteins very specifically after lysines and arginines. It also produces quite small peptides, typically in the range of 600– 2500 Da, which can be eluted efficiently from the gel. MALDI-TOF is a relatively simple and easy-to-use form of mass spectrometry, and it is more tolerant to salts and other contaminants present in the sample than ESI. Small aliquot of the extracted peptide mixture can be directly deposited to the MALDI target plate for PMF, and the rest of the sample can be stored for later analysis by ESI-MS. If the total peptide sample after in-gel digestion is used in MALDI analyses, best results are obtained when the samples are desalted. Desalting can be performed with commercially available ZipTips (Millipore) or by making a small reversed-phase material column into a gel-loader tip [23,24]. Mass accuracy of MS measurements is especially important in PMF [25]. Modern MALDI-TOF instruments are delayed extraction reflector instruments, and the mass accuracy obtained with them is in the range of 10–30 ppm for peptides with internal calibration. With this kind of mass accuracy usually 4– 5 peptides are enough to identify a protein unambiguously. MALDI produces mostly singly charged ions, so the spectra are
easy to interpret and since the sample is in crystallized form, analysis time is seldom a limiting factor.
4. Automation of peptide mass fingerprinting For high-throughput protein identification it is necessary to automate sample preparation together with MS measurements, data interpretation and database searches. There are already robots available for spot cutting, in-gel digestion, desalting of the samples and spotting for MALDI plates. In addition, MALDI measurements can be automated with modern MALDITOF-instruments. Data processing and database searching can also be automated, but in practice when very small amounts of protein digests are analyzed, the spectra have to be checked manually before database searches. In proteome analyses it would be tempting to identify all the proteins in the gel without cutting individual spots from the gel. There have been several attempts to do this, the most recent being called a molecular scanner [26,27]. In this approach the proteins in the gel are simultaneously proteolytically digested and electrotransferred onto a polyvinylidene difluoride membrane, and the membrane is directly scanned by MALDITOF. As promising as the molecular scanner is, serious problems still arise from the time needed to scan the whole membrane (for 16× 16 cm2 membrane 36 days
Fig. 1. The principles of: (A) MALDI; and (B) electrospray ionization. (A) The sample is mixed with a UV-absorbing matrix and crystallized to the MALDI target plate. The matrix absorbs energy from the laser causing the matrix and sample to be volatilized; (B) In ESI ions are formed from a liquid sample by applying a potential which causes the sample to spray and produce charged droplets. These droplets will form usually multiply charged ions upon solvent evaporation.
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quence information are used compared to searching with peptide molecular weights only. With sequence tag information also EST-databases can be used in database searches [28]. Sequence tag from one peptide is usually sufficient to identify the protein. Compared to MALDI measurements, nano-ESI-MS/MS is time consuming and laborous, and samples have to be desalted prior ESI-analysis. The analysis time is also limited since the liquid sample is consumed all the time during the MS-measurement. The problems of nano-ESI can be solved to some extent by coupling HPLC with the ESI-MS. ESI-analysis is concentration sensitive, and the use of modern micro- and nano-HPLC-methods provides high local peptide concentrations, and the total LC-separated sample can be introduced to the mass spectrometer without splitting. LC-separation before mass spectrometry is especially useful when complex mixtures are analyzed, because it is possible to do data-dependent experiments (DDE). In DDE all the ions coming into the detector are measured, and when there is a peak coming from the HPLC to the MS (i.e. an increase in the total ion current), the software will switch automat-
Fig. 2. Peptide mass measured is critical in calibrated. If trypsin cleave also itself into
fingerprinting. Mass accuracy of the peptides PMF, therefore the spectra has to internally be is used for in-gel digestion, the enzyme will fragments, which can be used for calibration.
of continuos scanning and more than 40 Gb of raw data).
5. Creating sequence tags by ESI-MS/MS Peptide mass fingerprinting can only be successful when the digested protein exists in the protein or genomic databases, and when four or more peptides are obtained in MALDI analysis. If this is not the case, or if the protein is only in EST-databases as partial sequences, it is necessary to obtain sequence information from the peptides. The most common way to do this is by ESI-MS/MS (Fig. 3). The advantage of creating sequence tags is that database searches are much more specific when both peptide molecular weight and se-
Fig. 3. Product ion scan. First, quadrupoles are set to pass through the whole peptide mixture, and this gives the molecular masses of the peptides present. In ESI, peptides are usually doubly or triply charged. One of the peptides, parent ion (seen in the inset), can then be selected by letting only this peptide to pass through Q1. It is then fragmented in the collision chamber, Q2, and the resulting fragments are detected in the last quadrupole (OR TOF). Partial amino acid sequence can then be read from the spectra.
