Elucidating Tissue- and Plasma-Specific Proteomic Alterations in Health and Disease

Summary

Proteomics is a high-throughput assessment of the interactions, function, composition, and structures of proteins and their cellular activities. Here, we present a protocol to assess total proteomic changes in various bio-samples, including plasma and tissues. The methodology reported herein is applicable to all bio-samples containing proteins.

Abstract

Proteomics is the large-scale study of expressed proteins, focusing on their structure, function, interactions, abundance, and post-translational modifications within a biological system. For example, phosphoproteomics, a vital subset of total proteomics, is the study of phosphorylation patterns and alterations on proteins. The methodology described herein illustrates a total proteomics approach to elucidate alterations in overall protein expression profiles, including phosphorylation patterns, in the plasma, brain, lung, spleen, and liver tissues collected from research animal models (i.e., ferrets and mice). This technique is applicable for other tissue types, as well as almost any bio-sample containing proteins (e.g., cultured cells). Following dissection, tissues of interest were flash frozen and stored at -80 °C. Tissues were then homogenized using a mortar and pestle with liquid nitrogen to preserve protein integrity and phosphorylation changes. Total protein was extracted from the tissue homogenates using a lysis buffer with a universal nuclease and protease/phosphatase inhibitors. Proteins extracted from tissue and equivalent plasma were converted to total peptides by controlled protease digestion using a commercial mass spectrometer sample preparation kit. Peptides were directly analyzed using ultra-high performance liquid chromatography/Orbitrap-tribrid tandem mass spectrometry. Identities of constituent proteins were reconstructed from the peptide mass spectral data, using proteomics bioinformatics software, as matched to a species-specific amino acid sequence library. Following this, the analysis should include strict cutoff criteria, especially using only high-confidence protein identifications. Statistically significant proteomic differences (>2-fold change; p < 0.05) can be determined between control and experimental groups.

Introduction

Proteomics, as the combination of proteome experimentation and data analysis, focuses on identifying and quantifying proteins, characterizing them based on composition, interaction, pre-translational and post-translational modifications (e.g., phosphorylation), sub-cellular localization, and structure under physiological conditions¹. Expression proteomics deals with quantitative comparisons of proteins that differ under changing biological conditions². It also holds a high importance in profiling a map of the complex, interconnected pathways, networks, and molecular systems, which directly control the major biological functions such as cell proliferation, differentiation, senescence, and cell death mechanisms. With the substantial advancement of experimental technology in recent years, proteomics methods have evolved from conventional approaches—including immunohistochemistry and enzyme-linked immunosorbent assay—to high-throughput techniques such as liquid chromatography^3,4. Proteomics-based approaches are gaining momentum in determining unique disease phenotypes and subtypes, as well as identifying new target interactions for therapeutic drugs (pharmacoproteomics). For example, patient stratification has primarily focused on genomic approaches to define homogeneous groups and guide treatment regimens. Transcription status alone, however, does not predict the fate of the translated proteins afterwards (e.g., stability or modification); and thus, non-genomic mechanisms contribute more to disease development, progression, severity, and resistance to therapeutics^5,6,7.

One such research area that has greatly benefited from quantitative comparisons of protein expression and activity is biomarker discovery in the field of neurodegenerative diseases and traumatic brain injury (TBI). The basis for biomarker discovery is to develop diagnostic techniques that facilitate early detection and potential targets for intervention. LC/MS-MS-based quantitative proteomics coupled with bioinformatics has aided the discovery of key protein biomarkers, including glial fibrillary acidic protein (GFAP) and ubiquitin carboxyl-terminal hydrolase isozyme L1 (UCH-L1), with a severe impact or penetrating TBI^8,9. For example, Military personnel frequently suffer TBI incidents during their missions, due to exposure to bomb explosions (i.e., blast-TBI). However, the underlying pathobiology of neurological deficits caused by blast-induced mild TBI remains unclear. Hence, understanding the molecular and ultrastructural mechanisms of blast-induced brain injury requires better identification of specific biomarker surrogates. With observation of proteomic changes, studies have demonstrated possible mechanisms of blast-induced oxidative stress in the brain to be associated with mitochondrial dysfunction, including impaired fission-fusion dynamics, decreased oxidative phosphorylation, and compensated respiration-relevant enzyme activities¹⁰. Insights on the early pathogenesis of blast-induced dysregulation of post-translational modifications such as acetylation, nitrosylation, deamination, and phosphorylation can also provide further characterization of its effects on the protein function implicated in axonal damage, inflammation, and blood-brain barrier disruption^11,12. With the advancement of systems biology approaches, mapping total proteome information from the brain and peripheral tissues in different animal species, including rodents and ferrets, has become a key procedure in blast injury research.

In this article, we focus on the methodology workflow for assessing quantitative aspects of expression proteomics and its post-translational modifications beginning from tissue or plasma, which has been extensively used for monitoring both physiological phenomena and pathological conditions. Utilizing the below methodology allows for consistency and reliability in sample preparation and data acquisition. By having this standardized methodology, comparisons can be made to determine preserved proteomic patterns across tissue types and phosphorylation patterns unique to specific tissues. Along with this, analysis across several preclinical animal models and human clinical samples can be made to obtain conserved phosphorylation patterns that could potentially serve as biomarkers and to tailor drug treatment regimens.

Protocol

Research was conducted under an IACUC-approved animal use protocol in an AAALAC International-accredited facility with a Public Health Services Animal Welfare Assurance and in compliance with the Animal Welfare Act and other federal statutes and regulations relating to laboratory animals. All supplies required for this procedure are listed in the Table of Materials. The procedure is subdivided into seven components.

CAUTION. Ensure proper safety protocols are followed when working with liquid nitrogen to homogenize tissue. The laboratory personnel wore freezer gloves, a freezer apron, a surgical face mask, and a face shield during homogenization of frozen tissue. This homogenization technique can be utilized for any tissue type, including extremely tough substances like bone, simply by adjusting the size of the mortar and pestle accordingly. See Figure 1A for the initial workbench setup. Refer to the Table of Materials for sizes used when homogenizing mouse and ferret tissues.

