Ultra-Fast Sample Preparation for High-Throughput Proteomics

Chapter 8 Ultra-Fast Sample Preparation for High-Throughput Proteomics Daniel Lopez-Ferrer, Kim K. Hixson, Mikhail E. Belov, and Richard D. Smith Abstract Sample preparation oftentimes can be the Achilles Heel of any analytical process, and in the field of proteomics, preparing samples for mass spectrometric analysis is no exception. Current goals, concerning proteomic sample preparation on a large scale, include efforts toward improving reproducibility, reducing the time of processing and ultimately the automation of the entire workflow. This chapter reviews an array of recent approaches applied to bottom-up proteomics sample preparation to reduce the processing time down from hours to minutes. The current state-of-the-art approaches in the field use different energy inputs such as microwave, ultrasound or pressure to perform the four basic steps in sample preparation: protein extraction, denaturation, reduction/alkylation, and digestion. No single energy input for enhancement of proteome sample preparation has become the universal gold standard. Instead, a combination of different energy inputs tends to produce the best results. This chapter further describes the future trends in the field such as the hyphenation of sample preparation with downstream detection and analysis systems. Finally, a detailed protocol describing the combined use of both pressure cycling technology and ultrasonic energy inputs to hasten proteomic sample preparation is presented. Keywords Proteomics · Enzymatic digestion · High pressure and mass spectrometry Abbreviations ESI FDR HIFU HPP IAM Electrospray ionization False discovery rate High intensity focused ultrasound High pressure processing Iodoacetamide D. Lopez-Ferrer (B) Pacific Northwest National Laboratory, Biological Science Division, Richland, WA; Caprion Proteomics U.S., LLC, 1455 Adams Drive, Menlo Park, CA 94025, USA e-mail: [email protected] A.R. Ivanov, A.V. Lazarev (eds.), Sample Preparation in Biological Mass Spectrometry, DOI 10.1007/978-94-007-0828-0_8,  C Springer Science+Business Media B.V. 2011 125 126 IT MAPED MS MS/MS MW NA PCT RP TCEP TOF D. Lopez-Ferrer et al. Ion trap Microwave-assisted protein enzymatic digestion Mass spectrometry Tandem mass spectrometry Molecular weight Not assigned Pressure cycling technology Reversed phase Tris(2-carboxyethyl) phosphine Time-of-flight 8.1 Introduction We are entering an age where systems biology approaches, including proteomics, transcriptomics, metabolomics, and other – omics techniques are being measured with greater depth and frequency. These large interconnected databases, relating networks and macromolecular associations, represent the burgeoning industrialization of biology. At the heart of understanding the inner workings of life, identification and characterization of proteins is paramount. Proteins are the fundamental molecular machines of cells. They are essential to the structure, communication, metabolism, replication, catabolism, and incorporation or synthesis of all necessary chemicals needed to sustain life. The genome, which is essentially static, may only be able to provide us with a propensity towards a certain cellular state. The proteome, however, is dynamic, actually revealing what is occurring in the cell at a specific time under a specific condition. To this end, the rapid study of the proteome in a short time frame is crucial to solving most present day biological challenges, and from a clinical perspective it is vital in the development of rapid and accurate diagnosis and prognosis of disease (Qian et al., 2009). Currently most high-throughput proteomic strategies for rapid comprehensive protein profiling employ ultra-high performance and multidimensional peptide chromatography systems (HPLC) coupled with high resolution mass spectrometers (MS) for detection and identification of a previously digested protein sample (i.e., shotgun proteomics) (Aebersold and Mann, 2003; Domon and Aebersold, 2006; Washburn et al., 2001; Wolters et al., 2001). The first and most important step during this type of proteome analysis is the sample preparation. If this primary step fails by being irreproducible (e.g., incomplete protein digestion) or low in quality (i.e., inaccurate determination of peptide concentration or is contaminated with polymers, salts, or surfactants), the downstream peptide separation, detection, or data analysis will fail. In addition, with proteome experiments constantly evolving to be larger and more