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ically from MS to MS/MS mode and do a product ion scan. In this way it is possible to generate much more MS/MS data from one sample than in nano-ESI experiments. Yates et al. have reviewed the strategies for automated protein identification using microcolumn LC-MS/MS [29].
6. Recent developments in MS instruments The first step for protein identification by MS is usually PMF with MALDI-TOF, and if this method fails to identify the protein, the next step is to create sequence tags by ESI-MS/MS. In PMF only small part of the sample on the MALDI plate is consumed during the analysis, so there is sample left to create sequence tags from it also. It is possible to partially sequence peptides with a MALDI-TOF instrument with a method called ‘post source decay’. However, this method produces spectra, that are difficult to interpret, and the mass accuracy of the measured peptide fragments is poor. Recently, MALDI with a hybrid quadrupole-TOF mass analyzer [30,31] and a prototype of MALDI-TOF/TOF instrument [32] were introduced. Thus it is possible to do PMF as the first choice of protein identification and sequence tags by MS/MS when necessary from the same sample. MALDI-hybrid quadrupole-TOF has shown great promise for highthroughput protein identification [31]. Prototype of rapidly switchable MALDI/ESI-ion source for a hybrid quadrupole-TOF has also been introduced [33]. This improves protein identification strategies since it is possible to measure MS spectra with both ionization modes and combine MS/MS information from singly and multiply charged ions.
7. Post-translational modification analyses in proteome studies One of the key advantages of doing proteome analysis is that both protein expression levels and post-translational modifications, like phosphorylation and glycosylation, can be studied. Protein phosphorylation analyses are especially interesting since phosphorylation plays major role in e.g. many signal transduction pathways. The easiest way to detect differencies in protein expression levels is by metabolically labeling the samples with 35S-Met before 2-DE followed by autoradiography of the 2-DE gel [34–36]. Different protein phosphoand glycoforms usually apprear as a ‘train’ of spots in these gels, but more detailed analysis is needed to determine the type of modification. Changes in protein phosphorylation can be studied either by labeling cells with 32P or by western blotting with phospho-specific
antibodies. If the 32P-approach is used the 2-DE gels are run as usually, and after gel comparison the spots of interest can be cut out from the gel and identified by mass spectrometry [36]. Soskic et al. [37] have used the blotting method where two similar 2-DE gels were run in parallel, one was used for blotting with phospho-specific antibodies and the other one for normal staining and cutting spots for mass spectrometry. Glycoprotein detection from 2-DE gels can be done with similar type of blotting experiments [38,39]. More detailed characterization of protein phosphorylation and glycosylation can be done by mass spectrometry but this requires more starting material than just a spot in 2-DE gel. In addition, these analyses are not as straightforward methodologically and are technically more difficult compared to protein identification by MS. There are several ways to locate the phosphorylation sites in a protein. To find out which peptides are phosphorylated in the digestion mixture, one can perform PMF by MALDI-MS before and after phosphatase treatment: phosphopeptides will lose their phosphate group and thus their molecular weight will be 80 Da lower after phosphatase treatment. More detailed characterization can be done with nano-ESItriple quadrupole instrument with parent ion scans: peptides are fragmented like in product ion scan (Fig. 3), but in this case Q3 is set to pass only the fragmented PO− 3 moiety, m/z 79, and Q1 is scanned to find the parent ion for PO− 3 moiety. When the phosphopeptides are found they can be sequenced by normal product ion scans. One additional possibility is to isolate phosphopeptides with immobilized metal ion affinity chromatography (IMAC) and analyze only this fraction by MS [40,41]. The invention of MALDI-hybrid quadrupole-TOF will probably help protein phosphorylation studies: the peptide sample can be analyzed before and after phosphatase treatment, and the peptides which lose their phosphate group can be directly sequenced from the MALDI sample. Protein phosphorylation studies by MS, possibilities and limitations, have been recently reviewed by Neubauer and Mann [42]. In glycosylation studies the glycans are liberated from the protein, and the glycan structures are determined from the free glycans. The glycan structures can be characterized by combining MALDI-measurements to exoglycosidase digestions [43–46] or if more detailed structural information is wanted by ESI-MS with product ions scans [46–48]. So far there have been very limited amount of reports where glycan structures have been characterized from electrophoretically separated proteins; in one study the N-linked glycans were enzymatically released from the protein in one-dimensional SDS-PAGE gel followed by structural analysis with MALDI-MS and exoglycosidase digestions [49].
