Protocols and Reagents

Directions for in-gel tryptic digestions of coomassie-stained 1D Bands and 2D Spots.

NOTE: Although nearly any SOS-PAGE system can be utilized upstream of an LC-MS analysis, the DPCF recommends lnvitrogen's NuPAGE Bis-Tris mini-gel system. A good general purpose gel covering a large MW range (6-200 kDa) is the 4-12% gradient gel (Cat. No NP0321) using MES running buffer. These pre­ cast gels provide excellent resolution, fantastic staining sensitivity (10 ng BSA using colloidal blue staining kit, Cat. No. LC6025), and are highly compatible with downstream LC-MS analysis. 

Important things to avoid keratin contamination:

1. Any gel manipulation prior to trypsin digestion should be done in a BSC or laminar flow hood.
2. Wear nitrile (not latex) gloves.
3. Wear a lab coat and make sure there is no gap between your coat sleeve and the gloves (lab tape works well).

Procedure:

1. After aspirating off liquid that sample is stored in, cut into equal size pieces using either a scalpel (cutting plug on the side of the Epi tube) or pinch it between the tongs of small tweezers to break it apart.
    
2. Wash gel pieces with 200-500 µL (depending on the volume of the gel pieces) of a mix of 40% acetonitrile and 60% 50 mM ammonium bicarbonate ("AmBic"), pH 8, for 15-30 minutes on rotator. Repeat with a second or third wash if necessary to remove stain.
    
3. Discard washes and shrink pieces with neat acetonitrile (pieces will turn white and stick together). Discard acetonitrile and dry pieces with tubes open at S0°C.
    
4. Reduction and Alkylation: Add 100 µL of 10 mM dithiothreitol stock (15.4 mg OTT in 10 ml 50 mM AmBic). Incubate 30 min at 80°C. Add 100 µL of 55 mM iodoacetamide stock (102 mg IAA in 10 ml 50 mM AmBic). Incubate 20 min at room temperature in the dark. Use Sigma OTT (D5545) and IAA (11149).
    
5. Remove excess reduction and alkylation mix; wash gel pieces twice with 500 µL aliquots of 50 mM AmBic. Remove excess.
    
6. Shrink gel pieces with 200 µL neat acetonitrile (gel pieces will again shrink, turn white, and stick together). Remove acetonitrile. Swell in 200 µL 50 mM AmBic. Remove AmBic. Repeat shrink and swell step.
    
7. Shrink gel pieces with 200 µL neat acetonitrile, and remove acetonitrile. Dry gel pieces with tubes open at S0°C briefly.
    
8. Swell gel pieces in 50 mM AmBic containing 10 ng/ul trypsin. Use Promega sequencing grade modified trypsin (porcine). Prepare stock from 20 µg vial at O.lµg/µL by diluting with 50 mM AmBic. This can be stored at -20°C for 6 months. Dilute this stock 1:10 with 50 mM AmBic pH 8.0 to give 10 ng/µL. Gel pieces should be just covered, but not in a large excess of volume (for a single 2D gel spot, use 25-30 µL of 10 ng/µL trypsin).

9. Digest overnight for 16-18 hours at 37°C.
    
10. Following digestion, centrifuge condensate to bottom of vial. Add extraction solution of 1% formic acid, 2% acetonitrile in water. The amount needed will vary with the gel volume, but typically you can use the same amount as the volume of trypsin solution you added (25 to 30 µL for a 2D gel spot). Vortex pieces in extraction solution occasionally over 30 minutes (optional: after adding extraction solution, sonicate the gel for 5 minutes. Be careful of contamination).
     
11. Remove supernatant containing peptides and pipette into LC vial. Depending on amount of protein expected, more peptides can sometimes be recovered by shrinking gel pieces in ~so µL of neat acetonitrile, then adding this to the LC vial. Speed vac to dryness and resuspend in LC buffer (0.2% formic acid, 2% acetonitrile in water) if last acetonitrile shrinking step is used.

In-gel digest protocol is based on the procedure from M. Mann's lab: "Femtomole sequencing of proteins from polyacrylamide gels by nano-electrospray mass spectrometry." M. Wilm, A. Shevchenko, T. Houthaeve, S. Breit, L. Schweigerer, T. Fotsis, M. Mann. Nature. 379, 1996, 466-469.

