IJMS

您所在的位置：网站首页 › resolvex › IJMS

IJMS

2023-03-14 22:06| 来源: 网络整理| 查看: 265

Next Article in Journal Importance, Applications and Features of Assays Measuring SARS-CoV-2 Neutralizing Antibodies Previous Article in Journal The Roles of Secreted Wnt Ligands in Cancer Journals Active Journals Find a Journal Proceedings Series Topics Information For Authors For Reviewers For Editors For Librarians For Publishers For Societies For Conference Organizers Open Access Policy Institutional Open Access Program Special Issues Guidelines Editorial Process Research and Publication Ethics Article Processing Charges Awards Testimonials Author Services Initiatives Sciforum MDPI Books Preprints.org Scilit SciProfiles Encyclopedia JAMS Proceedings Series About Overview Contact Careers News Blog Sign In / Sign Up Notice clear Notice

You are accessing a machine-readable page. In order to be human-readable, please install an RSS reader.

Continue Cancel clear

All articles published by MDPI are made immediately available worldwide under an open access license. No special permission is required to reuse all or part of the article published by MDPI, including figures and tables. For articles published under an open access Creative Common CC BY license, any part of the article may be reused without permission provided that the original article is clearly cited. For more information, please refer to https://www.mdpi.com/openaccess.

Feature papers represent the most advanced research with significant potential for high impact in the field. A Feature Paper should be a substantial original Article that involves several techniques or approaches, provides an outlook for future research directions and describes possible research applications.

Feature papers are submitted upon individual invitation or recommendation by the scientific editors and must receive positive feedback from the reviewers.

Editor’s Choice articles are based on recommendations by the scientific editors of MDPI journals from around the world. Editors select a small number of articles recently published in the journal that they believe will be particularly interesting to readers, or important in the respective research area. The aim is to provide a snapshot of some of the most exciting work published in the various research areas of the journal.

Journals Active Journals Find a Journal Proceedings Series Topics Information For Authors For Reviewers For Editors For Librarians For Publishers For Societies For Conference Organizers Open Access Policy Institutional Open Access Program Special Issues Guidelines Editorial Process Research and Publication Ethics Article Processing Charges Awards Testimonials Author Services Initiatives Sciforum MDPI Books Preprints.org Scilit SciProfiles Encyclopedia JAMS Proceedings Series About Overview Contact Careers News Blog Sign In / Sign Up Submit Journals IJMS Volume 24 Issue 6 10.3390/ijms24065350

Submit to this Journal Review for this Journal Edit a Special Issue ► ▼ Article Menu Article Menu Academic Editors

Christof Lenz

Christine Carapito Subscribe SciFeed Recommended Articles Related Info Link Google Scholar More by Authors Links on DOAJ Duong, V. Lee, H. on Google Scholar Duong, V. Lee, H. on PubMed Duong, V. Lee, H. /ajax/scifeed/subscribe Article Views Citations - Table of Contents Altmetric share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up ... Endorse textsms ... Comment Need Help? Support

Find support for a specific problem in the support section of our website.

Get Support Feedback

Please let us know what you think of our products and services.

Give Feedback Information

Visit our dedicated information section to learn more about MDPI.

Get Information clear JSmol Viewer clear first_page settings Order Article Reprints Font Type: Arial Georgia Verdana Font Size: Aa Aa Aa Line Spacing:    Column Width:    Background: Open AccessReview Bottom-Up Proteomics: Advancements in Sample Preparation by

Van-An Duong ORCID

and

Hookeun Lee * ORCID

College of Pharmacy, Gachon University, Incheon 21936, Republic of Korea * Author to whom correspondence should be addressed. Int. J. Mol. Sci. 2023, 24(6), 5350; https://doi.org/10.3390/ijms24065350 Received: 19 January 2023 / Revised: 28 February 2023 / Accepted: 9 March 2023 / Published: 10 March 2023 (This article belongs to the Special Issue Advances in Mass Spectrometry-Based Proteomics) Download Download PDF Download PDF with Cover Download XML Download Epub Browse Figures Versions Notes

