Our Core Facility Resources

NIH Grant Materials

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Facilities and Resources (.docx)

Major Equipment (.docx)

Frequently Asked Questions

How much protein is needed?

What post-translational modifications can be analyzed?

Can the amount of each protein present be quantitated?

Where are protocols that should/will be used?

How should I store the sample?

What are the capabilities of each different machine?

How can I check on the progress of my analysis?

What is a “normal” time for each phase of data collection and analysis?


I.  How much protein is needed?

It depends on the question. Generally 10 pmol per protein is sufficient for identification.

To identify a purified protein requires 10 pmol of the protein.

To characterize an interactome you should start with 0.5-1 µg total protein IP’d from 5-10 mg lysate.

To determine post-translational modifications may require 1 µg purified protein depending on the abundance of the modification.

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II.  What post-translational modifications can be analyzed?

We can routinely determine PTMs such as phosphorylation, ubiquitination, methylation, acetylation, hydroxylation, Met/Cys(S-oxidation), and limited proteolytic processing. Detection of other modifications may be possible so be sure to ask.

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III. Can the amount of each protein present be quantitated?

Quantitative proteomics can be separated into two major approaches, i) the use of stable isotope labeling and ii) label-free techniques. Common labeling techniques involve modifying peptides with isobaric tags (TMT or iTRAQ), labeling proteins with isotope-coded affinity tags (ICAT), or in vivo metabolically labeling proteins by incorporation of stable isotope labels with amino acids in cell culture (SILAC). For label-free approaches, protein/peptide quantification can be achieved based on peptide ion intensities, or the total number of peptide spectral counts, whereas with labeling approaches quantification is performed by comparing the signal ratio of unlabeled versus isotopically labeled peptide standards or SILAC labeled proteins, We have extensively compared and developed a variety of quantitative technologies and have published several manuscripts based on these studies over the past five years. The advantages and disadvantages of these methods have been recently compared in a review article (Zhou JY, Hanfelt J, Peng J. Clinical proteomics in neurodegenerative diseases. Proteomics Clin Appl. 2007;1:1342-50).

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IV. Protocols that should/will be used.

  1. Tissue homogenization
  2. IP methods
  3. TMT labeling methods
  4. Electrostatic Repulsion-Hydrophilic Interaction Chromatography (ERLIC) Fractionation
  5. LC/MS analysis
  6. Identification and spectral count based relative quantitation with Proteome Discoverer
  7. Identification and label-free LFQ based relative quantitation with MaxQuant.
  8. TMT Data Analysis

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  1. Tissue homogenization

(METHOD 1):

The sample was vortex in 300 uL of urea lysis buffer (8M urea, 100 mM NaHPO4, pH 8.5), including 3 uL (100x stock) HALT protease and phosphatase inhibitor cocktail (Pierce). All homogenization was performed using a Bullet Blender (Next Advance) according to manufacturer protocols. Protein supernatants were transferred to 1.5 mL Eppendorf tubes and sonicated (Sonic Dismembrator, Fisher Scientific) 3 times for 5 s with 15 s intervals of rest at 30% amplitude to disrupt nucleic acids and subsequently vortexed. Protein concentration was determined by the bicinchoninic acid (BCA) method, and samples were frozen in aliquots at −80°C. Protein homogenates (100 ug) were diluted with 50 mM NH4HCO3 to a final concentration of less than 2M urea and then treated with 1 mM dithiothreitol (DTT) at 25°C for 30 minutes, followed by 5 mM iodoacetimide (IAA) at 25°C for 30 minutes in the dark. Protein was digested with 1:100 (w/w) lysyl endopeptidase (Wako) at 25°C for 2 hours and further digested overnight with 1:50 (w/w) trypsin (Promega) at 25°C. Resulting peptides were desalted with a Sep-Pak C18 column (Waters) and dried under vacuum.

(METHOD 2):

Each tissue piece was homogenized in 500 uL of urea lysis buffer (8M urea, 100 mM NaH2PO4, pH 8.5), including 5 uL (100x stock) HALT protease and phosphatase inhibitor cocktail (Pierce). All homogenization was performed using a Bullet Blender (Next Advance) according to manufacturer protocols. Briefly, each tissue piece was added to Urea lysis buffer in a 1.5 mL Rino tube (Next Advance) harboring 750 mg stainless steel beads (0.9-2 mm in diameter) and blended twice for 5 minute intervals in the cold room (4°C). Protein supernatants were transferred to 1.5 mL Eppendorf tubes and sonicated (Sonic Dismembrator, Fisher Scientific) 3 times for 5 s with 15 s intervals of rest at 30% amplitude to disrupt nucleic acids and subsequently vortexed. Protein concentration was determined by the bicinchoninic acid (BCA) method, and samples were frozen in aliquots at −80°C. Protein homogenates (100 ug) were diluted with 50 mM TEAB to a final concentration of less than 2M urea and then treated with 1 mM dithiothreitol (DTT) at 25°C for 30 minutes, followed by 5 mM iodoacetimide (IAA) at 25°C for 30 minutes in the dark. Protein was digested with 1:100 (w/w) lysyl endopeptidase (Wako) at 25°C for 2 hours and further digested overnight with 1:50 (w/w) trypsin (Promega) at 25°C. Resulting peptides were desalted with a Sep-Pak C18 column (Waters).  An aliquot equivalent to 20 ug of total protein was taken out of each sample and combined to obtain a global internal standard (GIS) use later for TMT labeling (the total was split into 8 aliquots equivalent to 70ug of total protein).  An addition aliquot of 10 ug was taken from each sample to test the digestion efficiency and overall peptide composition.  All samples were dried down to complete using a Savant SpeedVac (ThermoFisher Scientific, San Jose, CA).

