Abstract
To identify proteins by the bottom-up mass spectrometry workflow, enzymatic digestion is essential to break down proteins into smaller peptides amenable to both chromatographic separation and mass spectrometric analysis. Trypsin is the most extensively used protease due to its high cleavage specificity and generation of peptides with desirable positively charged N- and C-terminal amino acid residues that are amenable to reverse phase HPLC separation and MS/MS analyses. However, trypsin can yield variable digestion profiles and its protein cleavage activity is interdependent on trypsin source and quality, digestion time and temperature, pH, denaturant, trypsin and substrate concentrations, composition/complexity of the sample matrix, and other factors. There is therefore a need for a more standardized, general-purpose trypsin digestion protocol. Based on a review of the literature we delineate optimal conditions for carrying out trypsin digestions of complex proteomes from bulk samples to limiting amounts of protein extracts. Furthermore, we highlight recent developments and technological advances used in digestion protocols to quantify complex proteomes from single cells.
Keywords: Bottom-up proteomics, Mass spectrometry, Single cell proteomics, Trypsin digestion
Graphical Abstract
1. Introduction:
Mass spectrometry (MS)-based proteomics is the method of choice for the global analysis of proteins. Besides the identification of protein-protein interaction networks and the analysis of post-translational modifications, MS-based proteomics has enabled the identification of almost the complete human proteome [1, 2]. As depicted in Figure 1, typical workflows, which are termed bottom-up or shotgun proteomics, begin with the enzymatic digestion of proteins in a complex biological extract. The resulting peptides are separated by RP-HPLC and are then identified by MS/MS [3–6]. Trypsin is the most utilized protease in bottom-up MS proteomics due to its high catalytic activity and specificity in cleaving proteins into peptides at the C-terminal side of two basic amino acids, arginine (Arg) and lysine (Lys). Arg and Lys have a natural abundance in most proteomes that results in peptides that are an ideal length for both RP-HPLC, which is able to use the volatile solvents required for electrospray, and the peptide ionization process in MS [7, 8]. After a search algorithm such as Mascot [9] fragments the respective database in silico into theoretical tryptic peptides, the primary peptide sequences and modifications can be deduced by matching the theoretical and experimental fragmentation spectra [10]. In silico digests indicate that trypsin yields unique, identifiable peptides that represent >98% of all human genes [7]. While trypsin is the most extensively used protease in bottom-up MS proteomics, a literature review indicates that a wide variety of conditions have been used in these studies. Here we discuss the implications of some of these variable factors and make recommendations for carrying out trypsin digestions of complex proteomes from bulk samples to limiting amounts of protein extracts from single cells. We also highlight protein extraction methods linked to trypsin use as well as areas where additional research is needed to further optimize trypsin digestion parameters.
Figure 1:
Flow chart depiction of selected larger scale extraction and digestion protocols. Filter-aided Sample Preparation (FASP) [17]: Reduced tissue and cell lysates in 4% SDS are mixed with 8 M urea and subjected to ultrafiltration, washing, and alkylation in a Microcon YM10 device. After digesting with LysC in 8 M urea, the concentrate is diluted to 2 M urea and digested with trypsin. After collecting the resulting peptides and a wash by centrifugation, the combined filtrates are desalted by Solid Phase Extraction (SPE) with a C-18 disc prior to LC-MS/MS analysis. Suspension Trapping (STrap) [34]: Alkylated cell lysates in 5% SDS are acidified and then introduced into the suspension trapping tip that has a neutral pH methanolic solution on top of 11 quartz fiber plugs that are supported by C18 plugs. The instantly formed fine protein suspension is trapped in the depth filter stack of quartz fiber plugs. This crucial step separates the particulate protein matter in space. SDS and other contaminants are removed in the flow-through, and a protease is introduced. Following digestion, the peptides are cleaned up using the tip’s hydrophobic part. This methodology allows processing of protein loads down to sub-microgram levels. Single Pot Solid Phase-Enhanced Sample Preparation (SP3) [19]: Proteins in a reduced, alkylated cell extract are bound to carboxylate-coated paramagnetic beads through the addition of ACN in a manner similar to Hydrophilic Interaction Liquid Chromatography (HILIC). Immobilization on the bead surface permits washing and removal of contaminating substances such as the 0.5% SDS in the Lysis Buffer. The bound proteins are eluted by adding an aqueous digest buffer that contains a LysC/trypsin mixture. The resulting peptides are desalted by binding, washing, and then releasing from the SP3 beads prior to LC-MS/MS analysis. In-Stage Tip (iST) [20]: Cells or other protein materials are transferred into a StageTip and processed in three steps. 1) Lysis, denaturation, reduction, and alkylation occur in a 6 M GdnHCl Lysis Buffer in the StageTip. 2) The sample is then diluted 1:10 with Dilution Buffer that contains a LysC/trypsin mixture. 3) The resulting digest is then eluted and separated into 6 fractions by strong cation exchange (SCX) Stage-Tip chromatography prior to subjecting each fraction to LC-MS/MS. Pressure Cycling Technology (PCT) [60]: Cells or tissue samples are placed in PCT Tubes in a buffer containing 8 M urea. Sample lysis, protein extraction, reduction, alkylation, and LysC/trypsin digestion occur in a Barocycler instrument that subjects the samples to alternating cycles of 45,000 psi versus atmospheric pressure. The urea concentration is reduced to 6 M for LysC and to 1.6 M for trypsin digestion. The resulting peptide samples are desalted using C18 SEP-PAK cartridges prior to Sequential Windowed Acquisition of All Theoretical Fragment Ion Mass Spectra (SWATH-MS) analysis. Low Input [43]: Cells are FACS-sorted into collection microreactors that are gel loading tips packed with 3–5 punches of quartz (SiO2) mesh that provide a filter onto which cells accumulate. After washing the cells, the tip is then kinked right below the quartz membrane. Lysis, denaturation, reduction, alkylation, and digestion all happen in the microreactor. Digests are then loaded onto a C18 StageTip for desalting and on-column Tandem Mass Tag (TMT) labeling. After labeling, the samples are mixed and subjected to small-scale sulfonated polystyrenedivinylbenzene (bSDB) fractionation at pH 10. Each of the resulting fractions are then analyzed by LC-MS/MS.
2. Trypsin Is an Ideal Enzyme for Digesting Proteins for Downstream MS-based Proteomics:
In silico tryptic digestion of the ~21,030 reviewed protein sequences in the human proteome (UniProtKB/Swiss-Prot) predicts 2.3 million tryptic peptides of suitable size for MS detection (7–35 amino acids, up to two missed cleavages) [11], the length cutoff being important since peptides longer than 6 amino acids are much less likely to match to a decoy database [12]. Tryptic peptides have an average length of 14 residues [13] and comprise 9.9 million amino acid residues of the 11.5 million total, that is, 86% of the proteome’s sequence [11]. Most tryptic peptides are sufficiently hydrophobic to be retained on the C-18 supports that are typically used for RP-HPLC-MS/MS and sufficiently hydrophilic to be released from these supports with increasing concentrations of acetonitrile (ACN) in the volatile ACN/water mobile phase that is commonly used for RP-HPLC [14]. Since tryptic peptides contain basic sites at their N- and C-termini, they are easily protonated and ionized under the acidic conditions that are used for RP-HPLC. In addition, the balance in basicity between the free amine at the N-terminus of a tryptic peptide and the C-terminal Arg or Lys results in good MS/MS fragmentation as predicted by the mobile-proton hypothesis [15].
Data supporting the use of trypsin for bottom-up proteomics comes from a wide range of studies. Swaney et al. (2010) compared the number of identified proteins following trypsin (3,313), AspN (3,183), LysC (3,030), GluC (2,813), and ArgC (2,708) digestion and SCX fractionation of a yeast extract and found that a trypsin digest identified the most proteins [16]. Similarly, using filter aided sample preparation (FASP) to digest duplicate lysates of 2 ×105 HeLa cells, Wisniewski et al. (2009) obtained the following average number of proteins identified by two peptides: trypsin (1,722), LysC (1,513), ArgC (1,388), GluC (828), and AspN (907) [17]. More recently, Sinitcyn et al (2023) [11] carried out digestions with 6 proteases on 6 human cell line extracts and subjected the resulting digests to off-line fractionation followed by RP-LC-MS/MS. Together, the 6 enzymes (i.e., Trypsin, LysC, LysN, AspN, chymotrypsin, and GluC) generated 7.4 million peptides that covered 99% of the human proteome. For each of the 6 cell line extracts, an average of 539,325 unique peptides were identified that were derived from ~16,000 proteins. The tryptic peptide data contributed the largest number of protein identifications and unique sequences, totaling 17,631 proteins with 56.5% median sequence coverage, and 99.5% of the 17,717 proteins identified with all 6 proteases [11].
Although “trypsin represents the gold standard in proteomics for protein digestion”, it nonetheless has some shortcomings [18]. Because 56% of yeast tryptic peptides are ≤6 residues, which are generally too short to be identified by RP-HPLC-MS/MS, only a restricted part of the yeast and other proteomes is covered [16]. Indeed, when processing MS/MS data from HeLa cells both Hughes et al. [19] and Kulak et al. [20] used a filter that required a minimum peptide length of 7 amino acids. In addition, since tryptic peptides derived from the C-termini may have any terminal residue at either end, they often have low charge and are not compatible with RP-HPLC-MS/MS analysis and search engines [18]. Lastly, splice site sequences are inherently biased for lysine codons such that trypsin digestion results in under-representation of junction-spanning peptides [11].
3. Cell Lysis and Protein Extraction:
3.a. General Considerations:
Achieving high and reproducible protein extraction efficiency from cells and tissues is key to the success of the subsequent enzymatic digestion. While a universal sample preparation protocol should quantitatively isolate all proteins from any given sample, attaining this goal is extremely challenging as proteins vary tremendously in their physicochemical properties and other sample characteristics, such as rigid cell walls, tissues that are difficult to lyse, and interfering cellular components (e.g., lipids, nucleic acids, etc.) all impact isolation efficacy [21]. Since it is more feasible to develop optimum procedures for extracting and solubilizing proteins from a limited range of sample types, most of the focus of this review is on model organisms such as E. coli and HeLa cells.
3.b. Lysis Buffer Composition
Approaches for preparing biological samples for MS/proteomics analyses generally couple physical disruption such as grinding, homogenization, sonication, high/low pressure (i.e., PCT), or heat with the use of chaotropic denaturants such as urea or guanidine hydrochloride (GdnHCl), or surfactants such as SDS, sodium deoxycholate (SDC), or MS-friendly agents like RapiGest [22] to help solubilize proteins [23–26]. Masuda et al. (2008) [27] evaluated the relative ability of 27 reagents to extract and solubilize proteins from an E. Coli membrane fraction. As shown in Table 1, 0.1% SDS or 0.1% RapiGest had the most ability to solubilize E. coli membrane proteins [27]. While 40% ethanol (EtOH) or 40% ACN did not significantly improve solubilization, 6 M urea provided an almost 2-fold increase and 10% octylglucoside (OG) or 10% SDC provided >3-fold increase [27]. The study by Glatter et al. (2015) [28] carried out on HeLa cells also found that RapiGest and SDC were more effective than urea at extracting proteins. This study also determined that 6 M GdnHCl is more effective than 8 M urea at solubilizing proteins (Table 1).