complex, it has become increasingly necessary to optimize the time-consuming sample preparation strategies conventionally used in today’s high-throughput proteomic laboratories. The typical proteome sample preparation process incorporates cell lysis, protein denaturation, reduction of disulfide bonds, prevention of the reformation of disulfide bonds by alkylation with iodoacetamide, digestion with a protease (trypsin being the most commonly used), and, finally, desalting/clean-up. 8 Ultra-Fast Sample Preparation for High-Throughput Proteomics 127 Typically, the largest bottleneck in sample preparation is the time required for the proteolytic digestion of the protein sample. A traditional proteome digestion calls for several hours of incubation time to produce a thorough and reproducible digest. Here we highlight recent advances in the acceleration of protein digestions down to just a few minutes using an array of different energy inputs to accelerate the kinetics of the digestion reaction. Additionally, we present a protocol that provides a rapid and thorough digestion of proteins using modified sequencing grade trypsin under high hydrostatic pressure. 8.2 Analysis Workflow Life comes in many forms (e.g., bacteria, animals, fungi, plants) that can be found in vastly different environments (i.e., growth media, soil, bodies of water, and other organisms). For this reason, there is no single universal proteome sample procedure for the initial protein extraction. After protein extraction, most workflows follow similar paths consisting of protein solubilization, protein reduction, alkylation, and protein digestion (Lopez-Ferrer et al., 2004; Manza et al., 2005; Wang et al., 2005; Wisniewski et al., 2009). A detailed schematic of a traditional proteome sample preparation workflow in comparison to new improved workflow implementations is illustrated in Fig. 8.1. 8.2.1 Protein Extraction and Solubilization The first step of sample processing in proteomic experiments is to obtain high quality and adequate protein amounts from the sample to be studied. The protein extraction varies significantly depending on where the sample comes from, the relative abundance of the proteins of interest as well as the presence of high abundance proteins that may obscure dissolution, detection and analysis of lower abundance proteins. Apart from working with body fluids, ground water, and secreted protein extracts, most proteome analyses target some sort of cell mass that requires an approach to lyse, homogenize, or solubilize cells in order to extract the proteins. During this process, it is critical to keep the samples ice cold and to use protease inhibitors in the lysis medium if the sample is particularly susceptible to endogenous proteases. Not doing so may increase the likelihood of undesirable side reactions, which could translate into irreproducible results. The most commonly used techniques for cell lysis/tissue disruption are: (a) manual homogenization, (b) vortexing, grinding, or bead beating with beads such as zirconia/silica beads, (c) freeze/thaw using liquid nitrogen or dry ice/ethanol (alongside manual grinding if working with plant tissues or yeast) and (d) sonication or (e) pressure. The combination of these approaches with the use of detergents is also becoming very popular, but this use needs to be compatible with the overall analytical strategy (i.e., detergents typically need to be removed prior to mass spectrometric analysis). In addition to the cell lysis methods 128 D. Lopez-Ferrer et al. Traditional Workflow Improved Workflows Frozen tissue Cell Pellet Body Fluid Protein Protein Extraction Extraction Combination of Pressure Cycling technology technology and liquid - and liquidextraction extraction (~30min) (~30min) Protein Protein Precipitation Precipitation ~2−3h) ((~2-3h) NA NA Solubilization Solubilization ofofProteins Proteins Protein Denaturation/ Protein Denaturation/ Reduction and Alkylation (~2h) Combined use of ultrasound, TCEP and IAM ( 3min) Protein ProteinDigestion Digestion ~4−12h) ((~4-12 h) Use of Microwaves, Ultrasound or Pressure for protein digestion (~1−20 min) LC-MS LC-MS analysis analysis TCEP and IAM (~3min) ~ Microwave Sonoreactor BarocyclerTM Fig. 8.1 Proteomics workflow. Proteins are extracted from different biological materials, precipitated to eliminate salts and other possible artifacts. Then the proteins are solubilized and completely denatured. The proteins are then digested and submitted for LC-MS analysis listed above, pressure cycling technology in