1. MALDI Sample Preparation
  2. In-gel Digestion and Crystallization

MALDI Sample Preparation

In-gel Digestion and Crystallization

Protocol was developed by J DeGrasse, M. Kalkum, A. Krutchinsky, J.C. Padovan and W. Zhang

Necessary reagents

  • Electrophoresis solutions, such as running buffers and sample loading buffers, may be prepared with milliQ-grade water. All other solutions should be made with HPLC-grade solvents.
  • 100mM ammonium bicarbonate. 3.95g ammonium bicarbonate into 500ml water. Filter with a 0.22µm size filter. Store at 4°C.
  • 200mM dithiothreitol. 30.9mg dithiothreitol into 1ml of 1mM aqueous HCl solution. Aliquot. Store at -20°C.
  • 1M iodoacetamide. 46.3mg into 250µl water. Aliquot. Store at -20°C.
  • Destaining solution. 50mM ammonium bicarbonate in 50% methanol / 50% water. Store at 4°C.
  • Trypsin. Add 25µl of 1mM HCl to the lyophilized trypsin vial (Roche #11418025001). Separate into 1, 2 or 5-µl aliquots. Store at -20°C.
  • POROS bead slurry (50µg/µl in 20% ethanol) (POROS 20 R2, Applied Biosystems). Dry stock POROS beads are washed in equal volumes of (1) methanol, then (2) 80% acetonitrile, then (3) 20% ethanol and then (4) resuspended in 20% ethanol at 50µg/µl and stored at 4°C.
  • 10% (w/v) trifluoroacetic acid (from Pierce #28902).
  • Elution solutions
    • DHB elution solution: 20% acetonitrile, 50% methanol, and 0.1% trifluoroacetic acid.
    • 4-HCCA elution solution: 70% acetonitrile, 0.1% trifluoroacetic acid.
  • Calibrants. Bradykinin (904Da), Neurotensin (1672Da), oxidized Insulin ß-chain (3496Da). To a 10µl 50µM stock solution (stored at -80°C), add 90µl 50% acetonitrile, 5% ethanol, 45% water. Store the 5µM stock at -20°C.


  1. Electrophoresis

    1. Dilute the protein sample in gel-loading buffer. (We suggest bis-Tris gel systems, Invitrogen)
      1. 2.5µl NuPAGE® LDS (lithium dodecyl sulfate) Sample Buffer (#NP0007).
      2. 1µl 200mM dithiothreitol (DTT).
      3. Bring to 10µl with sample and/or water.
    2. Heat at 70ºC for 10 minutes (with agitation, if possible), allow to cool to room temperature.
    3. To alkylate the reduced cysteines, add 1µl of 1M iodoacetamide per 10 µL of gel loading buffer. React in the dark for 30 minutes at room temperature.
    4. After running the gel, stain with GelCode Blue (Pierce, catalog number 24592). After recording an image of the gel, destain thoroughly to remove most of the background.
    5. Excise the bands of interest using tweezers and a fine scalpel (fine-point diamond tweezers and 15- or 30-degree feather microscalpel, Electron Microscopy Sciences).
      • Alternatively, one could remove the lane from the gel and cut 2-4mm slices down the length of the lane. This allows for the detection of low-abundance proteins that may not be detected by staining.
    6. Dice each slice into 1mm3 using either the scalpel or the tweezers. Transfer the cubes to a microcentrifuge vial filled with 500µl of destaining solution.
  2. Washing (and destaining) gel slices

    1. Shake the vials in a microvial mixer (Microtube Mixer, MT-360, TOMY Tech USA) at 4°C.
    2. Aspirate the destaining solution and replace with fresh solution every 30 minutes and wash for 4 hours or until all traces of Coomassie have been removed. Residual amounts of Coomassie and detergents (e.g., SDS, MOPS, etc.) will partially inhibit trypsin digestion. Also, the Coomassie will co-extract with the peptides.
    3. After the final aspiration, add 100µl acetonitrile to dehydrate the gel pieces. Aspirate the liquid. Air-dry for 5 minutes.
    4. One can leave the dried gel pieces overnight at 4°C.
  3. Digestion

    1. Prepare trypsin aliquot
      • Add 19µl 50mM ABC to a 1µl trypsin aliquot (50ng/µl final concentration).
    2. Add 75-150ng of trypsin to the dried gel pieces and allow pieces to swell (~5min).
    3. Top off with 40µl of 50mM ABC in 10% acetonitrile. Be sure gel pieces are covered.
    4. Digest at 37°C for 4-6 hours. Four hours is normally sufficient.
  4. Extraction

    1. Prepare POROS beads
      1. 1 volume stock POROS bead slurry.
      2. 9 volumes 5% formic acid, 0.2% trifluoroacetic acid(aq).
    2. Add 40µl of slurry to each digestion vial (or the equivalent of the digestion volume).
    3. Incubate on the microvial mixer at the highest setting. 4 hours at 4°C.
  5. ZipTip Conditioning and Sample Loading

    1. Prepare ZipTips
      1. Wash ZipTips with:
        1. 2 x 10µl 0.1% trifluoroacetic acid.
        2. 4 x 10µl DHB or 4-HCCA elution solution.
        3. 4 x 10µl 0.1% trifluoroacetic acid.
      2. leave 10µl 0.1% trifluoroacetic acid to keep tips wet, and remove prior to use.
    2. Transfer the 80-ml digest and bead mixture to the cleaned ZipTips and spin the liquid through for 1-2min at 1000rpm. Discard the flowthrough.
    3. Wash gel pieces in microcentrifuge vial with 20µl 0.1% trifluoroacetic acid, transfer wash onto ZipTip and expel the wash solution as above.
    4. Thoroughly wash beads 2x with 20µl 0.1% trifluoroacetic acid. Expel wash solution as above.
    5. One can leave the ZipTips overnight at 4°C if the beads are kept wet with 0.1% trifluoroacetic acid.
  6. Elution

    Choose from two options: DHB or 4-HCCA

    1. Prepare matrix
      1. To solid DHB (fill a 1.5ml microcentrifuge vial to the 100µl mark), add 50µl of DHB elution solution. Vortex for 5min or until the solution is saturated. Spin at 14k rpm for 5 minutes.
      2. In a fresh vial, dilute the saturated solution 1 in 3 (33% saturated DHB) with DHB elution solution.
    2. Place 2.5µl of the 33% saturated DHB onto the top of the ZipTip behind the POROS beads. Slowly elute onto the sample plate.
  7. Calibration

    1. Dilute the 5pmol/µl stock 1 in 10 in 50% acetonitrile, 5% ethanol, 45% water to produce a 500 fmol/µL solution.
    2. Dilute the 500fmol/µl solution 1 in 5 with matrix (33% DHB or 50% 4-HCCA) and spot 1-2µl spots in close proximity to your sample. One calibrant spot for every 10 sample spots is sufficient.