Research

LIke ThisLIke ThisLIke This

Useful Protocols


Avoiding Contamination

The Keratin Issue

  • Keratin contamination is almost always observed as a background protein. To minimize contamination, wear only powder-free nitrile gloves and use HPLC grade water to rinse all trays, containers and surfaces that contact the gel (including gloves, staining trays, scalpels, razor blades, light box, and cutting area). Gels should be stored at 4°C in 1% acetic acid/HPLC or dd H2O, or in 0.02% sodium azide. Use previously unused storage containers/gel trays to store gels intended for analysis by mass spectrometry.
  • We strongly recommend using freshly prepared solutions/buffers when preparing samples to be subjected to mass spectrometric analysis.

Other Contaminations

  • Other commonly observed contaminating proteins are caseins and BSA from dry milk, media and/or sera commonly used in biology labs. These protein contaminations can be greatly minimized by extensive washing of cell pellets, glassware and trays.
  • Polymers from detergents, PPG and/or PEG are another common source of contamination. Thus, no detergents should be present in samples to be digested for mass spectrometric analysis. Many soaps and hand cremes contain PEG compounds and so touching tubes, trays or gels with freshly washed or creamed hands can give rise to polymer contaminations.

Back to the TOP

Protein Precipitation

Ethanol Precipitation with Glycogen as Carrier

  • Add glycogen to the sample at a final concentration of 20 ug/ml (glycogen does not interfere with the subsequent SDS-PAGE).
  • Add sodium acetate at a final concentration of 0.1 M and pH 5.
  • Add 3 volumes of ethanol.
  • Vortex.
  • Let it stand 2-3 hours at room temperature (RT).
  • There should not be any visible precipitate!
  • Spin 10min at RT at 14 krpm.
  • Wash pellet with 70% ethanol.
  • Dissolve pellet in loading buffer.

Note:
If there is visible precipitate in total mixture before centrifugation, it means: a. there was too much salt in the protein solution; b. some amount of the protein(-s) will be lost due to insolubility.

To avoid this problem:

  • Omit the sodium acetate.
  • Decrease salt molarity of the protein mixture to 300-400mM for NaCl.

TCA Precipitation with Deoxycholate as Carrier

  • Dilute protein to 500 ul with water in Eppendorf tube
  • Add: 50ul of DOC, 25 ul of Triton X 100, 80 ul of TCA (13 % TCA)
  • Incubate for at least 2 hours on ice.
  • Spin in a bench-top microfuge at 4 oC for 30 min at 14,000 rpm.
  • There will be a flaky pellet up the side of the tube.
  • Remove the supernatant and turn tube upside down on a KimWipe and let the tube stand to remove most of the liquid.
  • Add 500ul of ethanol/ether or cold aceton. Sonicate sample in a sonication bath to thoroughly disrupt pellet.
  • Incubate on ice for 30 min.
  • Repeat step 4.
  • There will be little to no pellet.
  • Repeat step 6.
  • Let air dry at room temperature.
  • Re-suspend samples in 5 ul running buffer first, then 20 ul sample buffer.
  • Incubate for 15 min at 65 °C.
  • Load sample and run gel.

Back to the TOP

In-solution Digest

For in-solution digests and subsequent mass spectrometric analysis, the sample should be free of ANY detergents (SDS, Triton X100, NP-40, etc.pp.). Once that is insured, the following protocol can be used:

  • The pH of the solution should be adjusted such that the pH lies between 7.2 and 8.5.
  • Trypsin is added to the sample solution such that the enzyme to substrate ratio is in the 1:50 to 1:10 range.
  • Incubate for 4 to 8 hours at 37°C (or overnight).