Table 1:
Relative Ability of Various Agents to Extract and Solubilize Proteins from an E. Coli Membrane Fraction and HeLa Cellsa
| Solubilizing Agent | E. Coli Membrane Fraction Masuda et al (2008) [27] | HeLa Cells Glatter et al. (2015) [28] | ||
|---|---|---|---|---|
| Concentration | Relative Solubilizing Ability | Concentration | Solubilizing Ability | |
| SDS | 0.1% | 3.92 | ||
| RapiGest/SDC | 0.5%/0.5% | 1.8 | ||
| RapiGest | 0.1% | 3.46 | 1.0% | 1.8 |
| SDC | 10% | 3.08 | 2.0% | 1.5 |
| Octylglucoside | 10% | 3.04 | ||
| CHAPS | 10% | 2.04 | ||
| Urea/GdnHCl | 6 M/2 M | 1.4 | ||
| Urea | 6 M | 1.92 | 8 M | 1.0 |
| 2-Propanol | 80% | 1.71 | ||
| Tween-20 | 10% | 1.66 | ||
| Methanol | 40% | 1.63 | ||
| Acetonitrile | 40% | 1.04 | ||
| Ethanol | 40% | 1.04 | ||
| GdnHCl | 1.5 M | 0.50 | 6 M | 1.4 |
| Triton X-100 | 5% | 0.42 | ||
In Masuda et al. (2008) [27] the solubilizing ability is the relative ability of the listed additive at the listed concentration compared to 50 mM ammonium bicarbonate (ABC) buffer to solubilize proteins from an E. Coli membrane fraction after heating at 95°C for 5 min followed by sonication for 10 min and centrifugation [27]. In Glatter et al. (2015) [28] frozen HeLa cell pellets were resuspended in the indicated agent in the presence of 5 mM Tris(2-carboxyethyl)phosphine (TCEP) in triplicate experiments. Following sonication, urea-containing samples were incubated for 1 hr at 37°C, whereas all other samples were incubated at 60°C for 30 min. After the extraction and reducing steps, all samples were incubated with 10 mM iodoacetamide (IAA) at 25°C for 30 min prior to determining the protein concentrations (μg/μl) cited above.
STrap was used to compare the relative ability of different detergents to lyse and extract proteins from HeLa cells that were then all digested with the same trypsin protocol [29]. Based on the mean number of proteins that were identified (number in parentheses) from 50 μg extract, the relative order of effectiveness was 4% SDS/0.1 M DTT (3,743), 5% SDS (3,553), Radioimmunoprecipitation assay (RIPA, 3,388), 8 M urea (3,112), and then 1% NP-40 (1,743) [29]. Varnavides et al. (2022) [21] used 16 MS sample preparation methods to extract and analyze the HeLa proteome. Based on the amount of extracted protein, the relative order of effectiveness of three commonly used lysis and extraction agents was 1% SDC (322 μg) > 8 M urea (308 μg) > 6 M GdnHCl (300 μg) [21].
In aqueous solutions urea dissociates over time and upon heating to give ammonium ions and isocyanate. In alkaline conditions (i.e., pH 7–9) isocyanate reacts with protein and peptide N-termini and the ε-amino groups of lysines [30]. Under harsh conditions (i.e., 8.0 M urea,100 mM Tris-HCl, pH 8.5; 61°C for 15 hr) 99% of peptide N-termini and 90% of Lys residues in a complex mixture of tryptic peptides had been carbamylated [31]. Therefore, it is important to use freshly dissolved urea and to avoid prolonged exposure to elevated temperatures that can result in significant carbamylation that can hamper trypsin digestion [31]. Hence, Wisniewski et al. (2009) [17] carried out their FASP protocol at room temperature (RT, 18–22°C). Under these and other conditions that are typically used for in solution digests in urea <0.5% of identified peptides had carbamylated arginine or lysine residues [17, 21].
Many cell lysis and extraction reagents inhibit trypsin activity (e.g., 6 M GdnHCl) and/or interfere with LC-MS/MS (e.g., SDS [32] and RapiGest [33]). They can be removed by bead-based purification (SP3) [19], filtration (FASP) [17], StageTips that consist of a pipette tip with an inserted C18 disc (iST) [20], Suspension Trapping (STrap) in a pipette tip packed with quartz fiber filters on top of RP membrane discs [34], or by protein precipitation with acetone [32, 35], ethanol [36], chloroform/methanol [32, 35], or trichloroacetic acid (TCA)/acetone [35]. Varnavides et al. (2022) [21] obtained recoveries of 58.8% and 81.1% of HeLa cell proteins using chloroform/methanol and acetone precipitation respectively from lysates in 8 M urea. Both Puchades et al. (1999) [37] and Botelho et al. (2010) [32] found that chloroform/methanol precipitation outperformed conventional acetone precipitation. While Botelho et al. (2010) [32] found that conventional acetone precipitation of a 2% SDS solution did not sufficiently lower the SDS concentration to permit LC-MS/MS analyses, one or two extra acetone washes provided satisfactory removal of SDS and a protein recovery of about 80%. Varnavides et al. (2022) [21] obtained slightly higher recoveries using acetone (88.5%) rather than chloroform/methanol (85.2%) precipitation to recover HeLa cell proteins from 1% SDC lysates. Since GdnHCl is not soluble in acetone, Varnavides et al. (2022) [21] used ethanol (80.4%) and chloroform/methanol (69.1%) to precipitate HeLa cell proteins from lysates that contained 6 M GdnHCl. While surfactants such as SDS are difficult to remove prior to RP-HPLC-MS/MS analysis [17], acid cleavable surfactants such as RapiGest improve extraction and digestion efficiency while also being easy to remove by acid hydrolysis [22, 33, 38].
More recently, the Sample Preparation by Easy Extraction and Digestion (SPEED) protocol uses trifluoroacetic acid (TFA) to extract and denature proteins [39]. Although TFA dissolves E. coli cells and mouse liver tissue in 1 and 10 min. respectively at RT, lysis of the Gram-positive bacteria S. aureus requires an additional step of microwave irradiation for 10 sec. All of the resulting clear lysates have low viscosity as DNA is rapidly degraded by TFA [39]. TFA does not hydrolyze peptide bonds nor does it introduce modifications [21]. While it is likely that TFA leads to loss of glycosylations, many other PTMs such as phosphorylation, methylation, acetylation, and ubiquitination are presumably stable at low pH [39].
The extraction performance of SPEED [39] was evaluated using preparations of different sample types compared with detergent-based methods (FASP [17], SP3 [19]) as well as with urea-based in solution digestion (ISD-Urea [40]). For easy to lyse E. Coli cells the relative order of effectiveness in terms of extracted protein was SPEED (836 μg) > ISD-Urea (744 μg) > FASP (734 μg) > SP3 (664 μg). For the easy to lyse HeLa cells, the order was SP3 (209 μg) > SPEED (202 μg) > ISD-Urea (187 μg) > FASP (176 μg). For difficult to lyse mouse lung tissue the order was SPEED (1,244 μg) > SP3 (738 μg) > FASP (691 μg) > ISD-Urea (520 μg). SPEED increased the yield of protein extraction from difficult-to-lyse sample types by 68% for mouse lung tissue and 54% for B. cereus compared with detergent or chaotropic agent-based methods.
3.c. Lysis Method
3.c.i. Protein Extraction from Larger Numbers of Cells and Tissues:
Conventionally, tissue homogenization of bulk samples uses glass homogenizers, pestles, bead millers or a probe or water bath sonicator that uses sound waves to strongly agitate the sample. More recently, adaptive focused acoustics ultrasonication has been used to disintegrate tissue by delivering highly controlled, localized, and reproducible energy to small sample volumes (e.g., 12–100 μl) in a 96-well format [41]. PCT uses an instrument called a barocycler to apply ultra-high pressure (i.e., up to 45,000 psi) to a sample [26, 42]. Alternation between high and ambient pressures results in physical disruption of complex tissue samples, extraction, and solubilization of proteins. Tables 2 and S1 summarize, and Figure 1 provides flow charts for selected approaches used to lyse, extract, and then digest proteins from cells and tissues.
Table 2:
Selected Larger Scale Extraction and Digestion Protocolsa
| Study | Year | Cell Type | #Cellsb | Approach | MS | Trypsin Digest Denaturant | Fractionation | Quantitation | Quantified Proteins | |||||||
|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|
| 10% ACN | 1–2 M Urea | 0.05 % SDC | None | SCX | bSDB | None | LFQ | TMT | SILAC | |||||||
| Wisniewski et al. [17] | 2009 | HeLa | 1,250 | FASP | DDA | 1,059 | ||||||||||
| 200,000 | 1,934 | |||||||||||||||
| Wisniewski & Mann [45] | 2012 | HeLa | 38,462c | MED-FASP | DDA | 3,350 | ||||||||||
| Hughes et al. [19] | 2014 | HeLa | 1,000 | SP3 | DDA | 2,811 | ||||||||||
| 50,000 | 3,875 | |||||||||||||||
| Kawashima et al [44] | 2022 | HEK293 | 80,000d | SP3 | DIA with FAIMS | 10.716 | ||||||||||
| Kulak et al. [20] | 2014 | HeLa | 133,333 | iST | DDA | 9,667 | ||||||||||
| Meier et al. [46] | 2018 | HeLa | 133,333 | iST | DIA BoxCar | 7,775 | ||||||||||
| Shao et al. [26] | 2015 | HeLa | 59,000 | PCT | DIA | 2,096 | ||||||||||
| Myers et al. [43] | 2019 | Mouse Immune | 300,000 | Low Input | DDA | 6,427 | ||||||||||
| Doellinger et al. [39] | 2020 | HeLa | 704,000 – 836,000e | SPEED | DDA | 3,580 – 3,908 | ||||||||||
The solid colors indicate which protocols used the indicated denaturant (green), fractionation (blue), and quantitation (red) approach.
The number of cells refers to the approximate number of cells that were digested not analyzed by LC/MS/MS.
The 25 μg protein/sample that was digested [45] was converted to number of HeLa cells by assuming that the lysate from 200,000 HeLa cells contains 130 μg protein [17].
The 20 μg protein that was digested was converted to number of HEK293 cells by using the estimate of 250 pg protein/mammalian cell by Marx et al. (2019) [48].
In the Low Input protocol, Myers et al. [43] used up and down pipetting in 8 M urea to provide the shear forces for lysing FACS-sorted cells. Other solution-based approaches include the use of SDS in the SP3 [19, 44] and FASP [17, 45] approaches, 6 M GdnHCl in the iST [20, 46] protocol, and TFA in the SPEED Protocol [39]. In the SP3 approach benzonase endonuclease [47] is used to degrade DNA and RNA. In addition to SDS, the FASP protocol homogenizes tissues, shears DNA by sonication, and heats the extracts at 95°C for 3–5 min [17]. The SP3 protocol lyses HeLa and other mammalian cells by heating at 95°C for 5 min in SDS without physical disruption [19]. Another approach that uses SDS and heating at 95° (5 min) to lyse cells and sonication to shear DNA is the STrap method [34]. In the iST protocol up to 20 μg protein samples are loaded into StageTips and the lysates are boiled in 6 M GdnHCl for 5 min and then sonicated for 15 min in a water bath sonicator to denature proteins, shear DNA and enhance cell disruption [20]. The protocol by Shao et al. [26] provides an aggressive approach that couples PCT with 8 M urea. Since PCT also accelerates proteolytic digestion, it can process milligrams of wet tissues and convert them into peptide samples in <6 hrs. The programmable barocycler can be used for semiautomated preparation of up to 16 samples that should provide lower variability than manual approaches. Shao et al. (2015) integrated PCT with SWATH-MS to carry out reproducible proteomic quantification of biopsy-level tissue samples [26]. The minimal sample requirements were ~50,000 human cells and 0.2–0.5 mg of wet mouse or human tissue samples [26].
Commercially available MS sample preparation kits have been developed for iST (PreOmics), STrap (ProtiFi LLC), and in-solution digests (EasyPep, Thermo Scientific) [21].
3.c.ii. Protein Extraction from Single Cells:
The Minimal ProteOmic sample Preparation (mPOP) procedure in the Single Cell ProtEomics by Mass Spectrometry (SCoPE2) approach [49] extracts proteins in pure water. However, 9 of the 11 Single Cell Proteomics (SCP) studies, which are summarized briefly in Table 3 and in more detail in Table S2, added either an MS-compatible surfactant [i.e., 0.05–0.2% n-dodecyl-β-D-maltoside (DDM) [50–53], 0.3% RapiGest [54]] or an organic solvent [i.e., 20% trifluoroethanol (TFE) [55, 56], 20% ACN [57], or DMSO [58]] to the lysis buffer to help extract and denature proteins. While Brunner et al. (2022) [57] and Budnik et al. (2018) [59] also used sonication, 8 of the 11 SCP studies in Table S2 used high temperatures (i.e., 70–95°C for 5–60 min) to help lyse cells and denature extracted proteins. Since the ScoPE2 protocol [49] eliminated the sonication step that was used in the SCoPE-MS protocol [59], the simple freeze/heat cycle (i.e., −80°C/5 min, 90°C/10 min) in the ScoPE2 protocol appears to be sufficient to lyse mammalian cells.