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  2. IP Methods:

Immunoprecipitations should be conducted, and the beads washed, according to the manufacturers Instructions. The IP beads are spun down and residual urea is removed.  Avoid any buffers with strong detergents, although little Triton X-100 is fine. Digestion buffer (200 ul of 50 mM NH4HCO3) is added and the bead solution is then treated with 1 mM dithiothreitol (DTT) at 25°C for 30 minutes, followed by 5 mM iodoacetimide (IAA) at 25°C for 30 minutes in the dark. Proteins are digested with 1ug of lysyl endopeptidase (Wako) at room temperature for 2 hours and further digested overnight with 1:50 (w/w) trypsin (Promega) at room temperature. Resulting peptides are desalted with a Sep-Pak C18 column (Waters) and dried under vacuum.

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  3. TMT Labeling Methods:

The samples were randomized over 4 TMT 10-plex batches.  In each batch, the GIS samples took up channels 1 and 10 (TMT-126 and TMT-131, respectively).  Labeling was performed according to the manufacturer’s protocol.  Briefly, the reagents were allowed to equilibrate to room temperature.  Dried peptide samples (70ug each) were resuspended in 100ul of 100mM TEAB buffer (supplied with the kit).  Anhydrous acetonitrile (41 ul) was added to each labeling reagent tube and the peptide solutions were transferred into their respective channel tubes.  Please see labeling table for exact sample to channel match.  The reaction was incubated for 1 hour and quench for 15 minutes afterwards with 8 ul of 5% hydroxylamine.  All samples per batch were then combined and dried down.

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   4.  Electrostatic Repulsion-Hydrophilic Interaction Chromatography (ERLIC) Fractionation:

Peptides were resuspended in 100ul of 90% acetonitrile and 0.01% acetic acid.  The entire sample was loaded onto an offline electrostatic repulsion-hydrophilic interaction chromatography (ERLIC) fractionation HPLC system and 40 fractions were collected over a time of 40 minutes.  The fractions were combined into 20 and dried down.

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  5. LC-MS/MS analysis:

Derived peptides were resuspended in peptide 10 uL of loading buffer (0.1% formic acid, 0.03% trifluoroacetic acid, 1% acetonitrile). Peptide mixtures (2 uL) were separated on a self-packed C18 (1.9 um Dr. Maisch, Germany) fused silica column (25 cm x 75 uM internal diameter (ID); New Objective, Woburn, MA) by a Dionex Ultimate 3000 RSLCNano and monitored on a Fusion mass spectrometer (ThermoFisher Scientific , San Jose, CA). Elution was performed over a 120 minute gradient at a rate of 300nl/min with buffer B ranging from 3% to 80% (buffer A: 0.1% formic acid in water, buffer B: 0.1 % formic in acetonitrile). The mass spectrometer cycle was programmed to collect at the top speed for 3 second cycles. The MS scans (400-1600 m/z range, 200,000 AGC, 50 ms maximum ion time) were collected at a resolution of 120,000 at m/z 200 in profile mode and the HCD MS/MS spectra (2 m/z isolation width, 30% collision energy, 10,000 AGC target, 35 ms maximum ion time) were detected in the ion trap. Dynamic exclusion was set to exclude previous sequenced precursor ions for 20 seconds within a 10 ppm window. Precursor ions with +1, and +8 or higher charge states were excluded from sequencing.

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  6. Data Analysis:

Spectra were searched using Proteome Discoverer 2.0 against human Uniprot database (90,300 target sequences).  Searching parameters included fully tryptic restriction and a parent ion mass tolerance (± 20 ppm).  Methionine oxidation (+15.99492 Da), asaparagine and glutamine deamidation (+0.98402 Da) and protein N-terminal acetylation (+42.03670) were variable modifications (up to 3 allowed per peptide); cysteine was assigned a fixed carbamidomethyl modification (+57.021465 Da). Percolator was used to filter the peptide spectrum matches to an false discovery rate of 1%.

