Sensitivity of mass spectrometry analysis depends on the shape of the filtration unit used for filter aided sample preparation (FASP)

Efficient protein solubilization using detergents is required for in‐depth proteome analysis, but successful LC‐MS/MS analysis greatly depends on proper detergents removal. A commonly used sample processing method is the filter‐aided sample preparation (FASP), which allows protein digestion and detergent removal on the same filtration device. Many optimizations of the FASP protocol have been published, but there is no information on the influence of the filtration unit typology on the detergents removal. The aim of this study was to compare the performance of conic and flat bottom filtration units in terms of number of proteins identified by LC‐MS/MS. We have analyzed 1, 10 and 100 μg of total cell lysate prepared using lysis buffer with different SDS concentrations. We compared the FASP protocol using conic and flat bottom filtration units to ethanol precipitation method. Subsequently, we applied our most performant protocol to single murine pancreatic islet, and identified up to 2463 protein using FASP versus 1169 proteins using ethanol precipitation. We conclude that FASP performance depends strongly on the filter shape: flat bottom devices are better suited for low‐protein samples, as they allow better SDS removal leading to the identification of greater number of proteins.     

C-STrap Sample Preparation Method—In-Situ Cysteinyl Peptide Capture for Bottom-Up Proteomics Analysis in the STrap Format

Recently we introduced the concept of Suspension Trapping (STrap) for bottom-up proteomics sample processing that is based upon SDS-mediated protein extraction, swift detergent removal and rapid reactor-type protein digestion in a quartz depth filter trap. As the depth filter surface is made of silica, it is readily modifiable with various functional groups using the silane coupling chemistries. Thus, during the digest, peptides possessing specific features could be targeted for enrichment by the functionalized depth filter material while non-targeted peptides could be collected as an unbound distinct fraction after the digest. In the example presented here the quartz depth filter surface is functionalized with the pyridyldithiol group therefore enabling reversible in-situ capture of the cysteine-containing peptides generated during the STrap-based digest. The described C-STrap method retains all advantages of the original STrap methodology and provides robust foundation for the conception of the targeted in-situ peptide fractionation in the STrap format for bottom-up proteomics. The presented data support the method’s use in qualitative and semi-quantitative proteomics experiments.     

Automated on-line ionic detergent removal from minute protein/peptide samples prior to liquid chromatography-electrospray mass spectrometry

An automated on-line ionic detergent removal pre-column system coupled to capillary liquid chromatography-electrospray mass spectrometry is described. The system involves two micro precolumns, composed of a specific ionic detergent trapping column and a preconcentration column, respectively, and a packed 300 μm I.D. analytical column. Sample loading to the micro precolumns and regeneration of the detergent trapping column were performed at a flow-rate of 50 μl/min, while the flow-rate through the analytical column was set at 5.0 μl/min. Ionic detergent-containing tryptic-digested protein samples were directly applied to the micro precolumns without sample pretreatment and were analysed by UV absorption detection and electrospray mass spectrometry. The presented system allows for the fully automated removal of SDS with virtually no loss in protein/peptides. Maximum SDS load and breakthrough have been determined. Excellent protein recovery and complete removal of SDS is found. The chromatographic separation after SDS removal was completely restored and equalled the reference chromatogram. Mass spectral data confirm these findings. Finally, this technique allows for SDS removal from minute protein samples without the need for any sample handling.     

Membrane‐Based SDS Depletion Ahead of Peptide and Protein Analysis by Mass Spectrometry

SDS interferes with both bottom‐up and top‐down MS analysis, requiring removal prior to detection. Filter‐aided sample preparation (FASP) is favored for bottom‐up proteomics (BUP) while acetone precipitation is popular for top‐down proteomics (TDP). We recently demonstrated acetone precipitation in a membrane filter cartridge. Alternatively, our automated electrophoretic device, termed transmembrane electrophoresis (TME), depletes SDS for both TDP and BUP studies. Here TME is compared to these two alternative methods of SDS depletion in both BUP and TDP workflows. To do so, a modified FASP method is described applicable to the SDS purification and recovery of intact proteins, suitable for LC/MS. All three methods reliably deplete >99.8% SDS. TME provide higher sample yields (average 90%) than FASP (55%) or acetone precipitation (57%), translating into higher total protein identifications (973 vs 877 FASP or 890 acetone) and higher spectral matches (2.5 times) per protein. In a top down workflow, each SDS‐depletion method yields high‐quality MS spectra for intact proteins. These results show each of these membrane‐based strategies is capable of depleting SDS with high sample recovery and high spectra quality for both BUP and TDP studies.     