recent years has also become available as a means to provide effective and efficient lysis of many types of cells and tissues. Usually the next step is to isolate proteins from most other cellular components (e.g., DNA, RNA, lipids, salts). The most popular technique is to precipitate proteins with either trichloroacetic acid (TCA) or TCA/cold-acetone precipitation (Isaacson et al., 2006). Other protocols have also been very successful, such as the original phenol-chloroform extraction method published by Chomczynsky and Sacchi in the late 1980s (Chomczynski and Sacchi, 1987). Phenol protein extraction followed by protein purification using an ammonium acetate precipitation also works well to extract and purify proteins, especially those coming from difficult samples (e.g., plant leaves containing high concentrations of polyphenols and carbohydrates) (Isaacson et al., 2006). Other recent developments in the realm of protein extraction 8 Ultra-Fast Sample Preparation for High-Throughput Proteomics 129 include the work by Gross et al. consisting in the combined use of fluorinated alcohols and aliphatic hydrocarbons with pressure cycling technology (PCT). The authors demonstrated that efficient protein extraction from a sample was obtainable and furthermore after lysis this emulsion could be separated into a pellet and two liquid phases (Gross et al., 2008). DNA, RNA and their associated proteins were found in the pellet at the bottom of the tube, while proteins and small molecules remained in the middle phase, and the upper layer contained the lipids. Using this strategy, most proteins could be easily extracted by isolating the middle layer and drying off the volatile solvent. Many protein extraction methods leave the protein as a precipitate which then requires solubilization in order to carry out the downstream reduction, alkylation and digestion. The chosen solubilization method will need to be compatible with subsequent analytical methods used in the protein analysis (i.e., LC-MS). Common chaotropes used include urea, thiourea, and guanidine-HCl. These chaotropes denature the proteins and prevent protein aggregation by disrupting hydrogen bonds as well as intra- and intermolecular forces. Depending on which kinds of proteins are present in the sample, solubilization may be more or less challenging and may require different solvent mixtures such as organic-aqueous solvents like TFE (Wang et al., 2005), MeOH (Blonder et al., 2006) or ACN (Russell et al., 2001) in concentrations varying from 20 to 60%. When using high concentrations of organic solvent, care must be given since this strategy may also precipitate out some other proteins. Another strategy is to use detergents like the non-denaturing surfactant CHAPS, which due to its zwitterionic behavior can be removed using a strong cation exchange (SCX) solid phase extraction (SPE) clean-up after protein digestion (Hixson et al., 2006). 8.2.2 Reduction and Alkylation Strategies Proteins naturally contain intra- and intermolecular covalent disulfide linkages (bridging cysteines with other cysteine residues) which serve the purpose of providing protein stability and an appropriate confirmation and activity in vivo. To elucidate the protein sequence using tandem mass spectrometry it is necessary to reduce these bonds so that the protein can fully unfold for a complete digestion and accurate detection of the cysteine-containing peptide sequences. Typical reducing agents used in proteome sample preparation schemes include dithiothreitol (DTT), dithioerythritol (DTE) or β-mercaptoethanol. These reagents, although widely used, have a very pungent odor associated with them. In the case of DTT and DTE, they have to be made up fresh because they lose their reducing ability relatively quickly. More recently, phosphine derivatives like Tris(2-carboxyethyl) phosphine (TCEP) and tributylphosphine (TBP) have become more popular to use for reduction since these chemicals are stable and can be kept in neutralized solutions for many months at room temperature. In addition, these reagents can be used in conjunction with alkylating agents such as iodoacetamide (IAM) (Lopez-Ferrer et al., 2008a). Reduction reactions are commonly achieved in 45 min at 60◦ C. 130 D. Lopez-Ferrer et al. After the disulfide reduction, the free sulphydryl groups need to be blocked to protect them from undesired side reactions by the use of an alkylating agent. Alkylating reagents, such iodoacetic acid (IAA), IAM and acrylamide can be used for this purpose. Traditionally IAM is