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8. Protein–protein interaction studies by mass spectrometry Classical proteomics is focused on studying when and where a protein is expressed. Since most cellular functions are performed by protein complexes and not by individual proteins it will be important to identify also the interacting protein components. This can be done by biochemical purification of the protein complex followed by MS identification of the different components, e.g. components of the human spliceosome [28], yeast nuclear pore complex [50] and the plastid ribosome [51,52] have been identified with this strategy. Protein complexes are most often purified with an affinity based method, like co-immunoprecipitation [53,54] or GST-pull down experiments [53,55]. DNAbinding proteins can be isolated upon their affinity to specific oligonucleotides and identified by MS [56]. Rigaut et al. have developed a generic purification method for protein complexes under native conditions based on tandem affinity chromatography [57]. In this approach a TAP-tag is fused to the target protein, which is then introduced to the host cell or organism, maintaining the expression of the fusion protein. The TAP-tag consist of a protein A—and a calmodulin binding peptide-tags, and between the tags there is a TEV-protease cleavage site. The fusion protein and its interacting protein components can be efficiently purified from the cell extract using tandem affinity chromatography. Rappsilber et al. have studied the topology of a yeast nuclear pore complex, Nup85p, by affinity purification, cross-linking and mass spectrometry [58]. After affinity purification it is usually sufficient to separate the components in the complex by using one-dimensional SDS-PAGE and thus the pitfalls of 2-DE can be avoided. It is also possible to analyze large protein complexes directly by LC-MS [59]. In DALPC (direct analysis of large protein complexes) a denaturated and reduced protein complex is digested into peptides, and the resulting peptides are separated by two-dimensional chromatography before mass spectrometry. Link et al. [59] showed that DALPC is capable of identifying more than 100 proteins in a single run.
9. Future prospects At the moment proteome analysis are normally carried out by 2-DE followed by mass spectrometric protein identification. Mass spectrometry has proven to be the method of choice for large-scale, high-throughput protein identification [31,60]. The main drawback of this methodology is in 2-DE: the low abundant proteins cannot be found from the gels even if multiple narrow-range immobilized pH-gradient strips are run in
Fig. 4. The principle of ICAT. The ICAT-reagents have a thiol spesific reactive group and a biotin group. This way it is possible to label all peptides containing cysteines and avidin affinity purify them. The cysteines in proteins in cell state 1 are labeled with normal ICAT-reagent, and the cysteines in cell state 2 with deuterated, heavy ICAT-reagent. Then the samples are pooled, digested with trypsin and ICAT-labeled peptides are isolated with affinity chromatography followed by analysis with LC-MS. Protein levels can be quantitated by measuring peak ratios between normal ICAT-derivatized peptide and deuterated ICAT-derivatized peptide. For protein identification peptides are sequenced by MS/MS.
parallel [61]. 2-DE is also laborous and reproducible results are difficult to achieve. Therefore, a lot of progress has to be made before a 2-DE based proteome analysis is as automated and high-throughput screening method as DNA-microarrays are already today. Molecular scanner [27,28] is a promising attempt towards automated proteome analysis, but the problems associated with 2-DE remain. Since it is already possible to identify complex protein mixtures directly by LC-MS/MS [59] it would be tempting to perform proteome analyses without 2-DE. There is however a fundamental problem for doing it with MS alone: MS measurements are not quantitative. Isotope coded affinity tags (ICAT) combined with tandem mass spectrometry [62] is a new method to overcome this problem. The principle of ICAT is shown in Fig. 4. This method is an important step towards a global, automated and quantitative proteome analysis. Since proteins are post-isolation labeled with stable isotopes this method is not limited to sam-
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ples that can be metabolically labeled. The ICAT approach can also be extended to other functional groups than thiol groups from cysteines [62]. The power of proteome analysis lies in its ability to study protein–protein interactions and post-translational modifications in addition to studying changes in gene expression levels. Mass spectrometry is a crucial tool in all these analyses, and the new instrumental developments together with new, improved data processing softwares will make MS even more powerful tool in these studies.