Preparation and application of a partially degradable gel in mass spectrometry-based proteomic analysis

In-gel digestion is an attractive route in mass spectrometry-based proteomic analysis, which, however, often suffers from a certain amount of sample loss mainly due to insufficient protein digestion and peptide extraction. To address this, herein we establish a partially degradable gel-assisted protein digestion and peptide recovery method by means of a simple replacement of bis-acrylamide (BA) with bis-acrylylcystamine (BAC). Concretely, the protein sample solubilized using high concentrations of sodium dodecyl sulfate (SDS) and urea were directly entrapped and immobilized into BAC-crosslinked gel by vacuum-dried gel absorption followed by fixation treatment. After removal of SDS and urea by repeated washing, the proteins were subjected to in-gel digestion and the gel was reductively treated. The tryptic peptides were recovered from the partial degradation of the gel and analyzed afterwards by capillary liquid chromatography coupled with tandem mass spectrometry (CapLC-MS/MS). Compared with conventional BA-crosslinked gel method, this new method increased the numbers of identified proteins and unique peptides by 20.2% and 20.4%, respectively. The further statistical analysis demonstrated that the method improved the recovery of tryptic peptides particularly larger and/or hydrophobic peptides, thereby significantly facilitating protein identification. Thus, the newly developed method is a promising alternative for BA-crosslinked gel-based shotgun workflows and has potential application in the related fields of protein chemistry and proteomics.     

Protein identification and quantification from riverbank grape,Vitis riparia: Comparing SDS‐PAGE and FASP‐GPF techniques for shotgun proteomic analysis

Protein sample preparation optimisation is critical for establishing reproducible high throughput proteomic analysis. In this study, two different fractionation sample preparation techniques (in‐gel digestion and in‐solution digestion) for shotgun proteomics were used to quantitatively compare proteins identified in Vitis riparia leaf samples. The total number of proteins and peptides identified were compared between filter aided sample preparation (FASP) coupled with gas phase fractionation (GPF) and SDS‐PAGE methods. There was a 24% increase in the total number of reproducibly identified proteins when FASP‐GPF was used. FASP‐GPF is more reproducible, less expensive and a better method than SDS‐PAGE for shotgun proteomics of grapevine samples as it significantly increases protein identification across biological replicates. Total peptide and protein information from the two fractionation techniques is available in PRIDE with the identifier PXD001399 ( http://proteomecentral.proteomexchange.org/dataset/PXD001399 ).     

Comparison of detergent‐based sample preparation workflows for LTQ‐Orbitrap analysis of the Escherichia coli proteome

This work presents a comparative evaluation of several detergent‐based sample preparation workflows for the MS‐based analysis of bacterial proteomes, performed using the model organism Escherichia coli. Initially, RapiGest‐ and SDS‐based buffers were compared for their protein extraction efficiency and quality of the MS data generated. As a result, SDS performed best in terms of total protein yields and overall number of MS identifications, mainly due to a higher efficiency in extracting high molecular weight (MW) and membrane proteins, while RapiGest led to an enrichment in periplasmic and fimbrial proteins. Then, SDS extracts underwent five different MS sample preparation workflows, including: detergent removal by spin columns followed by in‐solution digestion (SC), protein precipitation followed by in‐solution digestion in ammonium bicarbonate or urea buffer, filter‐aided sample preparation (FASP), and 1DE separation followed by in‐gel digestion. On the whole, about 1000 proteins were identified upon LC‐MS/MS analysis of all preparations (>1100 with the SC workflow), with FASP producing more identified peptides and a higher mean sequence coverage. Each protocol exhibited specific behaviors in terms of MW, hydrophobicity, and subcellular localization distribution of the identified proteins; a comparative assessment of the different outputs is presented.     

APols-Aided Protein Precipitation: A Rapid Method for Concentrating Proteins for Proteomic Analysis

Amphipols (APols) are a newly designed and milder class of detergent. They have been used primarily in protein structure analysis for membrane protein trapping and stabilization. We have recently demonstrated that APols can be used as an alternative detergent for proteome extraction and digestion, to achieve a “One-stop” single-tube workflow for proteomics. In this workflow, APols are removed by precipitation after protein digestion without depleting the digested peptides. Here, we took further advantage of this precipitation characteristic of APols to concentrate proteins from diluted samples. In contrast with tryptic peptides, a decrease in pH leads to the unbiased co-precipitation of APols with proteins, including globular hydrophilic proteins. We demonstrated that this precipitation is a combined effect of acid precipitation and the APols’ protein interactions. Also, we have been able to demonstrate that APols-aided protein precipitation works well on diluted samples, such as secretome sample, and provides a rapid method for protein concentration.     