If denaturing, reduction of the SS-bonds, and alkylation of the free thiol groups is necessary, we suggest the following protocol:

  • Precipitate the proteins (see above).
  • Re-dissolve the proteins in 8 M urea, 50 mM ammonium bicarbonate, 20 mM DTT.
  • Incubate for 30 min at 56?C.
  • Cool down to room temperature.
  • Add iodoacetamide solution to a final concentration of 100 mM and incubate for 20 min at room temperature in the dark.
  • Dialyze the solution against 10 mM ammonium bicarbonate using e.g. the Slide-A-Lyzer dialysis MINI units from Pierce (MWCO 3500) overnight.
  • Depending on the protein concentration, either directly add trypsin to the sample solution such that the enzyme to substrate ratio is in the 1:50 to 1:10 range, or speedvac the sample, then re-dissolve the proteins in 50 mM ammonium bicarbonate with the appropriate amount of trypsin present.
  • Incubate for 4 to 8 hours at 37°C (or overnight).

In-solution Reduction/Alkylation prior to SDS-PAGE

  • Add DTT to a final concentration of 20 mM to the sample buffer (make sure that the sample buffer does not contain additional DTT or bME).
  • Incubate for 20 min at 65°C.
  • Allow to cool down to room temperature.
  • Add iodoacetamide to a final concentration of 100 mM.
  • Incubate for 20 min at room temperature in the dark
  • If the color of the bromophenol blue changed to greenish/yellowish, add one microliter of 1M Hepes or Tris (pH 7.5 to 8.8) to adjust the pH.
  • Load and run the gel.

Alternatively, following protocol can be used:

  • Add DTT to a final concentration of 20 mM to the sample buffer (make sure that the sample buffer does not contain additional DTT or bME).
  • Incubate for 20 min at 65°C.
  • Allow to cool down to room temperature.
  • Add acrylamide solution to a final concentration of 1 % to the sample volume.
  • Incubate for 20 min at room temperature
  • Load and run the gel.

Back to the TOP

Gel Staining

There are numerous staining methods available that are compatible with subsequent proteolytic digestion and mass spectrometric analysis. Most of them are based on Coomassie staining, silver staining or using fluorescence dyes such as Sypro Ruby or Sypro Orange. Whereas Coomassie staining is simple and cheap, the sensitivity is sometimes insufficient. Silver staining overcomes the sensitivity limitations of Coomasssie staining, but is rather cumbersome. The ease of Coomassie staining and the sensitivity of silver staining are combined in fluorescence Sypro dyes. Those, however, have the disadvantage that they are rather expensive, a fluorescence scanner is a prerequisite for imaging, and excising the gel plugs/bands of interest requires portable UV sources. Irrespective of the staining method used, it is crucial to avoid irreversibly fixing the proteins in the gel during staining; this consideration especially applies to some 'traditional' silver staining protocols that use glutaraldehyde or formaldehyde for fixing and developing which will render the gels incompatible with subsequent mass spectrometric analysis. A variety of mass spec compatible silver staining protocols are given below.

We have good experience with numerous commercial Coomassie staining kits including GelCode Blue Stain from Pierce, Bio-Rad's Bio-Safe Coomassie stain and the Colloidal Blue Staining kit from Invitrogen. A general Coomassie staining protocol is given below. If commercial kits are used, we strongly suggest following the manufacturer's protocols.

Coomassie Staining

  • ALWAYS WEAR POWDER-FREE GLOVES! DO NOT touch gels with your bare skin.
  • ALWAYS use ddH2O or commercial HPLC grade water when preparing buffers for staining.
  • Remove the gel from the cassette and rinse off the buffer with ddH2O.
  • Fix the gel with 45% methanol, 10% acetic acid, and 45% ddH2O for 15-60 min.
  • Rinse the gel for 5 min with ddH2O, repeat twice.
  • Cover the gel with Coomassie blue solution.
  • Gently shake for up to 3 hours.
  • Rinse the gel for 5 min with ddH2O and repeat twice.
  • Destain overnight with 45% methanol, 10% acetic acid, and 45% ddH2O.
  • Rinse the gel for 5 min with ddH2O and repeat twice.
  • Wrap the gel with clean cling film. Acquire the gel's image with a benchtop scanner.
  • Store the gel in 1% acetic acid or in 0.02 % NaN3 (sodium azide) in ddH2O.