1. Cryogenic grinding of tissues with a mortar and pestle

1. Take the weight of an empty collection tube and record it. While in the collection tube, weigh the tissue using an analytical balance and record the weight in a laboratory notebook. Subtract the weight of an empty tube from the tissue plus tube weight to get the mg of tissue used.
   NOTE: The tissues that were assessed for proteomic studies include brain, lung, liver, and spleen. The whole tissue was cryogenically ground into powdered form using a mortar and pestle.
2. Pour liquid nitrogen into the cryopod, but ensure it is less than halfway filled.
3. Transfer the tube containing the sample and the empty, previously autoclaved 1.5 mL microcentrifuge tube(s) onto the tube rack located in the cryopod. This ensures the sample remains frozen while cooling down the mortar and pestle, preventing the sample from thawing when transferring the homogenized tissue into the empty 1.5 mL microcentrifuge tube(s).
4. Pour liquid nitrogen on the mortar and pestle to cool these instruments prior to use. The mortar and pestle sit outside the cryopod (Figure 1A,B).
5. Transfer the tissue into the cooled mortar containing liquid nitrogen (Figure 1B).
6. Repeatedly place pressure on the frozen tissue using the pestle and make a rotating, grinding motion with the pestle against the tissue until the tissue is ground into a fine powder (Figure 1C).
   1. Ensure excessive force is not applied, which runs the risk of samples spilling out of the mortar. Make sure to have firm control over the pestle.
   2. Add additional liquid nitrogen to the mortar while homogenizing large tissues to reduce the chance of the sample thawing while grinding.
   3. While grinding the tissue, check to make sure it is not thawing. If it is thawing, place additional liquid nitrogen in the mortar.
7. Dip the ends of the metal spatulas in the liquid nitrogen to chill them. This helps to easily transfer the homogenized tissue into the 1.5 mL tube. This prevents the tissue from thawing out and sticking to the spatula during the transfer.
8. Once the tissue is homogenized into a fine powder and the metal spatulas are chilled, carefully transfer the homogenized tissue powder into the new 1.5 mL tubes.
   1. Split the sample into aliquots dependent on the number of assays to be completed.
      NOTE: For example, four assays were to be completed for the samples in this study, so the homogenized tissue was equally aliquoted into four 1.5 mL tubes.
   2. Store any extra homogenized tissue in an autoclaved 5 mL snap-top collection tube.
9. Store the aliquoted tissues in a -80 °C freezer until future analysis, with a realistic use within 1 year to minimize air as well as light oxidation of proteins.
10. Weigh the empty collection tube.
11. Subtract the weight of the empty collection tube from the initial measurement (collection tube plus sample) to obtain the weight of the tissue sample.

2. Tissue lysis for protein isolation

1. Scoop dry ice into an ice pan container.
2. Place a tube rack in the dry ice and allow it to chill.
3. Remove homogenized tissue samples from the -80 °C freezer and place in the chilled tube rack. This prevents the samples from thawing, which could cause protein degradation and/or phosphorylation changes during the weighing process.
   1. Ensure the 1.5 mL tubes containing the sample remain frozen on the tube rack in dry ice.
4. Place a new 50 mL canonical tube in the dry ice.
5. Remove the cap from the 50 mL canonical tube and place disposable plastic spatulas in the 50 mL canonical tube to keep them cool. This prevents the sample from sticking to the spatulas during their transfer to the homogenization tubes.
6. For tissue lysis and protein isolation, use 2 mL tubes containing five 2.8 mm ceramic beads (referred to as homogenizing tubes in this protocol).
7. Label the cap and the side of the homogenizing tube with the sample ID using a permanent ink marking pen.
8. Place the labeled, homogenizing tubes in the tube rack located in the dry ice to keep the tubes cold and prevent sample thawing.
9. Place one homogenizing tube on an analytical scale and tare the scale.
10. Return the homogenizing tube to the tube rack on dry ice.
11. Remove the lid from the homogenization tube containing the beads, and using one of the cooled plastic spatulas, transfer 5 mg to 10 mg of powdered tissue sample to the respective homogenizing tube.
12. Place the lid back on the homogenizing tube.
13. Weigh the homogenizing tube on the analytical balance and record the amount of sample in the homogenizing tube.
    1. Use this weight to determine the amount of lysis buffer required for effective total protein extraction.
14. Once the weight is recorded, place the homogenizing tube containing the sample back in the tube rack on dry ice to keep it cold.
15. Dispose of the used plastic spatula.
16. Use a new, clean plastic spatula for each sample to be transferred and weighed to prevent cross-contamination. Repeat steps 2.9 to 2.15 for all samples.
17. After transferring all samples to individual homogenizing tubes, calculate the amount of Protein Lysis Master Mix needed.
    1. Prepare the required volume of Protein Lysis Master Mix, which is calculated based on the number of samples that need to be processed and the weight of each powdered sample. Add 100 µL of Protein Lysis Master Mix per 5 mg of tissue to each tube.
    2. Refer to company guidelines for reagent volumes to make the Protein Lysis Master Mix (refer to Table of Materials).
18. Place the homogenizing tubes containing the homogenized, powdered samples in a 4 °C cold block. The homogenizing tubes can no longer remain on dry ice. Placing the homogenizing tubes on the cold block prevents the lysis solution from freezing.
19. Remove the lid from the homogenizing tubes and transfer the calculated amount of Protein Lysis Master Mix needed to the respective homogenizing tube using the appropriate capacity pipette.
    1. Place the lid back on the homogenizing tubes and rotate the lids until they are on tight.
20. Place the sample tubes into the homogenizer and set it on a soft tissue program (speed 4500 RPM, 2 x 30 s per cycle, 30 s pause) at 4 °C for homogenization.
21. Once the homogenization cycle is completed, remove the tubes from the instrument and place them back in the 4 °C cold block.
    1. Visually inspect the sample tubes. Ensure no powdered sample is still visible. If the sample is still visualized throughout the Protein Lysis Master Mix, complete tissue lysis again on the instrument.
    2. Optimize the settings for the type of tissue being utilized. For example, powdered tissue derived from fibrous organs (e.g., heart) may need additional cycles or a higher RPM.
22. Transfer the homogenizing tubes into a centrifuge. Enter the following settings on the centrifuge: 16,000 g for 10 min at 4 °C, and press Start on the centrifuge.
23. Once centrifugation is complete, carefully transfer the homogenizing tubes back in the 4 °C cold block.
24. Label new 1.5 mL tubes on the lid and the side with the sample ID and any other pertinent information. Once labeled, place these 1.5 mL tubes in a 4 °C cold block.
    NOTE: The tubes were labeled with at least the following: animal ID, study time point, animal type, and tissue type.
25. Transfer the corresponding protein lysate from the homogenizing tubes to the respective labeled 1.5 mL tubes kept on a 4 °C cold block.
26. Store samples at -80 °C for further analysis. This is a safe stopping point.