Table 3:
Selected Single Cell Proteome Analyses
| Study | Year | Cell Type | Approach | MS | Digest Vol. (μl) | Trypsin Digest Denaturant | Quantitation | Quantified Proteinsa | ||||||
|---|---|---|---|---|---|---|---|---|---|---|---|---|---|---|
| 40% DMSO | 20% ACN | 0.03 – 0.1 % DDM | 10 % TFE | 0.3% RapiGest | None | LFQ | TMT | |||||||
| Budnik et al. [59] | 2018 | Jurkat, U-937 | SCoPE-MS | DDA | 10.0 | 767 | ||||||||
| Zhu et al. [50] | 2018 | HeLa | nanoPOTS | DDA | 0.20 | 211 | ||||||||
| Woo et al. [52] | 2021 | Murine cell lines | Nested nanoPOTS | DDA | 0.03 | 1,690 – 1,735 | ||||||||
| Specht et al. [49] | 2021 | U-937, HEK293 | SCoPE-2 | DDA | 1.0 | ~1000 | ||||||||
| Schoof et al. [55] | 2021 | AML | DDA | 2.0 | 987 | |||||||||
| Brunner et al. [57] | 2022 | HeLa | T-SCP | DIA | 2.0 | 2,083 | ||||||||
| Gebreyesus et al. [54] | 2022 | MEC-1 | SciProChip | DIA | 0.08 | 1,500 | ||||||||
| Huffman et al. [58] | 2023 | HEK293 | nPOP/pSCoPE | Prioritized | 0.02 | ~1,430b | ||||||||
| Matzinger et al. [51] | 2023 | HeLa | DIA | 1.0 | 1,208c | |||||||||
| Petrosius et al. [56] | 2023 | HEK293 | WISH-DIA | 2.0 | 1,972 | |||||||||
| Truong et al. [53] | 2023 | HeLa | nanoPOTS | WWA-DDA | 0.20 | 1,758d | ||||||||
Number of proteins quantified per cell without use of Match Between Runs (MBR).
Median number of proteins identified/cell in 97 single cells as estimated from Figure 2a in Huffman et al. [58].
Average number of proteins identified per HeLa cell using MBR was 1,543.
Average number of proteins identified from the analysis of 10 HeLa cells without using MBR. With MBR an average of 3,042 proteins were identified per HeLa cell.
4. Enzymatic Digestion Workflows:
Figure 1 and Tables 2 and S1 provide a summary of selected approaches for carrying out enzymatic digestions of larger numbers of cells and tissues.
Low Input: FACS-sorted cells are collected in a 20 μl gel loading tip that serves as a microreactor for lysis, reduction, and alkylation in 8 M urea and trypsin digestion in <2 M urea [43].
FASP: Reduced, SDS-containing protein lysates are mixed with 8 M urea to disrupt SDS micelles and are then subjected to ultrafiltration, washing, alkylation, and LysC/trypsin digestion in a Microcon Ym10 filter device (Sigma-Aldrich) prior to LC-MS/MS analysis [17].
Suspension Trapping (STrap): A filter-based technology that enables preparation of SDS-containing protein lysates for enzymatic digestion [34]. A fine protein particulate suspension, which is susceptible to trypsin digestion, is trapped in a stack of quartz fiber filters in a 200 μl pipette tip and residual SDS washed away prior to trypsin digestion in the filter stack.
SP3: Uses SDS to lyse cells [19]. Proteins in a reduced, alkylated cell extract are bound on SP3 beads and then subjected to LysC/trypsin digestion prior to LC-MS/MS analysis.
iST: Cells or protein extracts are transferred into a StageTip where lysis, denaturation, reduction, and alkylation occur in a 6 M GdnHCl lysis buffer [20]. After diluting with a LysC/trypsin mixture, the resulting digest is eluted and separated into 6 fractions by Stage-Tip SCX chromatography prior to subjecting each fraction to LC-MS/MS.
SPEED: Consists of 1) lysis and extraction with pure TFA for 2–10 min at RT, 2) neutralization with 10x volume of 2 M TrisBase and 3) digestion [39]. After neutralization the lysates become slightly turbid as proteins precipitate and form fine homogeneous suspensions that do not sediment because of the high salt content. After the addition of TCEP and 2-Chloroacetamide (CAA), the samples are incubated at 95°C for 5 min. Proteins are then quantified in the resulting dispersions by turbidity measurement at 360 nm and subsequently digested with trypsin at a 1:100 to 1:20 (w:w) ratio depending upon the protein concentration.
Ludwig et al. (2018) [61] compared the numbers of proteins identified from LFQ analyses of STrap, FASP, and in-solution digestions of proteins extracted from colorectal cancer cells in 5% SDS or 8 M urea-based lysis buffers. Proteins lysed with 8 M urea were digested using in-solution (in 1.5 M urea), FASP, and STrap methods, while SDS-lysed proteins were digested using FASP and STrap approaches. In each case 100 μg of reduced/alkylated cell extract was digested overnight at 37°C with a 1:50 (w:w) ratio of trypsin and the resulting peptides were separated into 5 fractions by high pH RP-HPLC prior to LC-MS/MS analysis. The numbers of identified proteins ranged from 3,757 for FASP with SDS extraction to 4,662 for STrap with urea extraction (Table S3) [61]. Both the STrap and the FASP protocols with urea identified more proteins than the corresponding protocols with SDS [61]. The trypsin efficiency increased from 43% for the in-solution digests to 61–69% for STrap and FASP [61]. This increase was ascribed to the smaller digestion volumes and consequently higher trypsin and substrate concentrations permitted by STrap and FASP.
The relative digestion performance of several workflows was evaluated by Doellinger et al. (2020) [39]. Based on the mean number of proteins identified, the relative digestion performance with E. Coli was ISD-Urea (1,949) > SPEED (1,899) > SP3 (1,883) > FASP (1,823). The relative digestion performance with HeLa cells was SPEED (3,874) > ISD-Urea (3,744) > SP3 (3,660) > FASP (3,548) and with mouse lung tissue was SPEED (3,302) > FASP (3,210) > SP3 (3,189) > ISD-Urea (2,705). Since the additional sample preparation steps that are required for FASP and SP3 resulted in increased CVs compared with ISD-Urea, reproducibility of protein quantification appears to benefit from fewer sample handling steps [39]. To compare the ability of different extraction and digestion approaches to study the human microbiome, a commercial standard consisting of 20 bacterial species (~10 μg) was subjected to SPEED [39] and two detergent-based methods, STrap [34] and iST [20]. Based on the numbers of quantified proteins, the relative order of effectiveness was SPEED (4,255), STrap (2,998), and iST (2,084) [39]. To determine the suitability of SPEED for phosphoproteomics, triplicate aliquots of HeLa cells (100 μg) were prepared using SPEED and EasyPhos-based phosphopeptide enrichment and compared to the original high-sensitivity EasyPhos protocol using SDC for lysis [62]. SPEED identified 8.3% more (i.e., 6,412 versus 5,923) phosphorylation sites (S,T,Y).
Varnavides et al. (2022) [21] used 16 MS sample preparation workflows to analyze the HeLa proteome, including in-solution digest (ISD) protocols based on urea, GdnHCl, and SDC-based buffers as well as SPEED [39], FASP [17], STrap [34] (ProtiFi), and SP3 [19] protocols and two commercial kits: iST [20] (PreOmics) and EasyPep (Thermo Scientific). As shown in Table S4, the numbers of proteins identified for these 16 methods varied from 3,734–3,816 for the 3 ISD-GdnHCl to the 4,421–4,473 proteins identified with the 3 ISD-SDC protocols. Of the 6 cleanup technologies tested, the largest numbers of proteins were identified by the EasyPep (4,375) and iST protocols (4,337). Since these were the only two procedures tested that used LysC/trypsin digestion, we agree with Varnavides et al. that the increased efficiency of these two technologies likely results from the improved performance of LysC/trypsin versus trypsin only digestion [63–65]. The increased number of proteins identified with the SP3 (4,235) versus SP3-SDC (4,125) approach is consistent with the data obtained by Masuda et (2008) [27] that SDS is a better protein solubilizing agent than SDC (Table 1). In this regard, the excellent performance of the 3 ISD-SDC protocols supports previous studies that found that tryptic digests carried out in SDC provided more protein identifications than those carried out in GdnHCl, urea, or RapiGest [23, 27, 66] and is consistent with SDC having very high protein solubility (Table 1) as well as trypsin enhancement activity (see Table 6 below) [27]. While acetone and chloroform/methanol precipitation of urea resulted in a ~3% increase in the number of proteins identified (i.e., from 4,108 for the ISD-U to 4,238 for the ISD-U/A and ISD-CM samples), precipitation did not significantly benefit either the ISD-GdnHCl or ISD-SDC samples [21].
Table 6:
Effect of Selected Additives on Trypsin Activitya
| Additive | Concentration at Maximum Activity | Activity Enhancement Factor | Maximum Sustainable Concentration |
|---|---|---|---|
| Octylglucoside | 0.1% | 6.79 | 10% |
| Ethanol | 20% | 5.81 | 40% |
| NP-40 | 0.1% | 5.58 | 5% |
| CHAPS | 0.1% | 5.36 | 10% |
| SDC | 0.01% | 5.12 | 10% |
| Triton X-100 | 0.001% | 4.82 | 5% |
| Tween-20 | 0.001% | 4.63 | 10% |
| Methanol | 40% | 3.88 | 40% |
| SDS | 0.01% | 3.48 | 0.01% |
| 2-Propanol | 20% | 3.36 | 80% |
| Acetonitrile | 20% | 2.48 | 40% |
| Urea | 3 M | 1.26 | 3M |
| RapiGest | 0.1% | 0.429 | N.A. |
| GdnHCl | 0 M | 0 | 0 M |
Data compiled from Table 1 and for RapiGest from Figure 1 in Masuda et al. (2008) [27]. Trypsin activity is based on hydrolysis of a model substrate, N-benzoyl-lysine p-nitroanilide (BLNA), in 50 mM ABC for 24 hrs at 37°C with a 1:20 (w:w) ratio of enzyme:substrate. The maximum sustainable concentration is the highest concentration at which >30% of the original trypsin activity without additives remains after 24 hrs.
5. Comparison of Chemically Modified Trypsin from Different Species and Vendors:
Trypsin from varying biological origins (e.g., porcine, and human) are commercially available in natural, recombinant, and/or modified forms, and can differ in structure, specificity, and relative activity. Trypsin is a globular protein (PDB accession code 1AQ7) that comprises two six-stranded Greek-key β-barrels [67–69] and a Ca2+ ion that is important for activity. Green and Neurath (1953) determined that 1 or 10 mM Ca2+ increased the esterase activity of trypsin by 28% and that the single bound Ca2+ ion has a Kd of 4 μM [70]. The 223 residue, single chain, “cationic”, β-trypsin is the predominant product of bovine trypsinogen activation [71] and is stabilized by 6 disulfide bonds [72]. Trypsin autolysis with a cleavage between Lys-131 and Ser-132 produces α-trypsin, which has two polypeptide chains connected by disulfide bonds, while a further cleavage between Lys-176 and Asp-177 yields pseudotrypsin (ψ-trypsin) that has 3 discrete chains [73].
Bunkenborg et al. (2013) determined that both bovine and porcine trypsin are tyrosine O-sulfated at multiple sites that should be considered when screening for trypsin autolysis products [74]. They also demonstrated that while bovine and porcine trypsin were equally specific with no missed Arg/Lys cleavages, one commercial preparation of bovine trypsin (i.e., entry #3, Sigma #T1426 in Table 4) had a much higher level of non-tryptic cleavage activity that apparently resulted from chymotrypsin impurities. Deng, et al (2018) found that while ~78% of all the tryptic cleavage sites in two standard proteins were efficiently cleaved by porcine trypsin, only ~47% and ~53% were efficiently hydrolyzed by bovine and human trypsin respectively [75]. They also determined that the total hydrolysis rate constant for beta-casein was 3-fold higher for porcine as compared to bovine or human tryptic hydrolysis. These differences were explained by variations in trypsin’s secondary specificity. Hence, while the rate of cleavage with porcine trypsin was decreased by the presence of an Asp/Glu residue that was just prior to the Arg/Lys, the rate of cleavage by human trypsin was also slowed by the presence of an Asp/Glu that was located immediately after the Arg/Lys residue [75]. In agreement, Walmsley et al. (2013) found that bovine trypsin produced more peptides containing missed cleavages than did porcine trypsin [76].