Silver Staining

  • ALWAYS WEAR POWDER-FREE GLOVES! DO NOT touch gels with your bare skin.
  • ALWAYS use ddH2O or commercial HPLC grade water when preparing buffers for staining.
  • Remove the gel from the cassette and rinse off the buffer with ddH2O.
  • Fix the gel with 45% methanol, 10% acetic acid, and 45% ddH2O for 15 to 60 minutes.
  • Leave the gel in ddH2O for one hour on a shaking platform. Extended washing time helps to eliminate the yellowish background often observed after overlong development of the gel (see step 6 of this protocol).
  • Sensitize the gel with 0.02% sodium thiosulfate for 1 - 2 minutes. Agitate gently to make sure that the gel slab is covered evenly. Discard solution and quickly rinse the gel slab with two changes of water (10 seconds each).
  • Incubate the gel in chilled 0.1% AgNO3 for 30 minutes at 4°C (fridge).
  • Discard silver nitrate solution and quickly rinse the gel with two changes of water (30 seconds per rinse).
  • Develop the gel with fresh 0.04% formaldehyde in 2% sodium carbonate*. Discard the developing solution as soon as it turns yellow and replace it with a fresh portion.
  • When a sufficient degree of staining has been obtained, quench the stain by discarding the developing solution and replace with 1% acetic acid. Wash the gel with 1% acetic acid several times, then store in 1% acetic acid/ddH2O. Alternatively, the gel can be stored in 0.02% NaN3 (sodium azide) in ddH2O.
  • NOTE: In some cases the color of the stained protein bands might slightly change over time. However, these changes do not affect the results of mass spectrometric sequencing. For more details, see Blum et al, Electrophoresis, 8, pp93-99, 1987 and Schevchenko, A. et al. (1996) Anal. Chem. 68, 850-858.

SYPRO Ruby Staining

  • ALWAYS WEAR POWDER-FREE GLOVES! DO NOT touch gels with your bare skin.
  • ALWAYS use ddH2O or commercial HPLC grade water when preparing buffers for staining.
  • Use a clean plastic tray as staining container.
  • Rinse the gel 3 times with ddH2O.
  • Fix the gel with 10% methanol, 7% acetic acid, and 83% ddH2O for 60 min.
  • Stain the gel in SYPRO Ruby solution which is at least 10 times the volume of the gel.
  • Gently shake for at least 3 hours, up to overnight for maximal sensitivity.
  • Wash the gel with 10% methanol, 7% acetic acid, and 83% ddH2O for 30-60 min to decrease background fluorescence.
  • Wash the gel 3 to 5 min with ddH2O.
  • Acquire gel image under a blue light transilluminator (image can be seen by hand-held UV source).
  • Store the gel in 1% acetic acid or 0.02 % sodium azide.
    NOTE: SYPRO Ruby is a suboptimal choice for glycol- and lipoproteins.

Back to the TOP

Excising Gel Bands

  • Rinse the gel with Milli-Q or HPLC grade water. Excise bands of interest with a clean scalpel. Cut as close to the edge of the band as possible in order to reduce the amount of 'background' gel.
  • Chop the excised bands into pieces of approximately 1 x 1 mm and transfer the gel particles into a 0.5 ml microcentrifuge tube. Do not macerate the gel bands/plugs.

