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General Overview
Equipment and Supply Modifications
Pre-Lab Preparation
Procedure Modifications
Trouble Shooting and Cleanup
Alternative Lab Ideas
Equipment and Supply Modifications
Tip: "To make the (electrophoresis) dyes more dense than the buffer, add 1 part glycerin to 10 parts dye. That will make them settle into the wells. For a totally different lab situation, I use 0.025 grams of dye to 10 ml. deionized water and 1 ml. glycerol. The point of this lab is to see separation of the dyes, but I think it would work for any situation."
-- Tricia Glidewell, Marist School, Atlanta, Georgia. 2/13/99
Question: "Does anyone know how long the E. coli, plasmid, and ampicillin will 'keep' when refrigerated? Years?"
Answer 1: "This year I tried using the plasmid DNA from last year, and it was a complete failure. I believe the Carolina transformation kit instructions say to use the plasmid within a month. On another note, the DNA that comes with Carolina's room temperature restriction enzyme kit for the electrophoresis lab keeps really well in the fridge. I used some from last year and it worked just fine. I would think the ampicillin would keep in the fridge, but you have to add the ampicillin to the agar below 55 degrees. So if you have already added the ampicillin to the agar, you cannot reheat the agar in the bottle to melt and pour new plates."
-- Franklin M. Bell, St. Mary's Hall, San Antonio, Texas. 2/11/00
Answer 2: "My experience is that Luria broth and CaCl2 will keep fine in the refrigerator, amp is fine only if in powder form. The DNA will probably knick and will be very hard to transform after that much time. Incidentally, the E. coli CAN be kept and will last a year if stored in vials 50 percent/50 percent with glycerol in the freezer. Just make sure that everything is sterilized."
-- Frank La Banca, Stamford High School, Stamford, Connecticut. 2/1/99
Answer 3: "The shelf life (of ampicillin) as a powder is pretty indefinite. I would check the concentration and make sure it was about 150 ug/ul. It is also sensitive to heat, so if you put it in molten agar at a temperature above 65°C it will denature. So, it can't be added to agar and then reheated."
-- Steve Plum, Bemidji High School, Bemidji, Minnesota. 2/02/01
Answer 4: "Heat breaks down ampicillin activity. Kept in a -20°C freezer, ampicillin should last a year. Kept in a refrigerator at 4°C, it should last many months. Repeated thawing and refreezing tends to speed up the breakdown. If you have a stock supply, I advise aliquoting it into a number of individual units so that you can thaw just what you need."
-- Pat Ryan, Carolina Biological Supply Company, Burlington, North Carolina. 2/02/01
Answer 5: "I did the Bio-Rad pGlo lab and I know the amp sat on a shelf unrefrigerated for two years! We had successful results."
-- G. Rad Mayfield, East Rutherford High School, Forest City, North Carolina. 2/02/01
Answer 6: "In my experience ampicillin as a powder lasts almost forever if it is kept in the dark. I have been told that it is light sensitive. Plates made with ampicillin last at least two months if they are refrigerated -- they go off if they are in the light, too. The disks are made for use by clinical labs so, by law, they have to have an expiration date on them. The ones I have from Difco were labeled as expiring six months after purchase, but I have used disks that were about a year old and they were still effective. They had been stored in the refrigerator."
-- Joan Kiely, SUNY, Stony Brook, New York. 2/02/01
Question: "Are there any good kits that can help with this lab?"
Answer 1: "For the past couple of years I have been unhappy with the results that I have obtained using Carolina's AP DNA fingerprinting and transformation labs. I have just finished these labs during a summer school class for students that fail our tenth grade class using Bio-Rad's bacterial transformation and DNA fingerprinting kits with excellent results. These kits are part of their Biotechnology Explorer program. I was using these kids as guinea pigs to see if they would be able to understand the procedures and collect decent data. The transformation lab had excellent results with a high degree of transformation efficiency. The transformants fluoresce under UV light and even students that failed our prerequisite class to AP Biology were impressed. The fingerprinting lab showed clearly visible bands with overnight staining that were easy to measure and interpret. For anyone that is looking for a kit that is well constructed, is complete, and gives you good results, you might want to check out Bio-Rad's product line (800-424-6723). I feel like I am a spokesperson for the company, but after the results of the last few years of mediocre data I am a happy customer."