Comparison of sample preparation techniques for large‐scale proteomics

Numerous workflows exist for large‐scale bottom‐up proteomics, many of which achieve exceptional proteome depth. Herein, we evaluated the performance of several commonly used sample preparation techniques for proteomic characterization of HeLa lysates [unfractionated in‐solution digests, SDS‐PAGE coupled with in‐gel digestion, gel‐eluted liquid fraction entrapment electrophoresis (GELFrEE) technology, SCX StageTips and high‐/low‐pH reversed phase fractionation (HpH)]. HpH fractionation was found to be superior in terms of proteome depth (>8400 proteins detected) and fractionation efficiency compared to other techniques. SCX StageTip fractionation required minimal sample handling and was also a substantial improvement over SDS‐PAGE separation and GELFrEE technology. Sequence coverage of the HeLa proteome increased to 38% when combining all workflows, however, total proteins detected improved only slightly to 8710. In summary, HpH fractionation and SCX StageTips are robust techniques and highly suited for complex proteome analysis.     

基于超滤辅助样品制备方法的优化简

目的：基于超滤辅助样品制备（FASP）方法的出现使得使用去污剂（如SDS）的蛋白质提取方法与溶液内酶切方法得以兼容,因此提高了难溶性蛋白的鉴定数量。然而,超滤膜的非特异性吸附作用依然会造成蛋白的损失。我们拟针对该方法存在的问题对其进行改进。方法：对FASP方法进行了蛋白酶切条件、洗脱液选择、洗脱次数的改进;为测试优化方法的有效性和适用范围,选择标准蛋白BSA、鼠肝和鼠脑等3种样品进行考察。结果：相比报道的FASP方法,采用改进后的FASP方法使BSA的回收率提高了20％;经高精度质谱检测,对鼠肝、鼠脑的蛋白质鉴定结果分别比采用未优化的FASP多鉴定到2086和3592条特异肽段。结论：通过对FASP方法的优化,蛋白鉴定数量得到较大提高,该方法为蛋白质组深度覆盖研究提供了可靠的技术手段。     

Assessment and Refinement of Sample Preparation Methods for Deep and Quantitative Plant Proteome Profiling

A major challenge in the field of proteomics is obtaining high‐quality peptides for comprehensive proteome profiling by LC–MS. Here, evaluation and modification of a range of sample preparation methods using photosynthetically active Arabidopsis leaf tissue are done. It was found that inclusion of filter‐aided sample preparation (FASP) based on filter digestion improves all protein extraction methods tested. Ultimately, a detergent‐free urea‐FASP approach that enables deep and robust quantification of leaf and root proteomes is shown. For example, from 4‐day‐old leaf tissue, up to 11 690 proteins were profiled from a single sample replicate. This method should be broadly applicable to researchers working with difficult to process plant samples.     

Quantitative filtration-blotting of protein in the presence of sodium dodecyl sulfate and its use for protein assay

It is thought that sodium dodecyl sulfate (SDS), an anionic detergent, binds to hydrophobic moieties of peptide to destroy the conformational structure of protein. Because of this property, it is involved in many biochemical procedures such as separations of protein and proteolytic digestion. In the course of our study on a solid-phase protein assay, we found that SDS acts as an effective reagent for protein blotting onto a hydrophobic membrane of polyvinylidene difluoride with a manifold dot-blot apparatus. At least 0.1% SDS in an acid-ethanol blotting solution, while reducing the bias of pronounced interferers for protein assay to protein-membrane interaction, quantitatively retains protein on the membrane. Presumably, protein denatures by SDS to become an unfolded state and adsorbs into the membrane by hydrophobic interaction, even in the presence of excess SDS. Therefore, bolts stained with a pyrogallol red-molybdate complex (Pyromolex) reagent unreactive to the membrane allowed a precise protein determination without significant interference of materials, especially detergents in the sample solution. The filtration-blotting with SDS would be a crucial procedure for quantitative analyses such as immunoblotting in detergent-containing samples, together with the solid-phase protein assay with limited sample volumes, such as 20 microL or less.     

