Mass Spec Analysis Technical Resources

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1. Use of high pH reversed-phase peptide fractionation to analyze proteins of medium to low abundance in complex mixtures
This whitepaper describes how Thermo Scientific Pierce High pH Reversed-Phase Peptide Fractionation Kit utilizes reversed-phase chromatography at high pH to separate peptides by hydrophobicity, providing excellent orthogonality to low pH reversed-phase LC/MS gradient. This simple, highly reproducible protocol enables deep proteome sequencing.
 

2. Efficient and convenient enrichment of multi-spanning membrane proteins for proteomic studies
This whitepaper describes how the Thermo Scientific Mem-PER Plus Membrane Protein Extraction Kit enables enrichment of integral and membrane associated proteins from cultured cells using a simple, reagent-based procedure and a benchtop microcentrifuge. This kit utilizes a simple, sequential detergent extraction method to isolate specific multi-spanning membrane proteins (≥2 transmembrane domains) for downstream analysis by mass spectrometry.
 

3. Subcellular protein fractionation to enhance proteomic coverage of cultured cells
This whitepaper describes how the Thermo Scientific Subcellular Protein Fractionation Kit enables stepwise lysis of cells into functional cytoplasmic, membrane, nuclear-soluble, chromatin bound, and cytoskeletal protein fractions using low speed microcentrifugation in less time than traditional methods and with high reproducibility, for optimized downstream analysis by mass spectrometry.
 

4. Sampling the human population—ultra-highthroughput plasma protein profiling (uHTPPP) sample preparation for translational proteomics
This application note presents a robust, precise, and reproducible high-throughput sample preparation workflow for mass spectrometry-based large population cohort studies. This workflow uses an automated script to process plasma or serum using an EasyPep 96 MS sample prep kit on the Hamilton liquid handling robotic platform as well as proper QA/QC operations which can be further customized by incorporating Thermo Scientific Tandem Mass Tag labeling and peptide fractionation into the workflow.
 

5. Sequential enrichment using metal oxide affinity chromatography (SMOAC) to enhance phosphoproteome coverage for quantitative proteomic analysis
This application note describes how to enable deep phosphoproteome sequencing. A novel approach called SMOAC was developed, in which phosphopeptides are sequentially enriched using Thermo Scientific High-Select TiO2 and Fe-NTA Phosphopeptide Enrichment Kits followed by fractionation using the Pierce High pH Reversed-Phase Fractionation Kit which reduced the overall complexity of the samples and provided greater coverage of the phosphoproteome for both unlabeled and TMT-labeled peptides. The methods describe in this application note provide a powerful tool for large-scale phosphoproteomic analysis.
 

6. Rapid and efficient sample preparation using EasyPep technology: Optimized sample preparation for MS-based proteomics applications
This application note highlights the EasyPep sample preparation technology which enables rapid and efficient processing of different samples, scales, and throughput for mass spectrometry–based proteomics. By utilizing a standardized workflow, identical peptide and protein identification rates, quality, and reproducibility can be achieved with several sample types, including cells (mammalian, yeast, and E. coli), tissues (fresh and FFPE), and plasma, as well as for downstream applications including TMT reagent labeling, high-pH reversed phase fractionation, and phosphopeptide enrichment.

1. Next-generation TMTpro reagents for increased sample multiplexing
This application note discusses unique advantages of higher sample multiplexing using TMTpro reagents, impacts of tag structure on chromatography, use of TMTpro reagents with the latest-generation Orbitrap Eclipse Tribrid mass spectrometer equipped with the FAIMS Pro and optimizing instrument settings and data analysis in Proteome Discoverer software interface to enable deep proteomic analysis of samples with the highest accuracy and precision for protein quantitation.

Q. What is the difference between Relative and Absolute Quantitation? Which type of quantitation is used for proteomic samples?
A. Relative quantitation compares the amount of an analyte in different samples. Absolute quantitation measures the actual amount of an analyte in a sample using a known standard at different concentrations. Label-free, SILAC and TMT workflows are used for relative quantitation of protein samples. Heavy (i.e. AQUA) peptides are used as standard for absolute quantitation of target proteins.

Q. What is the main purpose of Tandem Mass Tag (TMT) reagents?
A. Tandem mass tag technology is used individually label different protein samples so they can be combined into a single sample for LC-MS analysis. The major advantages of this workflow are higher sample throughput, less instrument analysis time, high precision of peptide quantitation among replicates, and fewer missing quantified proteins among different samples.

Q. Can TMT reagents be used in combination with TMTpro reagents?
A. TMT and TMTpro reagents are different chemicals with different masses. Therefore, we do not recommend mixing the two tags as the combined samples are significantly more complex resulting in fewer quantified proteins.

Q. What type of mass spectrometer can I use for Tandem Mass Tag (TMT) analysis?
A. We recommend using high resolution Orbitrap Tribrid (e.g. Fusion, Lumos, Eclipse), Orbitrap Exploris (e.g. 240, 480) or Q Exactive (e.g. Plus, HF, HFX) series of instruments.

Q. What sample types can be used with different quantitation systems?
A. Label-free, TMT reagents and TMTpro reagents can be used with any protein sample type. SILAC and Neucode can only be used with cell lines & model organisms that can be metabolically labeled.