-- David Knight, University High School, Irvine, California. 7/27/00.
Answer 2: "I have had success with the Bio-Rad kit, but I don't like the small petri dishes. Carolina now has a kit with glowing genes that has given me very impressive transformation rates the last two years. They call it the Green Gene transformation kit, cat #21-1082."
-- Israel Solon, Greenhill School, Dallas, Texas. 1/16/01
Answer 3: "I used the Glow-in-the-Dark Transformation Kit from Carolina Biological and got great results in November with sophomores. I'll be using it next week with my AP kids. It uses the pVIB bioluminescence plasmid; no x-gal plates are needed; it doesn't require an ultraviolet light like the Bio-Rad kits-the colonies just glow with the lights off. My students were amazed!"
-- Joni Driscoll, NW Cabarrus High School, Concord, North Carolina. 1/16/01
Answer 4: "It is interesting that you had success with the Carolina kit, yet not with Bio-Rad. I was contemplating a switch to Bio-Rad after hearing rave reviews at the NABT conference. My pAMP/XGal kit from Carolina was very expensive. The biggest disappointment was the refill kit I ordered this year. It only includes new perishables. The carbon fiber was only good for one lab (and was not replaced), and disposable inoculating loops, plastic petri dishes, etc. have to be ordered separately. Also, results were not very good with the dehydrated DNA...."
-- Anne Brewer, Mooresville High School, Bloomington, Indiana. 1/17/01
Answer 5: "I just finished the Bio-Rad kit, and I must say that it was EXCELLENT. Near perfect results! The booklet was easy to follow. Preparation was simple, and often very clever. Much better than Carolina's!"
-- Jon Calos, Emma Willard School, Troy, New York. 1/18/01
Answer 6: "I also have been attracted to the Bio-Rad kit. The transformation is much more obvious to the kids since the protein for the glow didn't belong to a bacterium from the beginning but instead belonged to another organism of a different kingdom. They can get a dramatic understanding of the transfer of a gene from one organism to another with this kit. Unfortunately, I too have been disappointed in the results. Sometimes there will be only one glowing colony on a plate. It makes me feel that I have spent my money for nothing. That's too bad since the kit is about $30 cheaper than the Carolina kits I usually use."
-- Tricia Glidewell, Marist School, Atlanta, Georgia. 1/17/01
Answer 7: "I have had great luck with all Bio-Rad products and kits every time. I have used their Transformation, Protein Purification, and Restriction Analysis kits for the past couple of years. I used Carolina kits previously but feel their quality and quality of service has dropped in recent years. Have a tendency to use Flinn Scientific more than in the past."
-- Paul Gardner, Holland Hall School, Tulsa, Oklahoma. 1/18/01
Answer 8: "...Those of you that mentioned disappointment with the Bio-Rad pGlo kit you had purchased a couple of years ago, most likely did nothing wrong. I have used them for years and about two years ago suffered the same fate. When I called them, they said they had had several folks say their kit didn't work and they thought they had a 'bad batch.' I've had great success with the kits ever since (except for the kids that don't read the instructions)."
-- Rene McCormick, Carroll High School, Southlake, Texas. 1/19/01
Answer 9: "I just finished a bacterial transformation lab from FotoDyne (Lumi-Kit 120). This kit transforms the plasmid containing the lux operon from Vibrio vischeri into bacteria. My kids got great results -- after letting our eyes adjust to the dark we saw bioluminescence. They were so excited! Today they isolated their bioluminescent colonies to create a whole petri dish of glowing bacteria."
-- Pebble Barbero, Highland Park High School, Dallas, Texas. 1/18/01
Question: "We do an electrophoresis lab with our first-year biology students to reinforce the sex-linked gene concept. I make up 40+ 1 percent agarose gels for the lab. The kids are running xylene cyanol and bromophenol blue dyes to represent the X-chromosomes with the normal dystrophin gene and with the DMD gene (dystrophin gene with a deletion). I usually cut off the well ends of the gels and save them to use next term as practice loading gels in my AP electrophoresis lab. As I was cutting off the remainder to dispose of them, I wondered if I could use that 'used' agarose -- melt it and recast it for next term's DNA lab. It is slightly blue from the dyes run through it on the other end, but I can soak that out if that is an issue. I can refrigerate it like I do my practice loading gels. What do you think? Can agarose be recast without losing something in the process?"