Acknowledgements I want to thank Dr Jari Helin and Dr Sampsa Matikainen for their critical reviewing of this manuscript.
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Review
The role of mass spectrometry in proteome studies Tuula A. Nyman * Turku Centre for Biotechnology, BioCity, P.O. Box 123, FIN-20521 Turku, Finland
Abstract Mass spectrometry (MS) is an important tool in modern protein chemistry. In proteome analyses the expression of hundreds or thousands of proteins can be monitored at the same time. First, complex protein mixtures are separated by two-dimensional gel electrophoresis (2-DE) and then individual proteins are identified by using MS followed by database searches. Recent developments in this field have made it possible to do automated, high-throughput protein identification that is needed in proteome analyses. MS can also be used to characterize post-translational modifications in proteins and to study protein complexes. This review will introduce the current MS methods used in proteome studies, and discuss their advantages and disadvantages. New instrumental MS developments are also presented that are useful in these analyses. © 2001 Elsevier Science B.V. All rights reserved. Keywords: Mass spectrometry (MS); Two-dimensional gel electrophoresis (2-DE); Proteome analyses; MALDI-TOF
1. Introduction Proteomics is the study of total protein complements, proteomes, e.g. from a given tissue or cell type. Proteome analyses are usually carried out by using two-dimensional gel electrophoresis (2-DE) followed by protein identification by mass spectrometry (MS) and database searches. 2-DE dates back to 1970s [1], but protein identification and characterization by mass spectrometry has become possible only during the last decade. The most important reasons for this are the new soft ionization methods, matrix-assisted laser-desorption ionization (MALDI) [2] and electrospray ionization (ESI) [3,4], together with developments in sample preparation techniques and MS instruments. Equally crucial for protein identification by MS is also the exponentially growing amount of sequence information available in databases. Compared to transcriptomics, where the DNA-microarray technology is a completely automated method, proteome analysis requires a lot of manual work and suffers from problems mainly associated with 2-DE * Tel.: +358-2-333-8027; fax: +358-2-333-8000. E-mail address: [email protected] (T.A. Nyman).
[5,6]. Despite the difficulties the importance of proteomics is well recognized. DNA-microarray technology is based on measuring steady-state mRNA levels whereas proteomics studies proteins, which are the active agents in cells. Recent studies have also shown that mRNA expression and protein levels do not always correlate [7,8]. Proteins can be post-translationally modified or present as different isoforms, and these modifications cannot be predicted from the corresponding DNA-sequences. Mass spectrometry has an important role in characterization of these modifications.
2. Protein identification by mass spectrometry Mass spectrometers consist of the ion source, mass analyzer, ion detector and data acquisition unit. First, molecules are ionized in the ion source, then they are separated according to their mass-to-charge ratio in the mass analyzer and the separated ions are detected. Mass spectrometry has become a widely used method in protein analysis after the invention of MALDI and electrospray ionization methods (Fig. 1). There are several options for mass analyzer, the most common combinations being time-of-flight (TOF) connected to
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MALDI and triple quadrupole, quadrupole-TOF or ion trap mass analyzer coupled to ESI. In proteome analysis electrophoretically separated proteins can be identified by mass spectrometry with two different approaches. The simplest way is a technique called peptide mass fingerprinting (PMF) [9– 13]. In this approach the protein spot of interest is in-gel digested with a specific enzyme, the resulting peptides are extracted from the gel and the molecular weights of these peptides are measured. Database search programs can create theoretical PMFs for all the proteins in the database, and compare them to the obtained one (Fig. 2). In the second approach peptides after in-gel digestion are fragmented in the mass spectrometer yielding partial amino acid sequences from the peptides (sequence tags), and database searches are performed using both molecular weight and sequence information [14,15]. PMF is usually carried out with MALDI-TOF, and sequence tags by nano-ESI tandem mass spectrometry (MS/MS) [16– 18]. The sensitivity of protein identification by MS is practically in the femtomole range, even though sensitivities at the attomole level have been reported [19].