3. Total protein quantification

1. Remove samples from the -80 °C freezer and thaw the samples on wet ice in an ice bucket.
2. Once thawed, vortex all samples and standards from the protein quantification kit for at least 3 s.
3. Add 10 µL of each replicate of standard, samples, and appropriate blanks into a 96-well microtiter plate based on the schematic layout (Figure 2A).
   NOTE: For each tissue sample, a 1:100 dilution of the protein lysate into preferably phosphate-buffered saline is used in the protein assay analysis. The schematic layout can be altered depending on the number of samples used in the protein assay. The samples, standards, and appropriate blanks are loaded into a 96-well microtiter plate in triplicate for assessing protein concentration accuracy, but if many samples need to fit on a plate, duplicates are acceptable.
   1. For the samples with a low amount of starting tissue (i.e., less than 5 mg), use only a single replicate or duplicates for the protein assay to maximize the amount of protein lysate available for the subsequent protein digestion.
   2. Ensure accurate pipetting and calibrated pipettes are used.
4. Invert the Protein Assay Reagent bottle several times to mix well, and then pour it into a 10 mL Reagent Reservoir.
5. Using a 200 µL maximum capacity multichannel pipette, add 150 µL of the Protein Assay Reagent to Column 1.
6. Pipette up and down several times to mix the protein assay reagent with the 10 µL of content already in the well to help reduce precipitate in the wells.
7. Discard the pipette tips and place new tips on the 200 µL maximum capacity multichannel pipette.
8. Repeat steps 3.5-3.7 until protein assay reagent is added to the entire 96-well plate.
   1. Use fresh pipette tips for each column of wells to prevent cross-contamination.
   2. Ensure that at least one well is left empty with no reagents or sample added to use as a blank absorbance value for protein quantification.
9. Once the protein assay reagent has been added to all the necessary wells, cover the 96-well plate with an aluminum foil plate seal.
10. Vortex the 96-well plate on a vortex instrument for 1 min at medium speed, and then incubate it at room temperature (RT) for 5 min before reading the plate.
11. Inspect the 96-well plate and aluminum foil seal. If the reagent remains stuck to the aluminum foil seal, place it with an appropriate balance inside a centrifuge.
    1. Press and hold the quick spin until it reaches approximately 200 g.
    2. Take the 96-well plate out and visually inspect the aluminum foil seal to ensure no liquid remains on the foil.
12. Slowly remove the aluminum foil seal and place the clear lid back on the 96-well plate (Figure 2B).
13. Read the 96-well plate on a spectrophotometer at 660 nm, after allowing the colorimetric reaction to develop for 5 min.
    NOTE: The absorbance readings using a UV/Visible spectrophotometer are measured at room temperature. The 660-nm Protein Assay is based on the binding of a proprietary dye-metal complex to protein in acidic conditions that causes a shift in the dye's absorption maximum, which is measured at 660 nm.
14. Use the absorbance readings of the known standards to create the standard curve using graphing software (Figure 2C-E), which is further applied to calculate the protein concentrations for all samples.

4. Protein digestion

CAUTION. Ensure proper safety protocols are followed when working with the hazardous chemicals utilized in the protocol below. The laboratory personnel wore personal protective equipment (PPE) and completed these steps in a chemical fume hood. Review all safety data sheets prior to working with these chemicals.

1. Label a new set of 1.5 mL microcentrifuge tubes with the sample IDs on the lid and side of the tubes in permanent ink. Include any other pertinent information.
   1. Based on the standard curve equation, the sample absorbance readings, and the dilution factor, calculate the protein concentration of each sample in micrograms per microliter (µg/µL). Use this value to calculate the required volume of sample lysate containing 100 µg of protein (Figure 2C-E). Transfer this calculated volume of sample lysate into a 1.5 mL microcentrifuge tube.
2. Make up the final volume to 100 µL with the lysis buffer; this is the starting volume needed for the digestion protocol.
   NOTE: The final volume of protein sample plus lysis buffer should equal 100 µL.
3. According to previously published literature, use 1.5 µL of the plasma for digestion, which is approximately equal to 100 µg of protein.
4. Add 50 µL of Reduction Solution (10 mM dithiothreitol) to each 1.5 mL microcentrifuge tube, close the tops, and vortex briefly for 3 s. Then add 50 µL of Alkylation Solution (30 mM iodoacetamide) to each sample, close the tops, and vortex for 3 s.
5. Incubate samples at 95 °C using a heat block for 11 min to reduce and alkylate, respectively, the protein sample. This cleaves all disulfide bridges and caps them with an acetamide group, preventing their reformation, and thus denatures the protein into a fully unfolded state and maximizes exposure of peptide digestion sites.
6. After incubation, remove samples from the heat block and allow them to cool to RT.
7. While waiting for the samples to cool, reconstitute the lyophilized Trypsin/Lys-C Protease Mix (1:1 ratio, 50 µg each) with Enzyme Reconstitution Solution by adding 515 µL of the liquid to 1 vial of the solid reagent, i.e., 0.1 µg/µL each. Avoid direct skin contact with the protease solution and all steps afterwards, since trypsin can be a severe irritant and allergen.
8. Once samples are cooled to RT, add 50 µL of reconstituted Trypsin/Lys-C Protease Mix (2.5 µg each) to the reduced and alkylated protein sample solution and vortex briefly.
9. Incubate on a thermomixer at 37 °C and set to 1000 rpm for 3 h to digest the protein sample.
   NOTE: The Lys-C protease is included to help cleave any sites that are resistant to digestion by trypsin, due to folding or amino acid sequence (e.g., Lys-Pro bonds). This not only reduces each protein into numerous knowable peptides of a reasonably small size and total charge but is also highly reproducible if the same digestion conditions are closely followed. The incubation time of 3 h during the digestion is the minimal time that we recommend for ensuring the complete cleavage of all proteins.
10. After the 3 h of incubation, remove from the thermomixer using a pair of forceps to avoid finger burns.
11. Add 50 µL of Digestion Stop Solution (6% trifluoroacetic acid or formic acid in water) to the sample and vortex briefly.
    NOTE: This inhibits the reaction, since the two proteases have maximum activities at a neutral pH 7-8.
12. Store samples in a -80 °C freezer as this also halts any residual protease activity. This is a safe stopping point.