Answer 1: "Yes you can remelt and reuse old agarose. Having worked in a research lab and remelted agarose stock flasks in which I made more 2 percent more than I needed at the time. I've never needed to trim the ends off a gel to remelt that portion (your tax dollars at work), but I see no reason why that wouldn't be the same idea. Running the gel should not change it chemically. I've not tried to run the DNA off the gel in order to reuse the entire gel, but if money is an issue, that might be another strategy. Be careful that the gel doesn't sit in the refrigerator drying out too much -- the new plastic storage containers by Ziplock (or other companies) might work well for storage. That will change its concentration and it won't match the other gels. Don't store it in a buffer solution. I don't know what that would do to the gel over time. We just put the agarose flasks covered with parafilm in the refrigerator when in the lab. If you just use it for a practice loading gel, it won't matter if it is more concentrated or even contains old DNA, however, I've never been in a position to try using old gels over -- it definitely is worth trying with the practice loaded samples if you have an extra gel apparatus not otherwise being used. Also, If you only have a few casting molds, you can cast them, wrap them in Saran wrap, or put them in a plastic food storage dish and store them overnight."
-- Barb Fuller, William H. Hall High School, West Hartford, Connecticut. 1/21/01
Answer 2: "I just got out of an Amgen-sponsered mobio training conference. Two items pertinent to you: 1. Amgen runs their DNA off the ends of their gels and reuses them frequently. Their gels are prepackaged and very uniform, and they apparently don't want to waste time opening new packages. And, yes, gels can be reused. Just make sure the buffer doesn't get too hot from running too many gels in a row and be sure to mix buffer after running a gel. 2. The kit we're being trained on was left overnight in buffer after being used to separate some dyes. The dyes diffuse into the buffer and all is like new the next day. We used the xylene cyanol and bromophenol blue dyes, as well as Orange G. By the way, an earlier series of posting was describing DNA that ran ahead of the dyes. Apparently Orange G stays ahead of even the smallest DNA. 3. Several teachers talked about remelting and using agarose. I've done it as well. No has noticed any dysfunction."
-- Rich Smith, Buena High School, Ventura, California. 1/21/01
Answer 3: "You can reuse agarose gels with out recasting them -- we do it a lot in laboratories. The only problem I can think of with melting and recasting is the loss of water during the heating and cooling. You may need to add some water to keep the salt and agarose concentrations correct."
-- Joan Kiely, SUNY, Stony Brook, New York. 1/22/01
Answer 4: "After your gels have been analyzed and possibly photographed, I suggest that you place them back into the electrophoresis chamber and ensure that all the DNA has been run off the gel before melting and recasting. High voltages during electrophoresis might bring about a slight change in the consistency of the agarose when recycling it. However, I do not think this would have much effect on subsequent gel results for the classroom. I would caution against recycling agarose more than twice. Do not recycle agarose that has ethidium bromide, CarolinaBLU, or any other 'blue' stain in the gel itself."
-- Pat Ryan, Carolina Biological Supply Company Burlington, North Carolina. 1/22/01
Question: "Can you reuse electrophoresis buffer?"
Answer: "....The electrophoresis running buffer can be reused (at least 5 to 6 times). Most all of our electrophoresis kits use a 1X concentration of TBE. Some labs use TAE and that should be in a 1X concentration also. These are basically salt solutions and, therefore, you should be sure to shake the buffer solution to dissolve any salts that might have precipitated out during storage."
-- Pat Ryan, Carolina Biological Supply Company, Burlington, North Carolina. 1/24/01
Pre-Lab Preparation
Question: "I really messed up. I poured all of my Luria broth agar plates Friday and just realized that I did not put the ampicillin in any of them. So I would like to know if I can use nutrient agar with amp just as successfully or if I need to melt the agar, autoclave, put in the amp, and repour the plates. No time to order new Luria broth agar and no extra here at my school."
Answer: "You don't need to make new LB plates. You can put the amp on after the fact. We do it all the time. It is less than ideal because you may not have an even field of amp but it is good enough for a simple transformation. Figure the volume of the LB in the plate is about 25 mL; you want the amp to be 50ug to 100 ug/mL. You want to put 100ul of amp solution on each plate, so make a sterile solution of amp in water at 0.0125 mg/mL and spread 100 ul per plate with a spreader (just as you do the bacteria later). You now have plates containing 0.05mg/mL ampicillin."
