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.
- Wear nitrile (not latex) gloves.
- Wear a lab coat and make sure there is no gap between your coat sleeve and the gloves (lab tape works well).
- 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.
- 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.
- Heat at 40°C while shaking for 10 minutes. Spin down condensate.
- 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.
- Heat solution at 80°C while shaking for 15 minutes.
- Remove from heat and cool for 5 minutes (to room temp). Spin down condensate.
- 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).
- Incubate the solutions in the dark at room temperature for 30 minutes.
- Add trypsin to each solution at 1:50 trypsin:protein concentration. Make trypsin stock in 50 mM AmBic.
- Digest for at least 4 hours or overnight at 37°C while shaking.
- Following digestion, centrifuge condensate to bottom of vial. Add TFA and MeCN to give 0.5-1.0% TFA and 2% MeCN by volume.
- 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.
- Shake samples for 2 hours at 60°C. Centrifuge at 15,000 rpm for 5 min, and pipette supernatant into an autosampler vial.