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Home > Sample Preparation
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Sample Preparation
| Keratin contamination will always be observed so extra precautions must be taken to minimize the amount of contamination! |
- Wear gloves at all times during sample preparation. Wash outside of gloves with water before using them to handle your samples and vials.
- Thoroughly wash anything that will come into contact with your sample (i.e. gel apparatus, staining trays, gel excising implements, gel storage equipment, and Eppendorf vials) to remove keratins and contaminants. Keratin contamination is more of a problem, and limits our protein identification, when there is a low amount of protein in the gel band (such as gel bands that are only visible by silver or Sypro stains). Eppendorf vials should be washed with HPLC grade Acetonitrile (or use a high grade methanol if no Acetonitrile is available) and then HPLC grade water. Sample vials should be free of plasticizers and contaminants that can compromise high sensitivity mass spectrometric analysis. Anything that touches the gel or sample is a possible source of contamination. Avoid storing gel in saran wrap or similar material and instead use new, cleaned plastic or glass gel trays.
- Avoid using molecular weight cutoff filters (ex. Centricon filters) and if they must be used, thoroughly wash them. We find these are often a source of synthetic polymer contamination in samples.
- Call to discuss your project before sending blood, plasma, or radioactive samples. Samples should not have any particular matter in them, it will clog our 10-15 micron tipped columns, and you risk paying the cost to replace the column.
- Avoid using surfactants and detergents such as CHAPS, DMSO, Triton-X, etc.. If these must be used please contact us to discuss acceptable concentrations. Minimize the salt and buffer concentration in the sample.
- Vials: For liquid samples use the smallest vial necessary, (as low as to the 0.5-0.6ml vials). For gel samples, use a washed 1.5 ml Eppendorf tube (the vial you send us will be used for the in-gel digestion procedure so please make sure it is clean). Don't use vials with rubber o-rings or gasket. Avoid colored vials and avoid using paraflim to seal vials.
- Vial label: each of the vials that contain your samples must be labeled with the your name, sample name (choose something unique but short and different than sample name of previously submitted samples), and date. Please label both top and side of vial.
- After the gel has been run, non-specific dust contamination can still be introduced so keep samples covered or protected as much as possible.
- All samples must be shipped with completed sample submission forms.
- We have found that samples with a significant color tend to cause chromatography problems and often require ZipTip cleanup before digestion and/or LC/MS/MS analysis. Before sending samples that have a significant color, please contact us.
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- Use standard SDS-PAGE or 2D gels. Maximize the amount of protein on as minimal amount of gel as possible. A concentrated gel band works a lot better than a large diffuse gel band.
- Gel thickness of 1mm is preferred. Gradient, denaturing and native gels are acceptable.
- Stain with Coomassie Blue using standard conditions. Stain only until your bands are visible. If the amount of your protein is too low for coomassie stains, use silver or Sypro stains. Generally we recommend Sypro (Ruby) stains over silver stains due to higher compatibility with mass spectrometry. Sypro stains require a UV source to visualize gel spots or bands. If silver stain must be used, we require you use the Invitrogen Silverquest stain kit or a mass spectrometry compatible silver stain. Standard silver stains are not compatible with our LC systems. Silver stained gel spots or bands have to be destained as much as possible as the silver has been reported to inhibit trypsin digestion of proteins.
If your band ends up being very darkly stained (dark black in color) when using silver stain, it will be difficult to fully destain the bands. The issue is that it is necessary to significantly destain silver stained bands or it will cause problems. The problems include limiting the effectiveness of the enzymatic (usually trypsin) digestion which results in a reduction in our sensitivity. In these cases we suggest using coomassie stain instead of silver stain as the amount of protein in a dark silver stain band is usually enough to be visible as a light coomassie band.
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- Excise band as precisely as possible. If there is any diffuse stained edges to the band, omit it and excise only the clearly stained band. Do not mince up the gel into very small pieces (too easily lost in pipetting steps of the in-gel digestion procedure)
- Destain gels as much as you can before sending them to us (you will need the gel stained though so you can cut out the bands). For coomassie stained bands it is not so important to fully destain as it is with silver stained bands.
- Excise a band from a blank area of gel, about the same size as your sample gel bands (cut in the step above), and send it to us to run as a gel blank. You will not be charged for analyzing this gel blank. Add this as a sample on the sample submission form and label it 'gel blank.'
- Place each gel slice into a washed, plain 1.5 ml Eppendorf tube. Do not use any tubes with o-rings or gaskets.
- Wash the gel slice with 50% HPLC Grade Acetonitrile/Water at least 2 times for 10 min with occasional vortex mixing. Follow this with a rinse with just HPLC grade water. If you don't have HPLC grade solvents, use a new bottle of the highest purity solvent you have for the washes. Discard the wash solution. Use plastic pipette tips to remove the solution as the gel slice tends to stick to glass Pasteur pipets. This step helps wash out any residual detergent or salt. If you don’t have any high purity water (don’t use tap water) or Acetonitrile then skip this step as we will repeat this step anyway when we receive the samples.
- Rinse the Eppendorf cap off with the 50% HPLC Grade Acetonitrile/Water solution (or just high purity water if you don’t have HPLC grade Acetonitrile) and make sure this solution is removed before you close the cap tightly and ship the samples to us. No additional solution is needed to cover the gel band. Do not parafilm the tube. The gel band will remain moist and can be stored in a -20 degree freezer or shipped to us. Gel slice can be stored frozen for at least 6 months without degradation. The gel slices should be sent frozen on dry ice unless it is not available. Put your samples in a bag (instead of loose in the dry ice as vial tops can come loose) or into a 50ml conical tube with padding (this helps as delivery companies have been known to make vials lids pop open in shipment) and send them to us. See this suggested packaging guide. If dry ice is not available then try to get the sample to us in 4 days or less and just put a small amount of HPLC grade water (or the cleanest grade water you have) in each tube. Use just enough water so it keeps the gel pieces moist but not so much that the gel pieces are floating in the water (there is a small chance proteins may diffuse out of gel). You can send the room temperature samples in a FedEx package. The idea is just to make sure the gel bands arrive to us either frozen on dry ice or wet (not dried out) and haven’t been at room temperature for more than 4 days. For local users, please try to drop off your samples in the early morning as this is when the in-gel digestion procedure for the day begins. For samples in microtiter plates, cover with just aluminum foil (do not use sticky plate seal film).
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Basic preparation methods