-- Joan Kiely, SUNY, Stony Brook, New York. 2/29/00
Procedure Modifications
Question: "Can I incubate my plates over the weekend rather than just overnight?"
Answer 1: "Is this the blue colonies/beta galactosidase lab? If so, it's BETTER to let them incubate longer than 24 hours, because you get some interesting growth of white 'satellite' colonies in the zone around each blue colony. There are, if you will, opportunists growing in the area in which the engineered, resistant E. coli have secreted beta lactamase (I think that's the name of the enzyme) that breaks down ampicillin. Thus, untransformed cells are able to grow in those areas. It's a nice bonus for the students to figure out!"
-- Barb Beitch, Hamden Hall Country Day School, Hamden, Connecticut. 1/14/99
Answer 2: "You should not have a problem, however, the plates may dry out. You can incubate the plates at room temperature instead of over the weekend; that will slow the growth and prevent dehydration."
-- Frank LaBanca, Stamford High School, Stanford, Connecticut. 1/14/99
Answer 3: "If you leave the plates in the incubator for three days, the ampicillin may break down. I have found if I leave the plates in the incubator an extra day or so, satellite colonies form. This is either an extra, enlightening bonus for your class, or a source of confusion, depending upon their level of academia. There are two options you may want to consider: 1. Leave the plates at room temperature for three days. They should get as much growth as in the incubator for 72 hours. 2. Put the transformation tubes in the refrigerator on ice. This will give the bacteria that will face the ampicillin some recovery time to generate extra plasmids. Pour on Tuesday, incubate for 24."
-- Israel Solon, Greenhill School, Dallas, Texas. 1/14/99
Answer 4: "Do you keep a large container (e.g., beaker) of water in your incubator? That has always prevented plates from drying out in my lab."
-- Barbara Beitch, Hamden Hall Country Day School, Hamden, CT. 1/14/99
Question: "When your students do the electrophoresis lab, do you have any 'tricks' that help them see the wells more easily? My students have a difficult time finding the wells, especially in the smaller gels. I have them put the DNA in the wells when the gels are covered with the buffer solution in the electrophoresis chamber -- this is according to the directions. Are there other alternatives available?"
Answer 1: "I have noticed two things that help students to visualize using Carolina equipment and Carolina-Blu stain. One is to run a class demonstration after they collect data. Years of practice make this 40- year-old hand steadier with the Labpettes than the hands of my youthful charges (I am careful to only modestly voice this satisfaction). The second is voltage. I run the setup at a very low voltage so that it takes at least six hours to move across the gel. This gives tighter bands with less distortion. You will have to practice this to find out your apparatus' best voltage. Also, notice that the blue leading band goes off edge when the smallest DNA fragment is still only about 75 percent (conservatively!) of the way. Keep running it (be brave!) to spread the bands out."
-- Jerry Burke, St. Mary's School, Medford, Oregon. 2/3/00
Answer 2: "I save the leftover agar from previous years, to make practice gels for the next year. I put two or three combs across a petri dish, and pour practice gels. Pour some buffer across the top and you have a bunch of practice wells to use. You can make up some practice dye by mixing blue food coloring with a little bit of glycerin to simulate loading dye."
-- Frank Bell, St. Mary's Hall, San Antonio, Texas. 2/4/00
Answer 3: "Try putting some black (or even better -- red) electrical tape on the pan under where the wells will be. It will contrast much better."
-- Frank LaBanca, Stamford High School, Stamford, Connecticut. 2/4/00
Answer 4: "I put the gels on my overhead projector, which shines light through them, and this makes the bands stand out."
-- Bonnie Polan, Beverly High School, Beverly, Massachusetts. 2/4/00
Answer 5: "I find that the students can see the wells more easily if they view them at about 45 degrees (instead of looking straight down on them) or if they have a light at about the same angle. Other students like to put blue paper under the chamber. I find the best way is to barely cover the wells with buffer, load the wells, and then gently top up the chamber with buffer, from the opposite end to the wells."
-- Peter Gardiner, St. Michaels University School, Victoria, British Columbia, Canada. 2/4/00
Answer 6: "We have lots of different kinds of agar donated by local hospitals that never gets used, so I use this as practice gels for electrophoresis and save the expensive agarose for the real DNA. It works great. Just be sure to rinse the trays and combs well afterwards."
