686 DOI 10.1002/pmic.200700617 Proteomics 2008, 8, 686–705 REVIEW Technologies and methods for sample pretreatment in efficient proteome and peptidome analysis Xiaogang Jiang1, 2, Mingliang Ye1 and Hanfa Zou1 1 National Chromatographic R&A Center, Dalian Institute of Chemical Physics, Chinese Academy of Sciences, Dalian, China 2 Suzhou University, Suzhou, China Although great progresses have been made in proteomics during the last decade, proteomics is still in its infancy. Extreme complexity of proteome sample and large dynamic range of protein abundance overwhelm the capability of all currently available analytical platforms. Sample pretreatment is a good approach to reduce the complexity of proteome sample and decrease the dynamic range. In this article, we present an overview of different technologies and methods for sample pretreatment in efficient proteome and peptidome analysis. Methods for isolation of rare amino acid-containing peptides, terminal peptides, PTM peptides and endogenous peptides are reviewed. In addition, two automated sample pretreatment technologies, i.e. automated sample injection and on-line digestion, are also covered. Received: June 28, 2007 Revised: October 26, 2007 Accepted: November 1, 2007 Keywords: MS / Peptidome / Sample pretreatment 1 Introduction Proteomics, the analysis of protein components in a cell or an organism, has experienced a rapid development in the last decade. However, proteomic analysis is still technically challenging because of extreme complexity of proteome samples. Besides tens of thousands of proteins coded by the genome of many cells, there are splicing variants, PTMs, and genetic variations among individuals. And the proteins expressed in a cell have a large dynamic range of protein abundance which further complicates the proteome analysis. There is no amplification method for low abundance proteins comCorrespondence: Professor Dr. Hanfa Zou, National Chromatographic R&A Center, Dalian Institute of Chemical Physics, Chinese Academy of Sciences, Dalian 116023, P.R. China E-mail: [email protected] Fax: 186-411-8437-9620 Abbreviations: CNBr, cyanogen bromide; ConA, Concanavalin A; Cys, cysteine; His, histidine; ICAT, isotope-coded affinity tag; Met, methionine; MW, molecular weight; PST, protein sequence tag; RAMs, restricted access materials; SAX, strong anion exchange chromatography; SCX, strong cation-exchange chromatography; Trp, tryptophan © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim parable to the PCR technique for amplifying DNA. Therefore, sample pretreatment is a practical way to enhance the detection sensitivity of low abundance proteins for comprehensive proteomic analysis. At present, there are two dominant approaches for largescale proteome analysis. The first is based on protein separation by 2-D PAGE. Protein identification is achieved by MS analysis of excised and digested protein spots. Until now, 2-DE is still a main-stream method for protein separation, which allows the resolution of more than 5000 proteins simultaneously. However, some classes of proteins such as very acidic or basic proteins, extremely big or small proteins and membrane proteins are difficult to be separate by 2-DE [1]. Also, 2-DE is notoriously difficult to automate, which limits throughput and results in greater experimental variability with manual intervention. To overcome these limitations as well as to improve the throughput of protein identification, the second approach, shotgun proteome analysis based on the chromatographic techniques, has been developed [2]. In this approach, complex protein mixture is digested first and then the resulting digests are analyzed by LC-MS/MS. Thousands of proteins can be easily identified in a single experiment by shot-gun proteomics if multi-dimensional chromatography, typically strong cation-exchange www.proteomics-journal.com Technology Proteomics 2008, 8, 686–705 chromatography followed by RP chromatography (SCX-RP), is applied for separating the peptide mixture resulted from the digests of proteins prior to MS analysis [3, 4]. Shotgun proteomics is a high-throughput proteome analysis approach. However, proteome sample becomes much more complex due to the digestion of proteins, as one protein can generate dozens of peptides. The complexity of the proteome sample is far beyond the capacity of modern analytical systems; the large dynamic range in protein abundance also largely exceeds the dynamic range of all available analytical platforms. In practice, the peptides derived from high abundance proteins seriously suppress the detection of peptides from low abundance proteins. Thus direct analysis of protein digests often leads to loss of protein information especially those of low abundance proteins. To overcome the problem of complexity as well as the dynamic range problem, sample pretreatment is a good approach, which can be performed either at protein level or at peptide level. At protein level, the complexity of the sample could be reduced by pre-fractionation of proteins, and the detection sensitivity for low abundance proteins could be improved by removing high-abundance proteins or enriching specific groups of interested proteins [5, 6]. At peptide level, the complexity of the sample could also be reduced by pre-fractionation of peptides. Furthermore, a more effective method to reduce the complexity of a peptide mixture is to enrich representative peptides such as peptides containing rare amino acids or terminal peptides. Isolation of PTM peptides from peptide mixture is also an effective way to circumvent the suppression of high-abundance unmodified proteins or peptides on the analysis of protein modification. Here we reviewed the technologies and methods for sample pretreatment at peptide level. As a subset of proteomics, peptidomics analyzes low molecular weight (MW) proteins (,10 kDa) or endogenous peptides from biological source. The methods to isolate endogenous peptides for efficient peptidome analysis were also reviewed. In addition, two automated methods of sample processing such as sample introduction for nanoflow LC-MS/MS system, on-line digestion with micro-enzyme-reactor were also covered in this review. 2 Selective capture of peptides containing rare amino acids In shotgun proteomics, proteins in the proteome sample are first digested by protease such as trypsin, and then the resulting peptides are analyzed by LC-MS/MS. Enzymatic digestion of proteins generate dozens of peptides per protein, which will result in an extremely complex peptide mixture. To reduce the complexity of the samples, multi-dimensional separation of peptides is often conducted prior to MS analysis [7]. However, the number of peptides still overwhelms the peak capacity of current multi-dimensional © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim 687 separation systems. Theoretically, one unique peptide would be sufficient to unambiguously identify each parent protein. As one protein can generate many peptides, there are a large number of redundant peptides presented in the digests of proteins. If one or a couple of representative peptides could be isolated for each protein, the complexity of the samples for proteome profiling would be reduced by one to two orders of magnitudes at least. Based on the theoretical digestion of proteins in human database, tryptic peptides containing rare amino acids such as cysteine (Cys), methionine (Met), tryptophan (Trp),and histidine (His) account for less than 31% of all tryptic peptides, however, they represent more than 91% of all human proteins [8]. Thus isolation of peptides containing rare amino acids is an effective way to reduce the complexity of proteome sample while keeping the integrity of the proteome. However, the distribution of these rare amino acid residues in different proteins is different, thus the method by isolating peptides containing these residues bias to proteins with high percentage of these residues. As we all know, a protein contains only one N-terminal and one C-terminal peptide after digestion. Terminal peptides account for only about 6% of all tryptic peptides while represents 100% of proteins in the proteome [8]. Therefore, isolation of terminal peptides represents the idea approach to reduce sample complexity. Up to now, several strategies have been successfully developed to target and isolate peptides containing rare amino acids (Cys, Met, Trp, His, etc.) and terminal peptides. 2.1 Cys-containing peptides Among these rare amino acids in peptides, Cys is one of the earliest targets for amino-acid-based peptide selection. The isolation of Cys-containing peptides is typically achieved by Cys-specific tags possessing iodoacetyl or vinyl functionalities which react with the thiol group of Cys [9]. Among all Cys-specific tags, isotope-coded affinity tag (ICAT) developed by Aebersold and co-workers [10] probably has the highest impact as it can not only simplify the complexity of peptide mixture but also provide quantitative information. The ICAT reagents included a Cys-reactive idoacetyl group, a differentially isotope coded linker region, and an affinity tag (biotin). The proteins with Cys residues were labeled by ICAT reagent through the Cys-reactive group. After digestion of the labeled proteins, Cys-containing peptides were then isolated from the digest by avidin affinity chromatography. Isolation of Cys-containing peptides using Cys-specific affinity tags typically included two steps: the protein or peptides containing Cys were first labeled with the tag in solution and then the labeled peptides (after digestion if the proteins are labeled) were isolated by affinity chromatography. To circumvent the additional chromatographic purification step, solid-phase capture approach has been introduced [11]. The tag with a Cys-reactive group was anchored on solid-phase beads for the capture of Cys-containing peptides. Similar to ICAT reagent, the tag immobilized on solid phase also included three parts. www.proteomics-journal.com 688 X. Jiang et al. Besides the Cys-reactive group and isotope-coded group, a photo-cleavage group, instead of biotin group in ICAT reagent, was included to release the captured peptides from beads. Cys-containing peptides were first captured by the beads and then released by UV irradiation. Thus, simplification of complex peptide mixtures could be achieved without avidin affinity chromatography. In addition to use of chemical tags, chromatographic approach could also be used to isolate Cys-containing peptides from complex peptide mixture. Wang et al. [12] described a procedure in which Cys-containing peptides from tryptic digests of complex protein mixtures were selected by covalent chromatography based on thiol-disulfide exchange. Following disruption of disulfide bridges with 2,20 -dipyridyl disulfide, all proteins were digested with trypsin and acylated with succinic anhydride. Cys-containing peptides were then selected from the acylated digest by disulfide interchange with sulfhydryl groups on a thiopropyl Sepharose gel. Their results indicated that by selecting Cys-containing peptides, the complexity of protein digest could be reduced. Another chromatographic approach to isolate Cys-containing peptides was diagonal chromatography which was reported by Gevaert et al. [13]. Cys in proteins were converted to hydrophobic residues by mixed disulfide formation with Ellman’s reagent. Proteins were subsequently digested with trypsin and the generated peptide mixture was fractionated first by RP-LC. Cys-containing peptides were isolated from each primary fraction by a reduction step followed by a secondary peptide separation on the same column, and the secondary separation was performed under identical conditions as that of the primary