Pouring and Running Protein Gels/Electrotransfer

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Pouring and Running Protein Gels/Electrotransfer

Pouring and Running Protein Gels/Electrotransfer

1) Clean five pairs of minigel plates well wearing glove and using soap with yellow sponge. Rinse well with deionized water, then squirt with ethanol or methanol, wipe off excess, then place on rack to dry.

2) Assemble pairs of plates with clean .75 mm spacers and place in multi-plate holder. Separate pairs with one piece of weighing paper. Add several pieces of spacer plastic (place behind next to last gel plate pair), as required. If pouring gradient gels, clamp in place without rubber wedge in bottom. Have rubber wedge for all other gels. Make sure gel plates are tightly clamped; if not, more plastic spacers may be necessary. Clamp outlet tube with a hemostat.

3) If pouring straight 6, 8 or 10% gels, combine reagents in order below in a 100 ml beaker or 50 ml conical. Mix well before and after addtion of ammonium persulfate and Temed. Add immediately to top of gel until level reaches just below black line. Immediately add ddH2O to the top of each gel: use 40 l at a time for a total of 200 l/gel.

Separating Gel 6% 8% 10% Acrylamide:bis (30:0.8) 5 ml 6.7 ml 8.3 ml 1.5 M Tris, pH 8.8 6.25 ml 6.25 ml 6.25 ml ddH2O 13.3 ml 11.6 ml 10 ml 10% SDS 250 l 250 l 250 l 10% Ammonium Persulfate 200 l 200 l 200 l Temed 25 l 25 l 25 l

4) For 5 -15% gradient gels, prepare 5% and 15% solutions without adding final Temed. Have 5% in first chamber of gradient mixer and 15% in second chamber. Have tube leading from gradient mixer (on magnetic stirrer with stir bars in each chamber) to peristaltic pump to tube of multigel holder (make sure this latter tube is free of polymerized acrylamide). Have hemostat ready. Check that channel between two chambers of mixer is open, then add Temed to 5% with mixing followed by Temed to 15% with mixing. Immediately open outlet and turn on pump to run gradient into gel plates up to black line. At end of run, carefully clamp with hemostat, and add 40 l of ddH2O/plate working from back to front on the left side, then right side; repeat until have about 200 l/gel.

Separating Gel-Gradient 5% 15% Acrylamide:bis (30:0.8) 2 ml 6 ml 1.5 M Tris, pH 8.8 3.05 ml 3.05 ml ddH2O 6.8 ml 1.7 ml glycerol 0 ml 1.1 ml 10% SDS 120 l 120 l 10% Ammonium Persulfate 60 l 60 l Temed 6 l 6 l

5) After gel has polymerized, unclamp apparatus and use a razor blade to carefully separate pairs of plates. Wrap in plastic wrap containing moist 3 MM paper; label plastic with date and % gel. Store at 4°C. To pour the stacking gel, remove plastic wrapping and use filter paper to blot off excess water. Clamp into gel apparatus and insert a clean comb. Mix stacking gel well and pour immediately. Comb can be removed in 15-20 min.

Stacking Gel Acrylamide:bis (30:0.8) 1 ml 1.5 ml 0.5 M Tris, pH 6.8 2.5 ml 3.75 ml ddH2O 6.2 ml 9.3 ml 10% SDS 100 l 150 l 10% Ammonium Persulfate 100 l 150 l Temed 10 l 15 l 10 ml 15 ml

6) To run gel, add running buffer to bottom of gel apparatus sufficient to bring above level of the lower portion of the gel. Add running buffer with 20 ml pipet to well behind gel, but not yet to wells of gel. Individually dry well of gel with narrow cut pieces of filter paper. Add approx. 5 l of sample (or molecular weight standard) to 10 l of sample buffer with or without 1/10 volume of 1 M DTT, boil for 5 min at 90°C, place 1 min on ice, spin for 1 min (RT), and load in wells of gel using a long narrow pipet tip. Add running buffer gently (add at 15 l/well) to fill remaining volume of wells and top portion of gel apparatus. Run at 25 mAmp/gel. Remove gel and fix/stain in 0.08% Commassie blue in 25% isopropanol/10% methanol for 2 hr. Destain in 10% acetic acid/10% isopropanol. Photograph and/or dry on gel dryer (place dry ice in trap; place gel on destain-wetted cellophane, sandwich with another piece of wetted cellophane; turn on pump and dry for 1 hr with 60°C heat).

