Abstract:
Learning the Polymerase Chain Reaction technique is best accomplished by performing it, and hand-cycling with water baths eliminates the need for an expensive thermal cycler. Teachers who want this experience for lots of students can do so most economically by ordering PCR beads, primers, and DNA; this article will give you the know-how.
Polymerase Chain Reaction (PCR) has rapidly become a commonplace biotechnology procedure with all kinds of variations: standard PCR, real time PCR, reverse-transcriptase PCR, AFLP PCR, Colony PCR. For the classroom, PCR is not a difficult concept to understand and students benefit from knowing something about this very important technique. Teaching PCR also reinforces learning of the base pairing rules, DNA strand directionality, and DNA replication. There are plenty of animations of PCR on the internet, and there are desktop activities also available for teaching PCR, but actually running the experiment is undoubtedly the best way to learn about it.
However, one barrier to a “wet” PCR lab is the instrumentation: thermal cyclers, even the classroom versions, are quite expensive. At our site, we have been using a water bath version of PCR. Using water baths eliminates the need for the expensive thermal cycler, but even more important, it is hands-on practice that reinforces learning the steps of PCR. Our website protocol for water bath PCR has evolved from the original version published by Bloom, Freyer and Micklos (1996). The method uses two “water baths”, one at 55 degrees C., the other can be a boiling beaker of water. Students cycle the PCR reaction tube by hand, 20 seconds at boiling, 1 minute at 55 degrees C., back and forth 25-30 times. This crude method for temperature cycling works because the target DNA is on the small genome (48,502 base pairs) of the bacteriophage Lambda.
Another barrier to doing PCR with students is the expense. In our Lambda PCR version, the DNA is inexpensive, primers don’t cost much, but the expensive item is the ready-to-go PCR bead. The PCR beads cost approximately $1.40 per reaction, sold in 100 reaction packages. If a teacher is going to run PCR with lots of students, our method is very cost-effective. A teacher could order the 100 PCR beads, the primers, and the DNA at a cost of about $1.50 per student while the cost of the same experiment in the form of a popular commercial kit would be $5.24 per student.
The PCR beads, while expensive, are key to the success of this protocol. The beads contain the dried form of Taq DNA polymerase enzyme, buffer, bases (dNTPs), a source of magnesium ions, and stabilizers. Beads contribute to the success of the experiment in two ways. First, the teacher does not have to assemble all the bead ingredients, one by one, into a master mix, freeing you from all the orders and calculations that would entail. Second, students do not have to pipet tiny amounts of Taq plus buffer plus dNTPs plus Mg ions; all these reagents are contained in the dry bead. So students’ chances of success are greatly increased as the opportunities for pipetting errors are decreased. The student adds only template DNA, primers, and sterile distilled water to the bead in the tube to make up a final volume of 25 microliters and the reaction tube is ready to go into hand or instrument cycling. Using these PCR beads with a final volume of 25 microliters, you get 2.5 units of Taq DNA polymerase, 200 mM dNTPs in 10 mM Tris pH 9.0 and 1.5 mM MgCl2. These concentrations are pretty standard and do not need to be adjusted for a Lambda PCR reaction. PCR beads can be purchased from GE HealthCare; see appendix for ordering details. We have found that even in the hands of novice high school students and teachers, PCR product is successfully made most of the time. Hence, high schools and college classes CAN do PCR without expensive instrumentation and with an excellent chance of success.
However, one barrier remains: how to order primers and DNA, then get these into the correct concentration for the PCR reaction -even an experienced teacher might be daunted by the prospect of ordering primers and DNA from a biotechnology company and preparing them for student use. Hence, this article will walk you through the steps.
TEMPLATE DNA FROM LAMBDA
The PCR beads work with any DNA template, but to introduce PCR to students, bacteriophage Lambda DNA is a good startup choice. It is inexpensive and its relatively small genome (48,502 base pairs) makes an easy target for PCR primers. You can purchase10 mg of lambda DNA from Carolina Biological for $13.75. It comes at a concentration of 0.1 mg/ml, which is plenty of DNA for PCR because you need less than 1 ng for each reaction tube.