separation. The reducing agent removed the covalently attached group from the Cys side chain, making Cys-containing peptides more hydrophilic. Thereby, such peptides could be specifically collected during the secondary separation. They showed that this procedure efficiently isolated Cys-containing peptides, making the sample mixture less complex for further analysis. By isolating Cys-containing peptides a significant number of low abundance proteins could be identified and a dynamic range for protein identification spanning several orders of magnitude could be achieved. For example, Wang et al. [14] reported a global proteomic approach by isolation of cysteinyl-peptide complemented with a global enzymatic digestion method for the characterization of the whole mouse brain proteome. A total of 48 328 different peptides were confidently identified (.98% confidence level), covering 7792 non-redundant proteins (approximately 34% of the predicted mouse proteome). A total of 1564 and 1859 proteins were identified exclusively from the cysteinyl-peptide and the global peptide samples, respectively, corresponding to 25 and 31% improvements in proteome coverage compared to analysis of only the global peptide or cysteinyl-peptide samples. Of particular interest are the ,2000 membrane proteins (26%) and ,1000 extracellular proteins that were identified without any specific enrichment of membrane fractions. © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Proteomics 2008, 8, 686–705 2.2 Met-containing peptides The Met-containing peptides account for only 25.5% of all tryptic peptides, while they represent 98.9% of human proteins. This indicates that isolation of Met-containing peptides is more efficient to reduce the sample complexity than isolation of Cys-containing peptides. However, very limited applications are reported to isolate Met-containing peptides probably because of the difficulty in isolation of these peptides. Only two approaches have been reported for the isolation and analysis of Met-containing peptides. The first one is based on solid phase approach and the second one is also based on diagonal chromatography. The solid phase approach first used Met-specific beads to covalently attach Met-containing peptides via bromoacetyl functional groups to a solid support [15]. Target protein populations were firstly digested under reduced and alkylated conditions, and resultant peptides selectively extracted via covalent attachment to Met residues by bromoacetyl reactive groups tethered to the surface of glass beads packed in small reaction vessels. After conjugation, reactive beads were stringently washed to remove nonspecifically bound peptides and then later treated with beta-mercaptoethanol to release captured Met-containing peptides in their nascent state, without complicating affinity tags. This approach had been applied to an Escherichia coli lysate model system and had demonstrated facility in reducing global digest complexity, enhancing sensitivity to low protein expression levels, and improving significant quantitative capability. In the second approach using diagonal chromatography, Met side chains on peptides were oxidized between chromatographic runs, resulting in increased polarity and therefore earlier RP-LC elution of the oxidized peptides in the second dimension away from the bulk of unmodified peptides [16]. 2.3 Trp-containing peptides For targeting Trp-containing peptides, a chemical tagging method has been developed where Trp residues were labeled with 2-nitrobenzenesulfenyl chloride [17]. The tagged peptides were enriched by a Sephadex column based on the increase of hydrophobicity upon modification of Trp residues. However, many non-labeled peptides were also isolated during isolation of tagged peptides by Sephadex column. To improve the selectivity of the isolation, a phenyl resin, which was generally used for hydrophobic interaction chromatography and RP-LC, was used to isolate labeled peptides[18]. This was because the 2-nitrobenzenesulfenyl-Trp moiety was not only hydrophobic but was also aromatic, p-electron interactions between phenyl groups of the column and the Trp moiety of the labeled peptides were more specific. Besides the approach by labeling target residue followed with chromatographic purification, an interesting solid phase approach was presented to isolate Trp-containing peptides recently [19]. It was based on the reversible reaction of Trp with malondialdehyde and trapping of the derivate Trp-pepwww.proteomics-journal.com Proteomics 2008, 8, 686–705 tides on hydrazide beads via the free aldehyde group of the modified peptides. After conjugation, the captured peptides were recovered in their native form by specific cleavage reactions using hydrazine or pyrrolidine. The applicability of this Trp-specific enrichment procedure to complex biological samples was demonstrated for total yeast cell lysate. Analysis of the processed fraction by 1-D LC-MS/MS confirmed the specificity of the enrichment procedure, as more than 85% of the peptides recovered from the enrichment step contained Trp. The reduction in sample complexity also resulted in the identification of additional proteins in comparison to the untreated lysate. 2.4 His-containing peptides Instead of using chemical tag, IMAC loaded with Cu21 was utilized to enrich His-containing peptides from complex peptide mixtures [20–23]. It had been shown that most Hiscontaining peptides could be captured from tryptic digests with little non-specific binding through the use of very hydrophilic IMAC columns and imidazole as a displacer [24]. To reduce non-specific interactions of the IMAC resin with peptides containing Cys, Trp and carboxyl groups, samples were alkylated and acetylated before affinity capture of the His-containing peptides by IMAC columns. Combinational use of covalent chromatography and IMAC was applied to enrich peptides containing both Cys and His [25], which further simplified the complexity of peptide mixture. 2.5 Terminal peptides Each protein has only two terminals, i.e. N-terminal and Cterminal. Isolation of either N-terminal peptide or C-terminal peptide could significantly reduce the complexity of proteome sample. However, isolation of C-terminal peptides from protein digests for proteome analysis was not reported up to now, probably because of the difficulty in specific isolation of these peptides. In contrast, several methods for selective isolation of N-terminal peptides from complex mixture of digested peptides for proteomic analysis have been reported. Two chromatographic techniques were developed to isolate N-terminal peptides. The first one was based on diagonal chromatography which also was reported by Gevaert et al. [26]. In the procedure, free amino groups in proteins were first blocked by acetylation and then digested with trypsin. Except N-terminal peptides, new primary amino groups were generated after digestion. After RP chromatographic fractionation of the generated peptide mixture, primary amino groups of internal peptides were blocked using 2,4,6-trinitrobenzenesulfonic acid; this displayed a strong hydrophobic shift and therefore segregated from the unaltered N-terminal peptides during a second identical separation step. N-terminal peptides could thereby be specifically collected for LCMS/MS analysis. The second one was based on SCX [27]. As N-terminally acetylated peptides lacked a positive charge at © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Technology 689 their N-terminal amino group, such peptides could be enriched in the non-binding fraction and in the early fractions under eluting conditions of low concentration of salt. Oesterhelt et al. [27] utilized the above two chromatographic methods for the large-scale identification of naturally N-terminal blocked peptides from prokaryotes. Combining all data, 606 N-terminal peptides from Halobacterium salinarum and 328 from Natronomonas pharaonis were reliably identified. McDonald et al. [28] and Yamaguchi et al. [29] reported two methods using the biotin-avidin technique for the isolation of N-terminal peptides of proteins, respectively. Though biotin-avidin technique was used in both methods, the target peptides of avidin chromatography were different. In the first method, the internal peptides were trapped by an immobilized avidin column, while in the second method Nterminal peptides were trapped. The schemes for both approaches are shown in Fig. 1. In the first method (Fig. 1A), all free amino groups on proteins were blocked by acetylation in the beginning. Subsequently, proteolysis of proteins generated new peptides, and all but the N-terminal peptides exposed a new free amino group that was subsequently biotinylated. These biotinylated internal peptides were removed by recovery onto immobilized avidin, leaving behind the set of N-terminal peptides. In contrast, a different strategy was applied in the second method (Fig. 1B). Instead of blocking all amino groups, only e- amino groups of lysine residues were blocked by O-methylisourea. The left free a- amino groups on N-terminal of protein then reacted with biotinylcysteic acid. After proteolysis, the N-terminal peptides were enriched by avidin resins. The advantage of the first method was that all N-terminal peptides (whether naturally free or blocked) could be enriched. However, the second method could not isolate N-terminal blocked proteins as there was no free a-amino group on N-terminal of these proteins. Recently, Mikami et al. [30] also developed a method for selective isolation of N-terminal blocked peptides from protein digests. The approach was based on a newly designed isocyanate-resin, resin-NCO, which specifically reacted with a-amino groups under weak acidic conditions. Because only N-terminal blocked peptides had no a-amino groups, all peptides except N-terminal blocked peptides would be captured when the protein digest was incubated with resinNCO. Thus the N-terminal blocked peptides could be recovered from the supernatant of the above solution. Obviously, this method could also be used to isolate all N-terminal peptides if acetylation of protein was conducted before proteolysis. Isolation of only N-terminal peptides from entire proteins is the most effective way to reduce the sample complexity in theory. However, identification of proteins by only one peptide is often not confident enough because a decent spectrum can not be obtained for every peptide. Furthermore, there are many variations of modifications for N-terminal blocked peptides which result in difficulty for identification of these peptides. The Protein Sequence Tag (PST) www.proteomics-journal.com 690 X. Jiang et al. Proteomics 2008, 8, 686–705 Figure 1. Strategies for isolation of N-terminal peptides using biotin-avidin technique. (A): Free aand e-amino groups are acetylated before proteolysis, which is followed by biotinylation of proteolytically exposed a-amino groups. Subsequent subtractive binding to immobilized streptavidin creates a preparation enriched in those peptides that were originally derived from the N-terminus, blocked by acetylation and therefore refractory to biotinylation. (B): All e-amino groups of a protein are guanidinated with omethylisourea. And the Na-amino group is specifically modified with biotinylcysteic acid (BCA). The derivatized protein is digested with trypsin, and the digests are loaded into avidin resins for the specific adsorption of the Nterminal peptide fragment modified with BCA. The symbols triangle, starburst, and cross represent an alkyl group, a guanidino group, and BCA, respectively. Reprinted from [28, 29]. © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.proteomics-journal.com Proteomics 2008, 8, 686–705 technology reported by Kuhn et al. [31–33] might circumvent this problem. It dealt with the isolation and MS/MS based identification of one N-terminal peptide from each polypeptide fragment generated by cyanogen bromide (CNBr) cleavage of a mixture of proteins. The first step in the PST isolation process involved chemical cleavage of the protein mixture with CNBr. Then after the free primary amino groups were blocked by amine-reactive Basic Mass Tag, the CNBr cleavage polypeptides were further digested by trypsin, which generated new amino groups in all of the non-N-terminal cleavage peptides. These non-N-terminal cleavage peptides were removed by scavenger resin and the pool of peptides that represented the N-terminal fragments from each CNBr cleavage peptide was recovered. As the proteins were identified by multiple peptides, the identification was much more confident. The number of representing peptides generated by this method for each protein depends on the distribution of methionine. The PST process obtained up to eight peptides per protein on average from Saccharomyces cerevisiae and 99% of proteins had at least one peptide available per protein [32]. In that way, almost all the proteins could be analyzed and the moderate degree of redundancy increased confidence in protein identification. Taking advantage of the solubilization step using CNBr, the procedure allowed the study of complex mixtures of hydrophobic proteins, particularly membrane proteins, which had been demonstrated by analysis of a crude mitochondrial fraction [33]. If isotope-coded Basic Mass Tag was used to block free primary amino groups in proteins, this method could also be applied for quantitative proteome analysis [31]. 