Optimized Workflow for Proteomics and Phosphoproteomics With Limited Tissue Samples

Proteomics and phosphoproteomics play crucial roles in elucidating the dynamics of post‐transcriptional processes. While experimental methods and workflows have been established in this field, a persistent challenge arises when dealing with small samples containing a limited amount of protein. This limitation can significantly impact the recovery of peptides and phosphopeptides. In response to this challenge, we have developed a comprehensive experimental workflow tailored specifically for small‐scale samples, with a special emphasis on neuronal tissues like the trigeminal ganglion. Our proposed workflow consists of seven steps aimed at optimizing the preparation of limited tissue samples for both proteomic and phosphoproteomic analyses. One noteworthy innovation in our approach involves the utilization of a dual enrichment strategy for phosphopeptides. Initially, we employ Fe‐NTA Magnetic beads, renowned for their specificity and effectiveness in capturing phosphopeptides. Subsequently, we complement this approach with the TiO2‐based method, which offers a broader spectrum of phosphopeptide recovery. This innovative workflow not only overcomes the challenges posed by limited sample sizes but also establishes a new benchmark for precision and efficiency in proteomic investigations. Published 2024. This article is a U.S. Government work and is in the public domain in the USA. Current Protocols published by Wiley Periodicals LLC.


INTRODUCTION
The study of the proteome involves a comprehensive assessment of protein function and structure to gain a nuanced understanding of protein expression in specific tissues or cells (Breitkopf & Asara, 2012, Al-Amrani et al., 2021).Phosphorylation of proteins, a critical mechanism regulating cell signaling processes, such as gene expression and membrane transport, plays a pivotal role in both physiological and pathological events.However, the detection and identification of phosphorylated peptides on a proteome-wide scale present significant challenges due to the low abundance of phosphorylated proteins and potential losses during sample preparation (Dunn et al., 2010).
The trigeminal ganglion, a critical component of primary sensory ganglia, is responsible for transmitting pain and itch sensations.Its intricate structure comprises trigeminal ganglion cell bodies surrounded by a single layer of satellite glial cells, while Schwann cells wrap both distal and central afferent neuronal processes.The ganglion also accommodates fibroblasts, collagen fibers, small blood vessels, and various immune cells (Messlinger & Russo, 2019).Additionally, each mouse trigeminal ganglion is ∼0.1 g in size.As a result, studying the proteome and phosphoproteome of tiny neuronal tissues, such as the trigeminal ganglion, presents challenges due to the limitations imposed by tissue size and cell type composition.
To address these challenges, our customized workflow employs a high concentration of 5% SDS as a lysis buffer for protein extraction from the trigeminal ganglion.Additionally, a three-step process incorporating Fe-NTA magnetic beads enrichment and TiO 2 enrichment, is implemented to enhance the yield of phosphopeptides.This protocol provides a comprehensive workflow outline facilitating high-quality phosphoproteomic analysis with limited neuronal tissue samples.Detailed procedures cover sample lysis, protein digestion, clearance, enrichment, and liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis.NOTE: All protocols involving animals must be reviewed and approved by the appropriate Animal Care and Use Committee and must follow regulations for the care and use of laboratory animals.

