One-Dimensional Acrylamide Gel Electrophoresis:

An Exquisite, Comprehensive Guide

 

 

Part 1: Solution Recipes

 

SDS-Electrophoresis Buffer (for upper and lower reservoir of gel rig):

-100 mL of 10X Tris/Glycine/SDS buffer (from Bio-Rad, product #161-0772EDU)

-900 mL of diH20

 

Coomassie Gel Stain:

-450 mL water

-450 mL methanol

-100 mL glacial acetic acid

2.5 g Coomassie Brilliant Blue (R-250)

Filter and store at Room Temp (RT)

 

Coomassie De-staining Solution:

-400 mL water

-500 mL methanol

-100 mL glacial acetic acid

Store at RT

 

SDS Sample Buffer/”Laemmli” Sample Buffer:

-5 mL H20

-0.5 mL 1M Tris-Cl pH 6.8

-2.5 mL glycerol

-2 mL 10% SDS solution

-0.05 mL 0.25% bromophenol blue solution

Filter, store at RT

ADD 50 mcL BETA-MERCAPTOETHANOL PER mL BEFORE USE!

 

-OR-

 

-order Bio-Rad #161-0737 “Laemmli Sample Buffer”

 

Part 2: Gel Recipes

 

12% Separating Gel:

-16.8 mL H20

-10.4 mL 1.5M Tris pH 8.8

-400 mcL 10% (w/v) SDS

-12 mL 40% acrylamide (make sure rest of specifications are same as label in fridge)

-400 mcL 10% (w/v) Ammonium Persulfate (APS; made fresh weekly)

-40 mcL TEMED

 

6% Separating Gel:

-22.8 mL H20

-10.4 mL 1.5M Tris pH 8.8

-400 mcL 10% SDS

-6 mL 40% acrylamide

-400 mcL 10% APS

-40 mcL TEMED

 

Stacking Gel:

-12.2 mL H20

-5 mL 0.5M Tris, pH 6.8 (NOTE DIFFERENT CONC. AND pH HERE)

-1.92 mL 40% acrylamide

-100 mcL 10% APS

-20 mcL TEMED

 

 

 

PROTOCOL FOR RUNNING A 1D SDS-PAGE GEL:

 

• Clean (meticulously, here) glass plates, spacers, comb, and syringe w/Hamilton cannula with mild soap and diH20. Once clean, rinse plates lightly with MeOH or EtOH and let air dry. Do NOT dry with KimWipes as dust will interfere with polymerization later.
• Assemble gel sandwiches (‘big’ plate, ‘small’ plate, two spacers, and two white plastic clamps) with small plate facing side of clamps that have perpendicular black arrows on them. Using white plastic ‘gel alignment card’ and fingernail on bottom of plates, make sure everything is aligned before tightening clamps. Good alignment here will reduce chance of leaking. Also, use liberal amounts of ‘gel sealant’ on spacers to reduce leaks.
• Place assembled sandwich in white plastic casting stand; turn clamps on sides to lock down gel sandwich (you should feel/see it push down into gray gasket). Pipette a little bit of water on each side of the sandwich along the contact with the gasket to prevent air from bleeding into sandwich during polymerization.
• Prepare separating gel solution, fill with syringe with long Hamilton cannula tip pressed against ‘big’ plate. Fill to roughly the level of the black arrow on the white plastic gel clamp. IMMEDIATELY and carefully overlay separating gel with roughly a centimeter of 1X electrophoresis buffer.
• Once separating gel has polymerized (~ 1 hour), pour off overlay and prepare stacking gel solution. Pour stacking gel nearly to top of sandwich, insert gel comb at an angle to avoid trapping bubbles. Wait for polymerization (~45 mins).
• Outline where wells are with a Vis-à-vis marker, remove gel sandwich from casting stand.
• Attach gel sandwich to central core of gel rig (transparent gray plastic with the two long tubes coming off of it). If only attaching one gel sandwich, attach buffer dam to other side.
• Fill upper buffer chamber with 1x electrophoresis buffer.
• Remove comb and load protein samples (that have already been mixed up and heated at 95C for 5 mins with “SDS Sample Buffer”/Laemmli Sample Buffer).
• Place central core in lower buffer tank (big clear plastic piece); fill lower buffer chamber with enough 1x electrophoresis buffer to cover lowest 4 cm of gel.
• Put on lid (make sure color-coded electrodes line up correctly).
• Hook up electrodes to power supply, circulate cool water through central core if desired (you will get less distortion of bands if you do). Run gel… couple options here. You can run it at 50V if you want best resolution, but it will take roughly 20 hours. I usually set up a gel to start running it in the afternoon, run it at 50V until I come in the next morning, then finish it off at 200V. If you want speed, just run the whole thing at 200V and it should only take 3 or 4 hours but your resolution will suffer.
• Once dye front reaches bottom of gel, turn off power supply, disconnect electrodes, and remove gel sandwich from central core.
• Pry open gel sandwich (lift up the exposed portion of one of the spacers), discard stacking gel, and place gel in big glass dish filled with enough Coomassie stain to submerge it. Shake/shimmy gel loose from plate into the stain. Stain 2 hours with gentle rocking on the ‘Belly-Dancer’.
• Transfer stained gel to a new glass dish of Coomassie de-staining solution. Destain with gentle rocking overnight. (If not completely clear yet, transfer to a new dish of destain for a ½ hour or so and it should be great).
• Image on lightbox, take picture, record/label position of any clipped bands for mass spec.