Abstract: Liquid chromatography–tandem mass spectrometry (LC–MS/MS)-based proteomics is a powerful technique for profiling proteomes of cells, tissues, and body fluids. Typical bottom-up proteomic workflows consist of the following three major steps: sample preparation, LC–MS/MS analysis, and data analysis. LC–MS/MS and data analysis techniques have been intensively developed, whereas sample preparation, a laborious process, remains a difficult task and the main challenge in different applications. Sample preparation is a crucial stage that affects the overall efficiency of a proteomic study; however, it is prone to errors and has low reproducibility and throughput. In-solution digestion and filter-aided sample preparation are the typical and widely used methods. In the past decade, novel methods to improve and facilitate the entire sample preparation process or integrate sample preparation and fractionation have been reported to reduce time, increase throughput, and improve reproducibility. In this review, we have outlined the current methods used for sample preparation in proteomics, including on-membrane digestion, bead-based digestion, immobilized enzymatic digestion, and suspension trapping. Additionally, we have summarized and discussed current devices and methods for integrating different steps of sample preparation and peptide fractionation. Keywords: proteomics; sample preparation; in-solution digestion; FASP; S-Trap; SP3; LC-MS/MS; automation 1. IntroductionProteomics is an analytical technique that examines protein expression, structures, functions, and interactions in a particular cell, tissue, body fluid, or organism [1,2]. Protein composition and abundance are currently analyzed to identify disease markers or treatment mechanisms, as all changes in proteomes indicate pathological or biological processes [3,4,5]. Over the last two decades, liquid chromatography–tandem mass spectrometry (LC-MS/MS)-based proteomics has been developed as an alternative to time-consuming and labor-intensive gel-based proteomics and immunoassays [6]. LC–MS/MS-based proteomics has high throughput because thousands of peptides and proteins can be analyzed in a short time [7]. Furthermore, it can be automated to improve high-throughput performance, precision, and repeatability [8]. Some liquid-handling workstations, such as Agilent AssayMAP Bravo [9] and Biomek NXP Span-8 [10], can perform most steps of proteomic workflows.The two analytical procedures frequently used in proteomics are top-down and bottom-up approaches. In top-down proteomics, intact proteins are directly separated and analyzed using LC–MS/MS to identify, characterize, and quantify proteoforms (distinct proteins generated from a particular gene owing to genetic variations), alternative RNA splicing, and post-translational modifications (PTMs) [11,12,13]. Conversely, in bottom-up proteomics, proteins undergo enzymatic proteolysis and the resultant peptides are analyzed and identified. This strategy has been widely used because peptides are easier to separate and identify than proteins [14]. The peptide mixture in bottom-up proteomics comprises thousands of peptides; thus, multidimensional separation is usually performed for in-depth proteome analysis [15].A typical LC–MS/MS-based proteomic workflow consists of three major steps: sample preparation, protein/peptide separation coupled with MS/MS analysis, and data analysis [16,17]. The two proteomic approaches differ mainly in sample preparation. Top-down proteomics include protein extraction from biological samples and sample purification [18]. However, bottom-up proteomics require additional steps including protein reduction, alkylation, and enzymatic digestion [14]. Each stage of the bottom-up proteomic workflow has tremendously developed over the last two decades. The development of high-performance liquid chromatography (HPLC) instrumentation over the past decade has facilitated proteomic research by simultaneously separating many peptides in a single run [19,20]. Reversed-phase liquid chromatography (RPLC) plays a critical role in protein/peptide separation prior to MS/MS analysis [21]. Other separation mechanisms, such as strong cation exchange chromatography (SCX), strong anion exchange chromatography (SAX), size exclusion chromatography (SEC), and hydrophilic interaction chromatography (HILIC), are frequently combined with RPLC to develop multidimensional separation platforms [15]. Advances in multidimensional separation of proteins/peptides have been demonstrated in various systems, including SCX–RPLC [22,23], SAX–RPLC [24], SEC–RPLC [25,26], HILIC–RPLC [27,28], RPLC–RPLC [29,30], SCX–RPLC–RPLC [31,32,33], SAX–RPLC–RPLC [34], SCX–HILIC–RPLC [35], and RPLC–RPLC–RPLC [36].The mass spectrometer has also been improved in the past decade with the development of new fragmentation techniques and substantial increases in scan speed and mass accuracy [37,38,39], enabling the first profiling of the human proteome draft in 2014 [37,38]. Some types of mass analyzers frequently used in proteomics are quadrupole, ion-trap, time-of-flight, orbitrap, and Fourier-transform ion cyclotron resonance [39,40]. In recent proteomic studies, data have been analyzed using high-throughput and time-efficient software [41]. Raw MS/MS data are searched against databases to identify and quantify peptides and proteins using search engines such as X!Tandem [42], Mascot [43], Sequest [44], Comet [45], Maxquant [46], Byonic [47], MSFragger [48], and Open-pFind [49]. The results are further subjected to statistical tests to identify differentially expressed proteins (DEPs), enrichment analysis to determine biological relevance, and network analysis to visualize protein–protein interactions and protein groups [41].Apart from these stages, sample preparation remains a difficult task and the main challenge of bottom-up proteomics with laborious steps [50]. Generally, sample preparation aims to create a less complex peptide mixture that is suitable for analysis. It requires pre-fractionation; depletion of most unnecessarily abundant proteins; removal of DNA, lipids, and small metabolites; and sample clean-up from impurities (salts and remaining solid particles) [51]. Therefore, typical sample preparation processes for bottom-up proteomics usually include lysis/homogenization, protein extraction/precipitation, reduction, alkylation, enzymatic digestion, fractionation, and desalting (Figure 1). Sample preparation is an essential stage affecting the overall efficiency of proteomic studies. However, it is error-prone and has low reproducibility and throughput [52]. The early stages of proteomics used in-gel sample preparation [53]. Gel-free sample preparation has been developed in the past decade and is widely used in proteomic studies. Typical strategies include in-solution digestion (ISD), filter-aided sample preparation (FASP) [54], suspension trapping (S-Trap) [55], and single-pot solid-phase-enhanced sample preparation (SP3) [56]. Many groups have reported novel methods to improve and facilitate the entire sample preparation process or some of its steps to reduce time, increase throughput, and improve reproducibility. Single-cell proteomics has been developed in the past decades to handle samples with low protein amounts [57]. Typical single-cell proteomic approaches are: nanodroplet processing in one pot for a trace sample (nanoPOTS) [58]; nanoliter-scale oil-air-droplet (OAD) chip [59]; an integrated device for single-cell analysis (iPAD-1) [60]; digital microfluidic isolation of single cells for -omics (DISCO) [61]; and integrated spectral library-based single-cell proteomics [62]. However, this review does not cover single-cell proteomics due to their unique characteristics. In this review, we have summarized the current methods for preparing different proteomic sample types. In addition, we have presented and discussed recent developments to enhance the sample preparation process and integrate different sample preparation steps as well as their applications to biological samples. 2. From Biological Samples to ProteinsThe first stage of a bottom-up proteomic workflow is to obtain a protein mixture from biological samples. This includes sample pretreatment, enzyme inhibition, homogenization, protein extraction/precipitation, and protein fractionation. The procedures vary depending on the sample type. They are also used for sample preparation in top-down proteomics. 2.1. CellsFor cell samples, it is necessary to break down the cell membranes and homogenize the samples with lysis buffer and sonication [63]. During this process, some enzymes that may affect the protein structure, such as proteases and phosphatases, are released. Therefore, these enzymes should be inhibited by maintaining the samples at a low temperature and adding an enzyme inhibitor cocktail. Typical protease inhibitors include pepstatin A, leupeptin, aprotinin, and chymostatin [64]. They are commercially available independently or in mixtures. Phosphatase inhibitors, including sodium fluoride, sodium orthovanadate, sodium pyrophosphate, and beta-glycerophosphate, are used to protect phosphorylated proteins [65]. These enzyme inhibitors are usually added to the lysis buffers prior