One of the key applications used to characterize single or complex protein mixtures via bottom up proteomics is liquid chromatography−tandem mass spectrometry (LC−MS/MS).
Recent technical advances allow for identification of >10 000 proteins in a cancer cell line. On the peptide level chromatography methods, like strong cation exchange (SCX)
and hydrophilic interaction chromatography (HILIC), as well as high-pH reversed phase chromatography have been employed successfully. Because of its robustness
and ease of handling, the classical and still widely used approach for protein fractionation prior to LC− MS/MS is gel-based separation under denaturing conditions (SDS-PAGE).
Hydrophobic interaction chromatography (HIC) is a robust standard analytical method to purify proteins while preserving their biological activity. It is widely used
to study post-translational modifications of proteins and drug−protein interactions. HIC is a high-resolution chromatography mode based on the interaction of
weakly hydrophobic ligands of the stationary phase with hydrophobic patches on the surface of the tertiary structure of proteins. By employment of high concentrations
of structure-promoting (“kosmotropic”) salts, proteins in HIC retain their conform
In a recent publication, HIC was used to separate proteins, followed by bottom up LC−MS/MS experiments (1). HIC was used to fractionate antibody species
followed by comprehensive peptide mapping as well as to study protein complexes in human cells. The results indicated that HIC−reversed-phase chromatography (RPC)
mass spectrometry (MS) is a powerful alternative to fractionate proteins for bottom-up proteomics experiments making use of their distinct hydrophobic properties.
An additional observation noted that tryptic digests of the antibody used in the study yielded a protein coverage of 56% for the light chain and 63.2% for the
heavy chain. A consecutive proteolytic digestion protocol combing on-filter trypsin and elastase digestion drastically improved sequence coverage of
both light (100%) and heavy chains (99.2%).
1. Rackiewicz, M. et al. (2017) Hydrophobic Interaction Chromatography for Bottom-Up Proteomics Analysis of Single Proteins and Protein Complexes. J.Proteome.Res.16, 2318–23.
Bottom-up proteomics focuses on the analysis of protein mixtures after enzymatic digestion of the proteins into peptides. The resulting complex mixture of peptides is analyzed by reverse-phase liquid chromatography (RP-LC) coupled to tandem mass spectrometry (MS/MS). Identification of peptides and subsequently proteins is completed by matching peptide fragment ion spectra to theoretical spectra generated from protein databases.
Trypsin has become the gold standard for protein digestion to peptides for shotgun proteomics. Trypsin is a serine protease. It cleaves proteins into peptides with an average size of 700-1500 daltons, which is in the ideal range for MS (1). It is highly specific, cutting at the carboxyl side of arginine and lysine residues. The C-terminal arginine and lysine peptides are charged, making them detectable by MS. Trypsin is highly active and tolerant of many additives.
Even with these technical features, the use of trypsin in bottom-up proteomics may impose certain limits in the ability to grasp the full proteome, Tightly-folded proteins can resist trypsin digestion. Post-translational modifications (PTMs) present a different challenge for trypsin because glycans often limit trypsin access to cleavage sites, and acetylation makes lysine and arginine residues resistant to trypsin digestion.
To overcome these problems, the proteomics community has begun to explore alternative proteases to complement trypsin. However, protocols, as well as expected results generated when using these alternative proteases have not been systematically documented.
In a recent reference (2), optimized protocols for six alternative proteases that have already shown promise in their applicability in proteomics, namely chymotrypsin, Lys-C, Lys-N, Asp-N, Glu-C and Arg-C have been created.
Data describe the appropriate MS data analysis methods and the anticipated results in the case of the analysis of a single protein (BSA) and a more complex cellular lysate (Escherichia coli). The digestion protocol presented here is convenient and robust and can be completed in approximately in 2 days.
Filter-aided sample preparation (FASP) method is used for the on-filter digestion of proteins prior to mass-spectrometry-based analyses (1,2). FASP was designed for the removal of detergents, and chaotropes that were used for sample preparation. In addition, FASP removes components such as salts, nucleic acids and lipids. Akylation of reduced cysteine residues is also carried out on filter, after which protein is proteolyzed by use of trypsin on filter in the optimal buffer of the enzyme. Subsequent elution and desalting of the peptide-rich solution then provides a sample ready for LC–MS/MS analysis.
Erde et al. (3) described an enhanced FASP (eFASP) workflow that included 0.2% DCA in the exchange, alkylation, and digestion buffers,thus enhancing trypsin proteolysis, resulting in increases cytosolic and membrane protein representation. DCA has been reported (4) to improve the efficiency of the denaturation, solubilization, and tryptic digestion of proteins, particularly proteolytically resistant myoglobin and integral membrane proteins, thereby enhancing the efficiency of their identification with regard to the number of identified proteins and unique peptides.
