Protein Sample Preparation for Mass Spectrometry

Mass spectrometry (MS) has become the method of choice for protein detection, identification and quantitation. The accuracy, sensitivity and flexibility of MS instruments have enabled new applications in biological research, biopharmaceutical characterization and diagnostic detection. With its many ionization and measurement (e.g., ESI, MALDI, FT-MS, ion trap, quadrupole) options, MS can be used to analyze proteins and peptides ranging in mass from 50 to 300 kDa and in attomole to nanomole quantities.

Proper sample preparation for MS-based analysis is a critical step in the proteomics workflow because it can be both variable and time consuming. The quality and reproducibility of sample extraction and preparation significantly impacts MS results. The most successful proteomics laboratories recognize that sample preparation, instrumentation and software are all critical to proteomics research success; therefore, all three components must be properly integrated into robust workflows for consistent, high quality results.

Successful proteomic analysis emphasizes sample preparation, instrumentation and software.

Proteins of interest to biological researchers are generally present in a complex mixture of other proteins, which presents two significant problems in MS analysis:

1. The ionization techniques used for large molecules work well when the mixture contains roughly equal amounts of constituents. However, the dynamic range of protein concentrations in biological samples can exceed 10 orders of magnitude. If such a mixture is ionized using electrospray ionization (ESI), for example, the more abundant species have a tendency to "drown out" or suppress signals from less abundant ones.
2. The mass spectrum of a complex mixture is very difficult to fully analyze due to its overwhelming number of components. This problem is exacerbated by enzymatic digestion of a protein sample into a large number of peptide products. The success of liquid chromatography MS (LC-MS) and tandem MS (LC-MS/MS) depends on clean samples with limited sample complexity to minimize the suppression of ionization by high-abundance species and prevent MS undersampling of eluting peptides.

Protein preparation for MS analysis can be accomplished by many methods, so it is important to understand the steps leading to analysis. While intact proteins are typically studied by gel electrophoresis, the most common mass spectrometry workflows for complex protein samples analyze peptides, which are easier than proteins to fractionate by LC. Peptides also ionize and fragment more efficiently than whole proteins, resulting in spectra that are easier to interpret for protein identification. Peptide preparation involves reduction and alkylation of cysteines, digestion of the sample into peptides, desalting and concentration of the peptides and final analysis of these peptides by ionization (e.g., ESI) plus orbitrap-based MS.

The decision to use MALDI-MS or LC-MS or LC-MS/MS for proteomic analysis depends on a number of factors, including the level of sample preparation and throughput potential. For example, because multiple samples can be dried onto a single MALDI matrix, 96 samples can be analyzed in an hour compared to only one sample by LC-MS or LC-MS/MS. Conversely, all reduction in sample complexity must be performed off-line with MALDI-MS, and therefore sample preparation is considerably more critical for this approach than for LC-MS or LC-MS/MS, which requires minimal sample preparation because of the in-line reverse-phase LC used to reduce sample complexity prior to MS analysis.

Sample preparation workflow. Lysate samples are prepared from biological specimens or cultured cells by a customized protocol that may include cell lysis, subcellular fractionation, depletion of high-abundance proteins, enrichment of target proteins, dialysis and desalting. In-solution digestion entails irreversibly breaking disulfide bonds via reduction and alkylation followed by protein digestion into peptide fragments. An alternative approach is to first resolve proteins by 1D or 2D electrophoresis (1DE or 2DE, respectively) and then collect gel slices that contain the desired band(s). The proteins in these gel plugs are then reduced, alkylated and digested in situ. After peptides are extracted from the gel matrix, the peptides are enriched and salts and detergents removed and prepared for MS analysis. While this diagram encompasses the most common steps of sample preparation, the protocol must be tailored to specific samples and MS techniques for optimal results.

Cell lysis is the first step in protein extraction, fractionation and purification. Numerous techniques have been developed to obtain the best possible yield and purity for different organisms, sample types (cells or tissue), subcellular structures or specific proteins. Both physical and reagent-based methods may be required to extract cellular proteins because of the diversity of cell types and cell membrane (or cell wall) composition .

Physical lysis is a common method of cell disruption and extraction of cellular contents. However, it requires specialized equipment and protocols that are difficult to repeat because of variability in the apparatus (e.g., different dounce pestles or sonication settings). Also, traditional physical disruption methods are typically not conducive to small sample volumes and high-throughput sample handling. Finally, physical lysis methods alone are unable to solubilize membrane-associated proteins. In contrast, reagent-based lysis methods using detergents not only lyse cells but also solubilize proteins. By using different buffers, detergents, salts and reducing agents, cell lysis can be optimized to provide the best possible results for a particular cell type or protein fraction.

Cell lysis disrupts cellular compartments, which can activate endogenous proteases and phosphatases. To protect extracted proteins from degradation or artifactual modification by the activities of these enzymes, it is necessary to add protease and/or phosphatase inhibitors to the lysis reagents.

When the goal of cell lysis is to purify or test the function of a particular protein(s), special attention must be given to the effects that the lysis reagents have on the stability and function of the target proteins. Certain detergents will inactivate the function of particular enzymes or disrupt protein complexes. Downstream analysis of extracted/purified proteins may also require detergent removal in order to study proteins of interest or maintain long-term stability of the extracted protein.

