This study was designed to introduce students to the principles of tissue and cell fractionation, including differential centrifugation and protein determination. As a sideline they are also able to prepare blood smears and observe red cells, platelets, and white blood cell types. Students take an intact tissue, namely whole blood, and obtain a fraction enriched in red cells. The erythrocytes themselves are taken apart and the membranes isolated. Finally, in a subsequent laboratory the membranes themselves are taken apart and their proteins analyzed by polyacrylamide gel electrophoresis.
Human blood is readily available, but since the beginning of the AIDS epidemic it has become too much of a liability for use in teaching labs. Rodent blood is acceptable, but adequate quantities can be difficult to obtain, although a rabbit might provide a sufficient amount. Dogs are a safe source of blood provided samples can be obtained from a research colony or veterinary hospital. If blood from pound animals can be used there is often an added bonus, namely the presence of heartworm microfiliariae.
For instructor/teaching assistant use:
At the side benches:
Graph paper for standard curves, or one or more computers with graphics software
Starting with three ml whole blood and diluting with 10 ml saline solution gives satisfactory results. Student should perform the fractionation directly in a centrifuge tube, marking the level in the tube after the first dilution then filling to the mark thereafter. Resuspension should be accomplished by thorough trituration of the pellet with a pasteur pipet.
Resuspension of the red cell pellet in 5 mM Na phosphate without NaCl should clarify the suspension. It remains red, but is a clear rather than cloudy red. The first high speed pellet may include unlysed cells, since as the cells release their contents the osmolarity goes up. The second resuspension with 5 mM Na phosphate will complete the lysis. The membrane pellet is the same color as the supernatant (membranes themselves are colorless), so care must be taken not to take up the pellet when removing the supernatant. If there is any doubt, the student should stop removing the SN as he/she nears the bottom of the tube, and resuspend. The next pellet will be easier to see as the color of the SN becomes less intense. The small, fibrous part of the pellet consists of clotted matter (platelets, fibrin, white cells) and can be removed any time.
The Bradford assay was chosen for this introduction to protein determination because of its simplicity, and because the color of hemoglobin apparently does not interfere with the assay. The assay can be quickly repeated if protein samples are out of range.
The first set of absorbances for unknowns may include values outside the range of the standard curve. Actual protein concentration depends on the students' efficiency in performing the fractionation, as well as the hematocrit of the blood sample and actual amount used. Recommended dilutions of unknowns for the assay should get the students into the ballpark. They should be made well aware that their concentrations could be too great or too small. The most variation is in the concentration of the membrane pellet. Since this is the most important sample and there may not be much of it, students must be made aware that they should have a reasonable amount left after doing the assay. Students should make it a practice to flick or vortex any tube prior to sampling, especially if the material has been previously frozen.
Microscopic examination of the blood sample (cytology) is not an integral part of the protein preparation. However examination of whole blood is a valuable experience, and this gives the students an opportunity to practice their skills with microscopy.
At benches, per pair students, Nikon Labophot microscope, equipped with phase contrast, dark and bright field modes, 4x, 10x, 40x, and 100x objectives, plus accesories including immersion oil and lens paper
At the side benches:
Improper use of anticoagulant can ruin the appearance of white blood cells, and the cells begin to deteriorate almost immediately regardless of treatment. The cells may last several days if the sample is sterile and refrigerated, however fresh blood is by far the best. Staining intensity may vary from batch to batch. The stain should be tried before giving to the students. The dipping procedure for WrightÕs stain seems to work best.
Students are required to work in pairs. One student should be primarily responsible for the protein assay while the other is primarily responsible for the fractionation. Both must keep track of work performed by the other, and record all procedures and observations in the notebook. Students should turn in samples at the end of the day, to be frozen for the following week.
If computers and a suitable graphics probram are available the protein standard curves can be run in lab and checked by the instructor and/or a teaching assistant. Students must have completed the assay and obtained protein concentrations before leaving lab.
Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) has become an indispensible technique for many disciplines, in fact, most researchers in the biosciences use some form of electrophoresis in their work. The object of this study is to introduce the students to the nature of protein gel electrophoresis and to learn to apply the technique to a well-studied system. This study is designed to follow the fractionation of erythrocytes.
At benches, per pair students:
At the side benches:
Students will have one set of samples/pair, of known protein content. Each pair should have calculated, as a homework assignment, the amount of undilute sample, dH2O, and 2x sample buffer to mix in order to obtain the desired concentration of sample (usually 1 mg/ml) in 1x buffer. Generally, 100 to 200 µl of sample should be prepared, so that 20 µl of the same sample can be loaded onto each of two different gels with plenty of sample to spare. The algorithm for the calculation is given in the on-line lab manual. Samples are prepared in Eppendorf tubes, mixed, heated 10 min. at 60ūC, and kept at room temperature until the gels are ready for loading. Samples can also be frozen and thawed later for running on gels, but repeated freeze/thawing of samples should be avoided.