-- Carole McRight, Springdale High School, Springdale, Arizona. 2/4/00
Question: "How long do you run electrophoresis gels?"
Answer: "I usually run at 110 to 120 volts for about 60 to 90 minutes. I'm not sure how long your gels are, but that seems to work. I can usually do two classes in a day that way. I stain them myself and they are ready the next day for the students."
-- Israel Solon, Greenhill School, Dallas, Texas. 2/5/99
Question: "Is it possible for the DNA fragments to run ahead of the loading dye? The last two gel sets we have run seem to have lost the lower bands. The upper bands are way down the gel. The first set we ran was for 72 hours (!!!) at 12 volts (not ideal, I know). The second was for two hours at 100 volts. In both cases the dye was still migrating on the gel. I thought 30 minutes was sufficient at 100 V, but the dye had only moved one-fourth the distance or so."
Answer 1: "In agarose gel electrophoresis it is possible for DNA fragments to run ahead of the dye. The migration rate of the bromophenol blue is determined by your electrophoresis conditions -- salt concentration, voltage. Under the most commonly used conditions, bromophenol blue appears to be 250 to 300bp so a 200bp fragment of DNA will travel faster. I think that the migration of the dye molecule is determined mostly by the charge on the molecule, not its size. In polyacrylamide gels for DNA the apparent size of bromophenol blue varies from 12bp to 100bp depending on the acrylamide concentration. I do not know how much the agarose concentration affects dye migration in agarose gels. In SDS-PAGE for protein analysis, the dye travels ahead of almost all peptides. But that has to do with the physics of the system."
-- Joan Kiely, SUNY Stony Brook, New York. 1/15/01
Answer 2: "According to DNA Science by Micklos and Freyer, bromophenol blue electophoreses at about the same rate as a 300bp piece of DNA. Xylene Cyanol electophoreses at about the same rate as a 9000bp piece of DNA."
-- Pat Ryan, Carolina Biological Supply Company, Burlington, North Carolina. 1/15/01
Answer 3: "I have units from edvotek (1800 edvotek), which have volts for 25 and 50. Using 50 volts and two hours, the samples turn out great and I've never lost DNA. Are you using 7x15 or 7x7 gels? That could make a big difference as to how long and how much could be lost. I also wanted to add that edvotek will fix and replace many items for nominal costs or free! We had a teacher exchange here a couple years ago and the exchange teacher ran the water bath dry and blew the heater coil. All I paid for was shipping."
-- Kim Armitage, Vicksburg High, Vicksburg, Michigan. 1/15/01
Trouble Shooting and Cleanup
Question: "How do you dispose of used petri plates?"
Answer 1: "Autoclaving is best. If your school cannot afford an autoclave, a good electric pressure cooker (a la Sears) will do the trick... 14# pressure... 15 minutes). You can soak them in 10 percent bleach but that does not kill spores. What if you had anthrax bacilli? These are 'lean, mean, spore-forming machines.' I don't mean to be so lightheaded about it but spore-forming organisms can be as dangerous as toxigenic E. coli."
-- Stu Schnell, John C. Freeman High School, Los Angeles, California. 11/30/99
Answer 2: "Just a suggestion for everyone worrying about used plates. Form a good association with your local hospital. Ours is a designated 'friend of education.' They not only provide us with agar plates and biohazard bags but I return them to the hospital in the biohazard bags and they dispose of them for me along with their waste."
-- Marla Vaughn, Oroville, California. 12/6/99
Answer 3: "I used to take my plates to the microbiology lab at a nearby college. They were kind enough to autoclave them. Some people also pour 10 percent bleach over them, then discard them. However, the laws are changing, and in Connecticut we now have an arrangement with a toxic waste outfit that supplies us with a container lined with plastic. When the container fills up, we call them, and they come pick it up -- for a charge (I think we've arranged something like $50 per pickup). It takes a small school like us a long time to fill up the container. As long as you close it thoroughly and contain the smell, it's not so bad. I found this outfit in the yellow pages. We had to make a few calls before we found a company willing to pick up what they call medical waste."