PROTEIN EXTRACTION AND DIGESTION
Protein extraction is a pivotal initial step in proteomics, particularly when dealing with limited tissue samples or total protein.Achieving efficient protein extraction from neuronal tissues, such as the trigeminal ganglion, requires careful selection of the lysis buffer to optimize results.This part of the protocol focuses on the use of the 5% SDS lysis buffer for conducting protein extraction (Hu, Doyle et al., 2022, Wang, Veth et al., 2023).
Protein digestion is a critical tool for identifying, characterizing, and measuring proteins in proteomic research (Switzar, Giera et al., 2013).Predominantly, proteomics experiments rely on digestion of the protein into peptides prior to mass spectrometric (MS) analysis.Basic Protocol 1 describes the use of dithiothreitol (DTT) and subsequent iodoacetamide (IAA) for the reduction and alkylation of proteins, respectively, which leads to their denaturation.This refined protocol has been adapted from the original S-trap digestion method.
If the tissue is dense, e.g., bone, the use of liquid nitrogen is preferred.
2. Pre-cool a grinder and pestle on ice.
3. Take the tissue out from the freezer.Transfer the tissue into a 1-ml homogenizer.Add 100 μl of 5% SDS lysis buffer and thoroughly homogenize the sample at room temperature.
This step should not be done on ice, as SDS may precipitate at low temperatures, potentially affecting protein extraction from the tissue.
Be cautious and ensure that the grinder and pestle match, and that the top of the pestle can reach the bottom of the grinder.Otherwise, the tissue may get stuck at the bottom, resulting in inhomogeneous samples and poor protein recovery.
4. Transfer the homogenized tissue and buffer into a 1.5-ml microcentrifuge tube with low protein binding properties, then boil the mixture for 2 min.
5. Centrifuge samples 10 min at 14,000 × g, room temperature, and collect the supernatants.Keep the samples.
6. Determine the protein concentration using a BCA protein assay kit.After testing the concentration, retain the sample for digestion as outlined in Basic Protocol 2.
7. Take 100 μg protein aliquot from the total protein extract supernatant from Basic Protocol 1 for each sample.
For example, if the current protein aliquot is 30 μl, add 1.2 μl of 50 mM DTT (diluted 10-fold from stock solution) to the protein sample.
9. Add iodoacetamide (IAA) to a final concentration of 5 mM and incubate at room temperature for 45 min.Keep the reaction in the dark.
This step is to alkylate free cysteines.
For example, if the current sample volume is now 31.2μl, add 3 μl of 55 mM IAA (diluted 10-fold from stock solution) to the sample.
10. Add 12% aqueous phosphoric acid to the sample at a 1:10 (v/v) ratio, achieving a final concentration of 1.2% phosphoric acid.After adding, vortex the sample for thorough mixing.
For example, if the sample volume is now 34.2 μl, add 3.4 μl 12% phosphoric acid.
This step is essential since the protein trap binds at pH ≤ 1.
If enough protein is present, solution should turn opaque, indicating colloidal formation.
12. Transfer to the top of an S-Trap column; if total volume is >200 μl at this point, repeat this step and step 7 with ∼200 μl each time.
13. Centrifuge the micro column 1 to 2 min at 4000 × g, 4°C, until all buffer has passed through the S-Trap column.
Protein will be captured and held in the spin column's protein-binding matrix.
14. Wash captured protein by adding 150 μl binding/wash buffer and repeating centrifugation three times.
During centrifugation, rotate the S-Trap micro units 180°after each centrifugation cycle, especially when using a fixed-angle rotor.Marking the outside edge of the unit can help to recognize 180°rotation for each cycle.
15. Move S-Trap micro column to a clean 2-ml sample tube for protease digestion.
Make certain that no air bubbles are present between the protease digestion solution and the protein trap.
17. Cap the S-Trap column loosely to limit evaporation loss.
Preferably use a water bath or non-moving thermomixer.
Do not centrifuge the digestion before applying TEAB.
Apply TEAB directly into the trap containing the digestion buffer that was incubated.
22. Pool eluted peptides and dry down in the SpeedVac.Eluted peptides are then resuspended for TMT labeling (Basic Protocol 3).

TMT LABELING AND PEPTIDE CLEANUP
After protein digestion, when dealing with multiple samples, tandem mass tags (TMT) can be employed for labeling.TMT reagents utilize the principle of isotopes.Different isotopic labels are conjugated to specific amino acid residues in peptides, allowing for the labeling of peptides from different sources.The TMT labeling reagents enable simultaneous identification and quantification of proteins in diverse samples using tandem mass spectrometry.The protocol is adapted from TMTpro label reagent set user guide.
Peptide cleanup is essential for the removal of secondary metabolite contamination produced during sample preparation (Waas, Pereckas et al., 2019).These compounds can negatively impact the efficiency of subsequent phosphopeptide enrichment processes.To ensure an optimal protein yield and to prepare high-quality protein samples for mass spectrometric analysis, Basic Protocol 2 second section outlines the procedure for peptide clean-up using the Thermo Fisher Scientific prep kit. 2. Add 100 μl of 50 mM TEAB to resuspend peptides for each sample.