Q. How many samples can be combined for simultaneous, multiplexed proteomic analysis?
A. SILAC can be used to multiplex 2 to 3 samples. TMT reagents can be used to combine 6 to 11 samples. TMTpro reagents can be used to measure up to 16 samples simultaneously.

Q. Is it necessary to remove free TMT tags after peptides are labeled? If so, which method is recommended?
A. Excess unreacted or quenched TMT tags in peptide samples can interfere with colorimetric-based peptide assays, LC-MS chromatography and affect peptide/protein identifications during LC-MS. In-line LC trapping columns can be used to remove some excess reagent. The Peptide Clean Up Columns in the EasyPep kits can also be used to remove the excess TMT tags. The Pierce Peptide Desalting Spin Columns (Cat. No. 89852) can used to clean up samples labeled using alternative sample preparation methods.

Q. In the EasyPep sample preparation workflow, it is mentioned that TMT can be added to peptides before or after clean-up step. Are there any differences between these two methods, regarding the labeling efficiency or relative quantitation?
A. The EasyPep kit chemistry is fully compatible with the TMT reagents. Most commonly, TMT reagents are used to label peptides immediately after digestion and before peptide clean up. This enables combining samples for a single cleanup step which reduces variability for more reproducible quantitative measurements. Another option is to label samples after peptide cleanup. This approach provides efficient labeling of the samples but may require the use of a peptide quantification assay to ensure samples are equally mixed before LC-MS analysis. Both the workflows should provide high labelling efficiencies using our recommended TMT to sample ratios.

Q. How do I bridge the data from different samples sets for TMT-based quantitation?
A. This is most often done with the utilization of a "pool channel" for each multiplex set. This reference channel typically contains an equimolar mixture of all samples which is labeled with one of the TMT tags that is shared among the sets. The pool channel can be used to then normalize relative protein abundance measurements across multiplexed sets.

Q. Do I have to apply correction factors for analyzing data for TMT labeled samples?
A. Applying correction factors is required for more accurate relative protein quantitation as the isotopic impurity is variable among the tags, ranging from 5-10%.

Q. I have a lower number of peptide/protein identification in my combined TMT-labeled samples compared to the unlabeled sample—what can I do?
A. We recommend fractionating the sample using the Pierce High pH Reversed-Phase Peptide Fractionation Kit (Cat. No. 84868) to reduce the sample complexity before LC-MS analysis which increases the number of quantifiable peptides/proteins for multiplexed samples.

Q. I have low levels (<95%) of TMT labeling for my samples. What should I check?
A. Labeling efficiency can be determined for individual labeled samples by searching peptides using TMT/TMTpro as a variable (i.e. dynamic) modification peptide amino terminus and lysines. The ratio of peptide to tag by mass (w:w) should be 1:4 to 1:8 for complete labeling of most samples. TMT/TMTpro reagents can also be hydrolyzed rendering them less reactive if handled or stored improperly.

Q. I have low incorporation of heavy amino acids in my SILAC proteins. What should I check?
A. Verify that SILAC media without light lysine (and/or arginine) and dialyzed FBS was used for cell culture. Also, verify that cultured cells are healthy, viable and actively growing in SILAC media.

Q. What MS software can I use for protein & peptide quantitation?
A. Proteome Discoverer, MaxQuant, Skyline (for targeted analysis) can be used for protein & peptide quantitation.

Q. I have good peptide identification numbers but the variation between sample replicates is high. What do you recommend to improve sample reproducibility?
A. Review your sample-prep workflow to ensure consistent protein extraction, reduction/alkylation, digestion and clean up. We recommend using EasyPep products (Cat. No. A40006) for high quality, reproducible sample preparation. We also recommend quantifying peptides using the Pierce Quantitative Fluorometric Peptide Assay (Cat. No. 23290) or Pierce Quantitative Colorimetric Peptide Assay (Cat. No. 23290) to ensure the same peptide amounts are analyzed for each LC-MS analysis. Poor reproducibility could also be related to the LC-MS system performance which may require recalibration using Pierce Calibration Solutions. System performance can be assessed using stand protein digests such as Pierce HeLa Protein Digest Standard (Cat. No. 88328) or Pierce TMT11plex Yeast Digest Standard (Cat. No. A40938) and peptide standards such as the Pierce Peptide Retention Time Calibration Mixture (Cat. No. 88321) or Pierce LC-MS/MS System Suitability Standard (7 x 5 Mix) (Cat. No. A40010).

Q. Do Thermo Scientific calibration solutions work on other vendor MS instruments?
A. Thermo Scientific calibration solutions are designed specifically for Thermo Scientific MS instruments. Check with your MS instrument vendor on recommended solutions for mass calibration.

Q. Which calibration solution is best for each Thermo Scientific instrument?
A. Different calibration solutions are recommended for different Thermo Scientific MS instruments and applications. A chart and description of each calibration solution and its recommended usage can be found here.

Q. Are MS calibration solutions stable at room temperature or stored in other containers?
A. MS calibration solutions should be stored at the storage temperature on the MS calibration datasheet or label for maximal shelf life. Calibration solutions must be kept in the original container to maintain solution stability and purity. It is not recommended to calibrate MS instruments using expired calibration solution.