5. Peptide clean-up, drying, and resuspension

NOTE: This procedure is mainly to remove the digestion reagents, especially non-volatile salts, which can readily damage the LC/MS-MS instrument, as well as help isolate and concentrate the produced peptides. The peptide clean-up is accomplished using kit-provided desalting spin columns, which are pre-packed with a size exclusion (gel-filtration) resin that initially captures the peptides and then allows especially salts to be washed through. Complete transfer of the kit reagents to the spin columns in a chemical fume hood, as they contain toxic organic solvents.

CAUTION. Ensure proper safety procedures, including the wearing of PPE and hazardous waste containment regulations, are followed when working with the hazardous chemicals in the protocol below. Complete these steps in a chemical fume hood and review all safety data sheets prior to working with these chemicals.

1. Remove samples from -80 °C freezer and allow samples to come to RT.
2. Remove the white cap at the bottom of the Peptide Clean-Up column, loosen the green top cap, and place it into a 2 mL collection tube.
3. Place these in the centrifuge, close the lid, press start, and centrifuge at 3000 g for 2 min to remove all the pre-shipment conditioning liquid.
4. Discard the flow-through in a hazardous waste container and place the peptide clean-up column in a new 2 mL collection tube.
5. Label the green lids of the peptide clean-up column with the Sample IDs, using a permeant ink marking pen.
6. Gently transfer the protein digest preparation, which is approximately 300 µL in volume, into the dry peptide clean-up column, by using a 200 µL pipette (e.g., 150 µL twice).
   1. Ensure no bubbles are observed at the top of the spin column's resin. If bubbles are observed, remove the bubbles by gently tapping the column or using a clean 10 µL pipette tip to disperse the bubble(s).
      NOTE: Trapped air bubbles can impede the sample's smooth transition into the resin, and thus, negatively affect the efficiency of the peptide clean-up and yield.
7. Place these in the centrifuge, close the lid, press Start and centrifuge at 1,500 g for 2 min.
8. Discard the flow-through in a hazardous waste container and place the peptide clean-up column in a new 2 mL collection tube.
9. Add 300 µL of the Wash Solution A (water, 0.1% trifluoroacetic acid / TFA) on top of the column resin, again using a 200 µL pipette.
   1. Ensure no bubbles are observed at the top of the spin column's resin. If bubbles are observed, remove the bubbles by gently tapping the column or use a clean 10 µL pipette tip to disperse the bubble(s).
10. Place these in the centrifuge, close the lid, press Start, and centrifuge at 1,500 g for 2 min.
11. Discard the flow-through in a hazardous waste container and place the peptide clean-up column in a new 2 mL collection tube.
12. Add 300 µL of Wash Solution B (5% methanol, 0.1% TFA) on top of the column resin, again using a 200 µL pipette.
    1. Ensure no bubbles are observed at the top of the spin column's resin. If bubbles are observed, remove the bubbles by gentle tapping the column or use a clean 10 µL pipette tip to disperse the bubble(s).
13. Place these in the centrifuge, close the lid, press Start and centrifuge at 1,500 g for 2 min.
14. Discard the flow-through in a hazardous waste container and place the peptide clean-up column in a new 2 mL collection tube.
15. Repeat steps 5.12-5.14 once more so that the column is washed twice with Wash Solution B.
16. Transfer the peptide clean-up column into a new 2 mL microcentrifuge tube.
    1. Ensure the 2 mL microcentrifuge tubes used in this step are the kit-provided low protein-binding collection tubes.
    2. To conserve the available amount of low-protein-binding collection tubes, use standard laboratory ones for the previous wash steps.
17. Add 300 µL of Elution Solution (50% acetonitrile, 0.1% TFA) on top of the column resin, again using a 200 µL pipette.
    1. Ensure no bubbles are observed at the top of the spin column's resin. If bubbles are observed, remove the bubbles by gently tapping the column or use a clean 10 µL pipette tip to disperse the bubble(s).
18. Place these in the centrifuge, close the lid, press Start, and centrifuge at 1,500 g for 3 min to collect the clean peptide sample in the new low protein binding 2 mL collection tube.
    1. If desired, transfer 125 µL (approximately half) of the eluted clean peptide solution into a 1 mL glass vial, using a 200 µL pipette, and store at -80 °C.
       NOTE: This aliquot can be used later for additional analysis, such as tandem mass tag (TMT) labeling of the peptides, as done by a commercially available kit, for more precise quantification across samples, especially independently prepared sets.
19. Store the remaining sample, approximately 175 µL, at -30 °C in the low protein binding 2 mL tubes until ready to proceed with drying and reconstitution. This is a safe stopping point.
20. To dry the sample, thaw the peptide sample from -30 °C to RT. If proceeding with drying and reconstitution right after peptide clean-up, ignore this step.
21. Open the 2 mL microcentrifuge tubes and place them into a vacuum centrifuge concentrator.
    1. Make sure to balance the vacuum centrifuge concentrator if an uneven number of samples is being run at the same time.
    2. Close the vacuum centrifuge concentrator lid and press Start.
    3. Allow samples to run on the vacuum centrifuge concentrator for 3 h, without application of heating.
    4. Inspect tubes periodically to ensure no liquid is remaining inside, as this can interfere with the binding of the peptides to the separation column of the LC/MS-MS instrument.
    5. If liquid is still observed, continue spinning the samples for additional time.
    6. Check between 30 min and 1 h to see if liquid remains in the tube.
       NOTE: The peptides are highly stable during this process as they are under vacuum conditions and will not be deteriorated by a short period of dryness or moderate warmth generated by the vacuum centrifuge concentrator operation alone.
22. Once dry, immediately resuspend the sample in 50 µL of 0.1% formic acid in MS-grade water (e.g., 5 µL of 100% formic acid in 5 mL of water).
    1. Add 50 µL of 0.1% formic acid to the bottom of the 2 mL low protein-binding tube containing the dried peptide sample, using a 200 µL pipette.
23. Close the caps, then vortex for 15 s.
24. Transfer the reconstituted samples into the respective 1 mL glass vials, using a 200 µL pipette.
25. Store the glass vials containing the reconstituted samples at -30 °C, and analyze them by LC-MS/MS-based approach as soon as possible.
    NOTE: The frozen peptide samples are stable at -30 °C for short-term storage and -80 °C for longer storage.