Materials
Mix and vortex thoroughly.
3. Add 40 μl TMT reagent, which is dissolved in 100% acetonitrile, to each buffered peptide sample.Allow this mixture to incubate at room temperature for 30 to 60 min.
When using TMTpro label reagent, apply 0.1 to 1 mg label reagent for every 10 to 100 μg protein digest.
4. To stop and acidify the labeling reaction, add 50 μl each of 5% hydroxylamine and 20% formic acid solution to each labeling reaction.
5. Take 1 μl drop from each tube to verify pH < 4 using pH paper.
If pH >4, then add dropwise more 20% formic acid to the sample.
6.After labeling, combine the samples into one tube and then proceed with the cleanup procedure.

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Current Protocols Since each sample has been labeled, combining the samples will save time in the subsequent procedures.

Peptide Cleanup
7. Take off the white cap located at the bottom of the peptide cleanup column.Gently loosen the green top cap and put it into a 2-ml microcentrifuge tube.
8. Centrifuge the column 2 min at 3000 × g, 4°C, to remove any remaining liquid from the column, and then dispose of the liquid that flows through.9. Transfer the protein digest sample, which has a total volume of ∼300 μl, into the dry peptide cleanup column.
You can continue using the same 2-ml microcentrifuge tube until step 10.
11. Add 300 μl wash solution A to the column.
13. Add 300 μl wash solution B to the column.
15. Repeat steps 7 and 8 one more time.
16. Transfer the peptide cleanup column to a fresh 2-ml microcentrifuge tube.17.Add 300 μl elution solution to the column.
19. Use a vacuum centrifuge to dry the peptide sample.After drying, take 10% of the protein sample for fractionation (Basic Protocol 8) and analysis of background proteome.The remaining sample is used to enrich phosphopeptides using Fe-NTA magnetic beads.
Set vacuum centrifuge temperature ∼30°C to avoid overheating during drying.
Depending on the centrifuge, this step could take ∼1 to 2 hr.
The dried peptide may be invisible.
The dried peptide can be stored frozen at −80°C overnight.

IMAC Fe-NTA MAGNETIC BEADS PHOSPHOPEPTIDE ENRICHMENT
Mass spectrometry is pivotal for identifying protein phosphorylation sites and quantifying changes in phosphorylation levels.Nonetheless, analyzing protein phosphorylation via MS presents challenges, such as low stoichiometry, high hydrophilicity, suboptimal ionization, and incomplete fragmentation of phosphopeptides.Due to the comparatively low occurrence of phosphorylation modifications in complex protein mixtures, it becomes essential to enrich phosphopeptides to enhance the effectiveness of MS analysis.
PTMScan phospho-enrichment IMAC Fe-NTA magnetic beads utilize immobilized metal affinity chromatography to capture phosphorylated peptides.The negatively charged phosphate groups bind to the positively charged metal ions on the beads.Combined with liquid chromatography-tandem mass spectrometry (LC-MS/MS), this method facilitates the isolation, identification, and quantification of numerous phosphorylated cellular peptides.It achieves a high level of specificity and sensitivity, offering a comprehensive snapshot of phosphorylation in cellular and tissue samples (Liu et al., 2022).Basic Protocol 5 describes the detailed steps for using magnetic beads to enrich phosphopeptides.
There may be a significant insoluble pellet, but this should not be a concern as most of the peptides will remain in solution.
3. Rotating the PTMScan IMAC Fe-NTA magnetic beads end-over-end on a rotator for 10 min to resuspend the beads.
This can be done at room temperature.
4. Cut off 2 mm from the tip of a 20-μl pipette tip using a razor blade or scissors.Use this tip to take 20 μl bead slurry and transfer to a 1.5-ml microcentrifuge tube.
5. Add 1 ml binding/wash buffer to wash the IMAC magnetic beads.Allow the beads to settle on a magnetic stand.Carefully remove the wash solution, taking care not to dislodge any beads.Wash 3 times.
6. Transfer the cleared peptide solution into the microcentrifuge tube containing IMAC beads.
Do not dislodge any pelleted material; pipet sample directly on top of the beads at the bottom of the tube to ensure immediate mixing.
7. Tighten the cap on the tube.Seal the top of the microcentrifuge tube with Parafilm to prevent any potential leakage.Incubate on the rotator 30 min at room temperature.
8. After incubation, allow the magnetic beads to settle on the magnetic stand, then discard the supernatant carefully; do not remove any beads.The depleted peptide solution may be frozen at −80°C and saved for later SMOAC enrichment (labeled W1). 9. Add 1 ml binding/wash buffer to wash beads, gently rotate the tube until the beads are completely resuspended.
10. Allow the beads to settle on the magnetic stand, then transfer the supernatant into a separate tube, labeled W2.