In a recent publication (5) traditional FASP and eFASP were re-evaluated by ultra-high-performance liquid chromatography coupled to a quadrupole mass filter Orbitrap analyzer (Q Exactive). The results indicate that at the protein level, both methods extracted essentially the same number of hydrophobic transmembrane containing proteins as well as proteins associated with the cytoplasm or the cytoplasmic and outer membranes.
The LC–MS/MS results indicate that FASP and eFASP showed no significant differences at the protein level. However, because of the slight differences in selectivity at the physicochemical level of peptides, these methods can be seen to be somewhat complementary for analyses of complex peptide mixtures.
Biomarkers in biological fluids in particular have the potential to inform regarding risk of disease or to allow early detection for more effective treatment. Plasma/serum is considered the universal source of biomarkers. This fluid is, indeed, easily collected, and the important point is that plasma collects proteins from each and every tissue, compared to other fluids such as urine or cerebrospinal fluid. Optimizing experimental conditions (i.e., use of trypsin for the digestion of target proteins) used to discover or monitor biomarkers in plasma is critical to successful detection of biomarkers.
In a recent publication by Proc et al., plasma denaturation/digestion protocols were compared using quanititation methods. In this reference 14 combinations of heat, solvent (acetonitrile, methanol, trifluoroethanol), chaotropic agents (guanidine hydrochloride, urea) and surfactants (sodium dodecyl sulfate (SDS)and sodium deoxycholate (DOC) with effectiveness in improving tryptic digestion. Digestion efficiency was monitored by quantitating the peptides from 45 moderate- to high-abundance plasma proteins using tandem mass spectrometry in multiple reaction mode with a mixture of stable isotope labeled analogues of these peptides as internal standards. In the results, Proc et al. noted that use of either DOC and SDS produced an increase in the overall yield of tryptic peptides. Since SDS is not compatible with mass spectrometry and DOC can be easily remove by acid precipitation, the overall recommendation was the use of DOC with a nine hour digestion procedure.
N-Glycosylation is a common protein post-translational modification occurring on asparagine residues of the consensus sequence asparagine-X-serine/threonine, where X may be any amino acid except proline. Protein N-glycosylation takes place in the endoplasmic reticulum (ER) as well as in the Golgi apparatus.
Approximately half of all proteins typically expressed in a cell undergo this modification, which entails the covalent addition of sugar moieties to specific amino acids. There are many potential functions of glycosylation. For instance, physical properties include: folding, trafficking, packing, stabilization and protease protection. N-glycans present at the cell surface are directly involved in cell−cell or cell−protein interactions that trigger various biological responses.
The standard method used to profile the N-glycosylation pattern of cells is glycoprotein isolation followed by denaturation and/or tryptic digestion of the glycoproteins and an enzymatic release of the N-glycans using PNGase F followed by analysis mass spec. This method has been reported to yield high levels of high-mannose N-glycans that stem from both membrane proteins as well as proteins from the ER.(1,2)
For those researchers interested in characterizing only cell surface glycans (i.e., complex N-glycans) a recent reference has developed a model system using HEK-292 cells that demonstrates a reproducible, sensitive, and fast method to profile surface N-glycosylation from living cells (3). The method involves standard centrifugation followed by enzymatic release of cell surface N-glycans. When compared to the standard methods the detection and quantification of complex-type N-glycans by increased their relative amount from 14 to 85%.
The complexity of biological samples places high demand on mass spec analytical capability. Adequate monitoring of instrument performance for proteomics studies requires equally complex reference material such as whole-cell extracts. However, whole-cell extracts available commercially are developed for general research (e.g., enzymatic or Western blot analysis) and contain detergents and salts that interfere with reverse phase liquid chromatography and mass spectrometry. Even after clean up, the extracts need to be digested, requiring time, labor and experience to generate samples for use in mass spectrometry. To address the need for complex protein material, we have developed whole-cell protein extracts from yeast and human cells. The yeast extract offers the convenience of a relatively small and well annotated proteome, whereas the human extract provides a complex proteome with large dynamic range. The human extract also serves as reference material for studies targeting the human proteome.
The extracts are free of compounds that interfere with reverse phase liquid chromatography-mass spectrometry (LC-MS), and have been reduced with DTT and alkylated with iodoacetamide then digested with Trypsin/Lys-C Mix and lyophilized. These digested extracts (tryptic peptides) can be readily reconstituted in trifluoroacetic acid (TFA) or formic acid and injected into an instrument. The same human and yeast whole-cell extracts also are provided in an intact (undigested) form for users who would like to develop an independent method for preparing protein mass spectrometry samples. For convenience, the intact extracts are provided as a frozen solution.