Q. Which standards are recommended for determining LC-MS system suitability?
A.Pierce Peptide Retention Time Calibration Mixture (Cat. No. 88321) is recommended for LC system and gradient optimization. Pierce BSA Digest, equimolar, LC-MS grade (Cat. No. 88342), Pierce 6 Protein Digest, equimolar, LC-MS grade (Cat. No. 88342) and Pierce HeLa Protein Digest Standard (Cat. No. 88328) can be used for "bottom up" LC-MS method optimization and LC-MS suitability. Pierce LC-MS/MS System Suitability Standard (7 x 5 Mix) is best suited to assess the gradient, sensitivity and dynamic range of LC-MS system for discovery and targeted peptide quantitative workflows. Pierce TMT11plex Yeast Digest Standard (Cat. No. A40938) should be used to assess system performance for TMT-based quantitative workflows. Pierce 6 Protein Intact Mix (Cat. No. A33527) is recommended for "top down" LC-MS method optimization and LC-MS suitability.

Q. How do I assess my LC-MS system performance?
A. The best way to assess the performance of an entire LCMS system is using a calibration solution (MS), peptide retention time standards (Pierce Retention Time Calibration Mixture (PRTC) and Pierce LC-MS/MS System Suitability Standard (7x5 mix)), and a known standard sample (e.g. HeLa digest, BSA digest, 6 proteins digest) specific to your application. It is recommended to frequently assess LC-MS system performance on a regular basis to compare results over time.

Q. What's the difference between Pierce Retention Time Calibration Mixture (PRTC) and Pierce LC-MS/MS System Suitability Standard (7x5 mix)?
A. PRTC is a mixture of 15 peptides with varying hydrophobicity used to determine the quality of nano and capillary flow LC chromatography and column performance. The 7x5 standard contains 7 of 15 PRTC peptides with five isotopologue versions of each peptide containing no heavy, 1 heavy, 2 heavy, 3 heavy, and 4 heavy isotope-labeled amino acids. The isotopologues for each peptide are provided at a distinct concentration to generate seven standard curves to assess system performance. In addition to evaluating of LC chromatography, this standard is also used to assess the dynamic range and sensitivity of the LC-MS system.

Q. How do you verify the LC-MS system after installation of a new column for proteomic applications?
A. Several technical replicate injections of a suitable standard sample (e.g. Pierce HeLa Protein Digest Standard, Pierce BSA Digest Standard, or Pierce 6 Mix Protein Digest Standard) should be used to equilibrate and compare the new column to reference chromatograms.

Q. How do you analyze the data for Pierce Retention Time Calibration Mixture (PRTC) and Pierce LC-MS/MS System Suitability Standard (7x5 mix) standard?
A. Create a new Skyline document containing 15 PRTC heavy isotope-labeled peptides. The Skyline document can used for the DDA data analysis using MS1 filter or for the PRM/SRM data analysis with targeted MS filters. The skyline document for 7x5 standard is available as a web download document. Please refer 7x5 standard label for detailed information. Alternatively, the 7x5 standard data can be analyzed in Chromeleon software using the pre-built report template (web download document).

Q. What is the Pierce Reserpine Standard used for?
A. The Thermo Scientific Pierce Reserpine Standard for LCMS is a standard for determining the performance and sensitivy of all Thermo Scientific MS instruments. This standard must be diluted before use according to the procedure specified in the corresponding instrument Getting Started Guide.

Q. Which syringe(s) do you recommend for use with your calibration solutions?
A. We recommend using a 500 µl Hamilton Syringe (Prod#81265) for direct infusion of calibration solutions. The syringe and tubing can be cleaned with LC-MS grade 50% MeOH/50% water between uses and should be stored dry. Avoid storing the syringe or tubing with the calibration solution.

Q. How do I use the Pierce TMT11plex Yeast Digest Standard to optimize my LC-MS acquisition parameters?
A. The Pierce TMT11plex Yeast Digest Standard (i.e. TKO standard) can be used to rapidly and accurately assess ion interference (co-isolation of multiple analytes of similar mass-to-charge) for TMT-based quantitative mass spectrometry-based proteomic analysis.

Q. I have a low number of protein identifications from my protein digest sample—what can I do?
A. Check your MS system performance using our Pierce HeLa Protein Digest Standard (Cat. No. 88328). Analysis of this standard can help determine if the problem is from the sample preparation or the LC-MS system.

Q. I observe peptide retention time shifts in my LC chromatogram—do you have a standard I can use to check my LC performance?
A. Yes, the Pierce Peptide Retention Time Calibration Mixture (Cat. No. 88321) or Pierce LC-MS/MS System Suitability Standard (7x5 mix) offers synthetic heavy peptides to diagnose and troubleshoot your LC system components/attributes (pump, flow meter, TRAP column, separation column and gradient).

Q. A database search of my peptide samples shows low and/or no identifications, but I have excellent signal in my chromatograph. What might be the problem?
A. Your instrument may require calibration. Recalibrate using one of our Pierce Calibration Solutions. Verify correct search parameters were used for database searching (e.g. species, enzyme, fragment ions, mass tolerance, etc.)

Q. Is there a way to evaluate the sensitivity of my triple quad MS instrument?
A. Yes, Pierce Reserpine Standard for LC-MS (Cat. No. 88326) can be used to assess instrument sensitivity.

Q. After running Pierce TMT11plex Yeast Digest Standard (Cat. No. A40938), I observe low signal to noise and poor quantitative accuracy for the TMT reporter ions. What do you recommend?
A. Clean and recalibrate MS instrument using one of our Pierce Calibration Solutions. Verify settings for LC acquisition methods.