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Current Protocols 11.Wash two more times, transferring each subsequent supernatant into distinct tubes labeled W3 and W4, respectively.
12. Add 50 μl elution buffer, containing 50% ACN and 2.5% ammonia, to elute the phosphopeptides from the beads.Tap the bottom of the tube to resuspend the beads and then allow the beads to settle.
13. Rinse a fresh microcentrifuge tube with 0.5-ml ACN to remove any potential contaminants, vortex it, and discard the rinse.
14. Transfer the supernatant containing the eluted phosphopeptides to the rinsed tube.
16. Repeat the elution step (steps 12 to 14) once more and combine the resulting eluates.17.Dry the eluted phosphopeptides and the combined washed supernatant (W1 to W4) in a SpeedVac at 25°C.
This takes ∼4 to 5 hr due to large volume.
18. Freeze the eluted phosphopeptides at −80°C.The W1 to W4 tubes will then be used for Basic Protocol 6.

TiO 2 ENRICHMENT
The Thermo Fisher High-Select TiO 2 phosphopeptide enrichment kit offers an effective method for isolating phosphorylated peptides from complex protein digests and fractions, facilitating their analysis through mass spectrometry.TiO 2 exhibits a strong affinity for phosphopeptides, as well as other acidic peptides, and is water insoluble.Phosphopeptides can be effectively captured by TiO 2 beads in an acidic loading buffer and subsequently released using an alkaline elution buffer.This carefully optimized procedure, along with the specific buffers, enhances phosphopeptide yield, enabling direct MS analysis without requiring extra graphite or C18 purification steps.This method is adapted from the Thermo Fisher High-Select SMOAC protocol.
3. Add 20 μl wash buffer provided in the kit and centrifuge 2 min at 3000 × g, 4°C.
5. Discard the flowthrough.Keep the microcentrifuge tube for step 10.
6. Transfer the prepared TiO 2 spin tip and adaptor into a new 2-ml microcentrifuge tube.
Vortex the tube to make sure all the sample entirely dissolved.Lyophilized peptide must be entirely dissolved for optimal results.
9. Reapply sample in the microcentrifuge tube to the spin tip.Centrifuge 5 min at 1000 × g, 4°C.
If needed, keep the flowthrough for analysis.This fraction is called TiO 2 -FT.
10. Transfer the TiO 2 spin tip and adaptor into the collection tube prepared from step 5.
13. Transfer the TiO 2 spin tip and adaptor into a fresh 2-ml microcentrifuge tube.
Reordering the wash column steps leads to a notably increased occurrence of non-specific peptide binding.
16. Eliminate surplus liquid by gently blotting the bottom of the spin tip using a clean paper towel, such as a Kimwipe.17. Place the spin tip and adaptor in a new 2-ml microcentrifuge tube.
Use the same 2-ml microcentrifuge tube to combine the flowthrough.
20. Promptly dry the eluate in a SpeedVac concentrator to eliminate phosphopeptide elution buffer.
It is important to note that eluates cannot be stored in phosphopeptide elution buffer due to its high pH, which may lead to the loss of phosphopeptides.
21. Suspend the eluate with 50 μl of 0.1% formic acid for direct MS analysis.
If the starting peptide sample amount is <1 mg, suspend dried elute using 25 μl of 0.1% formic acid.
22. Combine TiO 2 -FT from step 9, with the saved wash fraction 1 and 2 from step 12 and 15.
23. Dry the combined sample by using the SpeedVac.The dried sample can be stored at −80°C until the next procedure.
Do not over dry.It is typical to observe some translucent jelly-like material at the end of dry step.
The drying procedure can take around 2-4 hours.