Purchasing the Lambda DNA is straightforward; the real challenge is to get it into the right concentration for a PCR reaction. For small genomes like Lambda DNA, it is recommended to use an amount between 0.1 and 1 ng of template DNA in a total reaction mixture of 25 microliters. Too much DNA would actually hinder getting good results. Hence, your 10 mg purchase will support 10,000+ PCR reactions! If a protocol calls for using 5 ml of lambda DNA at a concentration of 0.12 ng/ml, which delivers a total of 0.6 ng DNA into the PCR reaction tube (5 ml x 0.12 ng/ml = 0.6 ng ), this is how the calculations would go.
Step one: Convert the 0.12 ng/ml to its equivalent in mg/ml = 0.00012 mg/ml so that the units in the proportion below are the same.
Step two: Lambda DNA 21-1408 from Carolina comes as 100 ml of DNA at a concentration of 0.1 mg/ml– use C1V1 = C2V2 C= concentration, V = volume
C1 = conc of stock, 0.1 mg/ml
V1 = X, how much of stock do we need to get a certain amount of V2
V2 = (let’s use 100 microliters – this is arbitrary)
C2 = final conc of 0.00012 micrograms/microliter to use in the PCR reaction tube.
Solve for X.
0.1 mg/ml ( X) = 0.00012 mg/ml (100ml )
X = 0.12 ml - this number is mathematically correct but not practical – even the best micropipettor will not measure such a small volume. So to get around this practical dilemma, we use serial dilutions. Below is one version of a serial dilution; there is more than one correct way to get to the same final desired concentration.
Step one: Dilute the stock lambda DNA that is at 0.1 mg/ml 1/100 – take 1 microliter of stock DNA, add 99 microliters sterile distilled water (or TE: 10mM Tris, pH 8.0 and 0.1 mM EDTA). Now the DNA is at concentration of 0.001 mg/ml or its equivalent, 1 ng/ml. Mix thoroughly before proceeding to step two.
Step two: Now use C1V1 = C2V2
C1 = conc of diluted stock from step one, 1 ng/ml
V1 = X, amount of diluted stock we need to get to desired final concentration
V2 = (let’s use 150 microliters – this would be sufficient volume at correct concentration to provide 5 microliters for each of 25 students with some extra)
C2 = final conc of 0.00012 mg/ml to use in the PCR reaction tube.
Solve for X.
1 ng/ml (X) = 0.12 ng/ml (150 ml )
X = 18 ml of diluted stock from step one.
Step three: take 17.5 ml of diluted lambda DNA from step one, and bring to final volume of 150 ml with sterile distilled water or TE; mix well – label this tube as lambda DNA at concentration of 0.12 ng/microliter and use for PCR reactions.
PRIMERS
Ordering primers can be daunting. There are many companies that will supply primers (oligonucleotides), and of course, these are custom-made for each order. When you order, you are asked not only for the sequence, but you also have to indicate the level of purity and the synthesis scale you need. What does this mean?
Oligonucleotides are chemically synthesized on a synthesis column (Devor and Behlke, 2005). The column contains a fixed amount of the 3’ nucleotide in your order attached to a solid or bead support in the column. Synthesis of your requested oligonucleotide occurs by adding each base of the sequence, one at a time, to be added in the 3’to 5’ direction. The efficiency of this synthesis process varies with which base is being added as well as the position of that base in the growing primer sequence. “Scale” refers to the amount of that first 3’ nucleotide that is present in the column – the more present on the column, the higher the yield of final product. Different sequences give different yields, some better, some worse than average. Companies typically promise a certain yield with each level of synthesis scale, so the more primer you need, the higher the synthesis scale you select. The other choice is purity level – the cheapest and basic level is HPSF (high purity salt-free) which is sufficient for this experiment: a small genome target DNA and moderate length (20 mer) primers.
Primer Ordering Choices
When you order primers, the first thing is the sequence. Note that the primer sequence is always given in the 5’> 3’ order. If both primers are already specified, you are all set – just enter the sequence. But if you are using the DNA template sequence to derive the sequences for primers, be mindful of this convention. See Figure one for an example. You get to choose a “name” for your primer – like l F (lambda primer forward direction) – exercise your creativity here!
