Recordkeeping, Writing,
& Data Analysis


Microscope studies

Flagella experiment
Laboratory math
Blood fractionation
Gel electrophoresis
Protein gel analysis
Concepts/ theory
Keeping a lab notebook
Writing research papers
Dimensions & units
Using figures (graphs)
Examples of graphs
Experimental error
Representing error
Applying statistics
Principles of microscopy

Solutions & dilutions
Protein assays
Fractionation & centrifugation
Radioisotopes and detection

Guide to the study

Lab part 1

Lab part 2

Lab part 3

Selected methods



Assembling, Loading, and Running Gels

Cassettes should be rinsed free of any excess liquid, leaving the combs in place. If casting stands are used, the clay is scraped off of the front cover and the cover removed. Gel cassettes are separated with the aid of a single edged razor blade if necessary (having beveled plates helps). After scraping off any excess stacking gel, the surfaces of the plates must be rinsed and dried, and the best gels selected. Small air spaces may appear between stacking gel and resolving gel or between the gel and the glass plates. As the outside pressure on the plates is relieved the glass expands, creating some spaces. As long as there is no continuous channel from the top to the bottom of the gel, the spaces will not influence protein migration.

The assembly of a gel running stand varies with the type of apparatus. The top of the cassette must be continuous with an upper buffer chamber and the bottom must be continuous with a lower chamber so that current will run through the gel itself. The cassette must be sealed in place using gaskets or a sealant such as agarose. In a teaching lab the assembly is best described by going through the procedure, using a film, and/or having a demonstration set up. We fill both the upper and lower buffer compartments with an electrode buffer (running buffer) consisting of 25 mM Tris, 192 mM glycine, 0.1% sodium dodecyl sulfate (electrode buffer composition is part of the Laemmli method). We do not adjust pH of the electrode buffer. We remove the comb from the gel before filling the upper buffer compartment.

Loading gels

Hamilton syringes work well for loading samples into the wells. Ideally, the glycerol in a sample causes it to sink neatly to the bottom of the well, allowing as much as 20 µl or even more to be loaded. If the combs do not fit well or the plates are not clean the sample often hangs up, and we are limited to 10 µl or so.

Running gels

The anode (+ electrode) must be connected to the bottom chamber and the cathode to the top chamber. The negatively-charged proteins will move toward the anode, of course. Gels are usually run at a voltage that will run the tracking dye to the bottom as quickly as possible without overheating the gels. Overheating can distort the acrylamide or even crack the plates. The voltage to be used is determined empirically. We run our gels at 150 volts.

Notes on gel assembly and running

  • Criteria for a good gel include straight spacers, top, bottom of separating gel parallel, straight wells, appropriate depth of stacking gel.
  • Agarose will not stick to wet surfaces, so plates and apparatus must be completely dry before sealing; bubbles in agarose will eventually cause leaks.
  • Agarose alone will not hold a gel in place - the cassette must be secured in place.
  • We have found that the lane dividers are less likely to be distorted if we remove combs before the upper chamber is filled.
  • Lanes become distorted if the samples sit in the wells for too long before running.
  • The gel can't be rescued if the voltage is run backwards for any significant length of time.
  • If the upper chamber leaks out, the gel can be 'rescued' provided samples have entered the gel - the cassette is removed, everything dried, cassette re-sealed in place, buffer re-added, and electrophoresis resumed.
  • The apparatus should be placed in a tray in case of leaks, and not touched while the voltage is on.

Disassembly and staining

When the dye front is nearly at the bottom of the gel it is time to stop the run. For low percent gels with a tight dye front, the dye should be on the verge of running off the gel. When the percent acrylamide is high the dye front may be diffuse, since the dye is not homogeneous. If you know the approximate position of the lowest protein band you can let the dye run off. Only experience will tell you when it is appropriate to stop the run. Before removing gels the power must be turned off and cables removed (using one hand, to avoid making a circuit).

Removal the gel from the cassette is better demonstrated than described. The plates are separated and the gel is dropped into a staining dish containing deionized water. After a quick rinse, the water is poured off and stain added. Staining usually requires incubation overnight, with agitation.

Staining protein gels

A commonly used stain for detecting proteins in polyacrylamide gels is 0.1% Coomassie Blue dye in 50% methanol, 10% glacial acetic acid. Acidified methanol precipitates the proteins. Staining is usually done overnight with agitation. The agitation circulates the dye, facilitating penetration, and helps ensure uniformity of staining.

The dye actually penetrates the entire gel, however it only sticks permanently to the proteins. Excess dye is washed out by 'destaining' with acetic acid/methanol, also with agitation. It is most efficient to destain in two steps, starting with 50% methanol, 10% acetic acid for 1-2 hours, then using 7% methanol, 10% acetic methanol to finish. The first solution shrinks the gel, squeezing out much of the liquid component, and the gel swells and clears in the second solution. Properly stained/destained gels should display a pattern of blue protein bands against a clear background. The gels can be dried down or photographed for later analysis and documentation.

The original dye front, consisting of bromphenol blue dye, disappears during the process. In fact, bromphenol blue is a pH indicator which turns light yellow under acid conditions, prior to being washed out. In low percentage gels, sufficient protein may run with the dye front so that the position of the bromphenol blue front is permanently marked with unresolved proteins, often forming a continuous "front" across the bottom of the gel. In higher % gels, a distinct dye front is usually not obtained.

Coomassie blue may not stain some proteins, especially those with high carbohydrate content. Stains such as periodic acid-Schiff (PAS), fast green, or Kodak 'Stain's all' may detect different patterns. Silver staining is generally used when detection of very faint proteins is necessary.

Routine staining with Coomasie Blue is straightforward - about the only ways to ruin a gel at this point are physical damage (ripping the gel, for example), letting dye pool and precipitate in the gel, forgetting the alcohol at some step, allowing protein to dissolve and diffuse out of the gel. If that happens, the information is lost.

What's next?

Now that you have a stained gel, what do you do with it? The next part of this study will focus on gel analysis. Before tackling the details you might review the material presented at the very beginning of part 1, on the cytoskeletal structure of the mammalian red blood cell membrane. The known structure is your starting point for identifying proteins on your gel. If you have been there already, then you can move on to the gel analysis section.

Copyright and Intended Use
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Created by David R. Caprette (caprette@rice.edu), Rice University 14 Aug 96
Updated 24 May 05