3 Specific isolation of PTM peptides Besides the amino acid-specific and the general tagging strategies presented above, methods specifically tailored to isolate PTM peptides for the analysis of protein PTM have received a great deal of attention over the past few years. Because of the low abundance of modified proteins and low stoichiometry of the modifications, modified peptides present with large amount of unmodified peptide derived from unmodified proteins and unmodified amino acid sequence in the modified proteins. Shotgun proteomic analysis of PTM proteins relies heavily on the efficiency of isolation and enrichment of modified peptides. Phosphorylation and glycosylation are two most important PTMs of proteins, the methods to isolate phosphorylated and glycosylated peptides are reviewed in detail as follows. 3.1 Isolation of phosphopeptides Protein phosphorylation is one of the most important regulatory events in cells, guiding primary biological processes, such as cell division, growth, migration, differentiation, and intercellular communication in eukaryotes. And it has received a great deal of attention in the scientific community © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Technology 691 for decades. Shot-gun proteomics is the dominate approach for large-scale analysis of protein phosphorylation. In this approach, protein mixtures containing phosphoproteins are digested first by trypsin, and then phosphopeptides are enriched from the resulting peptide mixture for LC-MS/MS analysis. The specificity for the isolation of phosphopeptides is crucial as the presence of small amounts of non-phosphopeptides will still seriously suppress the ionization of phosphopeptides in MS. To improve the specificity of phosphopeptide isolation, a series of technologies, as shown in Table 1, are developed to specifically isolate phosphopeptides from complex peptide mixture. 3.1.1 Immobilized metal affinity chromatography IMAC was first used to isolate phosphopeptides 20 years ago, and it is still the most widely used method [34–37]. In IMAC, metal ions such as Fe31, Ga31 were typically immobilized to beads, phosphopeptides were selectively captured by the beads because of high affinity of metal ions for the phosphate moiety [38]. In recent years, new materials such as nano-materials, monolithic materials have been adopted as the support for IMAC. For example, Fe31 immobilized mesoporous molecular sieves MCM-41 with particle size of ca. 600 nm and pore size of ca. 3 nm were synthesized to selectively trap and separate phosphopeptides from tryptic digest of proteins [39], and Fe31 immobilized silica monolithic capillary column with id of 75 mm was prepared to enrich phosphopeptides from minute samples [40]. However, the conventional IMAC with Fe31 lacked enough specificity. Some non-phosphorylated peptides especially acidic peptides were also strongly bound to the adsorbents, which resulted in serious interference for the analysis of phosphopeptides. Therefore, esterification of the acidic groups of peptides prior to IMAC purification was conducted to improve the enrichment specificity [41]. This approach has been successfully applied to large-scale analysis of phosphorylation in the proteome of yeast [41], rat liver [42], and HT-29 human carcinoma cell line [43]. However, because of incomplete reactions, the esterification might also increase sample complexity and so interfere with subsequent MS analysis. Recently, we have developed a new IMAC adsorbents by immobilization of Zr41 on phosphonate modified polymer beads to improve the enrichment specificity [44]. The chelating groups for the immobilization of Zr41 in the new adsorbents and the immobilization of Fe31 in the conventional IMAC adsorbents are shown in Fig. 2. In conventional IMAC, metal ion like Fe31 was immobilized by use of chelating groups such as iminodiacetic acid. However, in the new IMAC, the Zr41 was immobilized by using a phosphate group. The adsorbent surface was first activated to covalently link a phosphate group and then Zr41 was immobilized by incubation with ZrOCl2 solution. Compared with conventional Fe31-IMAC, Zr41-IMAC showed higher specificity for isolation of phosphopeptides. And the high specificity of Zr41-IMAC adsorbent mainly resulted from strong interacwww.proteomics-journal.com 692 X. Jiang et al. Proteomics 2008, 8, 686–705 Table 1. Summary of phosphopeptide enrichment methods Methods IMAC Fe31/Ga31/Al31-IMAC Esterification prior to IMAC Zr41-IMAC Metal oxide ZrO2 /TiO2 /Al(OH)3 Ion-exchange chromatography SCX SAX Chemical modification ß-elimination Phosphoamidate Strength and weakness Reference The conventional and most popular method; using iminodiacetic acid or nitriloacetic acid as chelating group; lacking enough specificity due to binding very acidic peptides. Specificity is improved, however sample complexity may be increased because of incomplete and side reactions. A new type of IMAC using phosphate group as chelating group; Specificity higher than conventional IMAC. [39, 40, 114, 115] Specificity higher than conventional IMAC; may have steric hindrance to bind big phosphopeptides due to the absence of space arm. [45-47] Able to enrich and fractionate phosphopeptides; relative low specificity; tend to loss of multiple phosphorylated peptides due to weak interaction. Able to enrich and fractionate phosphopeptides; relative high specificity and recovery; low resolution for fractionation. [48, 49] Only limited to phosphorylated serine or threonine residue; low specificity because the interference of O-linked glycopeptides. Applicable to all phosphopeptides; low yield because of multi-step derivatization. [116] [41] [44] [50, 51] [117] 3.1.2 Metal oxide particles Metal oxide microparticles, such as titanium oxide, TiO2, [45], zirconium dioxide, ZrO2, [46, 47] have been proved to have much higher selectivity for trapping phosphopeptides than conventional IMAC beads. Similar to IMAC, the mechanism of using metal oxides for phosphopeptide isolation is also based on the strong interaction between metal ions and phosphopeptides. As there is no spacer arm on metal oxide beads, one disadvantage of using metal oxide beads for phosphopeptide enrichment, compared with IMAC, is the presence of steric hindrance. The phosphate groups of some large size phosphopeptides may have difficulty in accessing the surface of metal oxide particles. Figure 2. Schematic illustration of (A) conventional Fe31-IMAC and (B) novel Zr41-IMAC adsorbents for binding of phosphopeptides tion between chelating Zr41 and phosphate groups on phosphopeptides. The high specificity of Zr(41)-IMAC adsorbent was demonstrated by effectively enriching phosphopeptides from the digest mixture of phosphoprotein (alpha- or betacasein) and BSA with molar ratio at 1:100. It was also successfully applied for the analysis of mouse liver phosphoproteome, resulting in the identification of 153 phosphopeptides (163 phosphorylation sites) from 133 proteins in mouse liver lysate. © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim 3.1.3 Ion-exchange chromatography Compared with unmodified peptides, phosphopeptides have extra negatively charged groups. Thus phosphopeptides could be enriched based on their different electrostatic interaction with stationary phase in ion-exchange chromatography. Recently, ion-exchange chromatography is applied to simultaneously enrich and fractionate phosphopeptides. Phosphopeptides have weak interaction with SCX because of the phosphate groups, and so they will elute in early fractions in SCX and thereby can be separated from non-phosphopeptides which are strongly retained on SCX [48, 49]. It was found that the first four SCX fractions were highly enriched www.proteomics-journal.com Technology Proteomics 2008, 8, 686–705 with phosphopeptides. Though some phosphopeptides could be enriched and fractionated by SCX, this approach was not very specific as strong acidic peptides were also eluted in early fractions. Because some phosphopeptides, especially those with multiple phosphate groups, have net negative charge and will not bind on SCX column, SCX approach will result in loss of these phosphopeptides. Using of strong anion exchange chromatography (SAX) is another approach to enrich and fractionate phosphopeptides. Nüshe et al. [50] used SAX chromatography based on salt gradient as a rough pre-fractionation approach before IMAC for the analysis of phosphopeptides from Arabidopsis, and found the 2-D separation decreased the complexity of the phosphopeptide sample and yielded a far greater coverage of phosphorylated peptides. Dai et al. [51] applied SAX column to enrich the phosphopeptides from the flow through of SCX column and the bound phosphopeptides were fractionated by pH step gradient. Recently, the performance of using SAX for enrichment and fractionation of phosphopeptides were systematically investigated [52]. It was found that nearly only phosphopeptides were retained on the SAX column and the majority of non-phosphopeptides were removed. The SAX column was further applied to enrich and fractionate phosphopeptides in tryptic digest of proteins extracted from human liver tissue adjacent to tumorous regions for large scale phosphoproteome analysis. This resulted in the identification of hundred of phosphorylation sites from phosphoproteins. Techniques such as using IMAC and metal oxide particles were only applied to enrich phosphopeptides when only one phosphopeptide fraction was obtained. The phosphopeptide mixture enriched from proteome samples was still very complex. In contrast, the distinct advantage of using ion-exchange chromatography for phosphoproteome analy- 693 sis was that it could not only enrich phosphopeptides but also fractionated phosphopeptides. Thus it is very suitable for large-scale phosphoproteome analysis. 