Directions for in-gel tryptic digestions of coomassie-stained 1D Bands and 2D Spots - abbreviated

NOTE: Although nearly any SDS-PAGE system can be utilized upstream of an LC-MS analysis, the DPCF recommends Invitrogen’s NuPAGE Bis-Tris mini-gel system. A good general purpose gel covering a large MW range (6-200 kDa) is the 4-12% gradient gel (Cat. No NP0321) using MES running buffer. These pre-cast gels provide excellent resolution, fantastic staining sensitivity (10 ng BSA using colloidal blue staining kit, Cat. No. LC6025), and are highly compatible with downstream LC-MS analysis. Additional product information

A couple of very important things to avoid keratin contamination:

1. Any gel manipulation prior to trypsin digestion should be done in a BSC or laminar flow hood.
    
2. Wear nitrile (not latex) gloves.
    
3. Wear a lab coat and make sure there is no gap between your coat sleeve and the gloves (lab tape works well).

Procedure:

1. After aspirating off liquid that sample is stored in, cut into equal size pieces using either a scalpel (cutting plug on the side of the Epi tube) or pinch it between the tongs of small tweezers to break it apart.
    
2. Wash gel pieces with 200-500 μL (depending on the volume of the gel pieces) of a mix of 40% acetonitrile and 60% 50 mM ammonium bicarbonate (“AmBic”), pH 8, for 15-30 minutes on rotator. Repeat with a second or third wash if necessary to remove stain.
    
3. Discard washes and shrink pieces with neat acetonitrile (pieces will turn white and stick together). Discard acetonitrile and dry pieces with tubes open at 50°C.
    
4. Swell gel pieces in 50 mM AmBic containing 10 ng/uL trypsin. Use Promega sequencing grade modified trypsin (porcine). Prepare stock from 20 μg vial at 0.1μg/μL by diluting with 50 mM AmBic. This can be stored at -20°C for 6 months. Dilute this stock 1:10 with 50 mM AmBic pH 8.0 to give 10 ng/μL. Gel pieces should be just covered, but not in a large excess of volume (for a single 2D gel spot, use 25-30 μL of 10 ng/μL trypsin).
    
5. Digest overnight for 16-18 hours at 37°C.
    
6. Following digestion, centrifuge condensate to bottom of vial. Add extraction solution of 1% formic acid, 2% acetonitrile in water. The amount needed will vary with the gel volume, but typically you can use the same amount as the volume of trypsin solution you added (25 to 30 μL for a 2D gel spot). Vortex pieces in extraction solution occasionally over 30 minutes (optional: after adding extraction solution, sonicate the gel for 5 minutes. Be careful of contamination).
    
7. Remove supernatant containing peptides and pipette into LC vial. Depending on amount of protein expected, more peptides can sometimes be recovered by shrinking gel pieces in ~50 μL of neat acetonitrile, then adding this to the LC vial. Speed vac to dryness and resuspend in LC buffer (0.2% formic acid, 2% acetonitrile in water) if last acetonitrile shrinking step is used.

In-gel digest protocol is based on the procedure from M. Mann’s lab: “Femtomole sequencing of proteins from polyacrylamide gels by nano-electrospray mass spectrometry.” M. Wilm, A. Shevchenko, T. Houthaeve, S. Breit, L. Schweigerer, T. Fotsis, M. Mann. Nature. 379, 1996, 466-469.

A couple of very important things to avoid keratin contamination:

1. Any sample manipulation prior to trypsin digestion should be done in a BSC or laminar flow hood.
    
2. Wear nitrile (not latex) gloves.
    
3. Wear a lab coat and make sure there is no gap between your coat sleeve and the gloves (lab tape works well).

Procedure:

1. Perform Bradford assay to determine protein concentration of each solution. Normalize concentrations for samples within a group using 50 mM AmBic. Aim for a protein concentration of between 0.1 and 1 μg/μL at the end of this protocol.
    
2. In order to maximize solubility of proteins, add calculated volume of Waters Rapigest to have 0.1-0.2% Rapigest in final concentration pre-digestion. Make up Rapigest with 50 mM AmBic.
    
3. Heat at 40°C while shaking for 10 minutes. Spin down condensate.
4. Make up 100 mM DTT in 50 mM AmBic for reduction step. Add DTT to each solution to make the final DTT concentration 10 mM.
    