Q. I have extra peaks in my calibration spectra. Is the calibration solution contaminated?
A. The extra peaks during MS calibration may be due to contaminated ion transfer tube, ion source, syringe, peak tubing or calibration solution. The ion transfer tube and syringe can be cleaned using sonication in 50% LC-MS grade MeOH/Water. All calibration solutions are tested for purity; however, contamination can be introduced through a contaminated syringe or sorting the calibration solution in a different container.

Q. After LC-MS analysis using the Pierce Retention Time Calibration Mixture (PRTC) or Pierce LC-MS/MS System Suitability Standard (7x5 mix), the chromatography is drifting over time. How do I fix the chromatography so that it is more reproducible?
A. Purge LC system and check the flow meter calibration. Check mobile phase solvents and LC gradient. Evaluate and replace column if needed. Rerun PRTC/7x5 mixture to verify chromatography is consistent in replicate runs.

Q. What should I do if the heavy labeled peptide doesn't co-elute with the endogenous peptide?
A. Verify the relative fragment ion intensities for the heavy and endogenous peptide are similar. If there is no issue with the relative fragment ion profile, then there may be an issue with the heavy peptide sequence or purity. Reorder the heavy peptide or select a new unique peptide for the target of interest.

Q. What kind of substances interfere with mass spectrometry analysis?
A. Almost everything! Salts, buffers, detergents and other small molecules can all interfere with liquid chromatography separation and mass spectrometer source ionization.

Q. What is the best way to remove small contaminants?
A. For protein samples, Pierce Polyacrylamide Spin Desalting Columns (Cat. No. 89849)can be used to remove salts and other small molecular weight contaminates. Acetone precipitation is recommended for low protein amounts provided you have at least 10 µg sample. For peptide samples derived from protein digests, perform sample clean-up using Pierce Peptide Desalting Spin Columns (Cat. No. 89852) or Pierce C18 Spin Tips (Cat. No. 84850).

Q. Can desalting columns used for protein cleanup be used for peptide samples?
A. No. Protein desalting columns (e.g. Zeba desalting columns) use size exclusion which have a molecular weight cut off that is typically too high for peptide samples. Peptide desalting uses reversed phase chromatography resins (e.g. C18) to bind peptides for salt removal during washing.

Q. What is the best way to remove detergents?
A. Detergents are more easily removed at the protein level using acetone precipitation, dialysis, or the Pierce Detergent Removal Resin.

Q. What kind of buffers and solutions are compatible with peptide clean up and LC-MS?
A. Most organic solvents or volatile buffers are compatible with LC-MS. Water and acetonitrile with 0.1% formic acid is the most common solvent system for separation of peptide samples for MS. Trifluoroacetic acid (TFA) is an alternative ion pairing agent for off-line peptide C18 clean up or LC-UV analysis. LC-MS grade reagents are available 0.1% TFA (Cat. No. 85172), LC-MS water (Cat. No. 85189), acetonitrile (Cat. No. 85188), and TFA (Cat. No. 85183).

Q. If I already removed small contaminants and detergents at protein level, do I still need to clean up my peptides after digestion?
A. Yes! It is recommended to always perform additional clean up after protein digestion to remove and residual salts or partially digested proteins. Use Pierce Peptide Desalting Spin Columns (Cat. No. 89852), Pierce C18 Spin Tips (Cat. No. 84850) or an in-line C18 trap column (Cat. No. 160434).

Q. How do I determine if my sample is compatible and ready for LC-MS?
A. Samples prepared using LC/MS grade reagents are suitable for LC-MS; however, particulates and other small molecules can all interfere with liquid chromatography separation and mass spectrometer source ionization. Visually inspect samples for particulate matter. Use Pierce Peptide Desalting Spin Columns (Cat. No. 89852), Pierce C18 Spin Tips (Cat. No. 84850) or an in-line C18 trap column (Cat. No. 160434) to remove non-volatile salt before MS analysis.

Q. What is the most efficient way remove excess TMT reagent from my samples?
A. Our EasyPep sample prep kits (Cat. No. A40006) have been specifically optimized for this purpose and can remove excess TMT reagents from protein digests prepared using the kits. To remove excess TMT reagent from samples prepared using other sample preparation methods, we recommend Pierce Peptide Desalting Spin Columns (Cat. No. 89852) or Pierce High pH Reversed Phase Fractionation kits.

Q. I desalted my sample using C18, but I lost most of my peptides. What happened?
A. Peptides do not bind well to reversed phase resins at neutral pH or in the presence of organic solvents (e.g. acetonitrile). Ensure no organic solvents are present before and after cleanup (dry down sample on speedvac or equivalent) and acidify protein digest samples using formic acid or trifluoroacetic acid (TFA) to pH <3 before desalting.

Q. Which peptide assay should I use for quantitation of my sample: the Pierce Quantitative Colorimetric Peptide Assay (Cat. No. 23275), or the Pierce Quantitative Fluorometric Peptide Assay (Cat. No. 23290)?
A. The choice of which peptide assay depends on the sample type and composition of the sample buffer. The fluorometric peptide assay cannot be used to measure peptides with chemically modified amines such as acetylated peptides or TMT-labeled protein digests. The colorimetric assay can measure a broad wider range of samples but is not as sensitive as the fluorometric assay, requiring more sample for accurate detection. Finally, both assays are susceptible to interfering compounds in the sample or buffer which should be avoided or removed for best results.