Benchmarking sample preparation/digestion protocols reveals tube‐gel being a fast and repeatable method for quantitative proteomics

Sample preparation, typically by in‐solution or in‐gel approaches, has a strong influence on the accuracy and robustness of quantitative proteomics workflows. The major benefit of in‐gel procedures is their compatibility with detergents (such as SDS) for protein solubilization. However, SDS‐PAGE is a time‐consuming approach. Tube‐gel (TG) preparation circumvents this drawback as it involves directly trapping the sample in a polyacrylamide gel matrix without electrophoresis. We report here the first global label‐free quantitative comparison between TG, stacking gel (SG), and basic liquid digestion (LD). A series of UPS1 standard mixtures (at 0.5, 1, 2.5, 5, 10, and 25 fmol) were spiked in a complex yeast lysate background. TG preparation allowed more yeast proteins to be identified than did the SG and LD approaches, with mean numbers of 1979, 1788, and 1323 proteins identified, respectively. Furthermore, the TG method proved equivalent to SG and superior to LD in terms of the repeatability of the subsequent experiments, with mean CV for yeast protein label‐free quantifications of 7, 9, and 10%. Finally, known variant UPS1 proteins were successfully detected in the TG‐prepared sample within a complex background with high sensitivity. All the data from this study are accessible on ProteomeXchange (PXD003841).     

Efficient Tandem LysC/Trypsin Digestion in Detergent Conditions

All shotgun proteomics experiments rely on efficient proteolysis steps for sensitive peptide/protein identification and quantification. Previous reports suggest that the sequential tandem LysC/trypsin digest yields higher recovery of fully tryptic peptides than single‐tryptic proteolysis. Based on the previous studies, it is assumed that the advantageous effect of tandem proteolysis requires a high sample denaturation state for the initial LysC digest. Therefore, to date, all systematic assessments of LysC/trypsin proteolysis are done in chaotropic environments such as urea. Here, sole trypsin is compared with LysC/trypsin and it is shown that tandem digestion can be carried with high efficiency in Mass Spectrometry‐compatible detergents, thereby resulting in higher quantitative yields of fully cleaved peptides. It is further demonstrated that higher cleavage efficiency of tandem digests has a positive impact on absolute protein quantification using intensity‐based absolute quantification (iBAQ) values. The results of the examination of divergent urea tandem conditions imply that beneficial effects of the initial LysC digest do not depend on the sample denaturation state, but, are mainly caused by different target specificities of LysC and trypsin. The observed detergent compatibility enables tandem digestion schemes to be implemented in efficient cellular solubilization proteomics procedures without the need for buffer exchange to chaotropic environments.     

A high‐throughput sample preparation method for cellular proteomics using 96‐well filter plates

A high‐throughput sample preparation protocol based on the use of 96‐well molecular weight cutoff (MWCO) filter plates was developed for shotgun proteomics of cell lysates. All sample preparation steps, including cell lysis, buffer exchange, protein denaturation, reduction, alkylation and proteolytic digestion are performed in a 96‐well plate format, making the platform extremely well suited for processing large numbers of samples and directly compatible with functional assays for cellular proteomics. In addition, the usage of a single plate for all sample preparation steps following cell lysis reduces potential samples losses and allows for automation. The MWCO filter also enables sample concentration, thereby increasing the overall sensitivity, and implementation of washing steps involving organic solvents, for example, to remove cell membranes constituents. The optimized protocol allowed for higher throughput with improved sensitivity in terms of the number of identified cellular proteins when compared to an established protocol employing gel‐filtration columns.     

Increased proteome coverage by combining PAGE and peptide isoelectric focusing: Comparative study of gel‐based separation approaches

The in‐depth analysis of complex proteome samples requires fractionation of the sample into subsamples prior to LC‐MS/MS in shotgun proteomics experiments. We have established a 3D workflow for shotgun proteomics that relies on protein separation by 1D PAGE, gel fractionation, trypsin digestion, and peptide separation by in‐gel IEF, prior to RP‐HPLC‐MS/MS. Our results show that applying peptide IEF can significantly increase the number of proteins identified from PAGE subfractionation. This method delivers deeper proteome coverage and provides a large degree of flexibility in experimentally approaching highly complex mixtures by still relying on protein separation according to molecular weight in the first dimension.     