5. Heat solution at 80°C while shaking for 15 minutes.
    
6. Remove from heat and cool for 5 minutes (to room temp). Spin down condensate.
    
7. Make up 200 mM iodoacetamide (IAM) in 50 mM AmBic for alkylation step. Add IAM to each solution to make the final IAM concentration 20 mM (2X molar excess of DTT).
    
8. Incubate the solutions in the dark at room temperature for 30 minutes.
9. Add trypsin to each solution at 1:50 trypsin:protein concentration. Make trypsin stock in 50 mM AmBic.
    
10. Digest for at least 4 hours or overnight at 37°C while shaking.
     
11. Following digestion, centrifuge condensate to bottom of vial. Add TFA and MeCN to give 0.5-1.0% TFA and 2% MeCN by volume.
     
12. Spike in an amount of internal standard, yeast alcohol dehydrogenase (ADH1_YEAST), that would yield approximately 50 fmols on column. Best practice indicates that spiking in at least volume of 5 uL of ADH (10 uL preferably) is optimal. Typical loading for a 75um i.d. column is 0.75-1.0 μg total digested protein.
     
13. Shake samples for 2 hours at 60°C. Centrifuge at 15,000 rpm for 5 min, and pipette supernatant into an autosampler vial.

Mitochondrial Purification via Percoll Gradient from Adherent Cells Grown In Culture

Base Reference: Kristian et al, J. Neurosc. Methods 152:136‐143 (2006)

Materials

1. Homogenization buffer (HB): 210mM mannitol, 70mM sucrose, 5mM HEPES pH 7.12 at 25oC. Works fine using HEPES diluted from 1M stock (Gibco cat # 15630)
    
2. Homogenization buffer supplemented (HBS) with EGTA at a final concentration of 1mM and Roche’s Complete Mini EDTA‐free protease inhibitor cocktail (cat # 11 836 170 001) at a final concentration of 1X (1 tablet dissolved in 10ml of buffer)
    
3. 50% Percoll: Percoll *(Sigma cat # P1644) diluted 1 to 1 with 2X HB.
    
4. 22% and 15% Percoll solutions: The 50% Percoll stock is used to prepare all other Percoll solutions; to that effect dilutions of the 50% Percoll solution are done in HBS.
    
5. EGTA 1M pH 7.5 stock

*Sigma Percoll solution is considered to be a 100% solution

Procedures:

1. Grow cells to approx 85% confluence in media of choice. For N2a cells the procedures are described for cells grown in 4 tissue culture plates with a diameter of 15cm2.
    
2. Transfer plates to a bed of ice
    
3. Wash each plate twice with 8 ml of ice cold HB.
    
4. Collect the cells by scraping with a cell culture policemen using 1ml of HBS per plate. Repeat the collection once more (some times it is necessary to conduct a third scraping, depending of how well the cells adhere to their substrate. A simple visual inspection indicates if a third collection is necessary). Combine all collected cells. It is best to collect directly in the Potter homogenizer.
    
5. Homogenizer using a Potter Homogenizer whose pestle is driven at an estimated 2400 rpm by a GT motor control (Glass‐Col, Terre Haute, IN) at 40% capacity and a Glass‐Col Continue Duty DC motor (cat # 099C_K44). Use 20 strokes of the pestle in a period of 3 min, maintaining the homogenizer tube in ice. Repeat once.
    
6. Spin the homogenate at 1,500g for 3 min
    
7. Collect the supernatant and spin it at 13,000g for 17 min.
    
8. Resuspend the pellet in 1.4 ml of HBS
    
9. Prepare two 5ml centrifuge tubes with a discontinued Percoll gradient as follows: 1ml of 50% Percoll solution as cushion followed by layering 3ml of a 22% Percoll solution.
    
10. To the resuspended pellet add 0.3 ml of 50% Percoll (final sample should have a concentration of Percoll of 15%.) and layer 1ml of the 15% Percoll sample on top of the described 50%‐22% gradient. Use 15% Percoll to help balance the centrifuge tubes before centrifugation.
     
11. Spin at 30,700g for 6min.
     
12. Recover the mitochondria from the 50% ‐ 22% Percoll interface
     
13. Wash the mitochondria by diluting 1 to 10 with HBS at 15,600g for 30 min. Repeat the wash step to decrease the amount of percoll, protease inhibitors and EGTA to 0.25X, or use the buffer of choice depending on the needs of the experiment at hand.