Table 4:
Selected Commercially Available Trypsin Preparations
| # | Manufacturer | Part # | Description | Biological Source | Reductively Methylated? | TPCK Treated | Units/mg Protein | Burkhart et al. (2012) [13] | |
|---|---|---|---|---|---|---|---|---|---|
| #Filtered Tryptic Peptides | #Proteins Identified | ||||||||
| 1 | Promega | V5111 | Sequencing Grade Modified Trypsin | Porcine | Yes | Yes | 12,064a | 7,630 | 709 |
| 2 | Sigma | T8658 | Trypsin from Bovine Pancreas - for Protein Sequencing | Bovine | No | No | ≥8,000 | 6,276 | 612 |
| 3 | Sigma | T1426 | Trypsin from Bovine Pancreas | Bovine | No | Yes | ≥10,000 | 5,952 | 605 |
| 4 | Sigma | T0303 | Trypsin from Porcine Pancreas | Porcine | No | No | 13,000 – 20,000 | 5,406 | 582 |
Specific Activity from a particular lot of this product.
In addition to generating peptides that can interfere with LC-MS/MS analyses, trypsin auto-proteolysis can generate pseudotrypsin that has decreased rates of cleavage of Arg/Lys and a chymotrypsin-like specificity [71, 77]. For this reason, many commercially available trypsin preparations are chemically modified by reductive methylation of Lys residues, which renders trypsin resistant to auto-proteolysis [78, 79]. Many vendors also treat their trypsin products with an irreversible chymotrypsin inhibitor, 1-chloro-3-tosylamido-4-phenyl-2-butanone (TPCK) [80]. Through the use of a full factorial design of experiments (DOE) approach [81], Loziuk et al. (2013) optimized 4 parameters (i.e., trypsin type, enzyme-substrate ratio, pH, temperature) [82]. Trypsin type had the second greatest influence on protein identification with dimethylated porcine trypsin being more favorable than unmodified, TPCK-treated bovine trypsin. This latter conclusion is consistent with studies described above by Walmsley et al. (2013) and Deng et al. (2018).
Multiple studies have compared commercially available trypsin products. Burkhart et al. (2012) compared the ability of 6 trypsin preparations to digest human platelets [13] and found considerable differences that did not fully correlate with their cost. Table 4 summarizes the results of their study on the 4 trypsin preparations that are still commercially available. The Promega Sequencing Grade Modified Porcine Trypsin digest identified the largest number of tryptic peptides (7,630), which corresponded to 41% of the MS/MS spectra that were searched, and proteins (709). More recently, Woessmann et al. (2023) [83] evaluated the impact of 8 commercially available trypsins across a range of protease concentrations and storage times. While no significant differences were found between the mean number of protein (~2,500) or peptide (~12,500) identifications that resulted from digests of HeLa cell extracts, the number of missed cleavages varied from 11.6% (Roche #3708985001) to 20.6% (Sigma #EMS0007). While the 3 Promega trypsins (V5111, V5280, VA9000) all had a consistent number of missed cleavages, significant variations were observed between the 3 Sigma trypsins (EMS0004, EMS0006, EMS0007).
5.a. Reconstitution and Storage of Trypsin Solutions:
Niu et al. (2020) demonstrated that reconstitution and storage conditions have a significant impact on the occurrence of nonspecific trypsin cleavages at Tyr, Phe, Trp, and Leu [84]. Semi-tryptic peptides are peptides cleaved at the carboxy-terminal side of Arg and Lys at one end but not at the other. The relative extent of semi-tryptic cleavage at two sites in a monoclonal antibody was assayed for 8 commercially available trypsin products after reconstituting and standing for 4 hrs at RT in HPLC grade water or 50 mM acetic acid as recommended by the manufacturers. The level of semi-tryptic cleavage ranged from <10% after reconstituting in both solvents for Roche bovine (#11418025001) and Sigma porcine (#T-6567) to 100% for Princeton Separations porcine (#EN-151) trypsin. While 3 of the trypsin products (Promega V5280 and V5111 porcine and Pierce 90057 porcine) resulted in <15% semi-tryptic cleavage after reconstituting in HPLC grade water and standing at RT for 4 hrs prior to digestion, the level of semi-tryptic activity increased to 30–73% after reconstituting in 50 mM acetic acid, and standing at RT for 4 hrs prior to digestion [84].
6. Sequential Digestion with Multiple Proteases:
With the goal of improving the yields and numbers of fully cleaved tryptic peptides and thereby reducing missed cleavages and increasing the number of identified proteins, Glatter et al. compared a tandem endopeptidase LysC/trypsin digest with a trypsin digest of a yeast lysate [63]. An advantage of using LysC is that it is more resistant than trypsin to high urea concentrations (6–8 M). Following a 6 hr LysC digest in 6 M urea, the concentration was decreased to 1.6 M where trypsin activity is retained. The sequential LysC/trypsin digest resulted in a 9% increase in the number of fully cleaved peptides, a 21% decrease in the number of miscleaved peptides, and a 2% increase in the number of identified proteins [63]. Detailed analyses showed that trypsin alone cleaves less efficiently after Lys than after Arg, particularly when flanked by acidic amino acids and that this bias can be overcome by using the tandem LysC/trypsin protocol. Using a 293T human renal epithelial cell extract, Qian et al (2017) carried out a similar tandem LysC/trypsin protocol in 100 mM ABC buffer, pH 8.0 at 37°C with the initial LysC digest (1:100 w:w) being carried out for 3 hrs in 8 M urea prior to diluting to 2 M urea for the subsequent 16 hr trypsin digest (1:50 w:w) [64]. The tandem digest resulted in an 11.9% increase (i.e., 2,317 versus 2,070) in the number of identified proteins as compared to the trypsin digest [64].
Wisniewski et al. (2009) carried out a FASP study on HeLa cell extracts to determine if any other combination of commonly used proteases might provide deeper proteome coverage. In decreasing order, the following combinations of two independent digests and LC-MS/MS analyses with one digest being carried out with trypsin provided the indicated numbers of total identified proteins: ArgC (3,145), LysC (3,042), AspN (2,931), GluC (2,912), chymotrypsin (2,787), trypsin (2,699) [17]. More recently, Sinitcyn et al. (2023) determined the median extent of coverage of 6 human cell line extracts using different combinations of 6 proteases. While trypsin alone provided a median coverage of 54%, the top 3 combinations of 2 proteases (i.e., trypsin + GluC, trypsin + AspN, trypsin + LysC) all provided median sequence coverage of 62% [11].
Hakobyan et al. (2019) carried out a sequential trypsin digestion study in two different MS-compatible detergents, SDC and sodium lauroyl sarcosinate (SLS) on previously frozen E. coli cell pellets [65]. This study found that tandem LysC/trypsin digests resulted in ~10% (SDC) and 11% (SLS) more spectrum counts, respectively, from fully cleaved peptides than the corresponding trypsin digest. The authors suggested that the improved cleavage from tandem digests results primarily from the complementary target specificities of the two proteolytic enzymes rather than from the higher denaturant concentrations that are possible when tandem digests are carried out in chaotropic environments such as urea. Seven of the 9 larger scale studies in Table S1 and 5 of the 11 SCP studies in Table S2 used LysC/trypsin digests.
7. Manipulating the Milieu Conditions for Improved Trypsin Digestion:
To achieve optimal digestion, any denaturing agent used must be able to unfold most proteins while not introducing any chemical modifications, maintain trypsin structure and activity, and not interfere with the subsequent LC-MS/MS. In regard to maintaining trypsin’s structure and activity, Shuford et al. (2012) used a fractional factorial design of experiments approach with a FASP [17] workflow to determine the relative importance of 6 parameters (i.e., digestion time, concentration of trypsin, substrate:enzyme ratio, and the presence/absence of urea, methanol, and Ca2+) when using a protein cleavage isotope dilution mass spectrometry (PC-IDMS) assay to quantify 25 proteins in a xylem extract [85]. This study, which evaluated 16 different digest conditions, found that the most important positive factor was the presence of 10 mM CaCl2 in the digestion buffer. Indeed, as noted above, the crystal structure of porcine trypsin includes a bound Ca2+ ion (UniProt: P00761 Tryp_Pig, [67]) that promotes a conformational change to a more compact structure that stabilizes trypsin against autolysis [86]. This is consistent with the finding by Sipos and Merkel (1970) that the optimum Ca2+ is >1 mM and recommendations by Riviere and Tempst (2001) to include 5 mM CaCl2 and by Swaney et al. (2010) to include 10 mM CaCl2 in trypsin digestion buffers [16, 86, 87].
7.a. Reduction and Alkylation:
Cysteine constitutes about 3% of all amino acid residues in proteins [88]. Indeed, 91% of proteins contain at least one cysteine residue and it is present in >24% of predicted tryptic peptides [89]. Therefore, it is commonly thought that reduction and alkylation must be carried out to help denature and prepare extracted proteins for enzymatic digestion. In most proteomic workflows, lysis is carried out before the reduction of disulfide bonds with dithiothreitol (DTT) and the alkylation of free cysteines with iodoacetic acid (IAC) or iodoacetamide (IAA). Medzihradszky (2005) indicates that an approximately 500-fold excess of DTT at 60°C for 1 hr will complete the reduction of disulfide bonds [90]. After adding an 1,100-fold excess of IAA, which is the alkylating agent that has been used most commonly in protein digestion protocols [91], the mixture is then incubated at RT in the dark for 1.5 hr [90]. However, IAA has been shown to form many undesired by-products [91–96] including a 2-acetamidoacetamide covalent adduct to Lys residues that has the same atomic composition as the diglycine adduct that remains after trypsin digestion at a site of ubiquitination [91]. Chloroacetamide (CAA) is another alkylating agent whose chemistry should be more selective towards cysteines [97], and Nielsen et al. (2008) found that it does not produce the diglycine-like artifact nor any other apparent adducts [91]. Since IAA reacts with DTT these two reagents and steps cannot be combined. In contrast, Kulak et al. (2014) found that the reducing agent TCEP is compatible with CAA, which allows these chemicals to be incorporated into the lysis buffer and eliminates the need to perform reduction and alkylation as separate steps [20].
Wisniewski et al. (2020) [88] found that reduction with DTT and alkylation of Cys with IAA did not significantly affect the depth of coverage of mouse brain, HeLa cell, and human plasma proteomes determined with the MED FASP procedure using LysC or trypsin digestion [88]. In all cases, similar numbers of peptides and proteins were identified in the 1) absence of reduction and alkylation as in the presence of 2) reduction, 3) alkylation, or 4) reduction and alkylation. While the numbers of peptides containing Cys were negligibly low in the non-alkylated samples, in the reduced and alkylated samples Cys-containing peptides accounted for 16% of peptides in brain and 14% in liver samples. The latter data are in reasonable agreement with in silico analyses of the IPI human protein database that indicate that 24% of predicted tryptic peptides contain a Cys [89]. Moreover, Wisniewski et al. found that compared to a standard procedure using IAA for thiol-alkylation, that sample preparation carried out under conditions protecting thiols from oxidation enabled identifying 10–20% more peptides and proteins [88]. Based on a study that compared the use of 4 alkylating agents [CAA, IAA, 4-vinylpyridine (4-VP), methyl methanethiosulfonate (MMTS)] prior to in solution trypsin digestion of HeLa/HepG2 cell lysates, Kuznetsova et al. (2021) [93] concluded that: 1) CAA alkylation resulted in the largest number of identified peptides and 2) CAA did not result in Met oxidation or any other abundant modifications. 3) In contrast, alkylation of reduced extracts with IAA resulted in a 36% reduction in the median number of identified tryptic peptides that was ascribed to IAA side reactions with the most abundant being carboxyamidomethylation of Met. 4) CAA causes fewer side reactions and yields the most identified peptides [93]. More recently, while Varnavides et al (2022) [21] detected typical reduction and alkylation artifacts like off-target alkylation with IAA and DTT adducts in several of the methods tested, they all occurred at low levels (<0.5% or mostly lower). All 9 of the selected larger scale extraction and digestion protocols summarized in Table S1 carried out reduction and alkylation with three of the procedures using DTT/IAA [17, 19, 45], three using TCEP/IAA [26, 43, 44], and two using TCEP/CAA [20, 39, 46]. Four of the 11 SCP protocols summarized in Table S2 carried out reduction and alkylation with Zhu et al. (2018) using DTT/IAA [50], Woo et al. (2021) using TCEP/IAA [52], and Schoof et al. (2021) and Gebreyesus et al. (2022) using TCEP/CAA [54, 55].