3. Peptide mass fingerprinting by MALDI-TOF Protein identification by in-gel digestion and peptide mass fingerprinting is nowadays a routine method in most MS laboratories. This method requires optimized in-gel digestion [20– 22] and desalting protocols before MALDI analysis. Trypsin is the most often used enzyme for in-gel digestion, since it cleaves proteins very specifically after lysines and arginines. It also produces quite small peptides, typically in the range of 600– 2500 Da, which can be eluted efficiently from the gel. MALDI-TOF is a relatively simple and easy-to-use form of mass spectrometry, and it is more tolerant to salts and other contaminants present in the sample than ESI. Small aliquot of the extracted peptide mixture can be directly deposited to the MALDI target plate for PMF, and the rest of the sample can be stored for later analysis by ESI-MS. If the total peptide sample after in-gel digestion is used in MALDI analyses, best results are obtained when the samples are desalted. Desalting can be performed with commercially available ZipTips (Millipore) or by making a small reversed-phase material column into a gel-loader tip [23,24]. Mass accuracy of MS measurements is especially important in PMF [25]. Modern MALDI-TOF instruments are delayed extraction reflector instruments, and the mass accuracy obtained with them is in the range of 10–30 ppm for peptides with internal calibration. With this kind of mass accuracy usually 4– 5 peptides are enough to identify a protein unambiguously. MALDI produces mostly singly charged ions, so the spectra are
easy to interpret and since the sample is in crystallized form, analysis time is seldom a limiting factor.
4. Automation of peptide mass fingerprinting For high-throughput protein identification it is necessary to automate sample preparation together with MS measurements, data interpretation and database searches. There are already robots available for spot cutting, in-gel digestion, desalting of the samples and spotting for MALDI plates. In addition, MALDI measurements can be automated with modern MALDITOF-instruments. Data processing and database searching can also be automated, but in practice when very small amounts of protein digests are analyzed, the spectra have to be checked manually before database searches. In proteome analyses it would be tempting to identify all the proteins in the gel without cutting individual spots from the gel. There have been several attempts to do this, the most recent being called a molecular scanner [26,27]. In this approach the proteins in the gel are simultaneously proteolytically digested and electrotransferred onto a polyvinylidene difluoride membrane, and the membrane is directly scanned by MALDITOF. As promising as the molecular scanner is, serious problems still arise from the time needed to scan the whole membrane (for 16× 16 cm2 membrane 36 days
Fig. 1. The principles of: (A) MALDI; and (B) electrospray ionization. (A) The sample is mixed with a UV-absorbing matrix and crystallized to the MALDI target plate. The matrix absorbs energy from the laser causing the matrix and sample to be volatilized; (B) In ESI ions are formed from a liquid sample by applying a potential which causes the sample to spray and produce charged droplets. These droplets will form usually multiply charged ions upon solvent evaporation.
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quence information are used compared to searching with peptide molecular weights only. With sequence tag information also EST-databases can be used in database searches [28]. Sequence tag from one peptide is usually sufficient to identify the protein. Compared to MALDI measurements, nano-ESI-MS/MS is time consuming and laborous, and samples have to be desalted prior ESI-analysis. The analysis time is also limited since the liquid sample is consumed all the time during the MS-measurement. The problems of nano-ESI can be solved to some extent by coupling HPLC with the ESI-MS. ESI-analysis is concentration sensitive, and the use of modern micro- and nano-HPLC-methods provides high local peptide concentrations, and the total LC-separated sample can be introduced to the mass spectrometer without splitting. LC-separation before mass spectrometry is especially useful when complex mixtures are analyzed, because it is possible to do data-dependent experiments (DDE). In DDE all the ions coming into the detector are measured, and when there is a peak coming from the HPLC to the MS (i.e. an increase in the total ion current), the software will switch automat-
Fig. 2. Peptide mass measured is critical in calibrated. If trypsin cleave also itself into
fingerprinting. Mass accuracy of the peptides PMF, therefore the spectra has to internally be is used for in-gel digestion, the enzyme will fragments, which can be used for calibration.
of continuos scanning and more than 40 Gb of raw data).
5. Creating sequence tags by ESI-MS/MS Peptide mass fingerprinting can only be successful when the digested protein exists in the protein or genomic databases, and when four or more peptides are obtained in MALDI analysis. If this is not the case, or if the protein is only in EST-databases as partial sequences, it is necessary to obtain sequence information from the peptides. The most common way to do this is by ESI-MS/MS (Fig. 3). The advantage of creating sequence tags is that database searches are much more specific when both peptide molecular weight and se-
Fig. 3. Product ion scan. First, quadrupoles are set to pass through the whole peptide mixture, and this gives the molecular masses of the peptides present. In ESI, peptides are usually doubly or triply charged. One of the peptides, parent ion (seen in the inset), can then be selected by letting only this peptide to pass through Q1. It is then fragmented in the collision chamber, Q2, and the resulting fragments are detected in the last quadrupole (OR TOF). Partial amino acid sequence can then be read from the spectra.