Address question and/or comments to

Mirta Mihovilovic, Ph.D.
Senior Scientist
Duke University Medical Center
mihov001@mc.duke.edu

Mass Spectrometry Appropriate Protocol for Cell Lysis using Probe Sonication

1. Wash cell pellet 3x with 10 volumes of 50 mM Ammonium Bicarbonate. Pellet cells gently between washes by centrifuging at ~5000 rcf for 3 minutes. Carefully pipette supernatant off after final wash, so as to not disturb the cells.
    
2. Add 5 volumes of lysis buffer, and vortex to suspend. The minimum volume for probe sonication is 100 uL in a 0.5 mL eppendorf tube. Standard lysis buffer for proteomics applications is 0.1%‐0.25% w/v Rapigest SF (www.waters.com) in 50 mM Ammonium Bicarbonate. To increase coverage for membrane‐bound and integral membrane proteins, substitute with 0.5% w/v PPS Silent Surfactant (www.proteindiscovery.com) in 50 mM Ammonium Bicarbonate.
    
3. Place samples on ice.
    
4. Perform probe sonication with 3 cycles of 15 seconds on, 5 seconds off, at 20% power. If possible, keep samples on ice during sonication to prevent excessive heating.
    
5. Centrifuge samples at 15,000 rcf for 5 minutes, and measure protein concentration of the supernatant using a mini Bradford (www.bio‐rad.com) or micro BCA (www.piercenet.com) assay, depending upon the lysis buffer composition.

Protocol for Post Capture Sample Processing of SNO RAC and Acyl RAC Proteomics Samples

This protocol is utilized in the procedures of the following manuscripts, and has been optimized for use with ThiopropylSepharose 6B (GE Healthcare) resin:

The procedure is scaled for approximately 20 uL of TPS resin. Scale volumes appropriately if using more or less resin.

After incubation to capture free thiols:

1. Wash resin 3x with 1 mL 50mM Ammonium Bicarbonate , pH 8 (AmBic), removing the supernatant each time with spinning at 12000 rcf in a micro swinging‐bucket centrifuge for 2 minutes.
    
2. Add 100 uL of 0.25% ALS‐1 AmBic.
    
3. Add 5 uL of 0.1 ug/uL Promega Sequencing Grade Trypsin (0.5 ug).
    
4. Incubate overnight at 37C, 900 rpm on Thermomixer R. The speed may need to be increased depending on the quantity of resin and volume of liquid. The goal is to keep the resin suspended during trypsinization.
    
5. Spin at 12000 rcf in a micro swinging‐bucket centrifuge and remove supernatant in the morning and set aside. Supernatant may be analyzed directly by LC‐MS/MS in cases where the non‐Cys‐containing peptides from the bound proteins are of interest.
    
6. For all of the following wash steps, add 250 uL of listed buffer, vortex for a few seconds, spin at 12000 rcf in the swinging bucket centrifuge for 2 minutes, and then discard supernatant before next wash, but use caution not to pipette too close to resin bed. A gel‐loader tip with vacuum suction can be an alternative way to decant the supernatants (again, be careful not to suction up the resin!).
   • a. Wash resin 3x with HENS buffer (250 mM HEPES pH 7.7, 1mM EDTA, 0.1mM Neocuproine, 1%SDS).
   • Wash resin 3x with 50 mM AmBic, pH 8.
   • Wash resin 3x with 80% ACN/0.1% TFA.
   • Wash resin 3x with 50% MeOH.
   • Wash resin 3x with 50 mM AmBic, pH8.
      
7. Elute thiol‐containing peptides by adding 100 uL of 20 mM dithiothreitol in AmBic and incubate at 37C on Thermomixer R (900 rpm) for 45 minutes. 
    
8. Spin to pellet the resin, then pipette off peptide‐containing supernatant into low bind Epi tube.
    
9. Rinse resin with 20 uL of 50 mM AmBic, vortex, spin, and add supernatant from wash to peptides in step 9.
    
10. Alkylate with 100 uL of 80 mM iodoacetamide by incubation at room temperature in the dark for 30 min.
     
11. Reduce the volume of the samples in speed vac to approximately 20 uL, then perform a C18 Zip Tip (Millipore) cleanup of the peptides, following the manufacturers protocol.
     