In-gel Reduction/Alkylation

  • Wash the gel particles with 100-150 ?L of Milli-Q or HPLC grade water for 5 minutes. Vortex briefly, spin down the gel particles and discard the liquid.
  • Add acetonitrile (corresponding to approximately 3 - 4 times the total volume of gel pieces) and incubate for 10 - 15 minutes until the gel pieces shrink, become opaque and stick together. Spin down the gel particles and discard all liquid.
  • Swell the gel pieces in 10 mM DTT/50 mM ammonium bicarbonate and incubate for 30 minutes at 56°C to reduce the protein.
  • Spin down gel particles and discard all liquid. Shrink the gel pieces with acetonitrile.
  • Replace the acetonitrile with 55 mM iodoacetamide in 50 mM ammonium bicarbonate. Incubate for 20 min at room temperature in the dark.
  • Discard iodoacetamide solution. Wash the gel particles with 150-200 ?L of 50 mM ammonium bicarbonate for 15 minutes. Spin down gel particles and discard all liquid. Shrink the gel pieces with acetonitrile. Proceed with trypsin digestion (see below).

In-gel Digestion

  • Rehydrate gel particles in chilled digestion buffer (50 mM ammonium bicarbonate in HPLC grade water) containing 12.5 ng/?L trypsin at 4?C (on ice or in cold room).
  • Incubate for 30 - 45 min at 4°C. After 15 - 20 min, check the samples and add more digestion buffer containing 12.5 ng/?L trypsin to cover the gel particles if they have absorbed the previous aliquot.
  • Remove and discard remaining supernatant. Add sufficient (5-50 uL) 50 mM ammonium bicarbonate in HPLC grade water (without trypsin!) to cover the gel particles and to keep them wet during enzymatic cleavage. Incubate the gel particles at 37°C for at least 6 hours (preferably overnight).
  • After incubation, recover supernatant in a fresh Eppendorf tube.
  • Add 50 ?L of 0.4% acetic acid, 0.02% heptafluorobutyric acid (HFBA) and extract for 15 minutes with agitation. Repeat twice.
  • Pool all supernatants and store digest at -20 °C or proceed to filtration, desalting and concentration using StaGE tips.

Back to the TOP

Filtration, Desalting and Pre-concentration Using Micro-scale Reversed Phase Columns (StaGE tips)

Filtration, Desalting and Pre-concentration Using Micro-scale Reversed Phase Columns (StaGE tips)

  • Punch a little disk out of 3M Empore C18 membranes using a blunt 20G needle.
  • Transfer the reversed phase disk into a pipette tip of appropriate size such as GELoader tips from Eppendorf, P10 tips or P200 tips (in our hands the Corning tips with tapered tip has proven to be very useful). This can easily be done using a fused silica capillary inserted into the needle. Make sure that the disk is properly positioned such that all liquid has to be go through the disk.
  • If larger amounts of material have to be desalted and concentrated, several reversed phase disks can be inserted into the pipette tip.
  • Wet the membrane with 10 ul of methanol.
  • Condition the C18 disk with 10 ul of StaGE buffer A (0.4% HOAc and 0.02% HFBA in water).
  • Re-dissolve the lyophilized digest in StaGE buffer A or add 10x buffer A (4% HOAc and 0.2 % HFBA in water) to the digest to ensure final concentrations of HOAc and HFBA identical to 1x StaGE buffer A.
  • Force the liquid through the 3M Empore membrane by applying air pressure (e.g. disposable syringe).
  • Wash the disk with 10 to 50 ul of StaGE buffer A.
  • Elute the sample with 10 ul of StaGE buffer B (0.4% HOAc and 0.02% HFBA in 80% acetonitrile) into an appropriate sample vial.
  • Lyophilize the eluted sample and re-dissolve it in loading buffer for subsequent LC/MS analysis.
  • Alternatively, the sample can be eluted with matrix solution directly onto a MALDI target.

Back to the TOP

NOTES

  • Suggested ammonium bicarbonate stock solution: 1 M, store at 4 °C.
  • Suggested DTT stock solution: 1 M, store at -20 °C.
  • Suggested iodoacetamide stock solution: 500 mM, store at -20 °C in the dark.
LIke ThisLIke ThisLIke This
Close