Sometimes students over-dilute the membrane fraction. If the concentration of protein in the undilute sample is less than 2 mg/ml have the student simply mix a volume of sample with an equal volume of 2x buffer, and record the final concentration and amount loaded. All previously frozen samples should be thoroughly mixed prior to removal of contents, since freeze-thawing can result in layering of solutions of differing concentrations. Some insoluble material may result from freezing of samples.
At benches, per pair students:
In each fume hood:
At the side benches:
Vacuum pumps with traps & tubing to one-hole stopper to fit 125 ml flask
Incorrect cassette assembly usually results in leaks. The risk of contamination is reduced by keeping the casting stand on the bench paper. If the resolving gel is poured too high there isn't sufficient room for the stacking gel. The well former sometimes fits snugly between the plates, and students need to practice inserting it before pouring stacker. Failure of a gel solution to polymerize is the most common problem. It results from improper mixing of catalyst, failure to add catalyst, or adding the wrong amount of catalyst. It is especially important to remind students that since the concentration of TEMED is critical, they should inspect the pipet tip to ensure that the proper volume is drawn up. If old acrylamide solution is poured out to be replaced by a fresh batch of acrylamide, plates should be held in place by a gloved hand or they will fall out and break.
Polymerized acrylamide left in flasks should be rinsed and pried or shaken out before it dries up.
At benches, per pair students:
At side benches:
For instructorÕs use:
Samples should be prepared by one student, and cassettes assembled and gel solutions prepared by the other. Both students share responsibility for all procedures, however by delegating responsibility they can finish quickly. Each pair of students is to prepare several gels, and the pair will use the best of them, with remaining gels kept for those whose preparations failed or for demonstration purposes. Gels saved for demonstration of how to take them apart should be immersed in water to keep them from drying out.
Each pair of students will share a gel with the pair opposite them. One pair pours a low density gel, the second a high density gel. Both sets of samples and appropriate molecular weight standards will be loaded onto each gel. If samples are prepared right away, the faster pair can load their gel with both sets of samples, and start their run without delay.
It helps to have students put their names on a piece of tape and label the lid of the gel apparatus prior to running. The tape can then be transferred to the lid of the staining box in order to identify the gel later.
Power supplies should be distributed conveniently around the room, each with at least two sets of cables. An instructor or teaching assistant should check out the connections before applying power, since inproper connections can result in failure of the run or, worse, damage to the gel apparatus or injury to a student.
Potential problems include delays due to one group being slower than another. This can be prevented by permitting each group to set up its own gel, and loading all samples. The samples are then left for the slower group, and that gel can be hooked in while the first is running. To make this work, the teaching assistants must ensure that all groups have samples prepared. Sample preparation is much quicker than gel preparation, but they often need help with the calculations.
Loading problems include shallow wells, which result from inadequate polymerization (see above) or incorrect well former position. Sometimes acrylamide will polymerize in the wells after removal of the well former. This happens when the well former is thinner than the spacers. It helps to leave the well former in place until the cassette is immersed in buffer, then removing it and allowing the buffer to fill the wells. Samples are loaded under the buffer with a Hamilton syringe. This same syringe can be used to straighten crooked wells.
The outsides of gel cassettes and insides of the upper electrophoresis chambers must be clean and dry before sealing in with agarose or leaks will occur. Agarose should be applied to the gap while the chamber is horizontal so that the liquid agarose doesn't run through, then sealing is completed with the chamber upright. The bottoms of the wells should be visable above the agarose layer, but the plates must be covered with buffer when the chamber is filled, without having buffer run out the hole for the cathode. The buffer level in the lower chamber must cover the bottom of the plates. Both anode and cathode must be in contact with buffer. If adding a gel to the power source does not increase the current, then there is an open connection someplace.
Leaks during a run can be annoying, but are readily corrected. If the chamber leaks during the run, the apparatus must be removed from the power supply, the upper chamber drained, dried, and the cassette resealed. Samples should be into the gel and o.k. by this time.
Run time is about 1 hour, during which the students can leave. They do need to return to take apart and stain the gels, meanwhile instructors must make sure that the dye fronts do not run off. Dye fronts for low density gels are straight and sharp. For high density gels they are more diffuse, and all indications of a dye front disappear during destaining. To avoid confusion, the staining trays should be labelled with both students' names and with the students' names with whom they were partnered.
When removing gels, the plates are separated by twisting with a spatula or spacer and the lower left corner of the gel is cut off to mark the first lane. Students should note the position of the dye front at this time. The plate is turned upside down over the staining tray, containing deionized water, so that the gel is parallel with the water surface. The gel is coaxed off of the plate with a spatula or spacer so that it drops in, then the gel can be rinsed, the water poured off, and stain added. The extra gels can be used to demonstrate the technique.
Any fingerprints on the gel will stain, so the gel should be handled with gloves only. Gels will slip out of the tray while pouring off solutions, thus they must be held in place while pouring. Staining/destaining protocol is 1 hr in staining solution (or overnight), 1 hr with destain I, then destain II until background is clear. Gels shrink considerably in destain I, but will swell up again in destain II. Destain II can be changed more than once, and gels can be kept indefinitely in this solution.