6. Data acquisition of peptide samples on a UHPLC/MS-MS instrument

CAUTION. Ensure proper safety protocols are followed when working with an Ultra High-Performance Liquid Chromatography/Orbitrap tribrid tandem Mass Spectrometer system. The laboratory personnel wore PPE and underwent formal, hands-on instrument training provided by the UHPLC/MS-MS manufacturer. The main cautions are proper handling of the acidic and organic solvent mobile phases for the UHPLC and high-voltage ionization source for the MS.

1. Perform peptide separations using an Ultra High-Performance Liquid Chromatography (UHPLC) system. The LC column is an electrospray capable C18 reverse phase, rapid separation (RSLC) design, of dimensions 2 µm, 100 Å, 50 µm x 15 cm, as placed in-line after a guard pre-column of the same material.
2. Use the following UHPLC mobile phases: 0.1% formic acid in water (solvent A), 0.1% formic acid in acetonitrile (solvent B), and 0.1% formic acid in 90% water and 10% methanol (solvent C).
   NOTE: All solvents as well as formic acid, used for preparing the mobile phases are certified LC/MS grade and made only as needed. The UHPLC has a filtration and degasser unit prior to the pumps, so the mobile phases are not manually filtered and purged of air.
3. Set programmed injections of the total peptides prepared for the plasma and tissue samples at 3 µL to begin with, which is equivalent to ~3 µg of peptide material as determined by the starting amount of protein used.
   NOTE: This should produce an ideal MS signal intensity for maximum protein identifications. Injection volume can be increased to as much as 15 µL to further boost the signal output or decreased to as little as 1 µL if overloading is apparent.
4. Maintain the column and autosampler at 40 °C and 10 °C, respectively.
5. Set the solvent flow rate for the proteomics acquisition at a constant 0.3 µL/min, using the UHPLC nano-pumps, with a linear separation gradient of the water (A) to acetonitrile (B) mobile phases.
6. Carry out washes of the injection syringe and sample loop (20 µL capacity) using the water and methanol mobile phase (C), with the UHPLC loading pump initially set at 5 µL/min.
7. During the sample separation over 80 min total, set the LC gradient (linear curves) as follows: Initial - 5% B; 0-60 min - 32% B; 60-61 min - 95% B; 61-65 min - 95% B; 65-66 min - 2% B; and 66-80 min - 2% B. Likewise, run the loading pump at a constant 100% of A, but adjust the flow rate linearly as follows: Initial - 5 µL/min; 0-5 min - 1 µL/min; 5-61 min - 5 µL/min; and 61-80 min - 5 µL/min (Figure 3A).
8. Inject unknown peptide samples as sequential duplicate runs.
9. To prevent carryover, carry out at least two sequential blank injections of 0.1% formic acid in water between samples using a truncated LC gradient (linear curves) over 35 min total as follows: Initial - 5% B; 0-23 min - 32% B; 23-24 min - 95% B; 24-30 min - 95% B; 30-31 min - 2% B; and 31-35 min - 2% B (Figure 3B).
   1. Ensure the settings for the loading pump LC gradient remain unchanged.
   2. For peptide samples having high levels of endogenous contaminants, such as liver (e.g., bile acids), include a third blank every 3 to 5 samples and at the end of the run to help clean out the LC column. This consists of injecting 1 to 2 µL of solution containing 25% each of acetonitrile, isopropanol, methanol, and water.
   3. Alternatively, or in combination with the above step, perform a percentage change enhanced LC gradient (i.e., see-saw) over 35 min total as a system washout as follows: Initial - 2% B; 0-3 min - 2% B; 3-4 min - 95% B; 4-10 min - 95% B; 10-11 min - 2% B; and 11-15 min - 2% B; 15-16 min - 95% B; 16-20 min - 95% B; 20-21 min - 2% B; 21-25 min - 2% B; 25-26 min - 95% B; 26-30 min - 95% B; 30-31 min - 2% B; and 31-35 min - 2% B (Figure 3C).
10. Carry out the detection of the peptides, i.e., corresponding molecular masses, as they are separated by and eluted from the LC column, using an Orbitrap and an ion trap combination tandem quadrupole mass spectrometer (MS-MS) system.
    1. Before performing the sample runs, plug the LC column directly into the face of the MS instrument's interchangeable nano spray ionization (NSI) source, which is located on the upper left front corner of the device.
       NOTE: This exposes the inbuilt electrospray needle of the LC column that delivers the peptides, as suspended inside charged micro-droplets of LC mobile phase solvent, into the inlet of the detector's ion transfer tube.
    2. Also, to improve the electrospray volatilization efficiency, turn on the built-in heating unit of the NSI source and set it to 40 °C.
11. Also, before the sample run, check that there is a satisfactory external input of compressed nitrogen and helium gas into the MS instrument. Achieve this using ultra-high purity grade, 300 size steel-cylinder tanks of each gas, which have the regulator gauges set at static flow pressures of 60 and 40 psi, respectively.
12. If the MS instrument issues warnings about instability of the nitrogen gas input, then raise the pressure up to a maximum of 100 psi. Make sure the tanks are at least 10-20% full, which is a top valve gauge reading of ~250-500 psi; and if not, change them out to fresh ones. Dropping below 10% of full capacity has a risk of drawing contaminants into the MS instrument, which can settle at the bottom of the gas tanks.