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Current Protocols

Fe-NTA PHOSPHOPEPTIDE ENRICHMENT
The Thermo Fisher High-Select Fe-NTA phosphopeptide enrichment kit offers a swift and effective means of isolating phosphorylated peptides, with a specificity >90%.The streamlined process allows for the enrichment of phosphopeptides from protein digests or peptide fractions, priming them for MS analysis.Each spin column, provided within the kit, is equipped with a specialized resin designed for phosphopeptide capture, boasting exceptional binding and recovery capabilities, accommodating up to 150 μg of phosphopeptides per column.This method is derived from the High-Select Fe-NTA phosphopeptide enrichment SMOAC protocol.

Materials
High To achieve the best outcomes, ensure the lyophilized peptide sample is fully dissolved in the binding/wash buffer.
If desired, use pH paper to confirm that the pH of the resuspended sample is <3.
8. Add 200 μl peptide solution to the prepared spin column and secure the screw cap.
9. Mix the resin with the sample by gently tapping the bottom plug while holding the screw cap for 10 s, ensuring the resin becomes fully suspended.
Avoid using a vortex or turning the column upside down to prevent resin from splashing against the column walls, as this could lead to an increase in nonspecific peptide attachment.
10. Incubate for 30 min at room temperature.
Gently mix the resin every 10 min as described in step 9.
11. Gently remove both the bottom plug and top screw cap.Do not compress the bottom plug, which could cause liquid to flow back into the column.
Current Protocols 12. Put the column into the microcentrifuge tube.Centrifuge 30 s at 1000 × g, 4°C.
13. Add 200 μl binding/wash buffer provided in the kit to the column to wash column.
14. Repeat wash step 13 two more times for a total of 3 washes.Discard the flowthrough.
16.To elute column, first place column in a new microcentrifuge tube.Add 100 μl elution buffer provided in the kit to the column.Centrifuge 30 s at 1000 × g, 4°C.Keep the flowthrough.Repeat this step once.
It is normal for the resin to change to a brown color during this process.
17. Immediately dry the collected eluate using a SpeedVac vacuum concentrator to eliminate the elution buffer.
High pH elution buffer will lead to loss of phosphates on phosphopeptides.
18. Add 70 μl of 0.1% formic acid to suspend the dried eluate and then direct LC-MS analysis.
If starting peptide sample amounts is <1 mg, suspend dried eluate in 40 μl of 0.1% formic acid.

HIGH pH PEPTIDE FRACTIONATION
The Thermo Fisher Pierce high pH reversed-phase peptide fractionation kit is specifically designed to improve protein identification in complex samples via liquid chromatography-mass spectrometry (LC-MS) analysis, utilizing a high-pH reversedphase chromatography approach.This method efficiently separates peptides based on their hydrophobic properties, offering a strong alternative to low-pH reversed-phase LC-MS gradients and demonstrating its distinct advantage over strong cation exchange (SCX) fractionation by eliminating the need for an additional desalting process before LC-MS analysis.The kit's protocol is a refined version of the original method used in the Pierce kigh pH reversed-phase peptide fractionation kit.