the most used reagent, the main reason being that it was the most compatible with 2-DE (one of the earliest methods of choice for proteome analyses) because it does not change the protein pI. IAM to covalently block the sulfhydryl groups, thus preventing the reformation of disulfide bonds is referred to specifically as “carbamidomethilation” (or more generally “alkylation”) and can be carried out in about an hour. Since IAM is photosensitive, alkylations using this chemical are carried out in the dark to limit any unwanted side reactions. In recent years, ultrasonic energy or microwaves have been demonstrated to be effective at accelerating reduction and alkylation steps down to just a few minutes (Rial-Otero et al., 2007). Capelo and coworkers used indirect ultrasound energy by means of a sonoreactor which is ∼30 times less powerful than an ultrasonic probe, but ∼50 times more powerful than a regular bench top ultrasonic bath. The authors show that both steps, reduction and alkylation, could be effectively accelerated to just 3 min for each step. Lopez-Ferrer and coworkers have recently demonstrated that the two processes of reduction and alkylation can be coupled by using TCEP and IAM at the same time, reducing the time for complete reduction and alkylation to only 3 min total (Lopez-Ferrer et al., 2008a). In addition, the authors showed that by reducing the time of exposure to the ultrasonic energy, little or no carbamylation occurred. More recently, Basile and coworkers, showed that these reactions could also be accelerated by the use of microwave energy (Hauser and Basile, 2008; Hauser et al., 2008). 8.2.3 Enzymatic Digestion Typical protease digestion protocols consist of the addition of a specific protease (i.e., one that cleaves a polypeptide or a protein at specific amino acid residues). For proteomic purposes several sequencing grade proteases are available. Trypsin, a serine protease that cleaves exclusively after arginine (Arg) and lysine (Lys), is by far the most widely used because it is economical (compared with other proteases), has been modified to be very robust, and is well characterized. Additionally, Arg and Lys are typically spread fairly evenly throughout most protein sequences, thus a trypsin digest usually provides an array of detectable peptides for each protein present (Kiser et al., 2009). Other proteases commonly used in proteomic research but with less frequency include endoproteinase Lys-C (cleaves at lysine (Boyne et al., 2009)), S. aureus V8 (also known as endoproteinase Glu-C, and cleaves at glutamic acid (Glu) and aspartic acid (Asp) residues (Xiang et al., 2004)), and proteinase K (Wu et al., 2003; Wu and Yates, 2003) (cleaves aromatic, and aliphatic residues, but with less specificity for the latter). As an additional note, cyanogen bromide (although not an enzyme) has been commonly employed to cleave proteins at methionine residues, especially in applications that are focused on proteome characterizations within membranes (van Montfort et al., 2002). 8 Ultra-Fast Sample Preparation for High-Throughput Proteomics 131 In proteomic experiments, enzyme digestions are typically the most timeconsuming step within the sample preparation process because it is often recommended to digest a sample for many hours to ensure complete digestion. It is well known that the enzyme activity increases with temperature, only up to a point where the temperature becomes high enough to denature the protease enzyme resulting in a dramatic decrease in activity. Some researchers have investigated the use of thermo stable trypsin. Other pursuits at enhancing trypsin activity include efforts to modify the pH or the buffer composition, such as adding organic solvents or salts like CaCl2 to stabilize the enzyme-substrate complex (Russell et al., 2001; Wang et al., 2005). Despite these efforts, digestion times still usually require at least 30 min to be effective and complete (Havlis et al., 2003). Unconventional methods for enhancement of enzyme kinetics such as microwave, ultrasound and, more recently, pressure have been explored in the last 5 years as an alternative way for applying energy to a digestion reaction in order to accelerate the enzymatic hydrolysis reaction. 