13. Make the mass spectrometer method beforehand using a controlling software provided template that is designed for the optimal detection of peptides and its post-translational modifications (Figure 3D-H), and enter the following standard conditions:
    1. Set the total detection period to 80 min.
    2. Set the general detection mode to peptides.
    3. Set the NSI source to the positive mode (i.e., charge of the peptide ions).
    4. Set the nano-spray needle voltage to 2000 V and keep it as static.
    5. Set the nitrogen gas input to the NSI and set the detectors to static.
    6. Set the ion transfer tube temperature to 275 °C.
    7. Detect the masses of the whole peptides (i.e., parent ions; MS1) by setting the scan range to 350-2000 m/z, RF lens to 30%, maximum injection time to Auto, and the orbitrap mass analyzer to a 120,000 resolution.
       NOTE: The alternative ion trap capability of instrument is not used here to screen for the peptide masses, as it is less sensitive for monitoring this type of molecule compared to the orbitrap.
    8. Set the ion filters to discriminate between those peptides having multiple charge states in a range of 2-7. This allows the instrument, while recording the data, to automatically convert the inherently detected mass over charge ratios (m/z) of the molecules into their exact masses. Likewise, a peptide with just a single charge state (m/z = 1) does not require any conversion factor to obtain its exact mass.
    9. Set the dynamic exclusion gates for the ions to 1 pass at 60 s duration, ±10 ppm mass tolerance, and to drop out any molecules with natural isotopic atoms present (e.g., ¹⁴C).
    10. Detect the masses of identity confirmation fragments generated for the peptides (i.e., daughter ions; MS2) by setting the instrument to the data-dependent acquisition (DDA mode).
    11. Capture the whole peptides (i.e., parent ions) by quadrupole isolation, using a setting for the mass variability tolerance window of m/z = 1.6.
    12. Perform the fragmentation of the captured peptides by setting the electron impact mode to HCD (i.e., higher-energy collisional dissociation using nitrogen and helium gas as the electron transfer agent), collision energy to 30%, scan range to Auto, maximum injection time to Auto, and the Orbitrap mass analyzer to a 30,000 resolution.
    13. Acquire the final data (MS1 and MS2 ions) by setting the signal peak shape to a centroid format.
14. Maintain accurate mass determinations for detected peptides by routinely calibrating the instrument, usually prior to each sample batch run, as described below.
    1. Place the MS instrument in an ion source power off status, using the controlling software, and switch out the NSI source to the device's alternative internally heated electrospray ionization (HESI) source. This is the source used for applications that can require very large sample injection volumes and a corresponding high flow rate (1-2500 µL/min) for the LC pump, e.g., metabolomics and drug pharmacokinetics experiments.
    2. Select the full calibration check option on the controlling software.
    3. Fill a 500 µL glass syringe having a Teflon-coated plunger and stainless-steel blunt needle with a commercially available MS standard solution, and then connect the PTFE capillary tube line to the HSEI source (while powered off).
       NOTE: The standard used here contains a mixture of natural and synthetic molecules (16 total, e.g., caffeine) that span a mass range of m/z = 50-2800 and cover a wide range of possible chemical bond types, especially those that are associated with peptides (e.g., amide groups).
    4. Initiate a constant infusion of the MS standard into the HSEI source (while powered on), using a rate of 3-5 µL/min as done using a syringe pump. Once a stable pattern of standard peaks is noted on the controlling software live view window, activate the automated calibration check.
       NOTE: If no issues are found with the sensitivity and accuracy of the mass determinations for the standard, a successful full calibration typically takes < 40 min. Likewise, only if the instrument passes all the MS hardware parameters that are checked is it considered reliable for running samples.
    5. Confirm the success of the calibration by resetting up the NSI source and LC column, and then running a standard solution of human [Glu 1]-Fibrinopeptide B (Glu-Fib) at total injected amounts covering 1-10 ng (e.g., 1, 2, 6, and 10 ng). Use the same LC separation and MS detection methods as those described above for the unknown samples.
    6. Check the Glu-Fib standard for appropriate LC retention time (~20 min), presence of MS1 and MS2 ions (m/z = 1285.54, 813.39, 785.84, 445.12, 333.19, and 175.12), and linearity of increasing peak areas with amount injected.
    7. Then generate sequence tables, using the controlling and data capture software, for running the unknown peptide and blank samples that contain columns for entering the sample type, file name, sample ID, data path, LC/MS instrument method, vial position, and injection volume (Figure 3D-E).
    8. Save the resulting MS data files (extension: RAW; 500 - 800 GB each) in the designated path folder on the instrument computer's hard drive. Then transfer these files to a workstation computer that is powerful enough to drive the bioinformatics software, as described later, that converts the extremely extensive and complex peptide mass spectral data (i.e., MS1 and MS2 signals) into corresponding total protein composition profiles.