Materials
Pierce high pH reversed-phase peptide fractionation kit (Thermo Fisher, cat.no.84868) ACN, LC-MS grade (Thermo Fisher, cat.no.51101) 0.1% TFA (see recipe) H 2 O, LC-MS grade (Thermo Fisher, cat.no.51140) 2-ml protein LoBind tubes (Eppendorf cat.no.022431102) or Pierce low protein binding microcentrifuge tubes, 2-ml (Thermo Fisher, cat.no.88379 or 88380) Microcentrifuge with adjustable rotor speed up to 7000 × g SpeedVac (see Basic Protocol 2 for details) 1.Take off and discard the protective white tip from the bottom of the column supplied in the peptide fractionation kit.Then, place the column into a 2-ml tube.Centrifuge 2 min at 5000 × g, 4°C.Discard the flowthrough.
Do not exceed recommended centrifugation speeds.
2. Take off the top screw cap and add 300 μl ACN into the column.Secure the cap, place the spin column back into a 2-ml microcentrifuge tube and centrifuge 2 min at 5000 × g, 4°C.Discard flowthrough.

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Current Protocols 3. Repeat wash step 2 one more time.
Discard flowthrough.The column is now ready for use.
5. Prepare elution solutions as outlined in Table 1.
Allocate 300 μl for each solution per sample.Adjust the volume of elution solutions based on the number of samples to be fractionated if processing >3 samples.
Ensure the peptide is fully dissolved and does not contain organic solvents like ACN or DMSO.
7. Transfer the spin column to a fresh 2-ml tube.Add 300 μl sample solution onto the column, reapply the cap on top.Centrifuge 1 min at 3000 × g, 4°C.Keep eluate flowthrough fraction as "flowthrough." 8. Place the column into a new 2-ml tube.Add 300 μl of water onto the column.Centrifuge 1 min at 3000 × g, 4°C.Retain eluate as "wash" fraction.
For TMT-labeled samples, perform an extra wash using 300 μl of 5% ACN with 0.1% triethylamine (TEA; supplied with the kit) to remove any unbound TMT reagent.
10. Repeat step 9 for each elution step, using new 2-ml tubes and the respective elution solutions detailed in Table 1 for each fraction.
11. Use a SpeedVac or similar vacuum concentrator to evaporate all the liquid from each sample tube until dry.

LC-MS/MS ANALYSIS AND DATABASE SEARCH
In a typical LC-MS/MS analysis, peptides are separated based on hydrophobicity before entering mass spectrometry.The mass spectrometer operates in a "data dependent" mode where eluted peptides are isolated and fragmented inside the mass spectrometer.Accurate mass of the intact peptide and its fragments are recorded in the mass spec raw data.The raw data are then processed by a search engine to find the best match between each mass spectrum with a peptide from the designated protein database.After applying a false Current Protocols

A B C
Step 1: CST Fe-NTA 1433 Step 2: TiO2 Illustrates that all 10 samples are abundantly and evenly labeled with TMTpro labeling reagents.(C) A Venn diagram depicting how each step of the 3-step phosphopeptide enrichment process captures different quantities and groups of proteins.(D) A volcano plot presents the quantitative analysis of the global phosphopeptides dataset from the trigeminal ganglion, as identified by MS.In the treated group, the analysis revealed 61 upregulated and 33 downregulated peptides within the total proteome dataset.Peptides with fold changes >1.5 (p <.01, t-test) were identified as being significantly expressed in the treatment group.The x-axis represents the log2fold change, and the y-axis represents the -log10 FDR-adjusted p-value.
discovery rate (FDR) control, peptides with high confidence are inferred to proteins.A parsimony rule is applied so that the smallest list of proteins is reported to represent all peptides identified.Peptide quantification is based on abundance of TMT reporter ions.