Gel absorption-based sample preparation for the analysis of membrane proteome by mass spectrometry

A gel absorption-based sample preparation method for shotgun analysis of membrane proteome has been developed. In this new method, membrane proteins solubilized in a starting buffer containing a high concentration of sodium dodecyl sulfate (SDS) were directly entrapped and immobilized into gel matrix when the membrane protein solution was absorbed by the vacuum-dried polyacrylamide gel. After the detergent and other salts were removed by washing, the proteins were subjected to in-gel digestion and the tryptic peptides were extracted and analyzed by capillary liquid chromatography coupled with tandem mass spectrometry (CapLC-MS/MS). The results showed that the newly developed method not only avoided the protein loss and the adverse protein modifications during gel embedment but also improved the subsequent in-gel digestion and the recovery of tryptic peptides, particularly the hydrophobic peptides, thereby facilitating the identification of membrane proteins, especially the integral membrane proteins. Compared with the conventional tube-gel digestion method, the newly developed method increased the numbers of identified membrane proteins and integral membrane proteins by 25.0% and 30.2%, respectively, demonstrating that the method is of broad practicability in gel-based shotgun analysis of membrane proteome.     

Purification of membrane proteins in SDS and subsequent renaturation

A prerequisite for the purification of any protein to homogeneity is that the protein is not non-specifically associated with other proteins especially during the final stage(s) of the fractionation procedure. This requirement is not so often fulfilled when nonionic detergents (for instance Triton X-100) are used for solubilization of membrane proteins. The reason is that these detergents are not efficient enough to prevent the protein of interest from forming aggregates with other proteins upon contact with chromatographic or electrophoretic supporting media, which, due to their polymeric nature, have a tendency to induce aggregation of other polymers, for instance, hydrophobic proteins. The aggregation can be avoided if sodium dodecyl sulfate (SDS) is employed as detergent. We therefore suggest that membrane proteins should be purified by conventional methods in the presence of SDS and that the purified proteins, which are in a denatured state, are allowed to renature. There is good change to renature internal membrane proteins since they should not be so susceptible to denaturation by detergents as are water-soluble proteins because the natural milieu of the former proteins is lipids which in fact are detergents. In this paper we present a renaturation method based on the removal of SDS by addition of a large excess of G 3707, a nonionic detergent. By this technique we have renatured a 5'-nucleotidase from Acholeplasma laidlawii and a neuraminidase from influenza virus. The enzyme activities were higher (up to 6-fold) after the removal of SDS than prior to the addition of SDS.     

Quantitative Assessment of In-solution Digestion Efficiency Identifies Optimal Protocols for Unbiased Protein Analysis