12. Dry Zip Tip eluent and resuspend in normal LC/MS buffer. If protocol was performed properly, the peptides in the SNO‐RAC or Acyl‐RAC sample should be 95+% Cys‐containing peptides.

Reagents

Arg‐ and Lys‐free Media: DMEM, RPMI‐1640, Phenol red‐free MEM and 1:1 DMEM:F12 are available from Pierce. SILAC media is also available from Sigma‐Aldrich and Invitrogen. Several companies also offer custom media services.

Fetal Bovine Serum: FBS that has been dialyzed using a 10 kDa cut‐off membrane (Sigma‐Aldrich: F0392) is available in 100and 500 ml bottles from the Duke CCF (http://www.cancer.duke.edu/ccf/).

Amino acids: 500 μl frozen aliquots of sterile‐filtered L‐Arg (25 mg/ml), L‐Lys (25 mg/ml), L‐Pro (10 mg/ml), ¹³C₆¹⁵N₄‐L‐Arg and ¹³C₆ ¹⁵N₂‐L‐Lys can be obtained from the Duke Proteomics Core Facility.

Antibiotics: Penicillin, Streptomycin and Fungizone are recommended and can be purchased from the Duke CCF

Preparation of Media

Light Media: To a 500 ml bottle of Arg‐ and Lys‐free media, add 50 ml of dialyzed FBS, 5 ml of Pen/Strep/Fungizone and 500 μl each of Arg, Lys and Pro.

Heavy Media: To a 500 ml bottle of Arg‐ and Lys‐free media, add 50 ml of dialyzed FBS, 5 ml of Pen/Strep/Fungizone and 500μl each of ¹³C₆ ¹⁵N₄ –Arg, ¹³C₆ ¹⁵N₂ –Lys and Pro.

**Aliquots of heavy and light amino acids can be obtained from the Duke Proteomics Core Facility “at cost”, which is $65 for a single (500μL) aliquot of heavy/light Lys and Arg, and 2 aliquots of Pro.**

Conditioning of Cells

1. Split cells (e.g. HEK‐293, 1:6) and plate on two 60 or 100 mm dishes containing light media. After cells have adhered (up to16h after plating), aspirate media from one dish and replace with heavy media.
    
2. Continue to subculture light and heavy cells with the appropriate media until heavy isotope is fully (>99%) incorporated, at least 6 doublings.

Notes

1. Fungizone is recommended in addition to Pen/Strep to prevent contamination of the media.
2. Cells may grow slower‐than‐expected in media containing dialyzed serum.
3. For each new heavy cell line generation, we recommend a check of isotope incorporation and a check for evidence of proline conversion in the heavy sample by mass spectrometry after the recommended number of doublings, prior to performing experimental challenge using the cells. This can be done by the DPCF using a small number of cells (~1e6) for $100.

Authored by Dr. Matthew Foster, edited by J. Will Thompson

Duke Department of Pulmonology and Duke Proteomics Core Facility.

This protocol should generate approximately 0.7 to 1.5 mg protein for analysis, depending composition of the tissue. This precipitation protocol is intended as a substitute for more traditional protocols utilizing SDS‐based or urea‐based tissue homogenization, which does not afford the harvest of RNA during the protocol. Sample should be kept at 4C (on ice) during this protocol unless otherwise stated.

1. Weigh tissue section immediately upon removal from ‐80C by taring a 15 mL round‐bottom polypropylene falcon tube and adding tissue. Add 1 mL ice‐cold TRIzol® per 0.1 g tissue weight to falcon tube.
    
2. Homogenize tissue section vigorously using 5 second bursts at max speed. Homogenate should be even consistency with no visible chunks.
    
3. Transfer desired quantity of homogenate for processing to 2 mL Lo‐Bind eppendorf tubes in 1 to 1.5 mL aliquots.Store remaining homogenate at ‐80C. According to manufacturer, this homogenate is good for at least 1 year at ‐80C.
    
4. Incubate TRIzol tissue homogenate on ice/at 4C for 30 minutes with mixing if possible. If not,vortex occasionally.
    
5. Incubate TRIzoltissue homogenate for 5 mins at 30C. Cool sample on ice. Spin at 12,000 rcf for 5 minutes at 4C; unlysed tissue debris and cells will form a pellet at the bottom of the tube and clear TRIzol lysate should form supernatant.
    