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Current Protocols 1. Resuspend dry samples in an appropriate volume of solvent A before LC-MS analysis.
2. Load 1 μg tryptic digest onto a trap column and desalt it using an HPLC system at a flow rate of 4 μl/min for 5 min for each analysis.
4. Perform chromatographic separation using a two-part solvent system, with solvent A being 0.1% formic acid and solvent B combining 0.1% formic acid with 80% acetonitrile, at a flow rate of 300 nl/min.
6. Follow this by washing the column with 80% solvent B for 5 min and re-equilibrating it at 1% solvent B for 10 min before introducing the next sample.
7. Detect precursor masses in the Orbitrap with a resolution of 120,000 (m/z 200).
8. Detect higher energy collisional dissociation (HCD) fragment masses in the Orbitrap at a resolution of 50,000 (m/z 200), employing data-dependent MS/MS with a cycle time of 2 s and a dynamic exclusion period of 20 s.
9. The proteomics and phosphoproteomics data are subsequently analyzed using Proteome Discoverer (V2.5) against human protein database (Uniprot) with the following search parameters:

COMMENTARY
In a global phosphoproteomic analysis, the use of a large number of cells, tissues, or other biological material is recommended due to the inherently low levels of phosphorylated proteins in most biological systems and the potential loss occurring across multiple steps of sample preparation.Limited sample size poses a challenge in achieving optimal protein and peptide recoveries.To address this, we present a detailed method for enriching phosphoproteins, allowing for the use of limited sample size tissues while maintaining high-quality phosphoproteomic data.This method is suitable for small tissue samples while ensuring high-quality phosphoproteomic data.Our approach involves a 3-step sequential enrichment process: initially using Fe-NTA magnetic beads from Cell Signaling Technology, followed by TiO 2 enrichment, and concluding with secondary Fe-NTA phosphopeptide enrichment.

Lysis buffer
In the lysis stage, selecting the appropriate lysis buffer is critical.This buffer aids in breaking down cells or tissues for protein extraction.Both the type and volume of the buffer might require adjustments based on the initial sample and the target protein concentration.Factors to consider include the cell or tissue type and size, along with the need for effective protein extraction.In our study, we evaluated three different lysis buffers for trigeminal ganglion tissue: T-per (Thermo Fisher Scientific, cat.no.78510) with protease/phosphatase inhibitors, EasyPep TM Mini MS Sample Prep Kit lysis buffer (Thermo Fisher Scientific, cat.no.A40006), and a 5% SDS solution.Following the application of these lysis buffers, we proceeded with the Basic Protocol 1 for protein digestion and then assessed the protein yield.Our findings indicate that in the case of neuronal tissue, a 5% SDS lysis buffer is most effective.Further details on these findings will be provided in the Understanding Results section.

Protein concentration test
Determining protein concentration is a critical step in preparing samples for various downstream applications.Although this protocol does not delve deeply into the specific method used for determining protein concentration, it acknowledges the variability in available methods.Researchers often use techniques such as the BCA assay to quantify protein content.

Protein cleanup
Protein cleanup is crucial for removing contaminants and substances that might interfere with subsequent analyses.The timing of this cleanup, either before or after TMT labeling, is flexible.However, performing cleanup after TMT labeling can improve efficiency by allowing the combination of labeled samples into a single tube.This step streamlines the process, simplifying downstream processing and analysis for a more efficient workflow.

Phosphopeptide enrichment
When considering the sequence of phosphopeptide enrichment methods, it is advantageous to employ the CST Fe-NTA magnetic Beads method before utilizing the TiO 2 and SMOAC methods.This is primarily because

Troubleshooting
Please see Table 2 a troubleshooting guide for proteomic and phosphoproteomics analysis for limited tissue.