to sonication.Homogenized samples are then subjected to a protein extraction step to obtain samples with high protein concentrations before digestion. Protein precipitation is the most common method for protein extraction, particularly for diluted samples. Organic solvents (such as acetone, methanol, or ethanol) and their mixtures with trichloroacetic acid or sodium deoxycholate are generally used to precipitate proteins. The protein pellets are then collected and washed with pre-chilled solvent to remove contaminants [66,67]. Protein precipitation with acetone is widely used because it rapidly dissolves non-polar contaminants (such as lipids). In addition, chromatography, electrophoresis, dialysis, ultrafiltration, lyophilization, and crystallization have also been used for protein purification [68]. 2.2. Biological FluidsBiological fluids (blood, urine, saliva, nasal fluids, tears, and aqueous humor) need to be diluted with an appropriate amount of buffer to facilitate the subsequent enzymatic digestion process [69,70]. In the case of urine samples, proteins are extracted using solvent precipitation, ultrafiltration, centrifugal filtration, dialysis, and lyophilization [71]. Previously, Jesus et al. developed an ultrasonic-based, membrane-aided sample preparation for urine proteomic analysis, which includes urine filtration through a membrane (to retain proteins and remove salts) and ultrasonic energy-mediated tryptic digestion in the membrane [72].Blood samples are usually centrifuged to collect plasma or serum. Plasma and serum are generally subjected to immunodepletion to remove high-abundance proteins, thereby allowing the identification of low-abundance proteins [73]. Approximately 10,000 proteins are present in plasma, ranging from albumin at 35 to 50 mg/mL to low-abundance proteins at pg/mL [74]. Immunodepletion (spin columns and LC) and magnetic beads can remove up to the 20 most abundant plasma proteins [75,76,77,78]. Depletion results in carry-over, low reproducibility, low throughput, and loss of many albumin-bound proteins [79]. Other fluids are centrifuged to remove debris before protein extraction [80]. 2.3. TissuesTissue samples are usually rinsed with ice-cold saline to remove blood, serum, and fat [81]. Subsequently, they are homogenized using different apparatuses.Manual tissue homogenization involves the use of a pestle and mortar. Pestles and mortars can be made of glass, glass teflon, or stainless steel. Glass mortars and pestles can generate heat, whereas glass Teflon mortars and pestles (with a nonstick Teflon surface) minimize heat generation and sample loss. The apparatus is particularly suitable for soft tissues, such as the brain and liver. After placing the tissues in a tube on ice (such as a Potter homogenizer) with lysis buffer, a pestle is used to manually homogenize the tissues until no large tissue pieces are observed [82]. Under the mechanical shear force, proteins and other molecules are released into the buffers. Mechanical rotor–stator grinders are used to vigorously mix, accelerate, and press samples through the narrow gap between the rotor and stator. They can be used for most tissue types, from soft to tough tissues [83]. Bead-beating homogenization is a flexible and efficient method for preparing soft or hard tissue in seconds. This involves shaking tissues with tungsten carbide, stainless steel, or glass beads in pre-chilled tubes [84,85]. Commercially available bead-beaters can process 4–24 samples simultaneously [86]. Sonication is effective for homogenizing soft tissues. It is typically equipped with a tip generator that causes cavitation effects that disrupt the tissue [87].Liquid nitrogen pulverization involves the use of liquid nitrogen to freeze tissues for a short time. It can preserve protein integrity without generating heat [88]. The frozen tissues are then disrupted using a pestle and mortar [89]. A pulverizer (such as Covaris CP02 cryoPrep automated dry pulverizer) can be used to disrupt the tissues placed in strong and flexible plastic tissue tubes [90]. The resultant powder is mixed with lysis buffer containing enzyme inhibitors and lysed. This method is applicable to all types of tissues, particularly hard tissues and those with tough connective fibers.Pressure cycling homogenizer (PCT) is a technique that involves the use of hydrostatic pressure and mechanical grinding. The instrument (Barocycler) compresses air to create high pressure inside the reaction chamber. The pressure rapidly increased from ambient pressure to ~45,000 psi over several cycles, resulting in tissue disruption. This method is only applicable to soft tissues, whereas some hard and large tissues remain incompletely lysed [91].Similar to cells, homogenized tissues can be subjected to a protein extraction step to obtain samples with high protein concentrations. 