-- Barbara Beitch, Hamden Hall Country Day School, Hamden, Connecticut. 12/1/99
Answer 4: "If you don't have an autoclave or pressure cooker, you can disinfect plates with a 10 percent Clorox solution. Cover and let them sit for 10 minutes."
-- Bruce Faitsch, Guilford High School, Guilford, Connecticut. 12/1/99
Answer 5: "Microwaving is not a safe alternative to autoclaving in my opinion. It doesn't produce enough heat to kill spores, and it often heats the subject unevenly. Use an autoclave or one of the commercial disinfectants that is designed to kill spores. There are professional products on the market that totally disinfect, including killing spores. Wavacide is one such product. The label says that it is a sporicide when used at the recommended strength. I have used it, and it works as far as I can tell. I limit its use to environment, i.e., countertops, floor, etc., if an accidental spill occurs. If you work with bacteria, you should be prepared for spills because they will happen in a classroom, guaranteed! If you are talking about household disinfectants, I agree -- they do not kill spores -- in fact, they often do not totally kill bacteria vegetative cells. Autoclaving is the preferred method of safe disposal. For teachers not trained in microbiology they probably should use a professional disposal service. I would appreciate any information anyone can provide about a law addressing this issue and, if so, is it a state law, federal law, or just guidelines, because it has been my professional opinion that correctly trained teachers with the proper equipment (autoclave) would be able to safely dispose of bacterial waste. In fact, waiting weeks or even months for a professional disposal is not recommended because the wait may increase the contamination risk."
-- Bruce Faitsch, Guilford High School, Guilford, Connecticut. 4/10/00 and 4/11/00
Question: "Can anyone give me the 'party line' on generating a HindIII standard curve. A few of my students noticed that the EcoRI fragment sizes are much more accurate when the standard curve is generated by just 'connecting the dots' as opposed to constructing a line of best fit. The first data point on the HindIII curve, the largest fragment, is usually way out of line with the other points. So what is it? Is it a 'line of best fit' or 'connect the dots'? On a few university sites, I've read that connecting the points gives more accurate results."
Answer 1: "When I did this same lab as a student in a graduate molecular biology course, I was told the function was parabolic and to use a best fit curve rather than a best fit line. I have my students in AP Biology do it with the curve and it works great."
-- Marcia Sloan, Cleburne High School, Cleburne, Texas. 1/10/01
Answer 2: "I was taught to drop the top point (the large fragment) in a HindIII curve when using a 1 percent gel because that gel concentration works best for mid-size fragments. To separate larger fragments, a less concentrated gel would be used; to separate smaller fragments, a more concentrated gel would be used -- thus the various protocols for gel preparation, depending on what size fragments are of the most interest. This gives good results for us. BTW -- a few years ago as graphing calculators were being 'pushed' as new technology, students would use those for a best fit curve and got good results. However, since they can't use those on the AP Biology Exam, I also made them use the graph paper. It was amazing to see how many very bright kids couldn't use the graph paper. Now I won't even let them get their calculators out!"
-- Ellen Mayo, Mills Godwin Specialty Center for Science, Mathematics, and Technology, Richmond, Virginia. 1/10/01
Alternative Lab Ideas
Other Electrophoresis Labs
Tip: "If you already have the electrophoresis equipment, your first- year biology students will love doing 'Rainbow Electrophoresis.' Use the buffers and gels exactly as for DNA electrophoresis, but fill the wells with food colors. Nothing needs to be added to the food color. The separations will be beautiful colored bands. I have my students wrap the gel in plastic and use colored pencils to sketch the gels. This must be done shortly after separation, as the dye will diffuse through the gel if you wait until the next day. This is also a cheap way to let your AP students learn to load a gel and demonstrate the principles of the apparatus. This activity is written up in the 1993 Woodrow Wilson biotechnology module."
-- Theresa Holtzclaw. 2/12/99
DNA Extractions
Question: "Can you help on DNA extraction protocol?"
Answer 1: "I use dog testes (for DNA extraction). I usually get a big batch and freeze them and they have lasted for years. Be sure to ask for non-preserved ones from your vet and from mature dogs (not puppies). I bought the EDTA and SDS the first year and still have it. It takes so little that I have a lifetime supply and, in fact, I keep the solutions refrigerated and often use them the next year with great results."