7. Conversion of raw MS data to representative total proteome results

NOTE: The raw MS data captured for peptide samples are processed into total protein composition profiles (proteome), using bioinformatics software. In brief, the bioinformatic software reconstructs the original proteins in the sample by matching the raw MS data to their known peptide MS1 and MS2 spectra found in a web-based species-specific, proteome reference library, through extensive and unique coverage of the protein's amino acid sequence. Relative abundances of each identified protein are determined by the program using the MS1 signal intensities of its matched unique peptides. Determined abundances can be reliably compared across samples, but only if they are run within the same preparation batch and using identical LC and MS settings. Otherwise, previously mentioned absolute quantitative techniques based on internal standard/reporter ion recoveries, like selective TMT labeling of the peptides within each sample, will have to be employed.

1. Download onto the workstation computer's hard drive the proteome reference file from the web-based library.
   1. Find this by using the database located at the Uniprot website for the sample species of interest (e.g., ferrets / Mustela furo). This is called a FASTA file. Use the Proteomes search tool and Table view to locate and then download the proteome reference file.
   2. To maximize the possible number of protein identifications generated, select both the committee peer-reviewed (Swiss-Prot) and unreviewed (TrEMBL) entries to the Uniprot database when making the FASTA file. Likewise, choose the option to include all reviewed isoforms for the proteins.
2. Open the bioinformatics software, i.e., Proteome Discoverer, to the Start Page tab, and then select the Administration tab.
3. Go to the Maintain FASTA files subtab and then download into the program the above species-specific FASTA file generated for your experiments.
4. Go back to the Start Page tab and click on New Study/Analysis, and in the dialog box, type a study name in the Study Name field and select a file folder for saving the study in the Study Root Directory field.
5. Leave the Processing Workflow and Consensus Workflow fields blank for now, as they will be selected for later.
6. Click OK to save and leave the dialog box, which will then bring up the Study Definition page.
7. Go to the Study Definition tab and type in the project's details in the Study Description box located on the middle-left side.
8. Go to the Input Files tab and click on Add Files.
   1. This will bring up a dialog box in which the required raw data files to be processed into total protein composition profile (proteome) results can be selected and downloaded from the computer's hard drive.
9. Click on the raw MS data files that are now listed under the Input Files tab, which can be selected and highlighted individually or all at once.
10. Drag them over and drop them into the Files for Analysis section of the Analysis box located on the middle-right side as part of the Processing Step sub-box.
11. Go to the Workflows tab and click on Edit, which is found next to the Processing Step header of the Analysis box located on the middle-right side.
12. Click on Open Common and from the ProcessingWF_Tribrid folder, select the processing workflow file that is designated specifically for phosphoprotein sub-identifications, as obtained under HCD fragmentation-based MS detection (Figure 4A).
    NOTE: A processing step diagram containing editable nodes will then appear in the Workflow Tree box. There will be two nodes, as described below, which are marked with exclamation points indicating that there is a required entry of experiment-specific information by the user.
13. In the processing workflow tree diagram, click on and open the Spectrum Files RC node.
    1. In the Parameters of Spectrum Files RC category,, find and select under the Protein Database parameter the FASTA file, which was previously generated using the Uniprot website, for the species-specific reference library of peptide MS spectra that will be searched to identify the proteins of experimental interest.
14. Click on and open the Sequest HT node and make the following three changes:
    1. Under the Protein Database parameter, select the same FASTA file, as described above.
    2. Under the Enzyme Name parameter, select Trypsin (full) or alternatively Lys-C, which are the main and backup proteases used to prepare the original peptide digest.
    3. Under the Dynamic Modifications parameter, make sure at least Phospho / +79.966 Da (S, T, Y) has been selected.
       1. Use the latter setting to reveal peptides associated with proteins that are post-translationally modified by phosphorylation, via looking for corresponding MS spectra losses of a phosphate group(s) (m/z = 79.97).
       2. Find the phosphate-occupied amino acid sites (i.e., Serine, Threonine, or Tyrosine) by the expected alterations in their MS2 fragmentation patterns.
       3. Add other protein post-translational modifications of routine interest in the same fashion to the Sequest HT node at this point, e.g., methylation, acetylation, myristylation, if the exact mass loss of the related groups is known. Likewise, expand the search for the phosphorylation sites to other less frequently reported amino acids, e.g., histidine.
          NOTE: The Spectrum Files RC and Sequest HT nodes of the workflow tree for the processing step are the only two that require any parameters to be adjusted.
    4. Leave the rest as their default settings, as these have already optimized by the manufacturer to produce the highest number of designated protein identifications of interest. Likewise, this applies to all the other nodes in the workflow tree.
    5. Make sure that Percolator and IMP-ptmRS nodes are included at the end of the workflow tree.
       NOTE: Percolator uses reiterative fitting to greatly refine the end results. IMP-ptmRS is essential for reporting the appropriate identification and site localization of all protein post-translational modifications, including that of phosphorylation.
15. Go back to the Workflows tab and click on Edit found next to the Consensus Step header of the Analysis box located on the upper-right side. This will bring up the workflow tree diagram for the consensus step.
16. Click on Open Common and from the ConsensusWF folder select the consensus workflow file that will produce the desired output format for the protein composition profile (proteome) results (Figure 4B).
    NOTE: A consensus step diagram containing editable nodes will open in the Workflow Tree box. The diagram utilized for processing the data is generated by modification of the basic consensus workflow, where in-line Feature Mapper and Precursor Ions Quantifier nodes were added as a branch off the initial MSF Files node, to facilitate the quantification of protein relative abundances. While all the nodes in the workflow already contain optimized parameters established by the manufacturer, there are some containing very important entries to double check for the proper selection of as described below.
17. In the workflow tree diagram, click on and open the Precursor Ions Quantifier node. This is the node that is essential for quantifying the relative abundances of each identified protein.
    1. Under the General Quantification Settings category find and select under the Peptides to Use parameter the Unique + Razor option.
       NOTE: Unique peptides are those that contain amino acid sequences that are exclusive to a single protein; and thus, are used to specifically determine its amount as based on the MS signal intensities. Razor peptides are those shared among groups of proteins; and thus, are used to pull the calculated relative abundances into agreement.
    2. Under the Normalization and Scaling category change both the Normalization Mode and Scaling Mode parameters to the None option, as this type of adjustment is too restricted for reliable application across the entire proteome.
       NOTE: These settings are only activated when it is desired to normalize the calculated relative abundances of select proteins of interest against those that are stably intrinsic to the cell, e.g., housekeeping genes, and then express the resulting values as a scaled ratio.
    3. Next, click on and open the Peptide and Protein Filter node and under the Peptide Filters category change the Peptide Confidence At Least and Keep Lower Confident PSM parameters to the High and False options, respectively. This sets the peptide matches used in making the protein identifications to the greatest level of fit significance (i.e., ≥ 95%).
       NOTE: Alternatively, the first parameter can be set to Medium to relax the stringency of the peptide matches and thus greatly increase the output of protein hits; but this conversely leads to a very high risk of generating many false identifications. Likewise, it should never be the case that a "Low" value is used as an acceptable option for both parameters.
    4. These are the only nodes of the consensus step that need to have parameters adjusted, leave the rest as using their default settings.
       NOTE: Overall, the consensus step workflow handles all the post-hoc analysis, i.e., determination of the protein relative abundances, identification confidences (i.e., False Discovery Rates and peptide spectral match scores), and mapped peptide coverage. Result statistics and data distributions are also provided as end tables.
18. Once the above processing and consensus step workflows are in place, save the entire study and then click on the Run icon (gears) located at the top of the Analysis box. If multiple raw MS data files are being processed at once, then prior to starting the processing run, click on and checkmark the nearby By File icon (square).
    NOTE: The processing of one MS raw data file into a total proteome result file typically takes 20-40 min, but it can yield a running list of many thousands of protein identifications ranging from low, middle, to high confidence matches (i.e., with a weighting towards high hits), along with the phosphate modifications that are present within each entity (Figure 4C). The major result output parameters for the identified proteins are FDR (confidence), Master (if the FASTA file / library entry was reviewed), Accession #, Description, Coverage %, Sequence Coverage, Peptide Spectral Matches (e.g., number of unique peptides), Physical Characteristics (e.g., MW in kDa), Relative Abundances, and Modifications (i.e., phosphorylation groups).
    1. Column arrange the total proteome results in ascending or descending order or entirely filtered, using the Funnel icon on the tool bar (Figure 4C), according to research interests, e.g., show only those with a high confidence identification (protein FDR) or a high degree of peptide matching (e.g., by coverage % and/or number of unique found).
    2. If post-translational modification information is specifically desired, filter out identified proteins having phosphorylation for further analysis.
    3. Save the filtered results, without a file name change, as they can be restored to the original output by simply removing the applied filters and refreshing the window.