Understanding Results
This protocol outlines a workflow for preparing limited protein tissue samples, applicable to other neuronal tissues.We conducted tests with various lysis buffers, as shown in Figure 1A and observed that different lysis buffers yielded varying protein quantities.When we employed T-per and added proteases, it resulted in a total recovery of 1995 proteins.By using a miniprep kit from Thermo Fisher, we successfully identified 1283 proteins.The utilization of a 5% SDS lysis buffer allowed us to identify a total of 3662 proteins.The Venn diagram illustrates the overlap and unique proteins identified with different lysis buffers.Consequently, for neuronal tissues, such as trigeminal ganglion or dorsal root ganglion, we recommend using a 5% SDS lysis buffer.
Following protein digestion and cleanup, TMT labeling is executed to consolidate the protein samples for subsequent workflow steps.This process is vital for ensuring efficient downstream analysis.Post-labeling, we routinely assess the effectiveness of TMT labeling.Typically, we analyze 1 μg of the labeled sample to confirm that each sample has been sufficiently and uniformly labeled, as illustrated in Figure 1B.Ensuring even labeling of samples is crucial, as it significantly impacts the accuracy and reliability of subsequent experimental stages, including quantitative analysis and the detection of subtle changes in protein expression across different samples.
In our protocol, we outline a threestep enrichment process aimed at improving phosphopeptide identification.We successfully identified a total of 4454

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Current Protocols phosphopeptides.Figure 1C details the number of phosphopeptides identified via each enrichment step.The initial Fe-NTA method led to the identification of 1433 phosphopeptides, followed by the TiO 2 step, which uncovered an additional 199 phosphopeptides.The third step, employing Fe-NTA, revealed 62 more phosphopeptides.There is a final fractionation step of the combined sample from the above three-step enrichments, resulting in a total of 4454 phosphopeptides.Each method uniquely captures different proteins, thereby enhancing the overall yield in phosphoproteomics.Furthermore, although not illustrated in the figure, we identified a total of 2923 phosphoproteins, corresponding to 13,460 peptide groups.
The volcano plot in Figure 1D displays the global phosphoproteome data from trigeminal ganglion.It plots log2-fold changes on the x-axis against p-values on the y-axis.Using high-sensitivity LC-MS/MS for proteomics and phosphoproteomics, we identified peptides and phosphopeptides with differential expression in the trigeminal ganglia.A peptide is deemed differentially expressed if it exhibits a fold change >1.5 and a p-value <.05.The plot, generated by Proteome Discoverer 2.5, shows the overall peptide profile, highlighting peptides with significant increases or decreases.In the treated group, analysis revealed 61 upregulated and 33 downregulated peptides within the total proteome dataset.

Time Considerations
Basic Protocol 1 takes ∼1 to 2 hr, including the BCA assay process.Basic Protocol 2 requires ∼4 to 5 hr, which includes the drying process.Drying times may vary depending on the equipment used.Basic Protocols 3 and 4 together take ∼1 to 2 hr.Basic Protocol 5 lasts 5 to 6 hr and incorporates the drying process.Basic Protocol 6 is completed in ∼2 to 3 hr, including the drying process.Basic Protocol 7 takes ∼4 to 5 hr including drying process.Basic Protocol 8 takes ∼6 hr.
Current Protocols

Figure 1
Figure1(A) Shows how different lysis buffers yield varying amounts of proteins from trigeminal ganglion tissue.(B) Illustrates that all 10 samples are abundantly and evenly labeled with TMTpro labeling reagents.(C) A Venn diagram depicting how each step of the 3-step phosphopeptide enrichment process captures different quantities and groups of proteins.(D) A volcano plot presents the quantitative analysis of the global phosphopeptides dataset from the trigeminal ganglion, as identified by MS.In the treated group, the analysis revealed 61 upregulated and 33 downregulated peptides within the total proteome dataset.Peptides with fold changes >1.5 (p <.01, t-test) were identified as being significantly expressed in the treatment group.The x-axis represents the log2fold change, and the y-axis represents the -log10 FDR-adjusted p-value.

Table 1
Preparation of Elution Solutions for Thermo Scientific TMT-Labeled Peptides

Table 2
Troubleshooting Guide for Proteomic and Phosphoproteomics Analysis for Limited Tissue