-- Charlotte Freeman, Girls Preparatory School, Chattanooga, Tennessee. 11/3/99
Answer 2: " Place a small piece calf thymus (sweetbreads from the store) into a mortar. Cut the sample with a pair of scissors into smaller pieces. Grocery stores carrying pigs feet, calves brains, and such usually have sweetbreads; in Texas, Fiesta Foods and HEB carry them.
- Add 10 mL of 0.9 percent NaCl solution and grind with the pestle for 2-5 minutes.
- Strain the solution through 3 to 4 layers of cheesecloth into a test tube. (0.0 g of NaCl in 100 mL of water). Keep the tube with the suspension.
- Add 1.5 mL of 10 percent sodium dodecylsulfate (SDS) to cell suspension and mix. Lacking SDS add 3 or 4 drops of Dawn dishwashing liquid (it contains SDS). The SDS lyses the cell membrane by dissolving the lipoproteins in the cell membrane.
- Measure the total sample volume, then measure out two times that amount of ice cold 95 percent ethanol. (I keep it in the freezer prior to using.) Down the side of the tube, gently add the ethanol to the suspension.
- Use a glass stirring rod to gently stir the mixture until the DNA begins to precipitate at the interface. It usually precipitates as you add the EtOH. Then twist the glass rod to spool the DNA onto the rod. The precipitated DNA can be transferred to a new tube containing 95 percent ethanol and stored indefinitely in the freezer. From Texas Biotechnology Teacher Enhancement Project -- Texas A & M."
-- Nancy Hein, Hawley High School, Hawley, Texas. 11/3/99
Answer 3: "If you are determined to try DNA extraction with thymus gland, try an ethnic market. In the Houston area, I buy them at Fiesta. The other day I was amazed to see it for sale at Kroger. (Under its pseudonym of 'sweetbreads,' of course.) I did a DNA extraction with thymus in my classes for several years, then switched to doing it with onions. It works better because you can always get fresh material. Thymus that has been around too long or has been frozen and thawed several times doesn't work well. Bacterial cultures can also be used as a source of DNA."
-- Alexa Noble, Oak Ridge High School, Conroe, Texas. 11/3/99
Answer 5: "I have this recipe thanks to Dr. Jeff Smith of the Indiana Academy: For each ripe banana, peel and place in blender. Cover with 15 percent NaCl solution, add two drops of dish soap, and blend to smithereens! Next, pour through several cheesecloth layers into a beaker. Add a pinch of meat tenderizer (contains enzymes) to the beaker to break down the proteins. Divide the solution in the beaker into test tubes (fill them halfway). Let the precipitate settle out (which takes 15 to 20 minutes). Use ice cold (put in freezer overnight) 95 percent ethanol. Layer the alcohol on top of the banana mixture and stir the area between the layers with a glass stirring rod or wooden splint. The DNA should stick to the stirring rod! Note that:
- Blending breaks up the cells.
- The salt water denatures the proteins that could digest (lyse) the DNA.
- The banana is triploid.
- The soap acts as a surfactant to get the membranes, etc. out of the way."
-- Julie Smiley, Winchester Community High School, Winchester, Indiana. 2/23/00
Answer 7: "There is a terrific and easy DNA extraction from wheat germ that I did even with an at-risk biology I class. The procedure is from the Hughes Undergraduate Biological Science Education Initiative. In a 50 mL test tube, add 20 mL hot water (50-60° C) to 1 gram raw wheat germ and rock it by hand for 3 minutes; then add 1 mL Lemon Fresh Joy or Woolite and rock gently every one-half minute for five minutes (to avoid foam -- remove any you form by pipette). Tilt the test tube and SLOWLY pour 14 mL 95 percent ethyl alcohol (rubbing alcohol can be used but precipitates out less DNA) so that it forms a layer on top of the wheat germ solution. After letting it sit for a few minutes, a stringy film of DNA should form at the interface."
-- Carolyn Schofield, Robert E. Lee High School, Tyler, Texas. 1/30/99
Answer 4: "I got a DNA isolation procedure from Scientific American: it works very well for spooling DNA (Scientific American Set 1998, The Amateur Scientist 1998). You could probably clean the DNA up with a salt and ethanol precipitation to run it on a gel. I am embarrassed to admit that I just realized this may not be possible in a high school laboratory because you need a medium speed (10,000xg) centrifuge. Scientific American had directions on how to make one from a blender, I don't know if a clinical centrifuge would work. I don't know how well this will work with spooled DNA -- I will be trying it -- but in research labs people precipitate DNA a lot to change the suspension solution or just to remove impurities. So, if you are still interested, to precipitate DNA:
- Take your spooled gob, air dry, and then dissolve it in water or saline. If you add 1mM EDTA and use a buffered solution that may stabilize the DNA, but it is probably not necessary. The DNA strands are long, so if you mix it vigorously the strands will break. I do not think this will be a problem, but it may cause a smear on the gel, so be gentle.