Results

Visualization of total proteome results by optional processing into volcano plots
The total proteome results for each sample can be readily searched by accession # or description for specific proteins of research interest and the associated relative abundances are averaged and then compared between treatment groups for significant differences (e.g., as by t-test derived p-values). Alternatively, it can be of great exploratory value to look for global changes across the entire proteome, including any ...

Discussion

The above method utilized tissues that were snap frozen in liquid nitrogen to immediately halt all biological activity before transferring to -80 °C for long-term storage. Once all samples were collected, tissue homogenization occurred. During homogenization, it is critical to keep all tissues and tools cold to prevent reactivation or degradation of the proteins. Also, during the initial extraction of protein from homogenized tissue, samples are kept cold to prevent loss of protein activity or protein degradation. W...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
660 nm Protein Assay Kit	Pierce / Thermo Scientific	22662	For BCA to determine protein concentrations of samples, contains Pierce 660nm Protein Assay Reagent and Pre-Diluted Protein Assay Standards
Acetonitrile, Optima LC/MS grade	Thermo Fisher Scientific	047138.K2	Ultra-pure acetonitrile used for liquid chromatography/mass spectrometry
Aluminum Foil Plate-Seal	Beckman Coulter	538619	Seal used during development of protein assay plate
BioTek Synergy Neo2 Multi-mode Reader / UV-Vis Spectrometer	Agilent	https://www.agilent.com/en/product/microplate-instrumentation/microplate-readers/multimode-microplate-readers/biotek-synergy-neo2-hybrid-multimode-reader-1623195	Spectrophotometer/assay plate reader instrument used to complete 660nm absorbance readings to determine protein concentrations
ChromaCare LC-MS Instrument Flush Solution	Thermo Fisher Scientific	T111101000	Solution containing 25% each of acetonitrile, isopropranol, methanol, and water use to help clean out the LC system after running samples
CK28 Precellys tube	Bertin Corp / Technologies	P000911LYSK0A.0	Homogenized tissues are placed in these tubes for tissue lysis and protein isolation using the Precellyis instrument. These are the tubes utilized by our lab referred to in the protocol as "homogenizing tubes."
Cryopod	Brooks / Azenta Life Sciences	https://www.azenta.com/products/cryopod-carrier	Container used to hold liquid nitrogen for chilling mortar during tisuse homogenization
EasyPep Mini MS Sample Prep Kit	Thermo Fisher Scientific	A40006	One kit can process 20 samples, contents include the lysis solution for protein extraction, universal nuclease supplemented in the extraction solution, and regeants needed for Steps 1.4 and 1.5
EASY-Spray C18 LC column (15 cm L, 50 µm Dia, and 2 µm particle size)	Thermo Fisher Scientific	ES901	Electrospray LC column used for the seperation of peptides prior to MS detection
FlexMix calibration solution	Pierce / Thermo Scientific	A39239	Mixture of 16 standard molecules used to calibrate the detector accuracy and sensitivity of MS instrument prior to sample runs
Formic Acid, LC/MS grade	Thermo Fisher Scientific	85178	Ultra-pure formic acid used to acifidy all LC solutions and samples at a final concentration of 0.1% (v/v)
Halt Protease & Phosphotase Single-Use Inhibitor Cocktail (100x)	Thermo Fisher Scientific	78442	Supplemented in the extraction buffer to prevent degradation of proteins and phosphoproteins in sample
Methanol, Optima LC/MS grade	Thermo Fisher Scientific	047192.K2	Ultra-pure methanol used for liquid chromatography/mass spectrometry
Porcelain Mortar, 150 mL	Thermo Fisher Scientific	FB961C	150 mL capacity, used for homogenizing smaller ferret tissues (i.e., brain)
Porcelain Mortar, 275 mL	Thermo Fisher Scientific	FB961D	275 mL capacity, used for homogenizing large ferret tissues (i.e., liver)
Porcelain Mortar, 50 mL	Thermo Fisher Scientific	FB961A	50 mL capacity, used for homogenizing mouse tissues
Porcelain Pestl, 50 mL	Thermo Fisher Scientific	FB961K	For use with 50 mL mortar
Porcelain Pestles, 150 mL	Thermo Fisher Scientific	FB961M	For use with 150 mL mortar
Porcelain Pestles, 275 mL	Thermo Fisher Scientific	FB961N	For use with 275 mL mortar
Precellys	Bertin Corp / Technologies	https://www.bertin-technologies.com/product/sample-preparation-homogenizers/precellys-evolution-homogenizer/	Instrument used to homogenize tissue samples. The following settings were used: 4500 RPM, 2 x 30 s per cycle, 30 s pause.
Protein Assay Reagent	Pierce / Thermo Scientific	22662	Protein Assay Reagent used in steps 3.4–3.8
Proteome Discoverer	Thermo Fisher Scientific	CSW0064764	Bioinformatics software for analysis of proteomics/phosphoproteomics data
SureSTART MSCERT screw cap LC injection vials (0.02–1.5 mL capacity)	Thermo Fisher Scientific	6PMCK39TR	High preformance application certified, screw cap (PTFE, pre-slit septa) glass vials used to contain samples for injection on the LC/MS instrument
Ultimate 3000 UHPLC / Orbitrap Fusion Lumos Tribrid MS (LC/MS) system	Thermo Fisher Scientific	LC: discountinued / MS: FETD2-10002, by special request	Ultra-high preformance liquid chromatography/mass spectrometery (LC/MS) instrument used to separate out and then detect by molecular weight the peptides in the samples
Universal Nuclease for Cell Lysis, 100kU, 400 µL	Pierce / Thermo Scientific	88702	Additional universal nuclease was purchased
Vacufuge Plus	Eppendorf	https://www.eppendorf.com/us-en/Products/Centrifugation/Concentrator/Vacufuge-plus-p-PF-25748	Vacufuge used to dry peptide samples before reconstitution.
Water, Optima LC/MS grade	Thermo Fisher Scientific	047146.K2	Ultra-pure water used for liquid chromatography/mass spectrometry