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ically from MS to MS/MS mode and do a product ion scan. In this way it is possible to generate much more MS/MS data from one sample than in nano-ESI experiments. Yates et al. have reviewed the strategies for automated protein identification using microcolumn LC-MS/MS [29].
6. Recent developments in MS instruments The first step for protein identification by MS is usually PMF with MALDI-TOF, and if this method fails to identify the protein, the next step is to create sequence tags by ESI-MS/MS. In PMF only small part of the sample on the MALDI plate is consumed during the analysis, so there is sample left to create sequence tags from it also. It is possible to partially sequence peptides with a MALDI-TOF instrument with a method called ‘post source decay’. However, this method produces spectra, that are difficult to interpret, and the mass accuracy of the measured peptide fragments is poor. Recently, MALDI with a hybrid quadrupole-TOF mass analyzer [30,31] and a prototype of MALDI-TOF/TOF instrument [32] were introduced. Thus it is possible to do PMF as the first choice of protein identification and sequence tags by MS/MS when necessary from the same sample. MALDI-hybrid quadrupole-TOF has shown great promise for highthroughput protein identification [31]. Prototype of rapidly switchable MALDI/ESI-ion source for a hybrid quadrupole-TOF has also been introduced [33]. This improves protein identification strategies since it is possible to measure MS spectra with both ionization modes and combine MS/MS information from singly and multiply charged ions.
7. Post-translational modification analyses in proteome studies One of the key advantages of doing proteome analysis is that both protein expression levels and post-translational modifications, like phosphorylation and glycosylation, can be studied. Protein phosphorylation analyses are especially interesting since phosphorylation plays major role in e.g. many signal transduction pathways. The easiest way to detect differencies in protein expression levels is by metabolically labeling the samples with 35S-Met before 2-DE followed by autoradiography of the 2-DE gel [34–36]. Different protein phosphoand glycoforms usually apprear as a ‘train’ of spots in these gels, but more detailed analysis is needed to determine the type of modification. Changes in protein phosphorylation can be studied either by labeling cells with 32P or by western blotting with phospho-specific
antibodies. If the 32P-approach is used the 2-DE gels are run as usually, and after gel comparison the spots of interest can be cut out from the gel and identified by mass spectrometry [36]. Soskic et al. [37] have used the blotting method where two similar 2-DE gels were run in parallel, one was used for blotting with phospho-specific antibodies and the other one for normal staining and cutting spots for mass spectrometry. Glycoprotein detection from 2-DE gels can be done with similar type of blotting experiments [38,39]. More detailed characterization of protein phosphorylation and glycosylation can be done by mass spectrometry but this requires more starting material than just a spot in 2-DE gel. In addition, these analyses are not as straightforward methodologically and are technically more difficult compared to protein identification by MS. There are several ways to locate the phosphorylation sites in a protein. To find out which peptides are phosphorylated in the digestion mixture, one can perform PMF by MALDI-MS before and after phosphatase treatment: phosphopeptides will lose their phosphate group and thus their molecular weight will be 80 Da lower after phosphatase treatment. More detailed characterization can be done with nano-ESItriple quadrupole instrument with parent ion scans: peptides are fragmented like in product ion scan (Fig. 3), but in this case Q3 is set to pass only the fragmented PO− 3 moiety, m/z 79, and Q1 is scanned to find the parent ion for PO− 3 moiety. When the phosphopeptides are found they can be sequenced by normal product ion scans. One additional possibility is to isolate phosphopeptides with immobilized metal ion affinity chromatography (IMAC) and analyze only this fraction by MS [40,41]. The invention of MALDI-hybrid quadrupole-TOF will probably help protein phosphorylation studies: the peptide sample can be analyzed before and after phosphatase treatment, and the peptides which lose their phosphate group can be directly sequenced from the MALDI sample. Protein phosphorylation studies by MS, possibilities and limitations, have been recently reviewed by Neubauer and Mann [42]. In glycosylation studies the glycans are liberated from the protein, and the glycan structures are determined from the free glycans. The glycan structures can be characterized by combining MALDI-measurements to exoglycosidase digestions [43–46] or if more detailed structural information is wanted by ESI-MS with product ions scans [46–48]. So far there have been very limited amount of reports where glycan structures have been characterized from electrophoretically separated proteins; in one study the N-linked glycans were enzymatically released from the protein in one-dimensional SDS-PAGE gel followed by structural analysis with MALDI-MS and exoglycosidase digestions [49].