8.2.3.1 Microwave Irradiation For many years now, researchers have utilized microwaves to hasten reaction rates in their protein chemistry and proteomics experiments (Lill et al., 2007; Zhong et al., 2004). The instrumentation used ranges from domestic microwave ovens to sophisticated laboratory microwave equipment. In both cases, it has been shown that the effect of microwave radiation applied to proteases can dramatically accelerate their enzyme kinetics. At this point, the field is divided by those who believe that the improvement of the reaction kinetics is due solely to the thermal effect produced by the efficient absorption of the radiation through the medium, and those who believe that the main reason for enhanced enzyme kinetics is through both thermal and nonthermal effects derived from both energy transfer from the electromagnetic field and vibration modes of the molecules altering their dipoles. The latter hypothesis was demonstrated when Pramanik and coworkers fixed the temperature of a trypsin digestion reaction at 37◦ C while controlling the level of microwave irradiation (Pramanik et al., 2002). Their results indicate that heat is not the main player driving the reaction. Microwave assisted protein digestion (MAPED) has been applied in proteomics for in-solution and in-gel digestion of proteins. MAPED has been optimized by using different organic co-solvents as well as by using immobilized trypsin immobilized on magnetic micro- or nanospheres. Microwaves have also been used to accelerate acid hydrolysis of peptides in order to generate peptide ladders, where the difference between the detected ladder masses is the mass of the amino acid that has been cleaved. This methodology provides a very accurate peptide sequence and is excellent for protein characterization. Its application for high-throughput proteomics is more limited because the absence of specificity on the peptide cleavage makes it extremely challenging to interpret data resulting from a complex protein mixture digest (Zhong et al., 2004). Recently, Fenselau and coworkers have also explored what seems to be a very promising approach that utilizes low concentrations of formic or acetic acid to produce specific chemical cleavages in proteins (Swatkoski et al., 2007a, b, 2008). It has been 132 D. Lopez-Ferrer et al. [M-3H+]3+ 592.984 X 10 [M-3H+]3+ 554.610 115 Da [M-2H+]2+ 510.828 [M-2H+]2+ 568.342 115 Da 38.33 amu 550 575 57 amu 600 m/z 540 115 Da 580 m/z 28.75 amu [M-4H+]4+ 1117.598 [M-4H+]4+ 1088.841 100 Relative Abundance 500 1088 1089.5 1091 m/z 1117 1119.5 1120 m/z 50 0 20 40 Retention Time (min) 60 Fig. 8.2 Chromatographic profile of a microwave D-digestion of four proteins. Detailed mass spectra show pairs of signals 115 Da apart. This indicates that cleavage was occurring before and after aspartic acid residues. Also shown are a variety of peptide masses ranging from 1000 up to 4500 Da demonstrated that these chemical cleavages under certain conditions can be very specific, preferring amino acid sites before and after Asp. This approach is particularly interesting because Asp residues are not as common as Arg or Lys, providing a completely different map of the proteins that can yield a better characterization of the protein primary structure. Additionally, because it cleaves before and after the Asp residues it is very common to see doublets of a certain peptide in the mass spectrum, which improves the forthcoming protein identification. Figure 8.2 shows the digestion of four protein standards using this methodology. Basile and coworkers have taken this method a step further by developing an on-line acid digestion system (Hauser and Basile, 2008; Hauser et al., 2008). 8.2.3.2 Ultrasound The use of ultrasound energy for accelerating enzyme kinetics in proteomics was introduced for the first time by Lopez-Ferrer and coworkers in 2005 (Lopez-Ferrer et al., 2005). The study demonstrated that acceleration of a trypsin digestion was feasible for both in-gel digestions of single proteins and for in-solution digestions of either single proteins or complex global proteomes. The authors were able to apply ultrasonic energy to small volumes (less than 50 µL) using an ultrasonic probe. It is hypothesized that the effectiveness of this method is based upon the formation of localized high pressure and temperature zones in which the enzyme-substrate 8 Ultra-Fast Sample Preparation for High-Throughput Proteomics 133 kinetics is enchanced. This process is called cavitation and it is visible when transient bubbles appear in the solution during the ultrasonic irradiation. Lopez-Ferrer and coworkers further demonstrated that the reaction time could be shortened even further if immobilized enzymes were used (Lopez-Ferrer et al., 2009). By using immobilized enzymes, the enzyme:substrate ratio increases dramatically and also