7.b. Digestion Temperature:
While most trypsin digestions are carried out at 37°C [7, 16, 87, 90], Cheison et al. (2011) investigated the influence of different temperatures (i.e., 25, 37.5, and 50°C) on 10 min bovine trypsin digests of β-lactoglobulin in water at pH 7.8 and found that digests at 25°C and 50°C were slow as compared to 37.5°C [98]. Analyses of the resulting tryptic peptides indicated that cleavage after Arg was generally faster than after Lys and that chymotrypsin-like cleavages occurred more frequently at higher (i.e., 50°C) rather than lower (i.e., 25 and 37.5°C) temperatures. An isothermal titration calorimetry (ITC) study determined that at pH 7.5 the kcat for casein digestion by unmodified bovine trypsin decreased from 9 × 10−3 sec−1 at 37°C to 4 × 10−3 sec−1 at 20°C [99].
Havlis et al. (2003) determined that reductive methylation increases the thermal stability of trypsin [100]. They found that the T50, which is the temperature at which trypsin loses 50% of its activity within 30 min in 80 mM Tris(HCl), pH 8, increased from 40.5°C for the unmodified porcine to 48°C degrees for the reductively methylated (modified), TPCK-treated porcine trypsin [100]. Although they determined that the optimum temperature for a 30 min in gel digest of BSA with 0.5 μM modified porcine trypsin was 50°C, the average peptide yield from this assay was only 53% of that from a control “overnight” digestion at 37°C with the unmodified bovine trypsin [100]. Using a 3 min fluorescence assay with TPCK-treated, modified porcine trypsin, Finehout et al. (2005) [101] determined that the kcat for a casein digest carried out at pH 7.8 increased from 0.44 sec−1 at 37°C to 0.48 sec−1 at 48°C and 58°C. However, they also observed that at 58°C trypsin activity rapidly declines so that <40% of the initial activity remains after 2 hrs. As described above, Loziuk et al. (2013) used a full-factorial experimental design to optimize 4 parameters (i.e., trypsin type, enzyme-substrate ratio, pH, temperature) based on the number of proteins identified [82]. Of the 4 parameters tested in a 16 hr digest in 50 mM Tris(HCl), 2 M urea; temperature had the greatest influence, with 37°C being better than 48°C. Loziuk et al. (2013) suggested that the significant decrease in peptide/protein identifications at 48°C resulted from thermal denaturation of the modified porcine trypsin and rapid loss of tryptic activity. While 7 of the larger scale extraction and digestion protocols summarized in Table S1 carried out trypsin digests at 37°C [19, 20, 39, 43–46], Shao et al. (2015) used 33°C [26], and Wisniewski et al. (2009) used RT (i.e., 18–22°C) [17]. Similarly, while 4 of the SCP studies in Table S2 carried out trypsin digestions at 37°C [49, 55–57], Gebreyesus et al. (2022) used 40°C [54], Budnik et al. (2018) used 45°C [59], Matzinger et al. (2023) used 50°C [51], and Truong et al. (2023) used Promega’s “Rapid Digestion Trypsin” that enabled them to carry out a one hour digest at 70°C [53].
Promega’s “Rapid Digestion Trypsin” (product VA1060) is used with a proprietary “Resuspension Buffer” that enables trypsin digestions to be carried out for 60 min at 70°C with a recommended enzyme:substrate (w:w) ratio of 1:10. In the case of more difficult to digest proteins or complex samples the digestion time can be increased to as long as 3 hrs. Based on correspondence with Promega Technical Support, the “Rapid Digestion Trypsin” is Promega’s “Sequencing Grade Modified Trypsin”. While “there are only minor differences in purity and quality control between the enzyme that is in each kit, the important difference between Sequencing Grade Modified Trypsin and Rapid-Digestion Trypsin is the reaction buffer used. Trypsin activity and stability is strongly affected by buffer composition.” Doellinger et al. (2020) [39] found that use of the Rapid Digestion Trypsin in their “filter aided SPEED” analysis of HeLa cells reduced the numbers of protein and peptide identifications compared with the original protocol by 4% and 9–13% respectively. Interestingly, identification rates remained quite similar after reducing the digestion time from the 1 hr that was recommended by Promega to 15 min. Quantitative reproducibility was excellent for all samples and it was independent of the total processing time.
7.c. Digestion pH:
Medzihradszky (2005) recommended carrying out trypsin digests at pH 7–8.5 [90] and Perutka and Sebela (2018) indicated that trypsin has an optimal pH of 8–9 [71]. However, kinetic studies with casein determined that bovine trypsin has a pH optimum near 7 [102] and modified (dimethylated), TPCK-treated porcine trypsin has an optimal pH that is slightly higher than 8.1 [101]. This latter finding is consistent with protocols that carried out modified porcine trypsin digests in 0.1 M ABC at pH 8.0 with up to 4 M urea [87] or in 50 mM Tris(HCl) pH 8.0 with 1.5 M urea [16]. Similarly, and consistent with Farmer and Yuan (1991) [102], Cheison et al. (2011) found that 10 min digests at 37.5°C of β-lactoglobulin in water that were carried out with bovine trypsin at alkaline pH (i.e., pH 8.65 and 9.5) were slow as compared to digests carried out at pH 7.8 [98]. Also, Hao et al. (2011) observed that 16 hr in gel digests with modified porcine trypsin of a rat liver tissue extract that were carried out at pH 6 identified 2.5% fewer unique peptides than digests carried out at pH 8 [103]. Although Finehout et al. (2005) [101] and Hao et al. (2011) [103] determined that modified porcine trypsin has an optimal pH that is about 8, the full factorial DOE study by Loziuk et al. (2013) found that a pH of 7 rather than 8 resulted in maximizing the number of proteins identified [82]. Loziuk et al suggested that this discrepancy may result from trypsin being more stable at pH 7 or from the optimal pH being substrate dependent. The pH range for the larger scale extraction and digestion protocols in Table S1 was from 8–9.0 with 5 of the digests being carried out at pH 8, 3 at pH 8.5 ([19, 20, 46] and the Doellinger et al. digest being carried out a pH 8–9 [39]. The pH range for the tryptic digests for the SCP studies in Table S2 that provided this information varied from ~7.4 for the Budnik et al. (2018) and Gebreyesus et al., (2022) digests that were carried out in phosphate-buffered saline (PBS) [54, 59] to ~8.5 for the 5 studies carried out in triethylammonium bicarbonate (TEAB) [49, 51, 52, 55, 56] and the Bruner et al. study that was carried out in Tris(HCl), pH 8.5 [57] and the Huffman et al. (2023) [58] study that was carried out in HEPES, pH 8.5. Since the Zhu et al. (2018) digest buffer contained both PBS and ABC, its pH probably was between 7.4–8 [50].
7.d. Digestion Incubation Time:
7.d.i. Conventional Trypsin Digestions:
Most conventional trypsin digests are carried out “overnight” [16, 104], which is consistent with the recommendation by Riviere and Tempst (2001) to carry out 15 hr trypsin digests [87]. Medzihradszky (2005), however, suggests carrying out trypsin digests for 2–18 hrs [90]. There may be advantages to decreasing the digestion time. For example, Hao et al. (2011) found that Asn and Gln deamidation proceed readily under typical trypsin digestion conditions (i.e., 25 mM NH4HCO3, pH 8.0, 1 M urea, 1:30 trypsin:protein (w:w) ratio at 37°C) such that after a 16 hr digest approximately 5% and 2% of the tryptic peptides from a rat kidney tissue digest contained sites of Asn and Gln-deamidation respectively [103]. This study indicated that the main causative factors were the mildly alkaline pH and prolonged incubations at 37°C during tryptic digests. Reducing the in-gel trypsin digestion time from 16 to 8 hrs reduced Asn deamidation in a rat liver tissue extract to ~1.6%, with the latter corresponding to a ~43% reduction in the level of deamidation, and it also provided a ~2.5% increase in the number of identified unique tryptic peptides [103]. Somiari et al. (2014) found that decreasing the digestion time of human serum from 24 hrs (107 proteins identified) to 8 hrs with a 1:20 trypsin:protein (w:w) ratio resulted in a ~19% increase in the number of identified proteins [105]. Since Somiari et al. (2014) found more chymotryptic-like peptides in the 24 hr as compared to the 8 hr digested sample, they ascribed the decreased number of identified proteins to loss of trypsin specificity with increasing time of digestion [105]. Proc et al. (2010) determined that with a 1:20 ratio of modified porcine trypsin:substrate and in the presence of 1% SDC that 9 hrs provided optimal digestion of 45 human plasma proteins [66]. As described above, Shuford et al. (2012) used a FASP digestion scheme at 37°C and a fractional factorial design (FracFD) of experiments approach to optimize six bovine trypsin digestion parameters that included digestion time, trypsin concentration and enzyme:substrate ratio [85]. Increasing the digestion time from 5 to 16 hrs significantly improved the digestion efficiency as determined by a targeted protein cleavage isotope dilution mass spectrometry (PC-IDMS) assay that quantified 25 proteins. Subsequent analyses that were carried out in an optimized digestion buffer (10 mM CaCl2, 2 M urea, 50 mM Tris(HCl), pH 8.0) found that the minimum time required for complete digestion based on accurate quantification of all target proteins was 8 hrs [85]. The digest times for the larger scale conventional digests in Table S1 range from 4 hr [17] to 14 hr [19], 18 hrs [45], overnight [20, 43, 44, 46] to 20 hrs [39]. Similarly, as shown in Table S2, conventional SCP trypsin digestion times span from 2 hr [51] to 16 hr [54] with the Truong et al. [53] digest with Promega’s Rapid Digestion Trypsin being carried out for only 1 hr at 70°C.
7.d.ii. Fast Trypsin Digestions:
Ultrafast trypsin digestions can be achieved at 25°C in microdroplets, under a sheath of nitrogen gas at 120 psi, in as little as 250 μsec. These digests had decreased extents of Asn and Gln deamidation as compared to overnight digestions at 37°C [106]. Other approaches to speed trypsin digestions include the use of organic solvents, elevated temperatures, surfactants, PCT, STrap, high trypsin concentrations, and ultrasonication. In the case of PCT, the 45 min LysC digest in 6 M urea (enzyme:substrate ratio of 1:40) was followed by a 90 min trypsin digest in 1.6 M urea (enzyme:substrate ratio of 1:20) [60].
Since the STrap digestion occurs in a limited volume, at an elevated temperature of 47°C, and with a high concentration of trypsin (33 ng/μl), it is complete within 30 min [34]. Liu et al. (2021) [107] showed that the digestion time for a new 0.5 μg/sample TMT protocol can be shortened to 30 min at 21°C by increasing the trypsin concentration to 625 ng/μl, which is almost 7-fold higher than the highest trypsin concentration of 90 ng/μl that was used by any of the protocols in Tables S1 and S2. Indeed, the 2.5 μg of trypsin was sufficiently high that the excess trypsin acted as a carrier protein to help prevent adsorptive sample losses. TMT 10-plex profiling of human cortex samples revealed that this protocol quantified 84% as many proteins (i.e., 9,116 versus 10,869) as the standard overnight protocol. Similarly, the nested nanoPOTS chip for SCP resulted in a ~45-fold increase in trypsin digestion kinetics because both the trypsin and substrate concentrations were increased by 6.67-fold [52].