Q. Are the EasyPep MS sample prep kits compatible with IP-MS samples?
A. EasyPep MS sample prep kits can be used to process samples after immunoprecipitation. However, the EasyPep Lysis buffer should not be used for immunoprecipitation as it is contains a denaturing detergent which interferes with antigen binding.

Q. Can I use protease inhibitors for MS sample prep?
A. We don’t recommend adding protease inhibitors as they can affect trypsin and trypsin/Lys-C activity. If protease inhibitors are present in the protein sample or cell, they should be removed by dialysis, diafiltration, desalting or protein precipitation prior to enzymatic digestion. Addition of phosphatase inhibitors to lysis solution is recommended before cell lysis for phosphopeptide enrichment workflows.

Q. Which sample types have been tested with EasyPep MS Sample prep kits?
A. EasyPep MS sample prep kits have been tested with mammalian cell lines (HeLa, A549, HEK293, CHO), fresh/frozen tissues (brain,heart, liver), human plasma and serum, bacteria (E.coli), yeast and FFPE sections.

Q. Can I use different lysis buffer for EasyPep workflow?
A. The EasyPep lysis buffer has been optimized to work with sequential chemistries, digestion and clean up used in the EasyPep MS sample prep kits. Other lysis buffers are not recommended as they may contain buffers or detergents which may interfere with reduction, alkylation, digestion or peptide clean up.

Q. How can I measure peptide yields?
A. Peptide yield can be measured using the Pierce Quantitative Colorimetric Peptide Assay (Cat. No. 23275), or the Pierce Quantitative Fluorometric Peptide Assay (Cat. No.23290).

Q. Should I use protein A/G or streptavidin beads for my affinity purification?
A. Both methods provide robust results for immunoprecipitation. Biotin tagged antibodies with streptavidin beads provide lower background but require more upfront sample prep. Protein A/G beads are robust and straightforward in use but result in higher background than streptavidin beads.

Q. At which step should I label my samples with TMT reagents?
A. TMT labeling can be performed after reduction, alkylation, and digestion, and prior to cleanup, if the sample is buffered at appropriate pH (8-8.5) in a suitable buffer without primary amines (e.g. Tris, glycine). TMT labeling can also be performed after peptide clean-up. Labeling peptides after clean-up enables measuring and normalizing the peptide samples for equal mixing.

Q. How do I measure the digestion efficiency of my sample?
A. Digestion efficiency can be measured assessing the number of missed cleaved peptides in the sample. The Pierce Digestion Indicator for Mass Spectrometry can also be added to the sample to monitor digestion efficiency.

Q. How can I improve the digestion efficiency of my protein sample?
A. Digestion efficiency can be improved by adding more enzyme, incubating for longer digestion times, and ensuring the correct pH for digestion. Avoiding protease inhibitors and strong denaturants is recommended for maximum digestion efficiency for most proteases.

Q. I want to minimize the number of missed cleaved peptides for maximal digestion. Which enzyme should I choose: trypsin only, LysC followed by trypsin or a trypsin-LysC mixture?
A. Trypsin cleaves at both lysine and arginine but has difficult fully digesting peptide sequences if these amino acids are followed by a proline. LysC only cleaves at lysine and can cleave sequences with lysines followed by prolines. Therefore, LysC is often used in conjunction with trypsin to decrease the number of missed cleavages. Sequential digestion with LysC followed by trypsin is the best method to minimize the number of missed cleaved peptides as each enzyme can be used under their optimal conditions. However, trypsin-LysC mixtures can also be used to concurrently digest samples with shorter incubation times (<3hrs).

Q. What software can I use for proteomic data analysis?
A. We recommend Proteome Discoverer for proteomic data analysis.

Q. Where do I find resources for SureQuant kits and workflow?
A. SureQuant Targeted Mass Spec Assay Kits manuals are available online on Thermo website. For instructions to download attachments and additional files, please visit thermofisher.com/surequantdocs and enter the code indicated with the kit label to access instrument method and data analysis information. SureQuant workflow solution is available at www.thermofisher.com/surequant.

Q. What sample prep method is available for low abundant proteins in blood?
A.High Select Top14 Abundant Protein Depletion Mini Spin columns (Cat. No. A36369) can be used to deplete top 14 abundant plasma/serum proteins. The depleted samples can be processed with EasyPep MS sample prep kit to generate digested plasma/serum samples. The clean digested samples can be processed by nanoLC-MS/MS analysis for discovery experiments or nanoLC-PRM/MS analysis for targeted experiments.

Q. Are there MS reagents for membrane proteins?
A.CHAPS (Cat. No. 28299), OTG (Cat. No. 28351) and Sodium Deoxycholate (Cat. No. 89904) are detergents often used to solubilize and extract membrane proteins from samples

Q. How do I process bacterial samples with EasyPep MS Sample Prep kits (Cat. No. A40006)?
A. Add 1-2uL of 50mg/ml lysozyme solution (Cat. No. 90082) to 200uL of the EasyPep lysis buffer provided in the kit and proceed with the sample prep. Add 200ul of the lysis solution 5 X 10^ 8 bacterial cells E. coli cells. (OD600 equal to 1, corresponding to 1 X 10^8 CFUs). Lyse by pipetting the sample repeatedly and centrifuge at 16,000xg for 5 minutes. Alternatively, E. coli cells can be lysed in lysis byffer using bead beating or sonication instead lysozyme. After lysis dilute samples to 1mg/ml using lysis buffer, proceed with the EasyPep protocol for reduction, alkylation, digestion and clean up as described in the product manual.