there is a better mass transfer achievable (Kim et al., 2009) which altogether aids in a significantly more efficient digestion in the presence of ultrasound. Most recently, a protocol was developed where the use of high intensity focused ultrasound (HIFU) was applied to perform quantitative proteomic experiments using an 16 O/18 O labeling strategy. In this application, the enzyme is used not in the digestion of the proteins but in the post-digestion exchange of oxygen atoms at the C-termini. This hydrolysis/oxygen exchange reaction creates a 2 or 4 Da shift depending on the degree of reaction completion per peptide. These mass shifts can then be used to identify peptide abundance changes in a simultaneous mass spectrometric analysis of two mixed samples (namely, one that is labeled with 16 O added in equal mass to one that is labeled with 18 O). 8.2.3.3 High Pressure For a number of years, MAPED and HIFU have been used in proteomic sample processing schemes and have proven useful for increasing the kinetics of proteolytic digestions. In HIFU, along with an increase in heat, short-lived micro-environments of high pressure are also experienced by the sample. In food and biotechnology industries, hydrostatic pressure have been used to speed-up or inhibit certain reactions (e.g., enhancement of the curd formation in the manufacturing of cheese, inactivation of food pathogens orinhibition of enzymes). Evidence also exists to suggest that low to moderate pressures (e.g., 100–400 MPa) can activate enzyme activity (Cano et al., 1997), whereas higher pressures tend to inactivate the same enzymes (Hernandez and Cano, 1998). Lopez-Ferrer et al. demonstrated that proteome samples subjected to increased pressures (i.e., up to ∼20,000 psi) could effectively hasten trypsin digestion (Lopez-Ferrer et al., 2008b). It was found that under these high pressures, globular sample proteins can become more denatured (the thermodynamic understanding of this process is currently unknown), allowing for the modified sequencing grade trypsin (trypsin that has been reductively methylated and enhanced by N-tosyl-L-phenylalanyl chloromethyl ketone (TPCK) treatment) to rapidly digest the proteins. 8.3 Future Trends Considering the current and growing demand for proteomic analyses, especially those that utilize shotgun proteomic methodologies, the development of in-solution methods that enhance and hasten the digestions carried out during proteome sample preparation still needs to be optimized, automated and integrated with mass spectrometric analyses. The uses of microfluidic devices that can presently integrate different steps in the proteome workflow are currently restricted to specialized labs. 134 100 225 75 150 50 75 25 0 Rep A Rep B 1 min Rep A Rep B 0 #Total Identifications b) # Unique Peptides % Coverage a) D. Lopez-Ferrer et al. 2 min Digestion Time Fig. 8.3 (a) Schematic of a sonoreactor being used in an on-line set-up. (b) Complete digestion of protein samples in the presence of the low pH compatible protease pepsin can be achieved in 1–2 min as is indicated in (b), where is shown the level of proteolysis obtained by using the on-line sonoreactor setup with a bovine serum albumin sample digested with pepsin However, we have proposed two simple alternatives that can be easily implemented into every proteomic facility based on the use of either ultrasounds or pressure. Ultrasound probes are usually commonplace in most molecular biology laboratories and these same probes can be used for effective digestion and/or 18 O labeling using trypsin to produce peptides in a matter of minutes. The direct hyphenation with LC-MS is also feasible as it is shown in Fig. 8.3. In this case, digestion of 1 pmol of bovine serum albumin was performed with pepsin (being that pepsin is already compatible with the low pH solvents used for the RP separations), and the flow rate was modulated in order to allow the sample to be under the ultrasonic field for 1 or 2 min. As it is shown in Fig. 8.3b, the digestion is extremely efficient providing a proteome coverage of over 70% in less than a minute. Initial demonstrations of enhanced digestions using pressure conducted by Lopez-Ferrer et al. were carried out using a BarocylcerTM , an instrument capable of subjecting samples to a temperature and pressure controlled environment which can be cycled between ambient pressure and 35,000 psi. This instrument, originally marketed for cell lysis conducted in enclosed disposable sample containers (Smejkal et al., 2006, 2007), is currently being