Studies by Hildonen et al. (2014) on digests carried out at pH 8 at 37°C determined that prolonged digests and high trypsin:substrate ratios result in decreased amino acid coverage that was caused by complete digestion by trypsin resulting in an increased number of small peptides that are not LC-MS/MS detectable [104]. Indeed, the highest average amino acid sequence coverage in their study was obtained with a 5 min tryptic digest using a 1:40 enzyme:protein ratio [104]. They concluded that it would be best to prevent trypsin digests from going to completion by reducing the digest time to 5 min and to then allow up to three missed cleavages during the database searches [104].
Other approaches that have been used to speed trypsin digestions include alternating electric fields as well as the use of infrared, microwave, and ultrasonic energy [108]. The most promising approach is the use of ultrasonic energy with a sonicator that is equipped with a cup or microplate horn (i.e., probe) that has a small ultrasonic bath that can accommodate individual tubes or a 96 well plate respectively. Typically, these “sonoreactors” are 50 times more powerful than a regular ultrasonic bath that is not sufficiently powerful to speed enzymatic digestions [108]. Santos et al. (2007) developed in-solution digestion methods that involved 5 min sonication at RT with trypsin in either ~1.2 M urea [109] or 50% ACN [110]. In both ultrasonically-assisted digests (UADs) the protein sequence coverage was similar to that for a conventional overnight trypsin digestion in ~1.2 M urea at 37°C [109, 110]. UADs have also been developed for trypsin digestion of Coomassie Blue-stained SDS gel bands [111] and 2D gel spots [112]. Lastly, Jorge et al. also developed a 4 min in-solution UAD and demonstrated that the efficiency of digestion was not affected by increasing the temperature from 20° to 30°C [112]. Similar in-solution UAD protocols have been developed for analyzing optimum cutting temperature (OCT)-embedded [113] and frozen [114] solid tumor biopsy samples in 96 well plates. In the Jorge et al. (2019) study LC-MS/MS analysis of a fresh frozen kidney biopsy sample that had been subjected to UAD in a 96 well plate identified 1,736 proteins [113]. Finally, Carvalho et al (2020) [115] developed a FASP version of a UAD and found that high intensity focused ultrasonic energy first speeds up the kinetics of trypsin digestion and then, after 5 min, inactivates the enzyme. The FASP UAD method was compared to the standard overnight digestion FASP protocol, and no statistical differences were found for >92% of the proteins identified in E. coli, mouse brain, and mouse liver extracts [115].
7.e. Enzyme:Substrate (w:w) Ratio:
Conventional trypsin digestion protocols typically use an enzyme:substrate (w:w) ratio that ranges from 1:10 to 1:100 [16, 87, 90, 104]. Since product formation from an enzyme catalyzed reaction is proportional to reaction time and the concentration of the enzyme:substrate complex, which in turn is proportional to the free enzyme and substrate concentrations; the best approach to determine the optimum trypsin and substrate concentrations and incubation time is by the use of a fractional or full-factorial DOE approach [81]. The only such study that we are aware of was carried out with modified porcine trypsin by Loziuk et al. (2013) and it determined that a 1:50 trypsin:substrate (w:w) ratio with 40 ng/μl trypsin (37°C, pH 7, 2 M urea, 16 hrs) is more favorable for global proteome analysis than a 1:5 ratio with 400 ng/μl trypsin [82]. The lower enzyme:substrate ratio of 1:50 maximized protein identification, number of unique peptides identified, sequence coverage, and the ratio of tryptic/chymotryptic cleavage sites. It was postulated that the higher trypsin concentration resulted in greater autolysis, even with modified porcine trypsin, as a result of the enzyme being more likely to encounter enzyme instead of substrate. Loziuk et al. (2013) also suggested that the higher concentration of trypsin may result in excess digestion and nonspecific cleavage of fragments into peptides that are too small to be sequenced with good sensitivity by mass spectrometry [82].
Other challenges occur when relying on w:w ratios to carry out tryptic digests on decreasing amounts of substrate protein [116]. In this regard, Stone et al. (1990) determined that a trypsin:substrate (w:w) ratio of 1:25 is insufficient when the substrate concentration falls below about 20 ng/μl [117]. Since Hughes et al. (2014) maintained the same trypsin concentration of 25 ng/μl (rather than the same w:w ratios) when carrying out trypsin digests on extracts from both 50,000 and 1,000 cells (Table S1), the corresponding [trypsin] × [sample] products only decreased by 50-fold for the latter digest and the number of quantified proteins in the extract from 1,000 HeLa cells was 73% of that from the extract from 50,000 HeLa cells [19]. The approach that Hughes et al (2014) took to optimize their trypsin digest on an extract from 1,000 Hela cells, which had a trypsin:substrate (w:w) ratio of 1:1.2 (Table S1), is similar to the approaches taken to enable SCP. To achieve the highest possible trypsin and substrate concentrations, the digestion volumes in the SCP protocols in Table S2 have been reduced to as little as 20 nl [58] and the trypsin concentrations have been increased to as high as 90 ng/μl [58] (Table S1).
Based on the number of identified peptides, Doellinger et al. (2020) [39] determined that the optimum trypsin:substrate (w:w) ratios for digesting 50, 250, and 900 ng/μl E. coli extracts are 1:100, 1:50, and 1:20 respectively. As shown in Table 5, if these data are merged with selected data from the larger scale extraction and digestion protocols in Table S1 and the SCP studies in Table S2, it is evident that as the substrate concentration is increased over an 18,000-fold range from 0.05 to 900 ng/μl the corresponding trypsin:substrate (w:w) ratios used extend over a 5,000-fold range from 50:1 to 1:100.
Table 5:
Trypsin:Substrate (w:w) Ratios as a Function of Substrate Concentration
| Reference | Cell Types | [Trypsin] (ng/μl) | Substrate (ng) | [Substrate] (ng/μl) | Trypsin:Substrate (w:w) | Proteins Identified |
|---|---|---|---|---|---|---|
| Doellinger et al. (2020) [39] | E. coli | 9.0 | 900 | 900 | 1:100 | 1,965 |
| Doellinger et al. (2020) [39] | E. coli | 5.0 | 250 | 250 | 1:50 | 1,953 |
| Doellinger et al. (2020) [39] | E. coli | 2.5 | 50 | 50 | 1:20 | 1,914 |
| Hughes et al. (2014) [19] | HeLa | 25.0 | 150 | 30 | 1:1 | 2,811 |
| Woo et al. (2021) [52] | C10 | 16.70 | 0.10 | 3.33 | 5:1 | 1,735 |
| Specht et al. (2021) [49] | U-937, HEK-293 | 10.00 | 0.50 | 0.50 | 20:1 | ~1,000 |
| Budnick et al. (2018) [59] | Jurkat, U-937 | 2.50 | 0.50 | 0.05 | 50:1 | 767 |
7.f. Protein Denaturants:
Harris (1956) determined that trypsin is reversibly denatured in 8 M urea, with 75% of its activity lost in 2 min at 25°C, pH 7.6 but it retains 48% of its activity in 4 M urea, and ~97% of its activity in 2 M urea [118]. Probably, ~2 M urea is the most commonly used chaotrope for carrying out trypsin digestions [90]. Riviere and Tempst (2001) determined that the optimum urea concentration for carrying out trypsin digests is 1.5 M [87]. Three of the selected larger scale extraction and digestion protocols in Table 2 carried out trypsin digests in 1.6–2 M urea [17, 26, 43].
Masuda et al. (2008) [27] evaluated the influence of 27 chaotropic agents, surfactants, and organic solvents reported as enhancers of trypsin activity. As shown in Table 6, 0.01% SDC and 0.01% SDS enhanced trypsin activity by 5.1 and 3.5-fold respectively, while 0.1% RapiGest decreased trypsin activity by ~57%. With the exception of 20% ethanol, which increased trypsin activity by 5.8-fold, surfactants generally provided greater enhancement of trypsin activity than organic solvents or chaotropic reagents. In another study, Proc et al. (2010) evaluated 14 combinations of heat, solvent (acetonitrile, methanol, trifluoroethanol), chaotropic agents (GdnHCl, urea), and surfactants (SDS, SDC) by quantitating by Multiple Reaction Monitoring (MRM) the production of proteotypic tryptic peptides from 45 moderate-to high-abundance plasma proteins [66]. Both 0.025% SDS and 1% SDC increased the overall yield of tryptic peptides as compared to similar digests carried out in 1 M urea. SDC and SDS with a 9 hr digestion time produced the highest average digestion efficiencies (~80%), with the highest average reproducibility. The finding by Riviere and Tempst (2001) that trypsin digests can be carried out in <0.1% SDS [87] is consistent with the use of 0.025% SDS by Proc et al (2010) [66]. Since SDS can interfere with LC-MS/MS while SDC can be easily removed from the samples by acid precipitation, the authors recommended the use of SDC [66]. This recommendation is consistent with the finding by Varnavides et al. (2022) that the 3 in-solution digest protocols that they carried out with SDC identified more proteins in HeLa cell extracts than any of the other 13 protocols [21]. In contrast to Medzihradszky (2005), Riviere and Tempst (2001) found that trypsin does not digest proteins in the presence of GdnHCl because guanidine is a competitive inhibitor of trypsin [87]. This finding is consistent with the observation by Proc et al. (2010) that 0.9 M GdnHCl resulted in significantly lower digestion efficiencies [66] and the finding by Varnavides et al. (2022) that the 3 in-solution digest protocols carried out with GdnHCl identified fewer proteins in HeLa cell extracts than any of the other protocols [21].
Based on a 5 min colorimetric assay at RT Yu et al. (2003) [33] determined that unlike SDS, RapiGest solubilizes proteins without inhibiting trypsin or other common proteases such as LysC. They found the following remaining trypsin activities after 5 min in the indicated solvents: 0.1% RapiGest (99%), 2 M urea (80%), 50% MeOH (21%), 0.1% SDS (20%). Since Masuda et al. (2008) [27] found that 0.1% RapiGest decreased trypsin activity by >57% after a 24 hr assay at 37°C (Table 6), it appears that in 0.1% RapiGest trypsin is stable for 5 min at RT but at 37°C it is gradually denatured over longer periods of time. Based on a tryptic digest of 5 μg of pancreatic cell lysate, Chen et al (2007) found that 0.1% RapiGest provided a 33% increase in the number of proteins identified (i.e., 440) as compared to a similar 16 hr digest carried out at 37°C in the presence of 2 M urea (i.e., 331) and a 37% increase in the number of proteins identified as compared to a digest carried out in 80% acetonitrile (i.e., 322) [38].
Leon et al. (2013) evaluated 6 trypsin digestion protocols that were based on in-solution (4) or spin filter-aided (2) digestion of 100 μg aliquots of rat mitochondrial fractions that were then subjected to RP-HPLC-MS/MS using DIA [23]. The other variables that were investigated included 3 trypsin digestion conditions (0.1% RapiGest, 1 M urea, 0.5% SDC) and 2 methods for removal of detergents before RP-HPLC (acid precipitation or phase separation with ethyl acetate). Compared to the tryptic digest carried out in urea, the in-solution digests in RapiGest and SDC provided ~11% and ~18% more protein identifications, respectively. Regarding digests carried out in SDC, there was no significant difference in the number of protein identifications when the SDC was removed prior to RP-LC by acid precipitation versus phase separation transfer with ethyl acetate as described by Masuda et al. [27]. Analysis of tryptic digests carried out by Leon et al. (2013) revealed that SDC-assisted trypsin digestion followed by phase separation removal of SDC provided the highest recovery, protein sequence coverage, and largest number of protein identifications over a 5–7 hr digestion [23]. Other detergents that have been used during trypsin digests are 2% CHAPS (3-((3-cholamidopropyl) dimethylammonio)-1-propanesulfonate), 2% NP-40, and 2% octyl glucoside (OG) [87] which Masuda et al. (2008) [27] found to be the best trypsin enhancement factor of the 14 agents tested (Table 6). One of the larger scale digests in Table 2 added a surfactant into the digest with Meier et al. (2018) using 0.05% SDC [46]. Five of the 11 SCP studies in Table 3 added a surfactant into the digest with Gebreyesus et al. using 0.3% RapiGest [54] and 4 studies using the MS compatible n-dodecyl-β-D-maltoside (DDM) [50–53].