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  7. MaxQuant Data Analysis for label-free quantitation:

RAW data for the samples was analyzed using MaxQuant v1.5.2.8 with Thermo Foundation 2.0 for RAW file reading capability. The search engine Andromeda, integrated into MaxQuant 1, was used to build and search a concatenated target-decoy Uniprot human reference protein database (retrieved April 20, 2015; 90,411 target sequences), plus 245 contaminant proteins from the common repository of adventitious proteins (cRAP) built into MaxQuant. Methionine oxidation (+15.9949 Da), asparagine and glutamine deamidation (+0.9840 Da), and protein N-terminal acetylation (+42.0106 Da) were variable modifications (up to 5 allowed per peptide); cysteine was assigned a fixed carbamidomethyl modification (+57.0215 Da). Only fully tryptic peptides were considered with up to 2 miscleavages in the database search. A precursor mass tolerance of ±20 ppm was applied prior to mass accuracy calibration and ±4.5 ppm after internal MaxQuant calibration. Other search settings included a maximum peptide mass of 6,000 Da, a minimum peptide length of 6 residues, 0.05 Da tolerance for high resolution MS/MS scans. Co-fragmented peptide search was enabled to deconvolute multiplex spectra. The false discovery rate (FDR) for peptide spectral matches, proteins, and site decoy fraction were all set to 1 percent. Quantification settings were as follows: requantify with a second peak finding attempt after protein identification has completed; match MS1 peaks between runs; a 0.7 min retention time match window was used after an alignment function was found with a 20 minute RT search space. Quantitation of proteins was performed using summed peptide intensities given by MaxQuant. The quantitation method only considered razor plus unique peptides for protein level quantitation. The full list of parameters used for MaxQuant are available as MSSM_Proteomics_PFC_SEARCHPARAMS.xml accompanying the public release.

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  8. TMT Data Analysis:

MS/MS spectra were searched against a Uniprot curated mouse database (downloaded on 03/06/2015 with 53289 target sequences) with Proteome Discoverer 2.1 (ThermoFisher Scientific, San Jose, CA).  Methionine oxidation (+15.9949 Da), asparagine and glutamine deamidation (+0.9840 Da), and protein N-terminal acetylation (+42.0106 Da) were variable modifications (up to 3 allowed per peptide); static modifications included cysteine carbamidomethyl (+57.0215 Da), peptide n-terminus TMT (+229.16293 da) and lysine TMT (+229.16293 Da). Only fully tryptic peptides were considered with up to 2 miscleavages in the database search. A precursor mass tolerance of ±20 ppm and a fragment mass tolerance of 0.6 Da were applied.  Spectra matches were filtered by Percolator to a psm fdr of less than 1 percent.  Only razor and unique peptides were used for abundance calculations.  Ratio of sample over the GIS of normalized channel abundances were used for comparison across all samples.

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V. How should I store the sample?

In general, cell pellets or solution samples should be stored frozen. Gel slices should be stored in destain buffer at 4˚C. For IPs or other affinity methods the beads should be stored frozen in wash buffer.

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VI. What are the capabilities of each different machine?

The most economical analysis is run on the Orbitrap XL. If you only need to identify a protein band, or analyze several thousand peptides to get your answer, this is the machine you want.

The Q-Exactive Orbitrap can sequence about three times more peptides per run and excels in quantitative analysis.

The top-of-the-line Fusion can sequence 25,000 or so peptides per run and has a precision that is ideal for quantification of isobaric tags and other isotopic labels.

Your individual consultation will include a recommendation on which machine would be optimal for your needs.

 

Orbitrap XL

Q-Exactive Plus

Orbitrap Fusion

TSQ Vantage

Fragmentation

CID, HCD

HCD

CID, HCD, ETD

Electrospray

Resolution

100,000 @ 200 m/z

140,000 @ 200 m/z

450,000 @ 200 m/z

7500 @ 508 m/z

Speed

4 peptides/sec

10 peptides/sec

19 peptides/sec

Sensitivity

Peptides/run (approx.)

5,000

15,000

25,000

Optimal Application

Routine Protein ID from gel bands, spots, and simple mixtures (IPs and cellular fractions) Stable PTMs from purified samples

High resolution identification and quantification. Label-free Quantitation

Highest resolution and  sensitivity, labile PTMs, complex mixtures (total cell or tissue lysates) and iTRAQ, TMT, SILAC, ICAT

Selective Ion Monitoring (SIM), Selective/Multiple Reaction Monitoring (SRM/MRM)

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VII. How can I check on the progress of my analysis?

When your sample is ready for analysis we will notify you of the scheduled run time and machine(s) that will be used. Total time required depends on the machine used (see below). A calendar available in PPMS will show the schedule for sample runs on each of the Core’s machines.

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VIII. What is a “normal” time for each phase of data collection and analysis?

We strive to return your results in 2-4 weeks. However, since machines break down, staff members get sick, go on vacation, etc., there may be times when we cannot meet this deadline. We will do our best to keep you updated on the progress of your sample.

Step

Days after sample submission

Orbitrap XL

Q-Exactive

Fusion

Phase 1, Digestion

7

7

7

Phase 1, Additional sample processing (if needed)

10

10

10

Phase 2, MS data collection

10

24

24

Phase 3, Data analysis

13

27

27

Phase 4, Report to the investigator

14

28

28

 

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IX. How much will this cost?

Our fees include routine sample preparation and data analysis. Isobaric labeling or enrichment of post-translational modifications will incur a separate charge, as will sophisticated data analysis and presentation. The current costs of all services are posted here.

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