3.1.4 Chip-based methods Chip-based methods for analysis of phosphopeptides is a rather new approach [53–55], which could decrease sample loss, simplify analytical procedures, and finally achieve highthroughput detection by MALDI-TOF MS. Xu et al. [53] prepared porous silicon wafer with immobilized Fe31 affinity surface to analyze phosphopeptides by MALDI-TOF MS. Complex peptide mixtures were spotted onto the Fe31 immobilized porous silicon wafer, and non-phosphopeptides were removed from the silicon surface by thorough washing. After addition of matrix, the porous silicon wafer was directly placed on the MALDI target for the analysis of the captured phosphopeptides. The phosphopeptide enrichment and analysis procedures were all performed on the Fe31-terminated silicon wafer, which greatly reduced the sample loss and simplified the analysis procedure. Target purification of phosphopeptides followed by MALDI MS analysis was also reported by Dunn et al. [55]. Above approaches were based on the interaction between chelated Fe31 ion and the phosphate group in phosphopeptides. Recently, Zhou et al. [54] prepared porous silicon wafer with surface immobilized with Zr41 for phosphopeptide analysis. Figure 3 shows the scheme for the preparation of Zr41-terminated silicon wafer to trap phosphopeptides for MALDI-TOF MS analysis. The excellent selectivity of this approach was demonstrated by analyzing phosphopeptides in the digest mixture of betacasein and BSA with molar ratio of 1:100. Figure 3. Scheme for preparation of zirconium phosphonate modified porous silicon (ZrP-pSi) to trap phosphopeptides for MALDI-TOF MS analysis. Reprinted from [54] © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.proteomics-journal.com 694 X. Jiang et al. Proteomics 2008, 8, 686–705 3.2 Isolation of glycopeptides The phosphate group is quite small and may be buried inside of proteins. The phosphopeptide enrichment methods are typically not applicable for enrichemnt of phosphoproteins because of steric hindrance. However, the situation for protein glycolation is completely different. The attached carbohydrate group is much bigger and is more accessible. Thus the methods to isolate glycopeptides are often equally effective for the isolation of glycoproteins. In practice, the sequential use of the same method to glycoproteins and glycopeptides is often applied to improve glycopeptide enrichment specificity. This is especially true for affinity chromatography. 3.2.1 Lectin affinity chromatography Lectin affinity chromatography has been reported numerous times as a classic and efficient method for capturing glycoproteins as well as glycopeptides [56]. As shown in Fig. 4, a protocol based on tandem lectin affinity chromatography purification on the protein and peptide level has been reported to improve the specificity for glycopeptide isolation [57, 58]. Glycoproteins were first enriched from complex protein mixture by a lectin affinity chromatography. The obtained glycoproteins were then proteolyzed with trypsin and the resulting peptide mixture was subjected to affinity chromatography with the same lectin column to enrich glycopeptides. After deglycosylation of the glycopeptides, the resuting peptides were subjected to LC-MS/MS analysis for the identification of glycopeptides. Concanavalin A (ConA) lectin chromatography has been used most widely for isolation of glycoproteins/glycopeptides with high mannose-type and hybrid-type oligosaccharides. Kaji et al. [57] applied ConA lectin affinity chromatography for the large-scale identification of N-glycosylated proteins from Caenorhabditis elegans. After the tandem enrichment step, the purified glycopeptides were treated with PNGase F (a glucosidase specifically cleaving N-linked glycans) to remove N-linked glycans. The resulting peptides were analyzed by LC-MS/MS which resulted in the identification of 250 glycoproteins with the simultaneous determination of 400 unique N-glycosylation sites. Besides ConA, several other kinds of lectins were also utilized to enrich glycoproteins/glycopeptides having distinct types of carbohydrate structures as shown in Table 2. As glycoproteins/glycopeptides could be isolated based on their different glycan moieties through lectin specificity, the ability of different lectins to recognize specific glycosylation motifs was utilized to build a multi-lectin affinity platform for comprehensively capturing glycoproteins/glycopeptides of the proteome sample. Yang et al [56] reported one of the first large-scale glycoproteomic experiments utilizing multi-lectin (ConA, WGA, jacalin) affinity chromatography, analyzing human serum plasma and identifying approximately 150 different glycosylated proteins. In this approach, glycopro© 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Figure 4. Tandem lectin affinity chromatography approach to generate glycopeptides for glycoproteomics analysis. teins were isolated by agarose-bound lectins immobilized within the column and eluted following purification using a solution containing simple sugars. 3.2.2 Chemical approaches Even though lectin affinity capture is the most widely used approach due to its ease of implementation, the binding selectivity of lectins to specific conformations of different carbohydrate moieties has limited the utility of lectin in global glycoprotein analysis [59, 60]. Chemical capture approaches, which allowed the capture of all types of glycoproteins or glycopeptides, were developed [61]. As shown in Fig. 5, two type of chemical capture approaches based on hydrazide chemistry were reported. The first one captured glycoproteins and the second one captured glycopeptides. The final products of these two approaches were the same: the deglycosylated glycopeptides. The glycoprotein capture strategy was initially developed by Zhang et al. [62]. In this approach, covalent conjugation of glycoproteins to a solid support via hydrazide chemistry was used. Glycoprotein oxidation was carried out in order to convert the cis-diol groups of carbohydrates to aldehydes. The aldehyde groups reacted with the hydrazide groups immobilized on the resin, forming covalent hydrazone bonds. Thus glycoproteins were immobilized on hydrazide beads through the carbohydrate group. After immobilization, the resins were subjected to thorough washing to remove non-glycosylated proteins. The immobilized glycoproteins were then proteolyzed on the solid support. Non-glycosylated peptides were again washed away while the glycosylated peptides remained on the solid support. The formerly N-linked glycosylated peptides were finally released from the solid support www.proteomics-journal.com Technology Proteomics 2008, 8, 686–705 695 Table 2. Different lectin for isolating glycoproteins/glycopeptides of distinct types of carbohydrate structures Lectin type Distinct types of carbohydrate structures References Concanavalin A (ConA) Wheat germ agglutinin (WGA) Anguilla anguilla agglutinin (AAA) Sambucus nigra (SNA) Aleuria aurantia lectin (AAL) Helix pomatia agglutinin (HPA) Peanut agglutinin (PNA) High mannose-type and hybrid-type N-acetyl-glucosamine and sialic acid type Broad specificity for L-Fuc-containing glycans a-2,6-linked sialic acid residues Broad specificity for L-Fuc-containing glycans N-acetylgalactosamine residues Specific to T-antigen found commonly in O-glycans [56, 57] [118, 119] [58, 120] [58, 121] [122, 123] [58] [122, 124] Figure 5. Schematic illustrations of (A) glycoprotein capture strategy and (B) glycopeptide capture strategy for glycoproteomics analysis using hydrazide chemistry. using PNGaseF. This strategy was successfully applied to the analysis of cell surface proteins and for serum proteome profiling [63]. The glycopeptide capture strategy was reported recently by Sun et al. [64], which was quite similar to the glycoprotein capture strategy. As shown in Fig. 5, in this strategy the glycoproteins were first digested into peptides and the glycosylated peptides in the resulting digest were then immobilized onto the solid support by hydrazide chemistry. Other steps were essentially the same to that of the glycoprotein capture © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim approach. Digestion of proteins into peptides improved solubility of large membrane proteins and exposed all of the glycosylation sites to ensure equal accessibility to external capture reagents. Therefore, more glycopeptides should be captured and the capture efficiency should be improved greatly. The glycopeptide capture strategy was validated using standard protein mixtures that resulted in close to 100% capture efficiency [64]. Using this approach, glycopeptide selectivity as high as 91% was achieved on the microsomal fraction of the ovarian-cancer cells. www.proteomics-journal.com 696 X. Jiang et al. 3.2.3 Other approaches Besides lectin affinity chromatography and hydrazide chemistry, different chromatographic separation techniques were also used to enrich glycopeptides by the diverse physical and chemical properties of glycopeptides [65–69]. Hydrophilic interaction chromatography was applied to enrich glycopeptides because the carbohydrate group was very hydrophilic. For example, Wada et al. [69] reported a simple method, utilizing hydrophilic binding of carbohydrate matrixes such as cellulose and sepharose to oligosaccharides, to the isolation of tryptic glycopeptides from some standard glycoproteins. Both peptide and oligosaccharide structures were elucidated by multiple-stage tandem MS (MSn) of the ions generated by MALDI. Hagglund et al. [67] developed a novel approach to identify N-glycosylation sites in complex biological samples by enrichment of glycosylated peptides through hydrophilic interaction chromatography followed by partial deglycosylation, which removed the major part of the glycan and thus simplified the MS/MS spectra of glycopeptides. Another interesting technique for enrichment of glycopeptides is the use of SEC [63]. An in-silico trypsin digest of all the human protein sequences in the NCBI database, over 90% of the peptides displayed masses that were smaller than 2000 Da. Considering that the mass of the smallest N-linked oligosaccharide was over 1200 Da, most N-linked glycopeptides should be significantly larger than the non-glycosylated peptides. Therefore, Alvarez-Manilla et al. [65] have investigated using SEC for enrichment of glycopeptides. Analyses performed on human serum showed that this SEC glycopeptide isolation procedure resulted in at least a three-fold increase in the total number of glycopeptides identified by LC-MS/MS, demonstrating that this method is an effective tool to facilitate the identification of glycopeptides. Besides, boronate affinity chromatography [70] and graphite chromatography [68] could also be applied to enrich glycopeptides. Although these chromatographic techniques do not involve multiple chromatographic steps and do not require derivatization of glycoproteins/glycopeptides compared with affinity chromatography and hydrazide chemistry, these approaches often lack enough specificity for glycoproteomic analysis. The application of these approaches is typically limited to analysis of simple samples where only one or several glycoproteins are present. 4 Isolation of endogenous peptides for peptidome analysis Endogenous peptides play a central role in many biological processes and some classes of such biologically active peptides, e.g. hormones, cytokines and growth factors, have been known and studied for years. The term ‘peptidomics’ was introduced in 2001 to define the quantitative and qualitative analysis of endogenous peptides in biological samples [71, 72], primarily by chromatography or biochip platforms cou© 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Proteomics 2008, 8, 686–705 pled with various forms of MS. The pacemakers for the development of peptidomic technologies are modern MS and bioinformatics. They are ideally suited for sensitive and comprehensive peptide analysis, especially in combination with the massive information content of today’s genomic and transcriptomic databases. However, the complexity and high dynamic range of biological samples make the characterization of endongenous peptides a challenging task. To reduce the supression of endongenous peptides by high abundance proteins, several methodologies are currently used to enrich peptides from biological samples. 