6. Pipet 0.8 mL clear TRIzollysate to 2.0 mL LoBindeppendorf tube for protein extraction.
    
7. Decant any remaining remaining clear TRIzol‐tissue homogenate and store at ‐80C.
   
   ***Steps 8‐9 for generating an additional “sonication‐soluble” fraction***. If not isolating sonication-soluble fraction, skip to step 10. 
    
8. Decant all clear trizol homogenate from the top of the pellet. Add 0.5mL of fresh, cold TRIzol reagent to the top of the pellet.
    
9. Sonicate the pellet/0.5mL TRIzol at Power = 3 with probe sonicator. Sonicate with two 10‐second bursts, cooling on ice between. Repeat steps 4‐6 above for isolation of clear lysate from sonication‐soluble fraction. At step 6, you will only be able to isolate 0.4mL of clear TRIzol lysate. Proceed from step 10, with reagent volumes cut in half for each step in the protocol.
    
10. Add 0.2ml chloroform to 2 mL epi tube containing 0.8 mL TRIzol lysate, and vortex to mix. Centrifuge at 12,000rcf x 5 min to perform phase separation. Remove upper aqueous layer (contains RNA, process if necessary).0.5 mL of RNA/aqueous layer should be able to be removed without disturbing the protein/phenol/chloroform layer. Approximately 200 uL will be available for sonication‐soluble fraction.
     
11. To the remaining phenol/chloroform layer, add 1.2 mL coldMeOH, and vortex. Store sample for 10 mins at 30C, then cool the sample on ice and subsequently sediment the protein by centrifuging at 12,000 rcf for 10 mins. (per Kline et. al. JPR2008 v7 p5055)
     
12. Remove supernatant and wash pellet with one 1.0 mL aliquot of cold MeOH. Spin at 12,000 rcf between washes if needed.
     
13. Add 500 uL cold MeOH, then sonicate the protein pellet using burst sonication (5 seconds at 30 percent power) to generate a fine protein powder.Use the same volume, 500 uL, for sonication-soluble fraction for this step. Use approximately 4 bursts, and store in ice between bursts.This step is critical for protein re‐solubilization.
     
14. Allow the protein pellet to settle or use low‐speed centrifugation to settle protein (2000 rcf x 1 minute). Remove supernatant MeOH. Do not allow protein pellet to dry, but allow all MeOH from above the pellet to evaporate.
     
15. 15. Add 200 uL0.25% Rapigest in 50mM ammonium Bicarbonate and resolubilize pellet with assistance from vortex and heat to 60C. If needed, probe sonication can be used. Utilize same volume (200 uL), for resolubilization of sonication‐soluble TRIzol lysate.
     
16. Protein concentration should be between 2.5 and 7.5 mg/mL; perform 1:10 dilution of a small aliquot (2 uL:18uL) in 50 mM Ammonium Bicarbonate, and use a Bio‐Rad mini‐Bradford Assay to quantify protein (BSA as standard).
     
17. Normalize protein content and follow standard in‐solution digestion protocol.

Crosslinking Primary Antibody to Protein A/G resin using dimethylpimelimidate (DMP), for MS-Compatible Immunoprecipitation

(As an alternative to disuccinimidyl suberate (DSS) crosslinking)

Reagents:

1. PBS
2. Binding buffer: 0.2M sodium borate (adjust 0.2M Boric Acid to pH 9 with NaOH)
3.  Crosslinking reagent: 20mM DMP (dimethylpimelimidate, Pierce # 21666) dissolved in 0.2M sodium borate – make immediately prior to use
4. “Quenching” reagent: 0.2M ethanolamine (pH 8.0 in 50 mM Ammonium Bicarbonate)
5. Acid Wash buffer: 0.58% v/v acetic acid with 150mM NaCl
6. Lower stringency lysis buffer for wash: 150mM NaCl, 50mM Tris, 10mM EGTA, 0.2% NP40
7. MS Wash buffer: 50mM ammonium bicarbonate
8. Elution buffer: 0.25% Rapigest SF Surfactant in 50mM ammonium bicarbonate

Procedure:

Coupling antibody with agarose beads:

1. Couple antibody/control IgG andProtein A/G beads (Pierce # 20421) in PBS:
   • Add 60ul (20 ug) X antibody to 30 ul Prot A/G beads in 1ml PBS
   • Add 50ul (20 ug) IgG rabbit to 30 ul Prot A/G beads in 1ml PBS (control)
   • Rock 2-3 hours or overnight at 4C
2. Wash beads 3X with 1ml of 0.2M sodium borate pH 9 – after each wash, spin at 300-500g to pellet beads, gently remove supernatant
3. Save small sample to run on gel to check efficiency (see step 15)
   
   Crosslinking:
    
4. Make fresh DMP:
   • Let solid warm up to room temperature ~20min
   • Weigh out 0.0259 g DMP
   • Immediately add DMP to 5ml of 0.2M sodium borate (pH 9) to make 20mM DMP solution
5. Immediately add 1ml 0.2M sodium borate + 20mM DMP to (1) beads + X Antibody; and (2) beads + Control IgG
6. Rock at RT for 40min to perform antibody crosslinking
7. Spin down and remove supernatant
8. Wash beads once in 0.2M ethanolamine (pH 8.0) – this removes/quenches any residual DMP
9. Resuspend in 1ml of 0.2M ethanolamine (pH 8.0)
10. Rock at RT for 1-2 hours
11. Save small sample to run on gel to check efficiency (see step 15)
12. To remove uncoupled IgGs, wash with 3X 1mL 0.58% v/v acetic acid + 150mM NaCl
13. Wash 3X with 1ml cold PBS - after each wash, spin at 300-500g to pellet beads, gently remove supernatant
14. Save small sample to run on gel to check efficiency (see step 15)
    
    Check the efficiency of immobilization:
     
15. Sample beads (1) before and (2) after crosslinking, and (3) after the acid wash – elute by boiling resin in 2x LDS loading buffer, then run SDS-PAGE/Coomassie stain . **Ensure equal volumes of beads are analyzed for each of these samples. For example, if you are going to take out 5 uL of beads for the check and then elute in 20 uL 2X LDS and load 10 uL for SDSPAGE, be sure to do it the same for all resin binding steps.
16. Evaluate the results of the immobilization process prior to proceeding. (please share images with us!!)
    
    Proceed with IP:
     
17. Quantity of input material will vary depending on the expression of the target protein/complex. Typically, between 0.5 and 5 mg input protein is utilized. Preclearing is recommended, by adding your lysate to ~25 uL of uncoupled Protein A/G resin, rock for 30 minutes, and reserve supernatant for IP.
18. Add 1ml (~1-5 mg/mL protein) of precleared lysate to approximately 25 uL coupled resin. Use same resin quantity for all samples.
19. Rock overnight at 4C (with end-over-end mixing, if possible)
20. Save supernatant for WB analysis.
21. Wash resin 3X with lysis buffer containing 150mM NaCl, 50mM Tris, 10mM EGTA, 0.2% NP40 (you may want to save these washes for troubleshooting purposes… a signal in WB at this step (but not step 20) would suggest target bound to antibody but was then washed away)
22. Elute with 50-100ul of 2X LDS loading buffer, boil 5 min at 95C, pellet beads by centrifugation, take supernatant for SDSPAGE analysis (below). ***see step 23 for MS compatible elution protocol
    
    Recommended Setup for Coomassie gel (or WB analysis):

• Ladder
• 50ug lysate
• X Beads pre-elution
• X Beads post-elution
• X Elution supernatant
• IgG Rab Beads pre-elution
• IgG Rab Beads post-elution
• IgG Rab Elution supernatant
  
  *Stain with Invitrogen Colloidal Blue Staining kit (LC6025)
  *Alternative elution steps once IP has been optimized

1. After step 21 wash, wash resin 3X with 1 mL 50mM ammonium bicarbonate

2. Add 50-100ul of 0.2% Rapigest SF Surfactant (Waters Corporation) in 50mM Ammonium Bicarbonate.

3. Boil @ 95C for 5min, pellet beads by centrifugation, take supernatant for delivery to Duke Proteomics Core Facility, or for SDS-PAGE analysis as above.

Buffer A: 80% MeCN, 1% TFA
Buffer B: 80% MeCN, 1% TFA, 1M Glycolic Acid
Buffer C: 20% MeCN, 5% aqueous ammonia

**Be sure to check tubes after each spin to make sure that entire volume has passed through spin column.

** Buffer B is only used for complex lysates and typically not from gel-band analysis or single protein solution samples. Buffer B will increase the specificity of the enrichment, but does result in a slightly lower total absolute amount of phosphorylated peptides. If you choose to use Buffer B, it can replace steps “a” and b” in the “wash” portion of either protocol.