Consistent extract protein composition is ensured by tight control over cell culture conditions and manufacturing process. Lot-to-lot consistency of extracts is monitored by various protein and peptide qualitative and quantitation methods, including LC-MS. (Quality control results are provided upon request.) Our manufacturing process assures compatibility with reverse phase liquid chromatography and mass spectrometry, minimal nonspecific protein fragmentation, nonbiological post-translational modifi cations and,for digested extracts, minimal undigested peptides. The extracts are optimized for a high number of peptide and protein identifications in mass spectrometry analysis.
The novel mass spectrometry compatible surfactant sulfonate-(sodium 3-((1-(furan-2-yl)undecyloxy) carbonylamino)-propane-1-sulfonate (i.e.ProteaseMAX) facilitates both in-gel and in-solution digestion applications by reducing the time required, enabling protein solubilization/denaturation and increasing peptide/protein identifications.
A new application was highlighted in a recent publication (1) which utilized ProteaseMAX to lyse cells prior to trypsin digestion and subsequent mass spec analysis. The composition of the buffer determines the overall efficiency of cell lysis, dissociation of protein complexes, protein solubility and ease of removal prior to LC/MS-MS analysis.
When compared to lysis buffers containing either urea or SDC, ProteaseMAX provided the optimal number of identified peptides/proteins.
In addition it can be easily removed from the lysate by acidic precipitation.
Pirmoradian, M. et al. (2013). Rapid and deep human proteome analysis by single-dimension shotgun proteomics. Mol. Cell. Prot.12, 3330–8.
Neonatal sepsis is a systemic infection prevalent in preterm and very low birth weight infants and causes high morbidity. Most cases of neonatal sepsis are caused by pathogenic bacteria that invade the bloodstream, triggering an abrupt and overwhelming infection in the target organs accompanied by a systemic inflammatory response. Testing for neonatal sepsis is challenging because it does not affect a specific organ and presents multiple symptoms that are often confused with other related conditions (1). Current diagnostic tests for sepsis include those that identify markers of the host response to infection (e.g., procalcitonin, C reactive protein, cytokines, etc.) and those that detect bacterial infection in blood (bacteremia) (2). The lack of specific diagnostic biomarkers for early and accurate detection of neonatal sepsis has spurred the quest for next-generation biomarkers using powerful mass screening technologies such as proteomics. Continue reading “Testing for Neonatal Sepsis: The Next Generation of Biomarkers”
Trypsin/Lys-C Mix, Mass Spec Grade, is a mixture of Trypsin Gold, Mass Spectrometry Grade, and rLys-C, Mass Spec Grade. The Trypsin/Lys-C Mix is designed to improve digestion of proteins or protein mixtures in solution.It is a little known fact that trypsin cleaves at lysine residues with lesser efficiency than at arginine residues. Inefficient proteolysis at lysine residues is the major cause of missed (undigested) cleavages in trypsin digests.
Supplementing trypsin with Lys-C enables cleavage at lysines with excepetional efficiency and specificity. Following the conventional trypsin digestion protocol (i.e., overnight incubation at nondenaturing conditions, reduction,alkylation, 25:1 protein:protease ratio [w/w], mix and incubate overnight at 37°C.) Replacing trypsin with Trypsin/Lys-C Mix in this conventional protocol leads to multiple benefits for protein analysis including more accurate mass spectrometry-based protein quantitation and improved protein mass spectrometry analytical reproducibility.
The ISOQUANT® Isoaspartate Detection Kit is intended for quantitative detection of isoaspartic acid residues in proteins and peptides, which can result from the gradual, nonenzymatic deamidation of asparagine or rearrangement of aspartic acid residues.
The ISOQUANT® Kit is designed to provide information regarding the global formation of isoaspartic acid residues at Asn and Asp sites, not at each site separately.
The deamidation of asparagine residues and rearrangement of aspartic acid residues is characterized by the formation of a succinimide intermediate that resolves to form a mixture of isoaspartic acid (typically 70–85%) and aspartic acid.
The rate and extent of isoaspartic acid formation can vary widely among proteins, depending on the amino acid sequence and size of the target protein. Deamidation of Asn residues has been observed most frequently at Asn-Gly and Asn-Ser sites within proteins.
The ISOQUANT® Isoaspartate Detection Kit uses the enzyme Protein Isoaspartyl ethyltransferase (PIMT) to specifically detect the presence of isoaspartic acid residues in a target protein. PIMT catalyzes the transfer of a methyl group from S-adenosyl-L-methionine (SAM) to isoaspartic acid. Spontaneous decomposition of this methylated intermediate results in the release of methanol and reformation of the succinimide.