2.4. Protein QuantificationProtein concentrations in the samples are determined using different methods before protein digestion. Colorimetric dye methods are based on the color change upon protein–dye binding, whereas fluorescent dye methods rely on the fluorescence associated with the dye after protein–dye binding. Colorimetric dye methods (such as Bradford assay) are straightforward, fast, and compatible with most solvents, buffers, reducing substances, salts, thiols, and metal-chelating agents [92]. Fluorescent dye methods (such as EZQ fluorescent and Qubit protein assays) are susceptible and suitable for low protein samples [93]. Biuret methods rely on protein–copper chelation in an alkaline environment to reduce Cu2+ to Cu+, which reacts with a reagent in a bicinchoninic acid assay to form a purple complex that is detected between 550 and 570 nm [94]. In Lowry assays, Cu+ reacts with Folin–Ciocalteu reagent containing phosphotungstic acid and phosphomolybdic acid [95]. Biuret methods are compatible with most detergents and are less likely to cause protein-to-protein variations than Bradford assays [94]. 3. From Proteins to PeptidesSample preparation from proteins to peptides includes various steps, such as detergent removal, buffer exchange, reduction, alkylation, digestion, optional peptide fractionation, and desalting. The major bottleneck among these procedures is protein cleanup and digestion, which considerably affects the accuracy and reproducibility of protein quantification. 3.1. Protein Digestion 3.1.1. In-Solution and In-Gel DigestionISD and in-gel digestion have been used since the beginning of proteomics research. The ISD can handle 100–1000 µg protein. Urea and thiourea are generally used to solubilize proteins and denature their three-dimensional structures. Proteins in 8 M urea solution are reduced and alkylated to reduce intra- and intermolecular disulfide bonds within and between protein molecules [96]. Typical reduction reagents include dithiothreitol, tris(2-carboxyethyl)phosphine (TCEP), tris(3-hydroxypropyl) phosphine, and 2-mercaptoethanol. The free sulfhydryl groups formed by reduction are alkylated with alkylation agents (iodoacetamide, iodoacetic acid, N-ethylmaleimide, and S-methyl methanethiosulfonate) to prevent disulfide bond reformation. The protein samples are then diluted to reduce the urea concentration before enzyme addition. Typical digestion enzymes are trypsin and LysC. The trypsin:protein ratio usually varies from 1:20 to 1:50. Trypsin cleavage sites are located at the amino acid residues arginine and lysine. The obtained peptides obtained after digestion are 800–2000 Da, which is appropriate for MS/MS sequencing [97]. ISD is associated with inevitable sample loss and cannot be used for detergent-containing samples. It requires a large starting amount of protein (typically > 100 µg) and long digestion time (overnight).A modification of ISD, which can be applied to samples containing sodium dodecyl sulfate (SDS), has been developed recently [98]. This is a simple, robust, and reproducible SDS–cyclodextrin-assisted sample preparation (SCASP) method that can be applied to different sample types. After reduction and alkylation, the samples are mixed with cyclodextrin solution. SDS molecules are quickly incorporated into the internal cavities of cyclodextrins. Trypsin is then added for digestion. The SDS–cyclodextrin complex is removed from the peptide mixtures during desalting. When SCASP was used to analyze 5000 HeLa cell samples, ~2500 proteins were identified. Another ISD modification combines lysis, reduction, and alkylation into a single step to reduce time and sample loss. It is performed using the reducing agent TCEP and alkylating agent 2-chloroacetamide (CAA), which are compatible and can be directly incorporated into the lysis buffer [99,100].Doellinger et al. have recently developed sample preparation by easy extraction and digestion (SPEED) [101]. The main feature of SPEED is the use of pure trifluoroacetic acid (TFA; pKa = 0.2) to dissolve cells and tissues within a few minutes and form clear lysates. After neutralization with a weak base (Tris(hydroxymethyl)-aminomethane, pKa = 8.1), the lysates become slightly turbid due to protein precipitation as fine particles. The proteins are then simultaneously reduced and alkylated using TCEP and CAA, respectively. Then, the samples are heated