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8. Protein–protein interaction studies by mass spectrometry Classical proteomics is focused on studying when and where a protein is expressed. Since most cellular functions are performed by protein complexes and not by individual proteins it will be important to identify also the interacting protein components. This can be done by biochemical purification of the protein complex followed by MS identification of the different components, e.g. components of the human spliceosome [28], yeast nuclear pore complex [50] and the plastid ribosome [51,52] have been identified with this strategy. Protein complexes are most often purified with an affinity based method, like co-immunoprecipitation [53,54] or GST-pull down experiments [53,55]. DNAbinding proteins can be isolated upon their affinity to specific oligonucleotides and identified by MS [56]. Rigaut et al. have developed a generic purification method for protein complexes under native conditions based on tandem affinity chromatography [57]. In this approach a TAP-tag is fused to the target protein, which is then introduced to the host cell or organism, maintaining the expression of the fusion protein. The TAP-tag consist of a protein A—and a calmodulin binding peptide-tags, and between the tags there is a TEV-protease cleavage site. The fusion protein and its interacting protein components can be efficiently purified from the cell extract using tandem affinity chromatography. Rappsilber et al. have studied the topology of a yeast nuclear pore complex, Nup85p, by affinity purification, cross-linking and mass spectrometry [58]. After affinity purification it is usually sufficient to separate the components in the complex by using one-dimensional SDS-PAGE and thus the pitfalls of 2-DE can be avoided. It is also possible to analyze large protein complexes directly by LC-MS [59]. In DALPC (direct analysis of large protein complexes) a denaturated and reduced protein complex is digested into peptides, and the resulting peptides are separated by two-dimensional chromatography before mass spectrometry. Link et al. [59] showed that DALPC is capable of identifying more than 100 proteins in a single run.
9. Future prospects At the moment proteome analysis are normally carried out by 2-DE followed by mass spectrometric protein identification. Mass spectrometry has proven to be the method of choice for large-scale, high-throughput protein identification [31,60]. The main drawback of this methodology is in 2-DE: the low abundant proteins cannot be found from the gels even if multiple narrow-range immobilized pH-gradient strips are run in
Fig. 4. The principle of ICAT. The ICAT-reagents have a thiol spesific reactive group and a biotin group. This way it is possible to label all peptides containing cysteines and avidin affinity purify them. The cysteines in proteins in cell state 1 are labeled with normal ICAT-reagent, and the cysteines in cell state 2 with deuterated, heavy ICAT-reagent. Then the samples are pooled, digested with trypsin and ICAT-labeled peptides are isolated with affinity chromatography followed by analysis with LC-MS. Protein levels can be quantitated by measuring peak ratios between normal ICAT-derivatized peptide and deuterated ICAT-derivatized peptide. For protein identification peptides are sequenced by MS/MS.
parallel [61]. 2-DE is also laborous and reproducible results are difficult to achieve. Therefore, a lot of progress has to be made before a 2-DE based proteome analysis is as automated and high-throughput screening method as DNA-microarrays are already today. Molecular scanner [27,28] is a promising attempt towards automated proteome analysis, but the problems associated with 2-DE remain. Since it is already possible to identify complex protein mixtures directly by LC-MS/MS [59] it would be tempting to perform proteome analyses without 2-DE. There is however a fundamental problem for doing it with MS alone: MS measurements are not quantitative. Isotope coded affinity tags (ICAT) combined with tandem mass spectrometry [62] is a new method to overcome this problem. The principle of ICAT is shown in Fig. 4. This method is an important step towards a global, automated and quantitative proteome analysis. Since proteins are post-isolation labeled with stable isotopes this method is not limited to sam-
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ples that can be metabolically labeled. The ICAT approach can also be extended to other functional groups than thiol groups from cysteines [62]. The power of proteome analysis lies in its ability to study protein–protein interactions and post-translational modifications in addition to studying changes in gene expression levels. Mass spectrometry is a crucial tool in all these analyses, and the new instrumental developments together with new, improved data processing softwares will make MS even more powerful tool in these studies.
Acknowledgements I want to thank Dr Jari Helin and Dr Sampsa Matikainen for their critical reviewing of this manuscript.
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