4.1 Centrifugal ultrafiltration Centrifugal ultrafiltration with accurate MW cutoff is the most widely used method to extract peptides and remove proteins with larger MWs based on a size-exclusion mechanism [73–77]. Zheng et al. [77] utilized the protocol of centrifugal ultrafiltration for identifying endogenous peptides from serum, and the identification of over 300 unique peptides with 2 ppm or better mass accuracy per serum sample was achieved. However, few peptides over 3000 Da were directly identified with the above approach. Therefore, Hu et al. [78] developed a method for comprehensive peptidomic analysis, which combined ultrafiltration with SEC chromatography pre-fractionation. The peptides in mouse liver were first harvested by ultrafiltration with 10 kDa molecular weight cut off, and then they were prefractionated by SEC. The low MW peptides (MW,3000 Da) in the collected fractions were directly analyzed by LC-MS/MS which resulted in the identification of 1181 unique peptides (from 371 proteins). The high MW peptides (MW.3000 Da) in the early two fractions from SEC column were first digested with trypsin and then the resulting digests were analyzed by LCMS/MS, which leaded to the identification of 123 and 127 progenitor proteins of the high MW peptides in the two fractions, respectively. The low MW peptides were separated from the high MW ones, which improved the sensitivity for identification of the low MW peptides and allowed in-depth investigation of the high MW peptides. However, as a result of high protein content in biological sample, the ultrafiltration time will increase sharply if a large amount of biological sample is applied. Furthermore, other low MW contaminants (for example, salts) will also be concentrated which may compromise the performance for peptidome analysis. Instead of using ultrafiltration, another option is to use adsorbents carrying charged or hydrophobic groups for peptide enrichment. 4.2 Functionalized adsorbents Functionalized magnetic particle is one of the most widely used adsorbents for peptide enrichment [79–82]. Recently, Fiedler et al. [83] developed a standardized protocol for reproducible urine peptidome profiling by means of extracting www.proteomics-journal.com Technology Proteomics 2008, 8, 686–705 peptides with magnetic beads with defined surface functionalities (hydrophobic interaction, cation exchange, and metal ion affinity) followed by MALDI-TOF MS analysis, and 427 different mass signals in the urine of healthy donors were detected. In addition to functionalized magnetic particle, conventional chromatographic packing materials such as RP polymers were also used to enrich peptides [73, 84]. Because of high specific surface, nanoporous material [85, 86] and nanomaterial [87, 88] are also good adsorbents for peptide enrichment. For example, Li et al. [89] described a peptidome analysis approach using multiwalled carbon nanotubes as an alternative adsorbent to capture endogenous peptides from human plasma. In total, 374 unique peptides were identified with high confidence by 2-D LC-MS/MS analysis. And comparative studies showed that multiwalled carbon nanotubes were superior to C18 and C8 silica particles for the capture of the smaller peptides. Although the peptides can be enriched by these adsorbents, some proteins may also be enriched which suppress the detection of peptides. A special type of chromatographic packing materials called restricted access materials (RAMs) [90, 91] could minimize the adsorption of proteins. These porous packings have a bimodal surface topochemistry and have two functions during chromatographic separation: (i) SEC, i.e. macromolecular sample components like proteins having a MW larger than, e.g. 1500 Da can be directly eluted to waste in the dead volume of such a RAM column and (ii) adsorption chromatography by RP, ion-exchange or affinity chromatography, i.e. low MW sample components such as drugs and peptides can be bound and extracted. RAMs, moreover, have a biocompatible outer surface tolerating frequent injections of raw biofluids without significant loss of their mixed mode chromatographic properties. Because of these characteristics, RAMs are especially suitable for on-line extraction of low MW proteins and peptides respectively, as well as depletion of abundant high MW proteins [92]. Unger et al. [93–95] described a rather complex system for on-line extraction and multidimensional separation of peptides in human hemofiltrate and cell lysates applying the SPE column packed with RAMs and coupled to a conventional separation column. Recently, a capillary SPE column packed with RAM exhibiting SCX and size exclusion (SCX/ SEC) properties was coupled with a nano-LC-MS/MS system for peptidomic analysis of serum sample [96]. The capillary SPE column excluded serum proteins having a molecular mass larger than 1500 Da by SEC and simultaneously extracts peptides by SCX mechanism. After injection of 2 mL of human serum to the 1-D nanoLC-MS system, around 400 peptides could be identified. Similar to using RAM, Tian et al. [97] attempted to use mesoporous materials with critical pore sizes to selectively enrich peptides from human plasma but exclude other proteins by an accurate MW cutoff like that of centrifugal ultrafiltration. It was found that MCM-41 with a pore size of 20.5 Å was effective for enriching peptides in human plasma with a © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim 697 wide MW range from 1 to 12 kDa (Figs. 6A and C), while repelling most other plasma proteins outside, as shown in Figs. 6B and D. The unique pore structure of this material made it superior for peptide enrichment when compared to both adsorbent- and ultrafiltration-based methods. 5 Some interesting on-line sample pretreatment approaches 5.1 Automated sample injection Although mHPLC-MS/MS is the dominant platform for proteomic analysis, automation of sample introduction onto analytical column without seriously compromising the separation performance is very difficult. In typical cases, proteomic sample size ranges from a few microliters to 100 microliters. It will take a long time for sample loading if the samples are directly loaded onto capillary analytical column in nanoflow HPLC. The automation could be realized by different instrument configurations by using a short and large id. C18 trap column coupled with a C18 analytical column for rapid sample loading. Briefly, protein digests of large volumes are firstly loaded onto the trap column at a high flow rate in a short time and after equilibrium the adsorbed peptides are eluted from the trap column to the RP analytical column. Two types of instrument configurations have been adopted for the automated sample injection in mHPLC-MS/MS using trap column. One is directly connecting the trap column and C18 analytical column by a nanoflow switching valve. The flow through from trap column is directed to waste or analytical column during sample injection or separation by the switching valve [98–100]. In this type of system, proteolytic digest without prior purification could be directly injected and cleaned up online, which is very attractive in comprehensive proteome analysis. But the void volume introduced by switching valve would degrade the separation performance greatly. The other is a vented column system in which trap and analytical columns are directly connected via a microcross or microtee with an open/ close switching valve [101–104]. In this type of system, the mobile phase for mHPLC separation does not pass through the switching valve, so a regular six-port switching valve could be used instead of using a nanoflow switching valve. In order to minimize the void volume resulted from the microcross or microtee, Licklider et al. [102] packed the open space of microcross with C18 particles, and Meiring et al. [103] drilled the microtee to 0.6 mm id to fit a single micro sleeve having a V-shaped cut as a waste outlet and the trapping and analytical columns were butt-connected in this modified sleeve. However, these systems largely depend on experience and are not widely used. Another disadvantage of vented column system is that proteolytic digests containing denaturing agents can not be loaded directly. This is because a small portion of the sample solution will also enter the analytical column and contaminate the column during loading www.proteomics-journal.com 698 X. Jiang et al. Proteomics 2008, 8, 686–705 Figure 6. MALDI-TOF MS analysis of human plasma (A, B) before and (C, D) after exposure to MCM-41 by using a-cyano-4-hydroxycinnamic acid as matrix. Analysis in the MW range of (A, C) 1–15 kDa and (B, D) 10–100 kDa. Reprinted from [97]. of sample onto the trap column. So proteolytic digests must be desalted and cleaned before the automated sample introduction. Although the above systems enable the automated sample injection for proteome analysis by mHPLC-MS/MS, void volume resulted from the connections between trap column and analytical capillary column which inevitably leads to the degradation of separation performance. As further decreasing the void volume is a technique challenge, a good solution is to develop a void volume insensitive automated sample injection system for mHPLC-MS/MS analysis. All above systems use C18 trap column. It was found recently that automation of sample injection using SCX trap column instead of C18 trap column could alleviate the influence of void volume on separation [101, 105]. In this approach, protein digest was first loaded onto a SCX trap column, the captured peptides were then eluted onto C18 analytical column by a high salt buffer for separation. Because the peptide sample was retained on the entrance end of the C18 analytical column before the gradient started separation, the void volume before the capillary column hardly affected the separation adversely. Recently, the influence of the void volume between © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim trap column and analytical column were systematically investigated and no obvious degradation of the proteome analysis performance was observed even with a void volume as large as 5 mL. Similar to using C18 trap column, the automation of the sample injection using SCX trap column could also be realized by the two configurations, i.e. directly using the nanoflow switching valve [105] and using the vented column configurations [101]. The schematic diagrams for these two instrument configurations are shown in Fig. 7. The first configuration allowed direct injection of detergent containing sample, while the second one had the advantage of overall short separation time because of its small void volume. Besides 1-D separation, the SCX trap column systems could also be conveniently applied for large scale proteome and peptidome analysis of complex samples with multi-dimensional separation [89, 97, 101]. To load sample at high flow rate, low back pressure of the trap column is preferable. Instead of using a packed trap column, a phosphate monolithic trap column was prepared and used for automatic sample injection in nanoLC-MS/MS system[106]. Because of its low back pressure, sample injection at a high flow rate of 40 mL/min could be easily realized. www.proteomics-journal.com Proteomics 2008, 8, 686–705 Technology 699 Figure 7. Schematic diagrams of on line sample injection systems using SCX trap column (A): directly using