up to 70–80 °C to shorten the incubation time to 5 min. Subsequently, tryptic digestion is performed similar to that in ISD. Using SPEED, the authors identified ~2700 and ~1900 proteins in Escherichia coli and Staphylococcus aureus, respectively, which were higher than those reported in previous studies [101]. The authors demonstrated that the SPEED was highly reproducible and performed better than FASP, ISD, and SP3 for quantitative proteomics [101]. These results suggest that SPEED is a potential method for future studies in bottom-up proteomics. However, further studies are needed to evaluate this method.In-gel digestion is time-consuming and more laborious than ISD. Proteins are first separated by one- or two-dimensional polyacrylamide gel electrophoresis. Subsequently, the gel spots containing the proteins of interest are collected, cut into small pieces (approximately 1 × 1 mm2), and destained with 100 mM ammonium bicarbonate/acetonitrile (1:1, v/v). The proteins are then reduced, alkylated, and digested. Peptides are usually extracted from gel pieces using 50% acetonitrile/5% formic acid [53]. ISD and in-gel digestion are less efficient, particularly for clinical samples, which require high sample processing throughput and reproducibility. Therefore, other methods have been developed to overcome these limitations. 3.1.2. On-Membrane Digestion: FASP and MSternThese methods utilize membranes to separate proteins from detergents and other small molecular impurities. The FASP was developed in 2005 by Liebler et al. [102] and in 2009 by Mann et al. [54]. This method has been widely used for bottom-up proteomic sample preparation over the past decade. All sample preparation steps in the FASP are performed on an ultrafiltration unit containing a membrane with a proper molecular cut-off (typically 3000 or 10,000 Da) [103]. The proteins are first solubilized in a strong denaturing lysis buffer containing SDS and transferred to an ultrafiltration unit. SDS and other contaminants are removed by ultrafiltration, whereas proteins are retained over the membrane (Figure 2). Reduction, alkylation, and digestion are performed on the membranes. After digestion, the peptides are separated from undigested materials via ultrafiltration and collected for peptide clean-up, fractionation, and MS analysis [104]. Compared to ISD, FASP can be used for samples containing detergents [105]. However, each centrifugal step of the FASP is time-consuming (typically 20–30 min). In addition, the performance of FASP reduces when processing low amounts of protein (such as 4000 proteins in slow- and fast-twitch muscle fibers. Notably, they found 237 and 172 DEPs in slow- and fast-twitch muscle fibers, respectively, after endurance exercise training for 12 weeks. In another study, 124 pairs of tumor and non-tumor esophageal tissue samples were digested using FASP, and the resultant peptides were labeled in 25 groups in a TMT 11-plex experiment [85]. The authors identified 14,252 proteins, with 784 upregulated and 747 downregulated proteins in tumors (fold-change > 1.5, adjusted p < 0.01). Subsequently, they stratified the samples into two subtypes based on proteomic analysis: low-risk S1 and high-risk S2. They identified ELOA and SCAF4 as subtype signatures and constructed a subtype diagnostic and prognostic model.In a recent study, a Resolvex A200 positive-pressure solid-phase extraction unit was combined with a Bravo liquid-handling platform to perform positive-pressure FASP in a 96-well plate [111]. The method exhibited high reproducibility (Pearson correlation coefficient: r = 0.9993) when analyzing 40 technical replicates of mouse heart tissue lysates.Zhang et al. developed a miniaturized FASP method (micro-FASP) for processing small amounts of samples [112]. The authors used a filter with approximately 0.1 mm2 surface area to reduce the total reagent volume to 0.99) when processing Xenopus laevis lysate (12 replicates, 1 μg each). Similarly, Sandbaumhüter et al. developed a flexible well-plate μFASP device suitable for a small amount of protein (1 μg) [113]. The filter area for the centrifugal filter units of the conventional FASP was reduced from approximately 119 mm2 to 0.785 mm2 for the μFASP device. The authors identified ~1300 proteins from 1 μg HeLa digest. From single islets of Langerhans, with 0.9). Adding 30% acetonitrile to the digestion buffer effectively increased peptide recovery and reduced the percentage of missed cleavage. This method demonstrates the applicability of RP beads in protein digestion. However, further studies are needed to evaluate this. 3.1.4. Immobilized Enzymatic