Q. How do I process yeast samples with EasyPep?
A. Add 0.5mm diameter glass beads to the pelleted sample in EasyPep lysis buffer provided in the EasyPep sample prep kit (Cat. No. A40006) and proceed as recommended in the EasyPep protocol. Vortex vigorously for few minutes and spin at 16,000xg for 5 minutes and extract supernatant for protein assay and further downstream sample preparation.

Q. How do I process formalin-fixed, paraffin-embedded (FFPE) samples for MS analysis?
A. FFPE sections must first be de-paraffinized using xylene and then sequentially washed with EtOH (100%, 95%, 80%) to rehydrate the tissue. Protein extraction is performed using manual homogenization or sonication followed by heating at 95°C for 2 hours to reverse protein crosslinks.

Q. What is a method for reduction, alkylation, and digestion of an antibody?
A. Prepare 1mg/ml antibody in 7.0 M Guanidine HCl, 100 mM Tris @ pH 8.3. For reduction, add DTT (Cat. No. A39255) solution at 10mM final concentration and incubate for 30 minutes at room temperature. For alkylation, add sodium iodoacetate at 20mM final concentration and incubate in dark for 20 minutes at room temperature. Perform buffer exchange followed by digestion using Pierce MS grade Trypsin (Cat. No. 90058) (1:10 enzyme to substrate ratio).

Q. I would like to reduce my sample before digestion. Should I use DTT or TCEP for reducing proteins when alkylating with iodoacetamide?
A. Both DTT and TCEP can be used to reduce disulfide bonds which helps denature proteins for better enzymatic digestion. Typically, 5-10 mM of reducing agent is used for protein reduction. Heating samples (>37°C) can help accelerate reduction reactions. Using 3-5 molar excess (10-30mM) iodoacetamide is recommended for most applications. If DTT is used as the reducing agent, additional DTT may be added after alkylation to quench excess iodoacetamide. This quenching step is not necessary when using TCEP. Finally, reduction and alkylation reactions should be performed at pH 7-9 so using buffered TCEP solutions such as Bond Breaker TCEP solution is recommended for reduction instead of TCEP-HCl.

Q. What is the best enzyme to use for protein digestion?
A. Trypsin (Cat. No. 90058)or Trypsin/LysC (Cat. No. A41007) mix are most commonly used for proteomic applications in order to assure reproducibility and complete digestion. Other commonly used enzymes for purified protein characterization and unique applications include Chymotrypsin (Cat. No. 90056), Pepsin (Cat. N. 20343), LysN (Cat. No. 90300), AspN (Cat. No. 90053), GluC (Cat. No. 90054).

Q. When I perform quantitation of my peptide sample using the Pierce Quantitative Colorimetric Peptide Assay (Cat. No. 23275), I get different results than when I use the Pierce Quantitative Fluorometric Peptide Assay (Cat. No. 23290). Which one is best to use for the most accurate quantitation?
A. Since the different the peptide assays use different chemistries to measure peptides, they may result in different results. Interfering compounds are the most common source of background and inaccurate measurements. The fluorometric peptide assay is not recommended for peptides which have been modified using TMT reagents.

Q. After cell lysis, my sample is very viscous and difficult to pipette. How do I reduce the sample viscosity?
A. High sample viscosity after lysis is due to release of DNA from the nucleus. Sonication or addition of a nuclease such as the Pierce Universal Nuclease (Cat. No. 88700) can be used to breakdown DNA and reduce sample viscocity.

Q. I want to enrich my sample for phosphopeptides. Should I use the High-Select TiO₂ Phosphopeptide Enrichment Kit (Cat. No. A32993), High-Select Fe-NTA Phosphopeptide Enrichment Kit (Cat. No. A32992), or both?
A. Each kit has different selectivity for different phosphopeptides. The High-Select TiO₂ Phosphopeptide Enrichment Kit enriches more multiply phosphorylated peptides. The High-Select Fe-NTA Phosphopeptide Enrichment Kit enriches more monophosphorylated peptides. In addition, the High-Select Fe-NTA Phosphopeptide Enrichment Kit has slightly higher phosphopeptides specificity and yield than the TiO₂ kit. For the best enrichment for all types of phosphorylated peptides, we recommend our SMOAC method, which utilizes both kits sequentially.

Q. When I use the SMOAC method, there is a small pellet of non-volatile material remaining in the wash/flow through sample from the TiO2 enrichment after drying. Is this normal?
A. Yes, the buffer used for TiO2 column binding contains a non-volatile acid which appears in the sample tube upon drying. The residual material does not interfere with subsequent Fe-NTA enrichment and is removed during before LC-MS analysis.

Q. I want to fractionate a complex sample, but my sample is different from the sample described in the instruction booklet for the Pierce High pH Reversed-Phase Peptide Fractionation Kit (Cat. No. 84868). Should I use a customized fractionation gradient?
A. Recommended gradients are available for typtic protein digests samples with or without TMT labeling. Use of other chemical labels, performing selective peptide enrichment, or using a different digestion enzyme other than trypsin may affect the elution profile of the peptides during fractionation. For best sample peptide coverage, an ideal elution profile would result in relatively equal amounts of peptides in each fraction.