the nanoflow switching valve [105]; (B): using the vented column [101] 5.2 On-line digestion by immobilized enzyme reactor Usually, tryptic digestion of protein is performed either in gel or in solution. However, these methodologies have several drawbacks, such as long digestion time (typically .5 h), auto-proteolysis of trypsin and the difficulty for automation. On the contrary, the trypsin immobilized on matrixes has the advantages of high efficiency, high stability and easy to automation [107–110]. Schriemer et al. [107, 108] demonstrated on-line, real-time tryptic digestion of proteins by a packed immobilized trypsin cartridge, directly followed with RP protein separation. Using test mixtures of standard proteins, peptide mass fingerprinting with high sequence coverage could be easily achieved at the 20 fmol level, with detection limits down to 5 fmol. With the rapid development of monolithic column technologies, immobilized enzyme monolithic reactors were prepared for on-line digestion of proteins. For example, trypsin-based monolithic col© 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim umn with dimension of 50 mm64.6 mm id was prepared and applied to automated digestion of protein and protein identification for proteome analysis [111]. However, the large size of the reactor was not suitable for use in a nanoflow LC-MS system. Therefore, Feng et al. [112] prepared a nanoliter trypsin microreactor, and coupled it on-line with mRPLC-MS/MS for the analysis of the total cell lysate of Saccharomyces cerevisiae. After database search, a total of 1578 unique peptides corresponding to 541 proteins were identified when 590 ng yeast protein was digested by the microreactor with an incubation time of only 1 min. Recently, a fully automated CE-Microreactor-CE-MS/MS system was presented for protein ananlysis [113]. In this system, proteins were first separated by CE. A monolithic pepsin microreactor was incorporated into the distal end of this capillary. Peptides formed in the reactor were transferred to a second capillary, where they were separated by CE and characterized by MS. www.proteomics-journal.com 700 6 X. Jiang et al. Conclusion and perspective As there are a lot of redundant peptides presented in a peptide mixture, isolation of a subset of interested peptides represented an efficient approach to simplify the complexity of proteome analysis. Therefore, the key to this approach is to significantly reduce the sample complexity and meanwhile keep the integrity of the proteome. At present, isolation of rare amino acid-containing peptides, especially Cys-containing peptides, has already been proved to be an effective method to reduce sample complexity. However, the disadvantage for isolation of rare amino acid-containing peptides is that the distribution of these rare amino acid residues among different proteins is not even. Therefore, some proteins have many representative peptides while others have a few or even no representative peptides, proteome analysis by this approach bias to proteins with high percentage of this type of rare amino acid residues. In contrast, terminal peptides are the most evenly distributed peptides as every protein has one N-terminal peptide and one C-terminal peptide. Reducing the sample complexity by isolating terminal peptides has drawn strong attention since Gevaert et al. [26] published the first paper in 2003 on isolation of N-terminal peptides for proteome analysis. Up to now, several approaches were developed to isolate N-terminal peptides. It should be mentioned that identification of protein by only one peptide is not always successful. This happens because the peptide may not be the right size for MS/MS analysis or the peptide has some unknown modifications. To solve this problem, PST technology reported by Kuhn et al. [24–26] is an alternative way. However, it also depends on the distribution of Met-residues among proteins. An ideal approach to reduce the sample complexity for proteome analysis may be to isolate both N-terminal and C-terminal peptides. However, there is still a long way to go before this approach is applicable. The specificity for isolation of N-terminal peptides should be improved and an effective method to isolate C-terminal peptides should be developed. Phosphoproteome analysis depends heavily on LC-MS/ MS analysis of phosphopeptides. Because of low stoichiometry of protein phosphorylation, phosphopeptides always co-exist with huge amounts of non-phosphopeptides which seriously depress the detection of phosphopeptides. To specifically isolate phosphopeptides is still one of the key issues for phosphoproteome analysis. Use of metal oxide particles such as ZrO2 and TiO2 has already proved to be more specific than conventional IMAC with Fe31. If Zr41 or Ti41 is immobilized onto chromatographic support via space arm, the resulting new IMAC may have higher performance for isolation of phosphopeptides than corresponding metal oxides because the steric hindrance will be reduced. We already demonstrated that the new IMAC with Zr41 had excellent specificity for isolation of phosphopeptides [44]. Also, the new IMAC with Ti41 has been under investigation in our lab and the preliminary results are promising. The affinity chromatography with immobilized Zr41 or Ti41 might © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Proteomics 2008, 8, 686–705 become a new generation of high performance IMAC for specific isolation of phosphopeptides. As the phosphopeptide mixture enriched by the majority of currently available phosphopeptide enrichment methods is still very complex, fractionation of phosphopeptides prior to LC-MS/MS analysis is beneficial for large scale phosphoproteome analysis. Although phosphopeptides could be fractionated by SAX or SCX, their performance is far from ideal. High resolution techniques for fractionation of phosphopeptides need to be developed for comprehensive phosphoproteome analysis. Glycoproteomics analysis also relies on analysis of glycopeptides. Up to now, using lectin affinity chromatography and hydrazide beads are the two most effective approaches to isolate glycopeptides. Both approaches have its strengths and weaknesses. Lectin affinity chromatography approach keeps the carbohydrate moiety intact which makes characterization of the carbohydrate moiety possible. However, the carbohydrate moiety will be destroyed during chemical reaction with the hydrazide chemistry approach. The advantage of hydrazide chemistry approach is its extremely high specificity as the glycopeptide selectivity as high as 91% could be achieved [64]. But the specificity for lectin affinity chromatography was relative poor even when a tandem purification approach was applied. Hydrazide chemistry approach can capture all types of glycopeptides and so can be applied to analyze all types of glycosylation in theory. However, majority of applications are limited in analysis of N-linked glycosylation because of the formally N-linked glycosylated peptides could be easily cleaved from the beads by PNGaseF. In order to analyze other types of glycosylation by hydrazide chemistry approach, the methods to specifically cleave the corresponding peptides from the beads should be developed. For peptidome analysis, one of the key challenges is to isolate endogenous peptides from biologic sample where huge amounts of protein are presented. As the only difference between peptide and protein is the size, it is preferable to use size based separation approach to isolate endogenous peptides. RAMs have a bimodal surface topochemistry and have functions of both SEC and adsorption chromatography, thus it is a good approach for isolation of peptides. Application of new materials such as mesoporous materials with critical pore sizes for selectively enriching peptides but exclude other proteins by an accurate MW cutoff represents a new trend to isolate peptides [97]. Similar to RAM, to further improve peptide isolation efficiency, derivatization of inner surface and outer surface of these materials with different functionary groups is required. The inner surface should be derivatized with stationary phase such as RP, ion-exchange, etc. to bind peptides while the outer surface should be derivatized with hydrophilic groups to minimize the non-specific adsorption of proteins. Although sample pretreatment technologies have made big progress in the last decade, more improvements are required to further overcome the complexity and dynamic range problem for proteomics analysis. For selective enrichment of a subset of interested peptides, specificity should be www.proteomics-journal.com Proteomics 2008, 8, 686–705 improved to enrich the target peptides. This field will still be a vital part of proteomics, which is available to reduce proteome complexity substantially and simplify peptide identification. Moreover, development of high efficient and automated platforms, including protein digestion, desalting of sample, introduction of sample, etc, is urgently needed for proteomic analysis. With the development of new sample pretreatment methods, proteome analysis could mine deeper and more insights for biological and medical research in the future. Financial supports from the National Natural Sciences Foundation of China (20735004, 20675081), the China State Key Basic Research Program Grant (2005CB522701), the China High Technology Research Program Grant (2006AA02A309), and the Knowledge Innovation program of CAS (KJCX2.YW.HO9, KSCX2-YW-R-079) and the Knowledge Innovation program of DICP to H.Z. and National Natural Sciences Foundation of China (No. 20605022, 90713017) to M.Y. are gratefully acknowledged. The authors have declared no conflict of interest. 7 References [1] Rabilloud, T., Two-dimensional gel electrophoresis in proteomics: Old, old fashioned, but it still climbs up the mountains. Proteomics 2002, 2, 3–10. [2] Steen, H., Mann, M., The ABC’s (and XYZ’s) of peptide sequencing. Nat. Rev. Mol. Cell Biol. 2004, 5, 699–711. [3] Washburn, M. P., Wolters, D., Yates, J. R., 3rd, Large-scale analysis of the yeast proteome by multidimensional protein identification technology. Nat. Biotechnol. 2001, 19, 242– 247. [4] Link, A. J., Eng, J., Schieltz, D. M., Carmack, E. et al., Direct analysis of protein complexes using mass spectrometry. Nat. Biotechnol., 1999, 17, 676–682. [5] Azarkan, M., Huet, J., Baeyens-Volant, D., Looze, Y. et al., Affinity chromatography: a useful tool in proteomics studies. J. Chromatogr. B 2007, 849, 81–90. Technology 701 [11] Zhou, H., Ranish, J. A., Watts, J. D., Aebersold, R., Quantitative proteome analysis by solid-phase isotope tagging and mass spectrometry. Nat. Biotechnol. 2002, 20, 512–515. [12] Wang, S., Regnier, F. E., Proteomics based on selecting and quantifying cysteine containing peptides by covalent chromatography. J. Chromatogr. A 2001, 924, 345–357. [13] Gevaert, K., Ghesquiere, B., Staes, A., Martens, L. et al., Reversible labeling of cysteine-containing peptides allows their specific chromatographic isolation for non-gel proteome studies. Proteomics 2004, 4, 897–908. [14] Wang, H., Qian, W. J., Chin, M. H., Petyuk, V. A. et al., Characterization of the mouse brain proteome using global proteomic analysis complemented with cysteinyl-peptide enrichment. J. Proteome Res. 2006, 5, 361–369. [15] Weinberger, S. R., Viner, R. I., Ho, P., Tagless extractionretentate chromatography: a new global protein digestion strategy for monitoring differential protein expression. Electrophoresis 2002, 23, 3182–3192. [16] Gevaert, K., Van Damme, J., Goethals, M., Thomas, G. R. et al., Chromatographic isolation of methionine-containing peptides for gel-free proteome analysis: identification of more than 800 Escherichia coli proteins. Mol. Cell. Proteomics 2002, 1, 896–903. [17] Kuyama, H., Watanabe, M., Toda, C., Ando, E. et al., An approach to quantitative proteome analysis by labeling tryptophan residues. Rapid Commun. Mass Spectrom. 