adopted by many laboratories as the way to control enzymatic digestion of protein samples. Moreover, Lopez-Ferrer et al., recently demonstrated that high-pressure digestion could potentially be automated by using a modified high pressure HPLC equipment (Lopez-Ferrer et al., 2008c). Since high-pressure is integral to an HPLC system, protein samples added with trypsin to a sample loop can be pressurized briefly followed by direct injection onto a reversed phase C18 column. This set-up demonstrated the proof of concept for future automated sample preparation devices for high-throughput proteome sample preparation. With developments such as these, perhaps in the near future we will be able to routinely obtain a full proteome analyses (from whole cells to MS data) or 8 Ultra-Fast Sample Preparation for High-Throughput Proteomics a) 135 b) 649.48 651.48 652.48 m/z 548.12 547.45 733.58 548.79 737.58 735.58 m/z m/z 0 10 5 15 Retention Time (min) 20 Fig. 8.4 Schematic showing the valve diagram and coupling to an LC-MS system for an on-line 18 O differential proteomics digestion system. (b) Chromatographic profile of a BSA digestion either with 18 O or regular water. Detailed mass spectra of different peptide pairs is shown for +2 and +3 ions even comparative (using automated pressure assisted digestions in an on-line system with 18 O/16 O labeling) proteome analyses in just a manner of minutes (Fig. 8.4). 8.4 Protocol for Rapid Sample Preparation of Peptides for Shotgun Proteomics 8.4.1 Materials 8.4.1.1 Reagents Sequencing grade trypsin was obtained from Promega (Madison, WI). Iodoacetamide (IAM), ammonium bicarbonate, formic acid, HPLC grade solvents, 18 O water, pepsin and bovine serum albumin were purchased from Sigma-Aldrich (St. Louis, MO). Tris[2-carboxyethyl]phosphine (TCEP) was purchased from Pierce (Rockford IL, USA). ProteoSolve-SB kit was obtained from Pressure BioSciences (South Easton, MA, USA). 8.4.1.2 Equipment BarocyclerTM NEP-3229 instrument, disposable polypropylene PULSE tubes FT500, PCT MicroTubeTM and a PCT MicroTubeTM adapter kit were obtained from Pressure BioSciences (South Easton, MA, USA). A UTR200 Sonoreactor was purchase from Hischler (Teltow, Germany) 136 D. Lopez-Ferrer et al. 8.4.2 Procedure 8.4.2.1 Cell Lysis • Pellet the cells at a defined growth phase. • Resuspend the pellet in 1100 µL of Reagent A of the ProteoSolve-SB kit, sonicate in the sonoreactor, if necessary. • Transfer to PULSE Tube, add 200 µL of Reagent B of the ProteoSolve-SB kit and cap the tube. • Place the tube into the Barocycler and initiate the lysis program. • The lysis program consist of 20 cycles of pressure at 35 kpsi for 30 s and 5 s at atmospheric pressure. • Transfer the sample to a 2 mL microcentrifuge tube and centrifuge for 10–15 min at 15,000 rpm. • Transfer the bottom phase to a new reaction tube (protein content) and dry it down in the speedvac. 8.4.2.2 Reduction and Alkylation • Resuspend the sample in 100 µL of 8 M urea, sonicate the pellet into solution, if necessary. (Do not vortex) • Measure the protein concentration at this point (Sapan et al., 1999) • Add 1 µL of Bond Breaker TCEP solution (Final concentration 5 mM) • Add 4 µL of 500 mM IAM solution (Final concentration 20 mM) • Sonicate for 3 min in the sonoreactor at 50% power 8.4.2.3 Protein Digestion Dilute the sample 4-fold to 400 µL with 50 mM ammonium bicarbonate. Add a 1:50 mass ratio of trypsin to the sample protein. Divide the sample between 4 and 100-µL BarocyclerTM tubes. Place into the BarocyclerTM and initiate the digestion program. The digestion program consist of 10 cycles of pressure at 20 kpsi for 20 s and 5 s at atmospheric pressure. • Evaporate the samples down to ∼150 mL in a SpeedVac concentrator. • Measure and adjust the concentration of the peptide sample to optimum concentration for LC-MS system used. • • • • • Acknowledgments Portions of this work were supported by the NIH National Center for Research Resources (NCRR, RR018522), NIH National Cancer Institute (R21 CA12619-01), and the Pacific Northwest National Laboratory’s (PNNL) Laboratory Directed Research and Development Program. This research was enabled in part by capabilities developed under support from the U.S. Department of Energy (DOE) Office of Biological and Environmental Research and the NCRR, and was conducted in the Environmental Molecular Sciences Laboratory, a DOE national scientific user facility located at the Pacific Northwest National Laboratory (PNNL) in Richland, WA. 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