Q. I only have 200ug of protein digest sample. Is this enough for Fe-NTA enrichment?
A. We recommend using at least 0.5mg-3mg of clean (i.e. C18 desalted) and dried protein digest for optimal results, with anticipated yield of 1-3%. Using lower amounts of protein digest sample is possible; however, there will be a significantly less phosphopeptide yield with potentially more non-phosphopeptides present after enrichment and extra care must be taken to not lose the small amount of sample remaining.

Q. Is C18 clean up required before phosphopeptide enrichment?
A. Yes! C18 is necessary to remove small molecules that may interfere with phosphopeptide enrichment. Drying after clean-up is also necessary to concentrate the dilute samples and remove C18 organic eluting solvent so samples can be dissolved in the optimized buffers for phosphopeptide binding. Alternative binding buffers are not recommended as phosphopeptide yield and specificity will be affected. For maximal performance, ensure the clean, dried peptide samples are completely solubilized in binding buffer with agitation for at least 10 min.

Q. How much starting material is needed to fractionate my samples using the Pierce High pH Reversed Phase Fractionation kit?
A. A minimum of 100ug of protein digest is recommend for fractionation using eight fractions. Less sample can be used if fewer fractions are used.

Q. After performing phosphopeptide enrichment using the High-Select Fe-NTA Phosphopeptide Enrichment Kit, I notice a yellow/slight brownish color material in my eluate or after drying my samples. Is this normal?
A. The yellow/slight brownish color is indicative of iron which has been released from the column during elution along with the phosphopeptides. After acidification of the elution buffer or resuspension of dried peptides in 0.1% formic acid, the material can be removed by centrifugation of the sample at 16,000xg spin for 30sec and transferring the supernatant into a new container for LC/MS injection or peptide quantification. Alternatively, the material can be removed using a Peptide Desalting column or C18 tip.

Q. I have poor phosphopeptide enrichment specificity using either the High-Select TiO2 Phosphopeptide Enrichment Kit (Cat. No. A32993) or the High-Select Fe-NTA Phosphopeptide Enrichment Kit (Cat. No. A329929). What should I do?
A. Check the expiration date in order to assure that all components are still good. Keep TiO2 tips protected from light during storage. Make sure you are using the correct buffer component for resuspension of your sample, and that the sample is very thoroughly dried and resuspended well. When incubating peptide samples with the Fe-NTA Kit, ensure that you avoid end-over-end rotation or vortexing; instead gently tap the column to disperse the resin. When removing plugs from columns, be sure to avoid pushing liquid back into the column from the plug reservoir. After elution, do not store the phosphopeptides in the elution buffer but immediately proceed to dry the sample before storing at -80°C until mass spec analysis.

Q. How do I chose between High Select HSA/Immunoglobulin Depletion Resin and High Select Top14 Abundant Protein Depletion Resin?
A. Most customers use the Top14 resin to remove more abundant proteins. Customers who use the Top 2 may be interested in this product it is deplete albumin and IgG which may be sufficient to identify specific proteins of interest which are less abundant in plasma.

Q. Which proteins removed by High Select Top 14 Abundant Protein Depletion Resin?
A. The High Select Top14 Abundant Protein Depletion Resin uses immobilized antibodies to remove human serum albumin (HSA), albumin, IgG, IgA, IgM, IgD, IgE, kappa and lambda light chains, alpha-1-acidglycoprotein, alpha-1-antitrypsin, alpha-2-macroglobulin, apolipoprotein A1, fibrinogen, haptoglobin, and transferrin from serum, plasma, or spinal fluids.

Q. I have a lot of plasma samples to prepare for MS analysis. Is there a way to perform depletion in a 96 well plate?
A. The bulk Top14 depletion resin is compatible with a 96 well filter plate such as the Agilent filter microplate (Cat. No. 200957-100). Add 600µL of resin slurry (50%) to each of the wells in the 96-well filter plate, add 10-20µL of sample, and incubate with gentle shaking. Depleted samples can be collected in a new 96 well polypropylene plate by centrifugation for 2 minutes at 100xg or using a vacuum manifold.

Q. How do you assess plasma abundant protein depletion efficiency and reproducibility?
A. ELISA and MS analysis are often used for evaluation of depletion performance. Protein yields after depletion level can be measured using the BCA assay.

Q. What is the percentage of albumin removed using abundant protein depletion columns?
A. More than 99% of albumin can be depleted when columns use the recommended sample to resin ratio.

Q. What is the maximum loading capacity for the High Select Depletion Columns?
A. Mini columns can deplete up to 10uL (600ug) of plasma or serum; midi columns can deplete up to 100uL (6000ug) of plasma or serum

Q. Can I use the High Select HSA/Immunoglobulin Depletion Resin to process mouse samples?
A. No. The depletion resin contains immobilized antibodies that only recognize human plasma proteins.

Q. Can the High Select HSA/Immunoglobulin Depletion Resin be reused?
A. We don’t recommend reusing the depletion resin as depletion efficiency may decrease after regeneration and subsequent depleted samples may be contaminated from prior ones.