2003, 17, 1642–1650. [18] Matsuo, E., Toda, C., Watanabe, M., Lida, T. et al., Improved 2-nitrobenzenesulfenyl method: Optimization of the protocol and improved enrichment for labeled peptides. Rapid Commun. Mass Spectrom. 2006, 20, 31–38. [19] Foettinger, A., Leitner, A., Lindner, W., Selective enrichment of tryptophan-containing peptides from protein digests employing a reversible derivatization with malondialdehyde and solid-phase capture on hydrazide beads. J. Proteome Res. 2007, 6, 3827–3834. [20] Ji, J., Chakraborty, A., Geng, M., Zhang, X. et al., Strategy for qualitative and quantitative analysis in proteomics based on signature peptides. J. Chromatogr. B 2000, 745, 197–210. [21] Ren, D., Penner, N. A., Slentz, B. E., Inerowicz, H. D. et al., Contributions of commercial sorbents to the selectivity in immobilized metal affinity chromatography with Cu(II). J. Chromatogr. A 2004, 1031, 87–92. [6] Lescuyer, P., Hochstrasser, D. F., Sanchez, J. C., Comprehensive proteome analysis by chromatographic protein prefractionation. Electrophoresis 2004, 25, 1125–1135. [22] Ren, D., Penner, N. A., Slentz, B. E., Mirzaei, H. et al., Evaluating immobilized metal affinity chromatography for the selection of histidine-containing peptides in comparative proteomics. J. Proteome Res. 2003, 2, 321–329. [7] Ye, M., Jiang, X., Feng, S., Tian, R. et al., Advances in chromatographic techniques and methods in shotgun proteome analysis. Trac-Trend. Anal.Chem. 2007, 26, 80–84. [23] Ren, D., Penner, N. A., Slentz, B. E., Regnier, F. E., Histidinerich peptide selection and quantification in targeted proteomics. J. Proteome Res. 2004, 3, 37–45. [8] Zhang, H., Yan, W., Aebersold, R., Chemical probes and tandem mass spectrometry: a strategy for the quantitative analysis of proteomes and subproteomes. Curr. Opin. Chem. Biol. 2004, 8, 66–75. [24] Noubhani, A. M., Dieryck, W., Bakalara, N., Latxague, L. et al., Evaluation of chromatographic recycling for imidazole used in the chromatographic purification of His-tag recombinant proteins. J. Chromatogr. B 2003, 790, 153–159. [9] Leitner, A., Lindner, W., Current chemical tagging strategies for proteome analysis by mass spectrometry. J. Chromatogr. B 2004, 813, 1–26. [25] Wang, S., Zhang, X., Regnier, F. E., Quantitative proteomics strategy involving the selection of peptides containing both cysteine and histidine from tryptic digests of cell lysates. J. Chromatogr. A 2002, 949, 153–162. [10] Gygi, S. P., Rist, B., Gerber, S. A., Turecek, F. et al., Quantitative analysis of complex protein mixtures using isotopecoded affinity tags. Nat. Biotechnol. 1999, 17, 994–999. © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim [26] Gevaert, K., Goethals, M., Martens, L., Van Damme, J. et al., Exploring proteomes and analyzing protein processing by www.proteomics-journal.com 702 X. Jiang et al. mass spectrometric identification of sorted N-terminal peptides. Nat. Biotechnol. 2003, 21, 566–569. [27] Aivaliotis, M., Gevaert, K., Falb, M., Tebbe, A. et al., Largescale identification of N-terminal peptides in the halophilic archaea Halobacterium salinarum and Natronomonas pharaonis. J. Proteome Res. 2007, 6, 2195–2204. Proteomics 2008, 8, 686–705 [42] Moser, K., White, F. M., Phosphoproteomic analysis of rat liver by high capacity IMAC and LC-MS/MS. J. Proteome Res. 2006, 5, 98–104. [43] Kim, J. E., Tannenbaum, S. R., White, F. M., Global phosphoproteome of HT-29 human colon adenocarcinoma cells. J. Proteome Res. 2005, 4, 1339–1346. [28] McDonald, L., Robertson, D. H., Hurst, J. L., Beynon, R. J., Positional proteomics: selective recovery and analysis of Nterminal proteolytic peptides. Nat. Methods 2005, 2, 955– 957. [44] Feng, S., Ye, M., Zhou, H., Jiang, X. et al., Immobilized zirconium ion affinity chromatography for specific enrichment of phosphopeptides in phosphoproteome analysis. Mol. Cell. Proteomics 2007, 6, 1656–1665. [29] Yamaguchi, M., Nakazawa, T., Kuyama, H., Obama, T. et al., High-throughput method for N-terminal sequencing of proteins by MALDI mass spectrometry. Anal. Chem. 2005, 77, 645–651. [45] Cantin, G. T., Shock, T. R., Park, S. K., Madhani, H. D. et al., Optimizing TiO(2)-based phosphopeptide enrichment for automated multidimensional liquid chromatography coupled to tandem mass spectrometry. Anal. Chem. 2007, 79, 4666–4673. [30] Mikami, T., Takao, T., Selective isolation of N-blocked peptides by isocyanate-coupled resin. Anal. Chem. 2007, 79, 7910–7915. [46] Kweon, H. K., Hakansson, K., Selective zirconium dioxidebased enrichment of phosphorylated peptides for mass spectrometric analysis. Anal. Chem. 2006, 78, 1743–1749. [31] Kuhn, K., Prinz, T., Schafer, J., Baumann, C. et al., Protein sequence tags: A novel solution for comparative proteomics. Proteomics 2005, 5, 2364–2368. [47] Zhou, H., Tian, R., Ye, M., Xu, S. et al., Highly specific enrichment of phosphopeptides by zirconium dioxide nanoparticles for phosphoproteome analysis. Electrophoresis 2007, 28, 2201–2215. [32] Kuhn, K., Thompson, A., Prinz, T., Muller, J. et al., Isolation of N-terminal protein sequence tags from cyanogen bromide cleaved proteins as a novel approach to investigate hydrophobic proteins. J. Proteome Res. 2003, 2, 598–609. [33] Prinz, T., Muller, J., Kuhn, K., Schafer, J. et al., Characterization of low abundant membrane proteins using the protein sequence tag technology. J. Proteome Res. 2004, 3, 1073– 1081. [48] Beausoleil, S. A., Jedrychowski, M., Schwartz, D., Elias, J. E. et al., Large-scale characterization of HeLa cell nuclear phosphoproteins. Proc. Natl. Acad. Sci. USA 2004, 101, 12130–12135. [49] Ballif, B. A., Villen, J., Beausoleil, S. A., Schwartz, D. et al., Phosphoproteomic analysis of the developing mouse brain. Mol. Cell. Proteomics 2004, 3, 1093–1101. [34] Feuerstein, I., Morandell, S., Stecher, G., Huck, C. W. et al., Phosphoproteomic analysis using immobilized metal ion affinity chromatography on the basis of cellulose powder. Proteomics 2005, 5, 46–54. [50] Nuhse, T. S., Stensballe, A., Jensen, O. N., Peck, S. C., Largescale analysis of in vivo phosphorylated membrane proteins by immobilized metal ion affinity chromatography and mass spectrometry. Mol. Cell. Proteomics 2003, 2, 1234– 1243. [35] Jin, W. H., Dai, J., Zhou, H., Xia, Q. C. et al., Phosphoproteome analysis of mouse liver using immobilized metal affinity purification and linear ion trap mass spectrometry. Rapid Commun. Mass Spectrom. 2004, 18, 2169–2176. [51] Dai, J., Jin, W. H., Sheng, Q. H., Shieh, C. H. et al., Protein phosphorylation and expression profiling by Yin-yang multidimensional liquid chromatography (Yin-yang MDLC) mass spectrometry. J. Proteome Res. 2007, 6, 250–262. [36] Lee, J., Xu, Y., Chen, Y., Sprung, R. et al., Mitochondrial phosphoproteome revealed by an improved IMAC method and MS/MS/MS. Mol. Cell. Proteomics 2007, 6, 669–676. [52] Han, G. H., Ye, M. L., Zhou, H. J., Jiang, X. N. et al., Large scale phosphoproteome analysis of human liver tissue by enrichment and fractionation of phosphopeptides with strong anion exchange chromatography. Proteomics 2008, in press. [37] Wagner, V., Gessner, G., Heiland, I., Kaminski, M. et al., Analysis of the phosphoproteome of Chlamydomonas reinhardtii provides new insights into various cellular pathways. Eukaryot. Cell 2006, 5, 457–468. [38] Nuhse, T. S., Peck, S. C., Peptide-based phosphoproteomics with immobilized metal ion chromatography. Methods Mol. Biol. 2006, 323, 431–436. [39] Pan, C., Ye, M., Liu, Y., Feng, S. et al., Enrichment of phosphopeptides by Fe31-immobilized mesoporous nanoparticles of MCM-41 for MALDI and nano-LC-MS/MS analysis. J. Proteome Res. 2006, 5, 3114–3124. [40] Feng, S., Pan, C., Jiang, X., Xu, S. et al., Fe31 immobilized metal affinity chromatography with silica monolithic capillary column for phosphoproteome analysis. Proteomics 2007, 7, 351–360. [41] Ficarro, S. B., McCleland, M. L., Stukenberg, P. T., Burke, D. J. et al., Phosphoproteome analysis by mass spectrometry and its application to Saccharomyces cerevisiae. Nat. Biotechnol. 2002, 20, 301–305. © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim [53] Xu, S., Zhou, H., Pan, C., Fu, Y. et al., Iminodiacetic acid derivatized porous silicon as a matrix support for sample pretreatment and matrix-assisted laser desorption/ionization time-of-flight mass spectrometry analysis. Rapid Commun. Mass Spectrom. 2006, 20, 1769–1775. [54] Zhou, H., Xu, S., Ye, M., Feng, S. et al., Zirconium phosphonate-modified porous silicon for highly specific capture of phosphopeptides and MALDI-TOF MS analysis. J. Proteome Res. 2006, 5, 2431–2437. [55] Dunn, J. D., Watson, J. T., Bruening, M. L., Detection of phosphopeptides using Fe(III)-nitrilotriacetate complexes immobilized on a MALDI plate. Anal. Chem. 2006, 78, 1574– 1580. [56] Yang, Z., Hancock, W. S., Chew, T. R., Bonilla, L., A study of glycoproteins in human serum and plasma reference standards (HUPO) using multilectin affinity chromatography coupled with RPLC-MS/MS. Proteomics 2005, 5, 3353– 3366. www.proteomics-journal.com Proteomics 2008, 8, 686–705 Technology 703 [57] Kaji, H., Saito, H., Yamauchi, Y., Shinkawa, T. et al., Lectin affinity capture, isotope-coded tagging and mass spectrometry to identify N-linked glycoproteins. Nat. Biotechnol. 2003, 21, 667–672. [73] Aristoteli, L. P., Molloy, M. P., Baker, M. S., Evaluation of endogenous plasma peptide extraction methods for mass spectrometric biomarker discovery. J. Proteome Res. 2007, 6, 571–581. [58] Drake, R. R., Schwegler, E. E., Malik, G., Diaz, J. et al., Lectin capture strategies combined with mass spectrometry for the discovery of serum glycoprotein biomarkers. Mol. Cell. Proteomics 2006, 5, 1957–1967. [74] Johnson, K. L., Mason, C. J., Muddiman, D. C., Eckel, J. E., Analysis of the low molecular weight fraction of serum by LC-dual ESI-FT-ICR mass spectrometry: precision of retention time, mass, and ion abundance. Anal. Chem. 2004, 76, 5097–5103. [59] Lis, H., Sharon, N., Lectins: Carbohydrate-specific proteins that mediate cellular recognition. Chem. Rev. 1998, 98, 637– 674. [60] Nilsson, C. L., Lectins: Proteins that interpret the sugar code. Anal. Chem. 2003, 75, 348A–353A. [61] Bond, M. R., Kohler, J. J., Chemical methods for glycoprotein discovery. Curr. Opin. Chem. Biol. 2007, 11, 52–58. [62] Zhang, H., Li, X. J., Martin, D. B., Aebersold, R., Identification and quantification of N-linked glycoproteins using hydrazide chemistry, stable isotope labeling and mass spectrometry. Nat. Biotechnol. 2003, 21, 660–666. [75] Tirumalai, R. S., Chan, K. C., Prieto, D. A., Issaq, H. J. et al., Characterization of the low molecular weight human serum proteome. Mol. Cell. Proteomics 2003, 2, 1096–1103. [76] Yuan, X., Desiderio, D. M., Human cerebrospinal fluid peptidomics. J. Mass Spectrom. 2005, 40, 176–181. [77] Zheng, X., Baker, H., Hancock, W. S., Analysis of the low molecular weight serum peptidome using ultrafiltration and a hybrid ion trap-Fourier transform mass spectrometer. J. Chromatogr. A 2006, 1120, 173–184. [63] Zhang, H., Liu, A. Y., Loriaux, P., Wollscheid, B. et al., Mass spectrometric detection of tissue proteins in plasma. Mol. Cell. Proteomics, 2007, 6, 64–71. [78] Hu, L., Li, X., Jiang, X., Zhou, H. et al., Comprehensive peptidome analysis of mouse livers by size exclusion chromatography prefractionation and nanoLC-MS/MS identification. J. Proteome Res. 2007, 6, 801–808. [64] Sun, B., Ranish, J. A., Utleg, A. G., White, J. T. et al., Shotgun glycopeptide capture approach coupled with mass spectrometry for comprehensive glycoproteomics. Mol. Cell. Proteomics 2007, 6, 141–149. [79] Hsieh, S. Y., Chen, R. K., Pan, Y. H., Lee, H. L., Systematical evaluation of the effects of sample collection procedures on low-molecular-weight serum/plasma proteome profiling. Proteomics 2006, 6, 3189–3198. [65] Alvarez-Manilla, G., Atwood, J., 3rd, Guo, Y., Warren, N. L. et al., Tools for glycoproteomic analysis: size exclusion chromatography facilitates identification of tryptic glycopeptides with N-linked glycosylation sites. J. Proteome Res. 2006, 5, 701–708. [80] Villanueva, J., Lawlor, K., Toledo-Crow, R., Tempst, P., Automated serum peptide profiling. Nat. Protoc. 