7.g. Organic Solvents:
Russell et al. (2001) found that rates of protein digestion by modified porcine trypsin at 37°C in organic solvents are increased relative to those carried out in aqueous solutions [119]. Since these solvents are volatile, they can be easily removed by evaporation in a SpeedVac. The optimum organic concentrations for carrying out 1 hr tryptic digests at pH 8 at 37°C on a non-globular protein (e.g. cytochrome C) that is readily digested were 60% methanol, 40% acetone, and 40% ACN [119]. Similarly, Li et al. (2009) determined that 50% methanol was the optimal concentration for the accelerated [30 min at 37° with a 1:50 (w:w) ratio of bovine trypsin:substrate protein] digestion of proteins in human plasma [120]. Blonder et al. (2004) showed that modified porcine trypsin activity measured with a N-benzoyl-L-arginine ethyl ester (BAEE) assay was reduced within 5 min to 20% of that in an aqueous buffer by the addition of 60% methanol. A separate study showed that even in 60% methanol the remaining trypsin activity was still sufficient for the thorough digestion of solubilized membrane proteins [121]. The Fractional Factorial Design of Experiments study carried out by Shuford et al (2012) is in agreement with the BAEE assays carried out by Blonder et al (2004) [121] in that the most negative factor identified by Shuford et al (2012) was that the addition of 50% methanol drastically reduced the efficiency of a 16 hr trypsin digestion [85]. The latter agrees with the colorimetric assay data of Yu et al. (2003) [33] that found that 5 min in 50% methanol at RT reduced trypsin activity by 79%. Based on the production of proteotypic tryptic peptides from 45 human plasma proteins, Proc et al. (2010) reported that while 20% methanol and 5% trifluoroethanol (TFE) produced >9 hr digestion profiles with modified porcine trypsin that were similar to those obtained with 1% SDC, a significant reduction in digestion efficiency was observed for all 45 analytes in 40% ACN [66]. Somiari et al. (2014) carried out trypsin digests of serum and plasma samples in approximately 10% TFE [105] while Riviere and Tempst (2001) reported that 2 hr trypsin digests could be carried out in 40% ACN or 40% isopropanol [87]. Four of the SCP studies in Table 3 carried out tryptic digests in an organic solvent with Schoof et al. (2021) and Petrosius et al. (2023) using 10% TFE [55, 56], Bruner et al. using 20% ACN [57], and Huffman et al. (2023) using 40% DMSO [58]. Schoof et al. (2021) found that using 20% TFE during lysis and 10% TFE during trypsin digestion provided an approximately 5% increase in the average number of identified proteins (i.e., 1,791 versus 1,873) in a quadruplicate determination [55]. In addition, one of the larger scale digestion protocols in Table 2 carried out a trypsin digest in 10% ACN [20].
8. Multistage Trypsin Digestion:
Tryptic digests of complex cell extracts are often biased by high abundance proteins that produce a corresponding excess of frequently identified, high abundance peptides. Fonslow et al. (2013) developed a novel approach to meet this challenge that involved carrying out two tryptic digests [122]. After the initial digest was carried out under trypsin-limited conditions (i.e., trypsin:protein w:w ratio = 1:25,000; 12 hrs) on a 10-fold larger amount of extract than normal (i.e., ~1 mg), approximately 85% of the high abundance proteins were removed as peptides by use of a 10 kDa cutoff Amicon spin filter. The remaining lower abundance proteins were then subjected to a “normal” trypsin digestion (i.e., trypsin:protein ratio was 1:100; overnight). When this approach was used on an HEK cell extract it resulted in an 18% increase in the number of identified proteins (i.e., 7,716 versus 6,513) [122]. In a follow-up letter to the Editor, Ye et al. (2014) with a response from Fonslow et al. (2013), it was noted that protein digestion with trypsin is independent of protein abundances and that multistage trypsin digestion likely works by removing small, easily cleaved peptides rather than by removing peptides from high abundance proteins [123, 124]. Consistent with this conclusion, Pan et al. (2015) added two more trypsin digestion and 10 kDa ultrafiltration steps that provided a 2.2-fold increase in the number of identified proteins [125]. As noted by the authors, this multi-stage digestion strategy separates tryptic peptides with different cleavage kinetics while RP–HPLC separates peptides with different hydrophobicities [125].
9. Kinetics of Trypsin Digestion of Complex Proteomes:
There is a wide range in the rates of trypsin cleavage of individual Lys/Arg-X bonds with some “missed cleavage” sites not being cleaved due to very slow kinetics [126, 127]. Specifically, the “Keil Rule” is that trypsin cleaves after Arg or Lys but not before Pro [128]; however, an analysis of 14.5 million tandem spectra revealed an unexpectedly large number of tryptic cleavages before Pro [129]. Indeed, the frequency of (Arg/Lys)-Pro cleavages was higher than that for (Arg/Lys)-Cys cleavages and was comparable to that for (Arg/Lys)-Trp cleavages [129]. Hence, an updated version of the “Keil Rule” is that trypsin cleaves (Arg/Lys)-Pro bonds but with less propensity than most other Arg/Lys-X bonds. Based on studies carried out on beta caseins, the rate constant for the hydrolysis of an Arg-X bond is about 8-fold higher than that for Lys-X which means that trypsin has a significant preference for cleaving Arg-X bonds [130]. This finding is consistent with the observation that trypsin has a higher frequency of missed cleavages after Lys than after Arg residues [131] and with the analyses carried out by Rodriguez et al. (2008). Notably, this latter study found that while the trypsin cleavage rate generally is higher after Arg than after Lys, Lys-Tyr, Lys-Phe, and Lys-His have higher cleavage efficiency than the corresponding Arg linkages [129].
Pan et al. (2014) investigated the kinetics of trypsin-catalyzed digestion of a Hela cell lysate [132]. After digesting for 1, 4 and 18 hrs with a 1:40 (w:w) trypsin:protein ratio, aliquots of the digest were removed and then labeled with 3 different stable isotope dimethyl labels. MS/MS analysis identified 10,483 unique peptides from 2,270 proteins. Four types of cleavage sites (i.e., very fast, fast, slow, and very slow) were identified. In agreement with previous studies noted above [129, 130], trypsin cleaved Arg-X faster than Lys-X linkages. In addition, while tryptic cleavage sites surrounded by neutral residues were rapidly cut, those with neighboring charged (i.e., Asp, Glu, Lys, or Arg) or Pro residues were cleaved more slowly [132]. The analyses also showed that digestion of individual proteins in a complex proteome is independent of their abundances and their physicochemical properties, such as MW, grand average of hydropathicity (GRAVY), aliphatic index, and pI [132]. Thus, limited tryptic digestion would not enrich or deplete for selective types of proteins.
Basic or negatively charged residues adjacent to cleavage sites as well as PTMs impair proteolytic cleavage, leading to missed cleavage sites and complicating phosphoproteome analyses [18]. The interfering effect of (R or K)n(X)(S or T) residues near cleavage sites was first proposed by Lehmann and co-workers in 2001 [133]. Schlosser et al. (2001) observed the incomplete tryptic digestion of protein kinase A (PKA) within the phosphorylated PKA consensus sequence R-X-pS [133]. Dickhut (2014) [134] also quantified a strong reduction in tryptic cleavage within phosphorylated PKA motifs I-R-X-pS/pT and also R-X-X-pT sequences, however, I-R-X-pY sequences were almost unaffected. Structural predictions implied the formation of salt bridges between R/K cleavage sites and phosphoamino acids pS/pT as the main reason for impaired tryptic digestion [134]. Methylation of Lys and Arg provides an important epigenetic and regulatory mechanism for protein-protein interactions and protein function in transcription, DNA repair and signaling. Methylated basic residues strongly impair tryptic digestion, generating missed cleavages that hinder identification of peptide methylation sites [131].
10. Single Cell Proteomics (SCP) Analyses:
Because analyzing tissues that are composed of diverse cell types obscures cell-to-cell differences, there has been considerable interest over the last few years in developing technologies to analyze the proteomes of a few or single cells. Since the amount of protein in each of the mammalian cells used in the SCP studies in Tables 3 and S2 is limited to 0.1–0.5 ng, two approaches have been used to increase the rate of trypsin digestion: 1) minimize the digestion volume as this decreases protein losses due to adsorption and maximizes the [trypsin] and [substrate] concentrations that determine the initial rates of the trypsin digest; and 2) increase the concentration of trypsin. Hence, the digestion volumes for 9 of the 11 SCP studies in Table S2 range from 20 nl [58] to 2.0 μl [55–57]. In addition, most of these digests used much higher than the conventional 1:10 to 1:100 trypsin:protein (w:w) ratios that have long been used in larger scale digests. Thus, 10 of the 11 studies in Table S2 used an excess of trypsin with the ratios varying from 1.7-fold [50] to 50-fold trypsin:protein w:w ratio [59]. We note that while 2 of the 11 SCP studies did not use any surfactant or organic solvent to aid protein extraction and denaturation [49, 59], 4 of the studies used DDM [50–53], 1 used RapiGest [54], and 4 used organic solvents (i.e., 10% TFE [55, 56], 20% ACN [57], and 40% DMSO [58]). The 11 SCP studies summarized in Tables 3 and S2 quantified from 211 to 2,083 proteins with 4 of the studies quantifying 1,500 or more proteins.
11. Conclusions and Recommendations:
11.a. General Considerations:
Because of the exceedingly large number of variables in sample type, amount of protein input, and method workflow (see Tables S1 and S2), it is not possible to use the numbers of quantified proteins as a guide to uncover the optimum conditions for carrying out single cell and larger scale tryptic digests. However, based on the studies covered in this review, it is possible to at least begin to delineate optimal conditions for carrying out trypsin digestions on limiting and bulk amounts of complex proteomes. A significant limitation of many studies is that they investigate one factor at a time (OFAT) which can result in a local minimum or maximum being reached for the specific parameter being studied that may not be the true optimum [81]. To avoid this inherent limitation of OFAT, a fractional or full-factorial DOE approach allow for two-way and higher level interactions of each parameter under investigation [81]. In the text below we recommend optimal trypsin digestion conditions that are reasonably well supported by published data and indicate important conditions that need more research to optimize.
11.b. General Recommendations:
Table 7 provides a summary of important variables and considerations that have been identified in this review for optimizing trypsin digests of complex proteomes. We recommend extracting proteins with SDS, TFA, or SDC with TFA preferred for difficult to lyse samples. We recommend precipitating protein extracts from 1) SDS with acetone, albeit using an acetone wash as suggested by Botelho et al. (2010) [32], 2) GdnHCl with ETOH, and 3) SDC or urea with either chloroform/methanol or acetone. Alternatively, one of the “cleanup” technologies can be used such as iST [20] for GdnHCl, STrap [34] or SP3 [19] for SDC, or FASP [17] or SP3 [19] for SDS. We recommend that porcine trypsin be used since it has a lower missed cleavage rate than either human or bovine trypsin. Since commercially available bovine and porcine trypsin preparations may be contaminated with small amounts of chymotrypsin, this challenge can be obviated by treating with the irreversible chymotrypsin inhibitor, TPCK. Trypsin also should be reductively methylated so it is resistant to auto-proteolysis [78, 79] that can generate pseudotrypsin that has decreased rates of cleavage of Arg/Lys and a chymotrypsin-like specificity [71, 77] [82]. Based on the work by Niu et al. (2020), we recommend that trypsin is reconstituted in HPLC grade water prior to carrying out the digests [84].