- Add sodium acetate (pH5.0) to get a final sodium acetate concentration of 0.3M -- most people use a 3M sodium acetate stock solution, buffered with acetic acid to pH5.0. Then if you have 1 mL of DNA, add 0.1ml of the concentrated acetate.
- Then, add ethanol to a final concentration of near 70 percent (2.5 mL for 1 mL of DNA). You will start to see the DNA come out of the solution and you can chill it in the freezer to improve the precipitation.
- Finally, centrifuge it at 10,000xg for five minutes or so -- maybe longer in a clinical centrifuge would work -- and you have a glossy pellet of clean DNA that you can re-suspend in a buffer for electrophoresis or enzyme digestion.
Total genomic DNA will often give you a smear because the DNA molecules are so long and broken randomly."
-- Joan Kiely, SUNY, Stony Brook, New York. 2/9/00 and 2/11/00
Tip: "Try doing your onion DNA isolation without chopping them in a blender -- if you did that. I use just a sharp knife and it works pretty well. Also you might try kiwi fruits. Smash them with a large spoon in the detergent solutions you use for onions and they work really well."
-- Doug Herman, East High School, Sioux City, Iowa. 1/29/99
Question: " Is there an 'over-the-counter' substitute for sodium dodecylsulfate (SDS) used in DNA extraction protocols?"
Answer: "I use regular old liquid dish soap. It does the trick." -- Bob Heun, Brooks School, North Andover, Massachusetts. 12/2/99
Answer 6: "According to the Genetic Science Learning Center at the University of Utah, these detergents should work (for DNA extraction): Lemon Fresh Joy, Woolite, Ivory, Shaper, Arm & Hammer, Herbal Essence shower gel by Clairol, Tide, Dish Drops, Kool Wash, Cheer, Sunlight Dish Soap, Dawn, Delicate, All, and Ultra Dawn."
-- Bob Heun, Brooks School, North Andover, Massachusetts. 2/23/00
Tip: "One of the tricks I've discovered is to use wood splints for spooling, rather than glass rods. We save it in capped vials in 50 percent ethanol for years. It stays in the refrigerator (although that probably is not necessary) and keeps looking cool. I think Doris Helms' lab manual says to use 60° C rather than 40 to 50."
-- Barbara Beitch, Hamden Hall Country Day School, Hamden, Connecticut. 1/29/99
PCR Techniques
Multipart Question: "My AP Biology classes just finished our first try with PCR. We amplified the PV-92 loci for the alu sequence using the kit from Carolina. How do you handle the data within the context of the AP curriculum? Do you use the Web site to look at other populations and their Hardy Weinberg information?
Answer: "I would suggest that you use the DNA Learning Center's Web site because of all the lab extensions that can be done there."
"On some of the gels we got 'streaking,' many bands blurred, with the 550 and 850bp bands showing more prominently. Any ideas on how we can clear up the extra bands? The lab recommends a 'hot start.' Does anyone else do that and how do you handle it?"
Answer: "Keep everything on ice and bring the ice containers with samples to the thermocycler. Have the thermocycler prewarmed to 94° C. Load all samples as quickly as possible and start the program."
"If we finish the PCR one day and want to run the gels the next, do we need to store PCR products at 20 degrees or -4?"
Answer: "Store on ice in the refrigerator (0° C)."
"Finally, what is the latest word on the mutagenic properties of ethidium bromide? I was very cautious about student contact. However, I still worry about the risks."
Answer: "The EtBr used in this lab is an extremely weak solution. However, I recommend that the teacher is the only one to handle anything dealing with EtBr. The teacher should wear gloves and perform good laboratory practice. If possible, take pictures of the gels for the students to analyze."
-- Pat Ryan, Carolina Biological Supply Company, Burlington, North Carolina. 2/26/ 01
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