2006, 1, 880– 891. [66] An, H. J., Peavy, T. R., Hedrick, J. L., Lebrilla, C. B., Determination of N-glycosylation sites and site heterogeneity in glycoproteins. Anal. Chem. 2003, 75, 5628–5637. [67] Hagglund, P., Bunkenborg, J., Elortza, F., Jensen, O. N. et al., A new strategy for identification of N-glycosylated proteins and unambiguous assignment of their glycosylation sites using HILIC enrichment and partial deglycosylation. J. Proteome Res. 2004, 3, 556–566. [68] Larsen, M. R., Hojrup, P., Roepstorff, P., Characterization of gel-separated glycoproteins using two-step proteolytic digestion combined with sequential microcolumns and mass spectrometry. Mol. Cell. Proteomics 2005, 4, 107–119. [69] Wada, Y., Tajiri, M., Yoshida, S., Hydrophilic affinity isolation and MALDI multiple-stage tandem mass spectrometry of glycopeptides for glycoproteomics. Anal. Chem. 2004, 76, 6560–6565. [70] Zhang, Q., Tang, N., Brock, J., Mottaz, H. et al., Enrichment and analysis of nonenzymatically glycated peptides: boronate affinity chromatography coupled with electron-transfer dissociation mass spectrometry J. Proteome Res. 2007, 6, 2323–2330. [71] Verhaert, P., Uttenweiler-Joseph, S., de Vries, M., Loboda, A. et al., Matrix-assisted laser desorption/ionization quadrupole time-of-flight mass spectrometry: an elegant tool for peptidomics. Proteomics 2001, 1, 118–131. [72] Schulz-Knappe, P., Zucht, H. D., Heine, G., Jurgens, M. et al., Peptidomics: he comprehensive analysis of peptides in complex biological mixtures. Comb. Chem. High Throughput Screen. 2001, 4, 207–217. © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim [81] Villanueva, J., Philip, J., Chaparro, C. A., Li, Y. et al., Correcting common errors in identifying cancer-specific serum peptide signatures. J. Proteome Res. 2005, 4, 1060–1072. [82] Villanueva, J., Philip, J., Entenberg, D., Chaparro, C. A. et al., Serum peptide profiling by magnetic particle-assisted, automated sample processing and MALDI-TOF mass spectrometry. Anal. Chem. 2004, 76, 1560–1570. [83] Fiedler, G. M., Baumann, S., Leichtle, A., Oltmann, A. et al., Standardized peptidome profiling of human urine by magnetic bead separation and matrix-assisted laser desorption/ ionization time-of-flight mass spectrometry. Clin. Chem. 2007, 53, 421–428. [84] Koomen, J. M., Li, D., Xiao, L. C., Liu, T. C. et al., Direct tandem mass spectrometry reveals limitations in protein profiling experiments for plasma biomarker discovery. J. Proteome Res. 2005, 4, 972–981. [85] Gaspari, M., Ming-Cheng Cheng, M., Terracciano, R., Liu, X. et al., Nanoporous surfaces as harvesting agents for mass spectrometric analysis of peptides in human plasma. J. Proteome Res. 2006, 5, 1261–1266. [86] Terracciano, R., Gaspari, M., Testa, F., Pasqua, L. et al., Selective binding and enrichment for low-molecular weight biomarker molecules in human plasma after exposure to nanoporous silica particles. Proteomics 2006, 6, 3243–3250. [87] Jia, W., Chen, X., Lu, H., Yang, P., CaCO3-poly(methyl methacrylate) nanoparticles for fast enrichment of lowabundance peptides followed by CaCO3-core removal for MALDI-TOF MS analysis. Angew. Chem. Int. Ed. Engl. 2006, 45, 3345–3349. [88] Zhang, Y., Wang, X., Shan, W., Wu, B. et al., Enrichment of low-abundance peptides and proteins on zeolite nanocrys- www.proteomics-journal.com 704 X. Jiang et al. Proteomics 2008, 8, 686–705 tals for direct MALDI-TOF MS analysis. Angew. Chem. Int. Ed. Engl. 2005, 44, 615–617. to peptide and protein analysis. J. Sep. Sci. 2002, 25, 557– 568. [89] Li, X., Xu, S., Pan, C., Zhou, H. et al., Enrichment of peptides from plasma for peptidome analysis using multiwalled carbon nanotubes. J. Sep. Sci. 2007, 30, 930–943. [104] Yi, E. C., Lee, H., Aebersold, R., Goodlett, D. R., A microcapillary trap cartridge-microcapillary high-performance liquid chromatography electrospray ionization emitter device capable of peptide tandem mass spectrometry at the attomole level on an ion trap mass spectrometer with automated routine operation. Rapid Commun. Mass Spectrom. 2003, 17, 2093–2098. [90] Desilets, C. P., Rounds, M. A., Regnier, F. E., Semipermeable-surface reversed-phase media for high-performance liquid chromatography. J. Chromatogr. 1991, 544, 25–39. [91] Hagestam, I., Pinkerton, T., Internal surface reversed-phase silica supports for liquid chromatography. Anal. Chem. 1985, 57. [92] Boos, K., Grimm, C., High-performance liquid chromatography integrated solid-phase extraction in bioanalysis using restricted access precolumn packings. Trac-Trend. Anal. Chem. 1999, 18, 175–180. [93] Machtejevas, E., Andrecht, S., Lubda, D., Unger, K. K., Monolithic silica columns of various format in automated sample clean-up/multidimensional liquid chromatography/mass spectrometry for peptidomics. J. Chromatogr. A 2007, 1144, 97–101. [94] Machtejevas, E., John, H., Wagner, K., Standker, L. et al., Automated multi-dimensional liquid chromatography: sample preparation and identification of peptides from human blood filtrate. J. Chromatogr. B 2004, 803, 121–130. [95] Wagner, K., Miliotis, T., Marko-Varga, G., Bischoff, R. et al., An automated on-line multidimensional HPLC system for protein and peptide mapping with integrated sample preparation. Anal. Chem. 2002, 74, 809–820. [96] Hu, L., Ye, M., Boos, K., Jiang, X. et al., Fully automated serum peptidome analysis by coupling a novel bifunctional capillary trap column with nanoLC-MS. In preparation. [97] Tian, R., Zhang, H., Ye, M., Jiang, X. et al., Selective extraction of peptides from human plasma by highly ordered mesoporous silica particles for peptidome analysis. Angew. Chem. Int. Ed. Engl. 2007, 46, 962–965. [98] Devreese, B., Vanrobaeys, F., Van Beeumen, J., Automated nanoflow liquid chromatography/tandem mass spectrometric identification of proteins from Shewanella putrefaciens separated by two-dimensional polyacrylamide gel electrophoresis. Rapid Commun. Mass Spectrom. 2001, 15, 50–56. [99] Masuda, J., Maynard, D. A., Nishimura, M., Uedac, T. et al., Fully automated micro- and nanoscale one- or two-dimensional high-performance liquid chromatography system for liquid chromatography-mass spectrometry compatible with non-volatile salts for ion exchange chromatography. J. Chromatogr. A 2005, 1063, 57–69. [100] Mitulovic, G., Stingl, C., Smoluch, M., Swart, R. et al., Automated, on-line two-dimensional nano liquid chromatography tandem mass spectrometry for rapid analysis of complex protein digests. Proteomics 2004, 4, 2545–2557. [101] Jiang, X., Feng, S., Tian, R., Han, G. et al., Automation of nanoflow liquid chromatography-tandem mass spectrometry for proteome analysis by using a strong cation exchange trap column. Proteomics 2007, 7, 528–539. [102] Licklider, L. J., Thoreen, C. C., Peng, J. M., Gygi, S. P., Automation of nanoscale microcapillary liquid chromatography-tandem mass spectromentry with a vented column. Anal. Chem. 2002, 74, 3076–3083. [103] Meiring, H. D., van der Heeft, E., ten Hove, G. J., de Jong, A., Nanoscale LC-MS(n): Technical design and applications © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim [105] Wang, F., Jiang, X., Feng, S., Tian, R. et al., Automated injection of uncleaned samples using ten-port switching valve and stronge cation exchange trap column for proteome analysis. J. Chromatogr. A 2007, 1171, 56–62. [106] Wang, F., Dong, J., Jiang, X., Ye, M. et al., Capillary trap column with strong cation-exchange monolith for automated shotgun proteome analysis. Anal. Chem. 2007, 79, 6599–6606. [107] Slysz, G. W., Lewis, D. F., Schriemer, D. C., Detection and identification of sub-nanogram levels of protein in a nanoLC-trypsin-MS system. J. Proteome Res. 2006, 5, 1959–1966. [108] Slysz, G. W., Schriemer, D. C., Blending protein separation and peptide analysis through real-time proteolytic digestion. Anal. Chem. 2005, 77, 1572–1579. [109] Ye, M., Hu, S., Schoenherr, R. M., Dovichi, N. J., On-line protein digestion and peptide mapping by capillary electrophoresis with post-column labeling for laser-induced fluorescence detection. Electrophoresis 2004, 25, 1319– 1326. [110] Duan, J., Liang, Z., Yang, C., Zhang, J. et al., Rapid protein identification using monolithic enzymatic microreactor and LC-ESI-MS/MS. Proteomics 2006, 6, 412–419. [111] Calleri, E., Temporini, C., Perani, E., De Palma, A. et al., Trypsin-based monolithic bioreactor coupled on-line with LC/MS/MS system for protein digestion and variant identification in standard solutions and serum samples. J. Proteome Res. 2005, 4, 481–490. [112] Feng, S., Ye, M., Jiang, X., Jin, W. et al., Coupling the immobilized trypsin microreactor of monolithic capillary with muRPLC-MS/MS for shotgun proteome analysis. J. Proteome Res. 2006, 5, 422–428. [113] Schoenherr, R. M., Ye, M., Vannatta, M., Dovichi, N. J., CEmicroreactor-CE-MS/MS for protein analysis. Anal. Chem. 2007, 79, 2230–2238. [114] Neville, D. C., Rozanas, C. R., Price, E. M., Gruis, D. B. et al., Evidence for phosphorylation of serine 753 in CFTR using a novel metal-ion affinity resin and matrix-assisted laser desorption mass spectrometry. Protein Sci. 1997, 6, 2436– 2445. [115] Posewitz, M. C., Tempst, P., Immobilized gallium(III) affinity chromatography of phosphopeptides. Anal. Chem. 1999, 71, 2883–2892. [116] Oda, Y., Nagasu, T., Chait, B. T., Enrichment analysis of phosphorylated proteins as a tool for probing the phosphoproteome. Nat. Biotechnol. 2001, 19, 379–382. [117] Zhou, H. L., Watts, J. D., Aebersold, R., A systematic approach to the analysis of protein phosphorylation. Nat. Biotechnol. 2001, 19, 375–378. [118] Guo, L., Eisenman, J. R., Mahimkar, R. M., Peschon, J. J. et al., A proteomic approach for the identification of cell-sur- www.proteomics-journal.com Proteomics 2008, 8, 686–705 face proteins shed by metalloproteases. Mol. Cell. Proteomics 2002, 1, 30–36. [119] Vosseller, K., Trinidad, J. C., Chalkley, R. J., Specht, C. G. et al., O-linked N-acetylglucosamine proteomics of postsynaptic density preparations using lectin weak affinity chromatography and mass spectrometry. Mol. Cell. Proteomics 2006, 5, 923–934. [120] Wu, A. M., Wu, J. H., Singh, T., Liu, J. H. et al., Lectinochemical studies on the affinity of Anguilla anguilla agglutinin for mammalian glycotopes. Life Sci. 2004, 75, 1085– 1103. [121] Shibuya, N., Goldstein, I. J., Broekaert, W. F., NsimbaLubaki, M. et al., Fractionation of sialylated oligosaccha- © 2008 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Technology 705 rides, glycopeptides, and glycoproteins on immobilized elderberry (Sambucus nigra L.) bark lectin. Arch. Biochem. Biophys. 1987, 254, 1–8. [122] Morelle, W., Canis, K., Chirat, F., Faid, V. et al., The use of mass spectrometry for the proteomic analysis of glycosylation. Proteomics 2006, 6, 3993–4015. [123] Kochibe, N., Furukawa, K., Purification and properties of a novel fucose-specific hemagglutinin of Aleuria aurantia. Biochemistry 1980, 19, 2841–2846. [124] Neurohr, K. J., Young, N. M., Mantsch, H. H., Determination of the carbohydrate-binding properties of peanut agglutinin by ultraviolet difference spectroscopy. J. Biol. Chem. 1980, 255, 9205–9209. www.proteomics-journal.com
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