Table 7:
General Recommendations for Extracting and Digesting Proteins for LC/MS/MS Proteome Analyses
| Description | Recommendations | References |
|---|---|---|
| Protein Extractants | SDS, TFA, or SDC | [21, 29, 39] |
| Protein Precipitants | Acetone (with an acetone wash) for SDS ETOH for GdnHCl Chloroform/methanol or acetone for SDC or urea |
[21, 32, 35] |
| Digest Protocols | ISD-SDC, ISD-SDC/A, ISD-SDC/CM, EasyPep, iST-GdnHCl, or FASP-SDS | [21] |
| Proteases | LysC/TPCK-treated, dimethylated porcine trypsin | [7, 11, 13, 15–17], [63–65], [75, 76, 82] |
| Solvent for Reconstituting Trypsin | Ultrapure water | [84] |
| Trypsin Digest Component | 10 mM Ca++ | [16, 85–87] |
| Reduction and Alkylation | Not Recommended | [88, 91–96], [107] |
| Trypsin Digestion Temperature | 37°C | [7, 16, 82, 87, 90] |
| Trypsin Digestion pH | 8 | 15, 16, 18, 25, 40, 82 [101] [103], |
| Trypsin Digestion Time | 8–9 hrs | [66, 85, 103, 105] |
| Trypsin:Substrate (w:w) Ratio | As the substrate concentration is decreased, we recommend that the trypsin:substrate (w:w) ratios be increased as shown in Table 5. | |
| Detergents for In-Solution Trypsin Digests | SDC (preferred) or RapiGest or DDM | [23, 27, 38, 66] |
| Organic Solvents for In-Solution Trypsin Digests | 5–10% TFE, 20% MeOH, or 10–20% ACN | [20, 55, 57, 66, 105] |
Based on the study by Varnavides et al. (2022) [21], we recommend using any of the 3 in solution digests with SDC, the EasyPep or iST-GdnHCl commercial kits, or the FASP-SDS protocols (Table 7) that all were within about 5% of the top performing ISD-SDC/CM protocol. However, we recommend that the descriptive matrix in Figure 5 in Varnavides et al. (2022) [21] be used as a guide to help choose the technology that is most likely to provide the best yield of specific proteins of interest. In the case of difficult-to-lyse samples (e.g., mouse lung tissue and B. cereus), we recommend using the SPEED approach [39].
Since LysC/trypsin digests provide improved proteolysis due to the complementary target specificities of the two enzymes [11, 63–65], we recommend that all proteome extracts be subjected to LysC/trypsin rather than to just trypsin digests. While some protocols carry out sequential LysC/trypsin digests [17, 26, 50, 52], others [19, 20, 54, 57] carry out simultaneous LysC/trypsin digests. Although a tandem digest can take advantage of the increased stability of LysC in 8M urea [17], we have not found data that compares the relative effectiveness of sequential versus simultaneous LysC/trypsin digests of complex proteome extracts. Until such data is available, we recommend carrying out sequential LysC/trypsin digests.
Digest buffers should include 10 mM Ca++ as it promotes a more compact trypsin structure that stabilizes against autolysis [16, 85–87]. In view of unwanted side reactions with alkylating agents [88, 91–96]; the finding that reduction and alkylation does not significantly affect the numbers of identified proteins [88, 93]; and the success of a microgram scale proteome profiling procedure [107] and 7 SCP protocols in Table 3 that did not carry out reduction and alkylation [49, 51, 53, 56–59]; we recommend that proteome extracts be digested without carrying out prior reduction and alkylation. Furthermore, sample preparation should be carried out under conditions protecting thiols from oxidation [88]. If, however, the decision is made to carry out reduction and alkylation, we recommend that it be carried out with DTT and CAA.
We recommend that modified porcine trypsin digestions generally be carried out at 37°C for 8–9 hrs. However, if it is important to significantly reduce the trypsin digestion time, then we recommend using a 15 min digestion time at 70°C with Promega’s Rapid Digestion Trypsin and the accompanying “Resuspension Buffer” as described by Doellinger et al. (2020) [39]. If the digest time must be shortened below 15 min, we recommend using ultrasonically-assisted digests (UADs) as several studies have shown that in-solution [109, 110, 112–114], in-gel [111, 112], and FASP [115] UADs can be carried out in as little as 2–5 min.
Although successful trypsin digests have been carried out in multiple buffers over the pH range extending from 7 to 8.5, we believe that additional research is needed to identify the optimum buffer and pH for carrying out trypsin digestions by modified porcine trypsin on complex cell and tissue extracts. Until additional data is available, we recommend carrying out modified porcine trypsin digests at pH 8 as suggested by kinetic studies [101], Hao et al. [103], and several protocols [16, 17, 19, 26, 43, 87].
Since the progress curve of a trypsin catalyzed reaction is a function of the concentrations of trypsin and substrate rather than their w:w ratios, the approach of using a fixed w:w ratio has significant limitations. We recommend therefore that as the substrate concentration in the trypsin digest is decreased that the w:w ratio of trypsin:substrate be increased as indicated in Table 5.
We recommend that when sample sizes are sufficiently large to permit acidification followed by centrifugal or phase separation removal of detergents that in-solution trypsin digest workflows use SDC. Otherwise, and especially for SCP, we recommend that digests be carried out with the SCoPE2 [49] approach or in RapiGest [54], DDM [50–53] or in one of the following limited levels of organic solvents: 5–10% TFE [55, 66, 105], 20% MeOH [66], or 10–20% ACN [20, 57]. Since Proc et al. (2010) [66] reported that in the presence of 20% MeOH and 5% TFE that modified porcine trypsin produced digestion profiles similar to those with 1% SDC, we recommend giving highest priority to these two solvents and concentrations.
11.c. General Conclusions:
In conclusion, and as noted by Vandermarliere et al. (2013) [7], there is still “an unmet need for a gold standard, verified, simple yet efficient general purpose digestion protocol”. In this regard, it is likely that at least two optimum trypsin digestion protocols will be needed – with one for single to few cells and the other for larger scale digests. It is also likely that the conditions for optimal trypsin digestion will be somewhat protein and/or proteome dependent. Since the precise proteome under study presents a unique and complex chemical milieu that may enhance or inhibit protein digestion, for long-term or large-scale biological studies, a fractional factorial design [81] is recommended to optimize the digestion conditions for the specific proteome. That is, especially for those factors (e.g., digestion time, trypsin and substrate concentrations, choice and concentration of surfactant or organic solvent) that are likely to be somewhat proteome dependent, the “optimal” conditions that are suggested above should be viewed as a “starting” point for further optimization.
In addition to the specific recommendations described above, it is important to also highlight what is still not known. For example, if the optimal conditions for carrying out trypsin digestions on mammalian cells were well defined, then it would have been expected that all 11 of the SCP analyses in Tables 3 and S2 would have used them to carry out such demanding digests. This review thus highlights the need for further experimentation to determine: 1) the optimum buffer and pH for carrying out tryptic digests, 2) if RapiGest or another surfactant (e.g., DDM) or one of the several organic solvents recommended in this review provide the most optimum conditions for SCP digests, and 3) the relative effectiveness of tandem versus simultaneous LysC/trypsin digests of complex proteome extracts. Finally, Hildonen et al. (2014) conclude that for bottom-up proteomic studies it would be beneficial to prevent trypsin digests from going to completion by reducing the digest time from the conventional several hours to just 5 min and to then allow up to 3 missed cleavages during the database searches [104]. We suggest that it would be worthwhile to 4) carry out the research needed to determine if such an approach does result in reproducibly increasing the numbers of proteins that are identified in complex proteome extracts.
Supplementary Material
Table S1: Selected Larger Scale Extraction and Digestion Protocols
Table S2: Selected Single Cell Proteome Analyses
Table S3: Comparisons of Numbers of Identified Proteins Using Different Extraction and Digestion Protocols
Table S4: Comparison of Bottom-Up Proteomics Sample Preparation Methods from Varnavides et al. (2022) [21]
Significance:
Currently, bottom-up MS-based proteomics is the method of choice for global proteome analysis. Since trypsin is the most utilized protease in bottom-up MS proteomics, delineating optimal conditions for carrying out trypsin digestions of complex proteomes in samples ranging from tissues to single cells should positively impact a broad range of biomedical research.
Highlights.
Enzymatic digestion is essential to identify proteins by the bottom-up MS approach.
Trypsin is the most commonly used enzyme for bottom-up proteomics.
Trypsin activity is highly dependent upon numerous factors.
This review delineates optimal conditions for trypsin digestion of proteomes.
Acknowledgements:
We especially thank Dr. David Muddiman (North Carolina State University) for careful reading of our manuscript and for highlighting the benefits of fractional and full-factorial design of experiments. We also thank the reviewers for the time and effort they devoted to reviewing this manuscript and for their insightful comments and detailed suggestions for improving it.
Funding Sources:
We acknowledge support from the NIH (Yale/NIDA Neuroproteomics Center grant DA018343). Research in the laboratory of ACN is supported by NIH (AG047270, AG062306, AG066508, DA018343) and the State of Connecticut Department of Mental Health and Addiction Services.
Abbreviations:
- ABC
ammonium bicarbonate
- BAEE
N-benzoyl-L-arginine ethyl ester
- bSDB
sulfonated polystyrenedivinylbenzene
- BLNA
N-benzoyl-lysine p-nitroanilide
- CAA
chloroacetamide
- CHAPS
(3-((3-cholamidopropyl) dimethylammonio)-1-propanesulfonate)
- CB
Coomassie Blue
- CVs
Coefficients of Variation
- DIA
data independent acquisition
- DDA
data dependent acquisition
- DDM
n-dodecyl-β-D-maltoside
- SDC
sodium deoxycholate
- DOE
Design of Experiments
- DTT
dithiothreitol
- EtOH
ethanol
- FASP
filter-aided sample preparation
- FDR
False Discovery rate
- FracFD
fractional factorial design
- GRAVY
grand average of hydropathicity
- GdnHCl
guanidine hydrochloride
- HEPES
(4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid)
- HILIC
Hydrophilic Interaction Liquid Chromatography
- hr
hour
- IAC
iodoacetic acid
- IAA
iodoacetamide
- iProChip
integrated proteomics chip
- ISD
in solution digest
- iST
in-Stage-Tip
- iTRAQ
Isobaric Tag for Relative and Absolute Quantitation
- ITC
isothermal titration calorimetry
- LFQ
Label-free Quantification
- MeOH
methanol
- mPOP
Minimal ProteOmic Sample Preparation
- MBR
match between runs
- MMTS
methyl methanethiosulfonate
- MRM
Multiple Reaction Monitoring
- MS/MS
tandem mass spectrometry
- MW
molecular weight
- nanoPOTS
nanodroplet processing in one pot for trace samples
- OCT
optimum cutting temperature
- OG
octyl glucoside
- OFAT
one factor at a time
- PBS
phosphate-buffered saline
- PC-IDMS
protein cleavage isotope dilution mass spectrometry
- PCT
pressure cycling technology
- ppm
parts per million
- PTMs
post-translational modifications
- RIPA
Radioimmunoprecipitation assay
- RP-HPLC
reversed phase high performance liquid chromatography
- RT
room temperature
- SCIProChip
single-cell proteomics chip
- SCoPE-MS
Single Cell ProtEomics by Mass Spectrometry
- SCP
single cell proteomics
- SCX
strong cation exchange
- SDC
sodium deoxycholate
- SLS
sodium lauroyl sarcosinate
- SPE
solid phase extraction
- SP3
Single-Pot Solid-Phase-enhanced Sample Preparation
- SPEED
Sample Preparation by Easy Extraction and Digestion
- STrap
suspension trapping
- SWATH-MS
Sequential Window Acquisition of All Theoretical Fragment Ion Mass Spectra
- TCA
trichloroacetic acid
- TCEP
Tris(2-carboxyethyl)phosphine
- TMT
Tandem Mass Tag
- TEAB
Triethylammonium bicarbonate
- TFA
trifluoroacetic acid
- TFE
trifluoroethanol
- TPCK
1-chloro-3-tosylamido-4-phenyl-2-butanone
- Tris(HCl)
Tris(hydroxymethyl)aminomethane hydrochloride
- T-SCP
true single-cell–derived proteomics
- UADs
ultrasonically-assisted digests
- 4-VP
4-vinylpyridine
Footnotes
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Conflicts of Interest:
The authors declare there are no conflicts of interest.
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Associated Data
This section collects any data citations, data availability statements, or supplementary materials included in this article.
Supplementary Materials
Table S1: Selected Larger Scale Extraction and Digestion Protocols
Table S2: Selected Single Cell Proteome Analyses
Table S3: Comparisons of Numbers of Identified Proteins Using Different Extraction and Digestion Protocols
Table S4: Comparison of Bottom-Up Proteomics Sample Preparation Methods from Varnavides et al. (2022) [21]