Daniel:Protocols/SlidePAGE

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Polyacrylamide Gels Adhered to Slides[edit]

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Materials[edit]


Protocol[edit]

Day 1-Bind Silane[edit]

Harvard Protocol

  1. Load slides (facing the same direction) into metal slide racks
  2. Sterilize the following items (15 minutes under UV light in PCR hood):
    1. two large glass Coplin buckets & lids
    2. metal slide racks with slides loaded
    3. 1 L dH20 in a plastic container
    4. stir bar
  3. Add 220 uL Acetic Acid to the 1 L dH20 (pH -> 3.5; no need to measure)
  4. Add 4 mL Bind-Silane
  5. Mix for 15 minutes using stir-bar.
  6. Place loaded slide racks in glass Coplin buckets
  7. Add stirred mixture to Coplin buckets (500 mL per bucket; enough to cover the slides)
  8. Cover buckets and incubate with slow shaking for 1 hour.
  9. Leaving racks in buckets, rinse 3 times with dH20
  10. Rinse once with 100% ethanol.
  11. Remove slide-racks from buckets and allow slides to dry (in racks) under AirClean hood.
  12. Transfer slides to plastic storage boxes (keep the Teflon coated sides of the slides facing in the same direction)
  13. Label boxes with initals and date of treatment.
  14. Place slide boxes in dessicator for long-term storage.

Day 2-Gel Prep[edit]

Harvard Protocol

  1. Remove 8 Bind-Silane treated slides from storage box in dessicator and place face up in AirClean hood.
  2. Turn on UV lamp in AirClean hood for 15 minutes.
  3. Label slides with numbers and date using a SHARPIE pen (other inks will wash off in hexane)
  4. Almost completely cover slides with coverslips (Fisherbrand, 18mm x 30mm, #1, untreated). Keep a small area of oval exposed for subsequent gel loading.
  5. Prepare fresh "ABD mix" in a 15 ml polystyrene conical:
    1. 9 mL IEF 40% Acrylamide
    2. 1 mL Acrylamide / Bis (19:1; 38%:2%)
    3. 200 mg DATD
  6. Using a 3-cc syringe and a 0.22 micron filter, filter ~1 mL of the ABD mix into a 1.5 mL microcentrifuge tube.
  7. Prepare fresh 5% APS. Notably, the APS bottle should be stored in some sort of room-temperature dessicator. We just store it inside a large plastic screw-top container.
    1. 0.5 g APS →10 mL dH20
  8. Prepare fresh 5% TEMED
    1. 2 uL TEMED → 38 uL dH20.
  9. Put a few drops of 30% BSA into a 1.5 mL tube (BSA should be stored at 4'C)
  10. Prepare the gel-casting mix (200 uL total volume). Do not add APS until immedietely prior to casting the gels. This recipe is for a 10% gel with sufficient mix for at least 8 (identical) slides.
    1. 50 uL A-B-D mix (FILTERED)
    2. 1.33 uL 30% BSA
    3. 138.66 uL dH20
    4. 4 uL 5% TEMED
    5. 4 uL 5% APS
    6. 0.5 uL Template (at appropriate concentration)
  11. Suck up 18 microliters of the mix and pipet it into the small exposed area of the oval, such that surface tension pulls the liquid into the space between the coverslip and glass to cover the full surface area of the oval. The liquid should distribute under the coverslip such that only a small amount (1 to 2 uL) cannot fit. Slide the coversip over such that the oval is completely covered.
    1. Load as many as 8 gels from a single mix (after adding APS)
    2. if >8, split master mix prior to adding APS (prevents over-polymerization)
    3. Split master mix prior to adding APS for different template on each slide
  12. Place the slides on a flat tray and load the tray into the argon chamber (and fill with argon). Allow gels to polymerize for ~30 minutes.
  13. Remove slides from argon chamber. Use a razor-blade cleanly remove cover-slips from polymerized gels (just stick the edge of the blade under the coverslip and gently "pop" it up)
  14. To wash off the excess acrylamide monomer, place slides in a dH20 filled plastic Coplin jar and incubate for 30 minutes at RT with slow shaking.
  15. Remove slides from Coplin jar and place face up in PCR hood.