For 600ug capacity tips:

1. Bring samples up in 150uL of Buffer A.
2. Pre-Elute
   • Add 100 uL Buffer C, centrifuge (1000 x g, 2min)
3. Condition
   • Add 100 uL Buffer A, centrifuge (3000 x g, 2min)
   •  Add 100 uL Buffer A, centrifuge (3000 x g, 2min)
   • Remove the 200 uL of effluent from the waste fluid tube.
4.  Bind
   • Add sample in 150uL Buffer A, centrifuge (1000 x g, 10 min)
   • Re-apply flowthrough, centrifuge (1000 x g, 10 min)
5. Wash
   • Add 100 uL Buffer A, centrifuge (1000 x g, 2min)
   • Add 100 uL Buffer A, centrifuge (1000 x g, 2min)
   • Add 100 uL Buffer A, centrifuge (1000 x g, 2min)
   • Add 100 uL Buffer A, centrifuge (1000 x g, 2min)
   • Switch to new collection tube (1.5mL Epi)
6. Elute
   • Add 100 uL Buffer C, centrifuge (1000 x g, 5min) 
   • Add 100 uL Buffer C, centrifuge (1000 x g, 5min) 
7. Acidify
   • Add ~6 uL of neat formic acid to elution fraction and check pH < 4.
8. Freeze and Lyophilize.

For 200ug capacity tips (exact protocol from GL Sciences):

1. Bring samples up in 65 uL of Buffer A.
2. Condition
   • Add 20 uL Buffer A, centrifuge (3000 x g, 2min)
   • Add 20 uL Buffer A, centrifuge (3000 x g, 2min)
   • Remove the 40 uL of effluent from the waste fluid tube.
3. Bind
   • Add sample in 65 uL Buffer A, centrifuge (1000 x g, 10 min)
   • Re-apply flow through, centrifuge (1000 x g, 10 min)
4. Wash
   • Add 20 uL Buffer A, centrifuge (3000 x g, 2min)
   • Add 20 uL Buffer A, centrifuge (3000 x g, 2min)
   • Add 20 uL Buffer A, centrifuge (3000 x g, 2min)
   • Add 20 uL Buffer A, centrifuge (3000 x g, 2min)
   • Switch to new collection tube (1.5mL Epi)
5. Elute
   • Add 50 uL Buffer C, centrifuge (1000 x g, 5min)
   • Add 50 uL Buffer C, centrifuge (1000 x g, 5min)
6. Acidify
   • Add ~3 uL of neat formic acid to elution fraction and check pH < 4.
7. Freeze and Lyophilize.

Our Bradford assay reagents are found in the Quick Start Bradford Protein Assay Kit 2 from BioRad, part number 500‐0202. The kit contains the BSA standard set and the 1x dye reagent.

When performing a Bradford assay, use a UV Flat Bottom Microtiter Plate from Thermo, part number 8404. The microtiter plates are in a 96‐well format.

For the calibration curve, pipet 5 uL in duplicate for each standard in the BSA standard set. Each 5 uL sample goes in a separate well. There are seven standards in the set, which are:

0.125 ug/uL
0.25 ug/uL
0.5 ug/uL
0.75 ug/uL
1 ug/uL
1.5 ug/uL
2 ug/uL

When pipetting out the calibration curve, make sure to include two blank wells for the zero point in the calibration curve.

Add 5 uL of each sample to a separate well. If you have enough sample, adding 5 uL in duplicate is preferable.

Add 245 uL of the 1x dye reagent to every well that contains a sample. Remember to add dye reagent to the blank (zero point) wells.

Read the measurements on a plate reader set to a wavelength of 595 nm.

Solutions to make:

2% formic acid in water

5% ammonium hydroxide in methanol

• Dilute samples to ~1 mL with 2% formic acid in water.
• Draw through 2 x 1 mL of methanol.
• Draw through 2 x 1 mL of 2% formic acid in water.
• Draw through diluted sample.
• Draw through 2 x 1 mL of 2% formic acid in water.
• Draw through 2 x 1 mL of methanol.
• Draw through 2 x 750 uL of 5% ammonium hydroxide in methanol, collecting eluate in a suitable device.
• Speed Vac to dryness