Sample complexity negatively affects the ability to detect, identify and quantify low-abundance proteins by MS, because peptides from high-abundance proteins can mask detection of those from low-abundance proteins. Therefore, the more that a sample can be simplified and the greater that the dynamic range of protein concentrations can be reduced, the greater will be the ability to detect proteins at very low concentrations.

Depletion and enrichment strategies have been developed to remove high abundant proteins or isolate target proteins in the sample, respectively. Depletion is more often used to reduce the complexity of biological samples such as blood or serum, which contain high concentrations of albumin and immunoglobulins. Depletion strategies utilize immunoaffinity techniques such as immunoprecipitation and co-immunoprecipitation (IP and co-IP, respectively), and commercial kits are available to remove these and other high abundant proteins from samples. A significant drawback to this approach, though, is that abundant proteins often bind to other proteins, which could result in the depletion of complexes with low-abundance proteins.

Protein enrichment encompasses numerous techniques to isolate subclasses of cellular proteins based on unique biochemical activity, post-translational modifications (PTMs) or spatial localization within a cell. Post-translational modifications such as phosphorylation and glycosylation can be enriched using affinity ligands such as ion-metal affinity chromatography (IMAC) or immobilized lectins, respectively. In addition, PTM-specific antibodies have been used. Other techniques entail metabolic or enzymatic incorporation of modified amino acids or PTMs to introduce unique protein chemistries that can be used for enrichment. Finally, proteins can also be enriched using various enzyme class-specific compounds or cell-impermeable labeling reagents that selectively label cell surface proteins.

Separation of distinct subcellular fractions is another method of enrichment and can be achieved through the careful optimization of physical disruption techniques, detergent-buffer solutions and density gradient methods. For example, with the phase-separating detergents, hydrophobic membrane proteins can be solubilized and extracted from hydrophilic proteins. Density gradient centrifugation is another technique and can be used to isolate intact nuclei, mitochondria and other organelles before protein solubilization.

Whether they are simple or complex, samples often need to be processed in several ways to ensure they are compatible and optimized for digestion and analysis by mass spectrometry. For example, because MS measures charged ions, salts—especially sodium and phosphate salts—should be removed prior to MS to minimize their detection.

Dialysis and desalting products allow buffer exchange, desalting, or small molecule removal to prevent interference with downstream processes. Protein assays help monitor protein concentration for consistent control of experimental loading or yield.

Sample proteins are denatured in one of two ways, depending on whether in-solution or in-gel digestion is performed. For in-solution digestion strategies, proteins are denatured with strong chaotropic agents such as urea or thiourea. This step is either followed by or combined with disulfide reduction using a reducing agent such as Tris(2-carboxyethyl)phosphine (TCEP) or dithiothreitol (DTT). The free sulfhydryl groups on the cysteine residues are then alkylated with reagents such as iodoacetamide or iodoacetic acid to irreversibly prevent the free sulfhydryls from reforming disulfide bonds. The denatured, reduced and alkylated proteins are then digested by endoproteinases, (e.g., trypsin, chymotrypsin, Glu-C and Lys-C), which hydrolytically break peptide bonds to fragment proteins into peptides.

Protein separation by 1- or 2-dimensional gel electrophoresis (1DE and 2DE, respectively) is an alternative to in-solution protein denaturation, reduction, alkylation and digestion. This approach employs SDS polyacrylamide gel electrophoresis (SDS-PAGE) to denature and separate proteins in a sample. After electrophoresis, protein bands or spots are visualized using Coomassie, fluorescence or silver stains. Protein bands or spots are then excised from the gel and destained, and the proteins in the gel plugs are reduced, alkylated and digested in situ. The peptides are then extracted from the gel matrix and prepared for MS analysis.

The decision to perform in-solution or in-gel digestion depends on a number of factors, including the sample amount and complexity. In-solution digestion is useful when the sample amount is small because peptide extraction from the gel matrix after in-gel digestion can result in significant peptide loss. In-solution digestion is also good for samples with low-to-moderate complexity and when detergents would negatively affect the sample. The benefit of in-gel digestion is that SDS-PAGE combines protein denaturation with separation, which visually indicates the relative abundance of proteins in the sample. Also, peptide extraction inherently removes much of the detergents and salts, although peptide recovery is affected. Regarding processing time, in-solution digestion can be performed more rapidly because SDS-PAGE is not required. It also has greater high-throughput potential because of automation of the entire process (although automated systems have also been developed for in-gel digestion and extraction).

Enrichment of specific target peptides and sample clean-up are required for successful analysis of low-abundance proteins or identification of post-translationally modified peptides. Enrichment for specific PTMs (e.g., phosphorylation, ubiquitination and glycosylation) is performed by affinity purification using PTM-specific antibodies or ligands. For example, phosphopeptides can be enriched by IP using anti–phospho-specific antibodies or by pull-down using TiO2, which selectively binds phosphorylated serine, tyrosine or threonine. After peptide enrichment, salts and buffers can be removed using either graphite or C-18 tips or columns, and detergents can be removed using affinity columns or detergent-precipitating reagents. Dilute samples can also be concentrated using concentrators of varying molecular weight cutoff (MWCO) ranges.

Purified peptide samples are then ready for the final preparation for MS analysis, which varies based on the type of analysis. For LC-MS or LC-MS/MS analysis, the correct choice of mobile phases and ion-pairing reagents is a required to achieve good LC resolution and analytical results.

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