DigestionImmobilized enzyme reactors (IMERs) are flow-through devices widely used in biological and chemical reactions. They have a high specific surface area, low reagent consumption, and a fast reaction rate [127]. In these devices, enzymes are adsorbed, entrapped, or covalently bonded to nanostructured materials or solid supports without losing their activity [128]. IMERs can process a small amount of protein ( 0.96) by label-free quantification [99]. The authors also developed an in-house iST 96-well device for high-throughput sample processing [99]. However, proteins can non-reversibly bind to the C18 materials in these iST devices, resulting in sample loss.These iST devices have been used in several studies. Adahi et al. applied C18-SCX StageTip with acid- and salt-based elution methods and identified >22,000 phosphopeptides in 7 SCX fractions from HeLa cells [174]. Similar to SP3, the iST method is suitable for low amounts of starting protein. Sielaff et al. found that iST, SP3, and FASP performed similarly when processing 20 μg HeLa cell lysate (concerning the number of protein identifications and reproducibility) [106]. However, when the sample amount was reduced, the performance of FASP decreased, whereas SP3 and iST showed similar proteomic coverage. From samples of 25,000 immune cells, the number of proteins identified using SP3 was the highest (3152), followed by that using iST (2343) and FASP (109). Geyer et al. used the iST protocol in a 96-well device to process capillary blood plasma samples [175]. Plasma was depleted before performing the iST protocol. From 1 µL plasma, sample preparation took 300 proteins within 50 min of MS using data-independent acquisition analysis [179].Later, they developed SISPROT combining 3D peptide fractionation (3D-SISPROT) [34]. The device was similar to the SISPROT, except that SCX was replaced with SAX. Protein digestion was performed on a SAX disk, followed by SAX fractionation and high-pH RP fractionation. This 3D-SISPROT device was used to generate 3 SAX fractions using 3 buffer solutions at pH 12, 6, and 2. Subsequently, the peptides in these SAX fractions were further fractionated into five, three, and three high-pH RP fractions, respectively. As a result, the authors identified >8000 proteins from 11 high-pH RP fractions with a starting amount of 30 µg HEK 293 cell lysate. Subsequently, they developed a mixed-mode SISPROT using a 1:1 SCX and SAX beads [32]. From 1 mL serum, they generated 4 mixed-mode SCX–SAX fractions and 12 mixed-mode high-pH RP fractions and identified 862 proteins within 12 h of MS. They have also developed SISPROT–Ti4+-IMAC enrichment for phosphoproteome profiling [180] and Glyco–SISPROT for glycoproteome profiling [181]. 5. Authors’ Outlook and Concluding RemarksSample preparation is the most crucial stage of a proteomic study, yet it can be time-consuming, labor-intensive, and error-prone [50]. Many methods, including on-membrane digestion (MStern, FASP, and fa-SPEED), bead-based digestion (proteomic reactor, SP3, and C4-tip), IMERs, S-Trap, and on-slide digestion, have been developed to replace ISD to reduce time, increase throughput, improve reproducibility, and integrate different steps in the process. The primary features of these methods are summarized in Table 1.Among these methods, FASP has been the gold standard over the past decade [54,102]. However, it requires a long centrifugation time for each step and shows reduced performance at low protein amounts [106]. FASP cannot be integrated with downstream peptide cleanup and fractionation. The filter membrane of the FASP spin device can be damaged during a long centrifugation time [115]. Micro-FASP [112] and μFASP [103] have been developed to make this method applicable to samples with 100 µg protein); long digestion time; low throughput and reproducibility[97]SCASPUse cyclodextrin to remove SDS before digestion[98]Simultaneous lysis, reduction, and alkylationUse TCEP and CAA; reduce time and sample loss[99]SPEEDUse pure TFA to dissolve cells and tissues without strong detergent; use tris(hydroxymethyl)-aminomethane to neutralize sample and precipitate proteins; reduce time and sample loss[101]In-gel digestionUse polyacrylamide gel electrophoresis to separate proteins; cut, digest, and analyze gel spots separately; low throughput and reproducibility[53]On-membrane digestionFASPUse a membrane (3000 or 10,000 Da) to separate proteins from detergents and contaminants; on-membrane digestion; tolerant to strong detergent; long centrifugal time; reduced performance with samples of low protein amount (

【本文地址】

IJMS

IJMS

今日新闻

推荐新闻