7) Instead of Coomassie blue staining, gels may be silver stained. At the end of a gel run, place gel in a plastic silver staining box containing a mixture of 250 ml methanol, 249.5 ml ddH2O and 0.5 ml of 37% formaldehyde. Fix overnight at RT. The next day, carefully pour off and replace with 500 ml of ddH2O containing 2.5 mg DTT. Incubate for 30 min. Carefully pour off and add 350 ml of ddH2O in which is dissolved 0.35 gm of AgNO3. Place on light box for 10 min, then remove to rotator for 25 min. Rinse gel 1 x with ddH2O (AgNO3 is discarded into AgNO3 waste bottle). Rinse gel 2 x with 100 ml each of 'developer' (400 ml of ddH2O contai'ing 12 gm of Na2CO3 and 200 l of 37% formaldehyde) quickly. Add 200 ml of 'developer' to gel and allow to go for 2 - 5 min. Stop development by adding 20 ml of 2.3 M citric acid (9.66 gm/20 ml; F.W. 210). Rotate for 1 hr, then wash gel for 30 min with three changes of ddH2O. Store in 0.03% NaCO3 to prevent bleaching or dry or photograph. Molecular weight marker load should be 1 l of 1/10 diluted stock/lane.

8) For electrotransfer from gel to Hybond-ECL nitrocellulose (Amersham#RPN 2020 D), as follows. Have blotting buffer prepared (can be reused a number of times; record each time used by marking label). Cut nitrocellulose to same size (or slightly larger) as gel. Have two pieces of Whatmann 2 MM paper cut to same size as gel. Have plastic tray containing transfer buffer and immersed in it in the following order: plastic cage ('+' side down), sponge (push out air bubbles), one piece of Whatmann paper and nitrocellulose filter. Turn off power to gel. Place gel plates on bench. Gently remove glass side. Notch bottom left corner. Discard stacking gel. Carefully invert gel onto nitrocellulose and use a spatula or gel spacer to encourage gel to separate from plate onto filter. Add onto gel in order: other Whatmann piece, sponge (remove bubbles), and other half of cage. Insert caged gel into blotting buffer and transfer (- from gel to + on filter) overnight at 60 mAmp (RT). Stop transfer, remove cage into transfer buffer. Use black marker to outline orientation of gel on filter, and remove filter.


1) Gel Sample Buffer 4% SDS 10% SDS ddH2O 5.8 ml 5.8 ml 0.5 M Tris, pH 6.8 3.2 ml 3.2 ml SDS 0.4 gm 1.0 gm glycerol 1.0 ml 1.0 ml bromophenol blue 0.2 mg 0.2 mg

2) Gel Running Buffer

Tris base 30 gm Glycine 144 gm SDS 10 gm - Make up to 1000 ml with ddH2O and pass through 0.2 m bottle top filter.

3) Transfer Buffer

Tris base 6.05 gm Glycine 28.8 gm - Make up to 1600 ml with ddH2O, dissolve, then add 400 ml of methanol.

4) 1.5 M Tris, pH 8.8

Tris base 91 gm - Make up to 450 ml with ddH2O, pH to 8.8 with approx. 10 ml of concentrated HCl, then make up to 500 ml and pass through 0.2 m filter.

5) 0.5 M Tris, pH 6.8

Tris base 30.3 gm - Make up to 450 ml with ddH2O, pH to 6.8 with approx. 25 ml of concentrated HCl, then make up to 500 ml and pass through 0.2 m filter.

6) Acrylamide:Bis (30:0.8)

Acrylamide 150 gm (wear mask) Bis 4 gm - Make up to 500 ml with ddH2O and pass through 0.2 m filter. Store in dark glass bottle.

7) 10 % SDS

SDS (gel electrophoresis grade) 20 gm - Make up to 200 ml with ddH2O, heat to dissolve and pass through 0.2 m filter.

8) 10 x PBS

KCl 2 gm, KH2PO4 2 gm, NaCl 80 gm ,Na2HPO4.7H2O 21.6 gm*, *or Na2HPO4(anhydr) 11.4 gm - Make up to 1000 ml with ddH2O. Autoclave.

Link: http://people.virginia.edu/~gwl6s/protocols/page.html