The majority of mass spectrometry-based protein quantification studies uses peptide-centric analytical methods and thus strongly relies on efficient and unbiased protein digestion protocols for sample preparation. We present a novel objective approach to assess protein digestion efficiency using a combination of qualitative and quantitative liquid chromatography-tandem MS methods and statistical data analysis. In contrast to previous studies we employed both standard qualitative as well as data-independent quantitative workflows to systematically assess trypsin digestion efficiency and bias using mitochondrial protein fractions. We evaluated nine trypsin-based digestion protocols, based on standard in-solution or on spin filter-aided digestion, including new optimized protocols. We investigated various reagents for protein solubilization and denaturation (dodecyl sulfate, deoxycholate, urea), several trypsin digestion conditions (buffer, RapiGest, deoxycholate, urea), and two methods for removal of detergents before analysis of peptides (acid precipitation or phase separation with ethyl acetate). Our data-independent quantitative liquid chromatography-tandem MS workflow quantified over 3700 distinct peptides with 96% completeness between all protocols and replicates, with an average 40% protein sequence coverage and an average of 11 peptides identified per protein. Systematic quantitative and statistical analysis of physicochemical parameters demonstrated that deoxycholate-assisted in-solution digestion combined with phase transfer allows for efficient, unbiased generation and recovery of peptides from all protein classes, including membrane proteins. This deoxycholate-assisted protocol was also optimal for spin filter-aided digestions as compared with existing methods.MS-based proteomics is an indispensable technology for the characterization of complex biological systems, including relative or absolute protein expression levels and protein post-translational modifications. The most popular method for analyzing medium to high complexity protein samples in large-scale proteomics relies on protein digestion by using the endoprotease trypsin. Analysis and sequencing of tryptic peptides by liquid chromatography-tandem MS (LC-MS/MS)¹ then enables identification and determination of protein expression levels based on the peptide ion abundance level or the (fragment) ion intensities of identified peptides. This peptide-centric approach thus strongly relies on efficient, unbiased and reproducible protein digestion protocols. Efficiency is required to maximize the number of detectable peptides per protein (coverage) to distinguish unique proteins within protein families with similar sequences and/or sequence variants, and to detect post-translational modifications. Unbiased generation of peptides is required for the resulting data set to most accurately reflect the relative (stoichiometry) and absolute protein abundance in a sample. A particular protocol should be unbiased with respect to abundance, molecular weight, hydrophobicity and protein class. Membrane proteins for example are often suspected to be underrepresented. For MS-based proteomics approaches several critical steps can be distinguished: (a) disruption and solubilization of cells and protein complexes, (b) protein denaturation and enzymatic proteolysis, (c) MS-compatible peptide recovery, which normally entails removal of reagent leftovers and desalting before MS analysis, (d) adequate peptide separation (achieved by liquid chromatography), and (e) MS peptide analysis and sequencing (MS/MS), including the chosen data acquisition strategy.Comparative evaluations of digestion protocols generally consist of qualitative studies using standard tandem mass spectrometry. These approaches may reveal efficiency (i.e. more identifications), but are unable to reveal digestion protocol induced bias with respect to peptide and protein abundance, including membrane proteins. In addition, most data-dependent acquisition workflows are intrinsically biased, which is detrimental for making comparisons. The aim of the present study was to systematically assess efficiency and bias of trypsin-based protocols applying both standard qualitative and label-free quantitative MS approaches.The in-gel digestion protocol for proteomics, established over 15 years ago (1), has been the cornerstone method affording robust protein identifications from many sample types. Although sodium dodecyl sulfate (SDS) interferes with trypsin digestion and hampers LC-MS analysis, this powerful detergent can still be used to achieve complete protein solubilization as gel-separation is an effective way to remove interfering substances. Gel-based approaches are however not optimal for protein samples of increasing complexity and dynamic range (2). Inherent and practical limitations include, for example, concentration-dependent, incomplete peptide recovery and error-prone handling procedures (3–6). This hampers throughput, reproducibility and unbiased protein analysis, which in recent years has prompted a shift toward the application and optimization of in-solution digestion procedures.Previous comparative studies revealed that for in-solution digestions, the acid labile and MS-compatible detergent RapiGest performed most favorably compared with buffer only, urea, other detergents and organic solvents (7–9). Sodium deoxycholate (SDC), naturally found in mammalian bile (10), has emerged as a cheaper MS-compatible detergent for in-solution digestion (11). Unlike other detergents, SDC was found to enhance trypsin activity almost fivefold at a concentration of 1% (12). Like RapiGest, SDC can also be removed by acidification, but potentially without detrimental peptide loss if a phase separation protocol involving organic solvent is applied (12).An alternative strategy is to perform protein digestion on spin filter devices, introduced a few years ago by Manza and co-workers (13), and further developed by Wisniewski et al. (14). This approach allows the use of SDS to first achieve complete protein solubilization followed by removal of the detergent through repeated washes with urea (14). This is an effective way to remove interfering chemicals and small molecules after protein solubilization, and before digestion, without substantial sample loss. Although this protocol is touted to be a highly effective and universal method for any type of sample, digestion is performed using urea or buffer only and has so far not been evaluated in combination with detergents such as SDC.For our comparative study we selected protocols and methods based on spin filter-aided and standard in-solution digestion that were previously reported optimal and we also report novel optimized protocols. We investigated several experimental parameters including reagents for protein solubilization and denaturation (SDS, SDC, urea), spin filter aided removal of SDS before digestion (urea, SDC, buffer), trypsin digestion conditions (buffer, RapiGest, SDC, urea), and methods for removal of detergents before analysis of peptides (acid precipitation or phase separation with ethyl acetate).Mitochondria are organelles carrying out key metabolic processes fundamental for cellular function (15). The mitochondrial proteome is predicted to contain up to a thousand proteins (16) and is very heterogeneous with a wide range of protein pI, molecular weight and hydrophobicity values (17). We selected mitochondrial preparations to serve as model sample of medium complexity, containing a favorable combination of peptide and protein classes, including soluble and insoluble membrane-anchored or integral proteins.Using standard qualitative as well as data-independent quantitative LC-MS/MS workflows we demonstrate that SDC-based protocols combined with phase separation are the most optimal for both in-solution and filter-aided tryptic digestion, yielding the highest efficiency and lowest bias. This workflow enabled quantitative and objective assessment of various protein digestion conditions, identifying optimal protocols for efficient and unbiased protein analysis.