Q. How do I proceed to downstream mass spec sample preparation after abundant plasma protein depletion?
A. After the depletion, samples are significantly diluted in buffer. Depleted samples should be concentrated by speed vac drying, solvent precipitation or diafiltration. Samples dried using a speedvac are directly compatible with the EasyPep kit chemistry after resuspension using the lysis buffer.

Q. After depleting, I still see albumin and other abundant proteins after LC-MS analysis. Did the depletion work?
A. Depletion does not remove all abundant proteins in the sample. Even with 99% depletion of a specific plasma protein, highly abundant proteins will still be the most abundant proteins in the sample detected by MS analysis. However, depletion significantly decreases the most abundant plasma proteins enabling identification of lower abundant proteins.

Q. How do I determined if a crosslinkers is cell permeable?
A. Hydrophobic crosslinkers with a high logP are better able to cross membranes than hydrophilic ones. In addition, crosslinkers which have a charged functional group (e.g. sulfo-NHS) are typically not cell permeable. Crosslinking of cytosolic or nuclear marker proteins can be used to assess crosslinker cellular permeability.

Q. What solvent is recommended crosslinker reconstitution?
A.DMSO (Cat. No. 85190) is the recommended solvent for most crosslinkers. Sulfonated crosslinkers can be reconstituted using DMSO or water.

Q. How do I remove excess crosslinker from my protein sample?
A. Diafiltration using a Protein Concentrator (Cat. No. 88514) or dialysis are common methods for removing excess crosslinker.

Q. What is the advantage of a crosslinker with a long PEG linker?
A. PEG linkers increase crosslinker solubility and the solubility of crosslinked protein complexes. However, these crosslinkers may not be membrane permeable and may interfere with proteolytic digestion. Amine-reactive crosslinkers with long PEG spacers include BS(PEG)5 (Cat. No. 21581) and BS(PEG)9 (Cat. No. 21582), but crosslinkers with alternate reactivity are also available with long PEG spacers.

Q. Are there crosslinking reagents available for non-lysine crosslinking?
A. There are a variety of crosslinkers available which react to many different amino acid functional groups including amine, sulfhydryl, carboxyl, carbonyl and non-specific (i.e. photo-reactive) crosslinkers. See more information here.

Q.DSBU (Cat. No. A35459) and DSSO (Cat. No. A33545) are both MS-cleavable crosslinkers. Which one should I choose?
A. Although the solubility and reactive of both crosslinkers is similar, DSBU has a longer linker than DSSO which may result in different crosslinks depending on the protein of interest. Also, the recommended collision energies for each crosslinker is different as DSBU requires a lower fragmentation energy for cleavage. Therefore, we recommend DSSO for MS2-MS3 methods, and DSBU for MS2 methods.

Q. There are multiple crosslinkers with different lengths. Which one should I chose?
A. The length of the crosslinker determines which crosslinks are possible and provides a distance constraint for protein structure determination. Some crosslinks are less rigid in structure than others, resulting in a wider variety of possible crosslinks. Crosslinker length should be chosen based on the availability of reactive amino acids and their positions in the structure of the proteins of interest. Multiple crosslinkers are likely to give complementary information and better elucidate protein structure.

Q. How do I separate mono-links (dead-end crosslink-modified peptides) from crosslinks?
A. Fractionation techniques are the most common method for separation of crosslinks from mono-links; strong cation exchange and size exclusion chromatography are often successful.

Q. How do I confirm that my protein sample is crosslinked?
A. Crosslinking is commonly confirmed on a protein of interest via SDS-PAGE. The crosslinked protein molecular weight should be larger as indicated by reduced gel mobility after crosslinking when compared to an unmodified control sample.

Q. Most crosslinkers are homobifunctional with the same reactivity at each end. What are the reasons to choose a heterobifunctional crosslinkers? 
A. Heterobifunctional crosslinkers have different reactive groups for protein conjugation and crosslinking. The different reactive groups can yield different crosslinks compared to homobifunctional crosslinkers. In addition, some heterobifunctional crosslinkers use chemistries which are activatable significantly increasing the chance of conjugation between proteins and minimizing intra-protein crosslinks.

Q. I do not observe any crosslinked proteins after adding my crosslinker. What went wrong?
A. Ensure that the buffer used for crosslinking is compatible with the crosslinker and is at the recommended pH for optimal reactivity. Titrate the crosslinker to determine the optimal molar excess of reagent to protein. Avoid amine-containing buffers for NHS-based crosslinking reactions. Depending on the protein, you may need to try a crosslinker with a different length or reactivity to be successful.

Q. I can see that my protein crosslinked by SDS-PAGE, but I don’t see many crosslinks in MS analysis. What do you recommend for improving my results?
A. First, verify that the correct crosslinker and modifications are chosen for database search software. Also, check your MS acquisition methods have the correct crosslinker mass for MS1 selection and MS/MS fragmentation. Lastly, try using a different crosslinker or one that is MS-cleavable crosslinker to increase the number of identified crosslinked peptides.

Q. I observed crosslinked peptides which are outside the distance constraints of the linker length. How is this possible?
A. If the protein is a homodimer or part of a multimer complex, there could be crosslinks between peptides of the same protein that are intra-protein crosslinks that could be outside of the distance constraints. Alternatively, the protein may have been denatured before or during the crosslinking reaction which could result in crosslinking between amino acids that are farther apart than what is found on the native protein structure.