After you enter your sequence, you will be asked to indicate if you need modification of the primer – for our lambda PCR, the response is NO. In the context of the lambda PCR, you also choose the basic purification level, HPSF, and standard documentation of the sequence. Next, you have to select the synthesis scale. Typical choices may be: 10 nmole (0.01 mmol), 25 nmole (0.025 mmol), (50 nmole (0.05 mmol), 0.2 mmol, 1.0 mmol, 10.0 mmol and up. The higher the synthesis scale is, the greater the yield, and the greater the cost per base. If you choose 25 nmole scale, the price should be about $.30/base and should yield enough primer for roughly 250 reactions, which is plenty for the typical classroom. Choosing a higher scale will cost more per base and deliver much more primer than you may ever use.
Using Your Primers
Primers are usually shipped in lyophilized (dry) form, but since the powder may land anywhere in the tube during shipping, it is important to SPIN the primer tube before you open it! This action will ensure that the primer is at the bottom of the tube and won’t fly out when you pop the cap. If you are going to use all the primer right away, you can dissolve in sterile distilled water and store at 4 degrees C. for a few days. More likely you will have an excess and will be storing some of the primer for future use. For longer term storage, resuspend in TE (10 mM Tris pH 8.0; 0.1 mM EDTA, pH 8.0) and divide stock into several aliquots for storage in the minus 20 degrees C. freezer. Then you can take out one aliquot for each use, and avoid repeated freezing and thawing of the entire stock, which can degrade the primers. But how much solvent should you use to get the primers into solution form?
To solve this challenge, you need to examine the synthesis report that accompanies your primer shipment. The documentation will typically verify the sequence, the length, the GC content, the scale of synthesis and purification level for each primer. Additionally, it will give you the actual yield of the synthesis in one of several forms: as OD units, in mg, and in nmoles. If you take the primer yield expressed in nmols and multiply times 10, you will get the volume in ml that you need to add to get your primer at a concentration of 100 pmol/ml, a convenient stock concentration. Most report sheets will also directly give you the volume to add to get to 100 pmol/ml. This volume will differ for each primer because the sizes and synthesis yields will be different for each primer. So, upon primer arrival, when ready to dissolve:
Getting primers into stock solutions
Step one: using synthesis report or certificate of analysis report, find the amount in nmol for the primer, and multiply x 10 to get volume of solute to obtain stock concentration of 100 pmol/ml.
Example: 40.1 nmol x 10 = 401 ml
Step two: spin tube or vial of primer before opening, then add 401 ml of sterile distilled water or TE to obtain stock concentration of 100 pmol/ml.
Step three: divide dissolved primer into several aliquots, label tubes, freeze all but one tube in -20oC freezer.
Getting primers into working concentrations
For our Lambda PCR, we want the primers at a concentration of 12.5 pmols/ml. Why 12.5 pmols/ml? This is somewhat arbitrary. The recommended concentration range for primers in a PCR reaction is 0.1-2.0 mM (equivalent to 0.1-2.0 pmols/ml). If we use 3 ml of each primer at 12.5 pmols/ml, we will have a final concentration of 1.5 pmols/ml of each primer in the 25 ml final volume in the PCR tube, which is within range.
Step one: use C1V1 = C2V2 to determine how to get the stock primer solution into a working concentration of 12.5 pmols/ml
C1 = conc of stock, 100 pmols/ml
V1 = X, how much of stock do we need to get a certain amount of V2
V2 = (let’s use 100 microliters – this is arbitrary)
C2 = final conc of 12.5 pmols/microliter to use in the PCR reaction tube.
100 pmols/ml (X) = 12.5 pmols/ml (100 ml)
X = 12.5 ml - which means take 12.5 ml of stock primer solution at 100 pmols/ml and add 87.5 ml to bring to final volume of 100 ml, the working concentration of primer.
Final reaction mixture as calculated above:
-take one PCR tube with bead, add 3 ml of forward primer and 3 ml of reverse primer, each at a concentration of 12.5 pmols/ml, add 14 ml sterile distilled water, and finally add 5 ml of lambda DNA at concentration of
0.12 ng/ml. Mix thoroughly to dissolve the bead completely and then immediately put into thermal cycler or begin rotation between water baths, as per protocol. A control tube with NO DNA template should also be run. When the cycling is complete, confirm the presence (or absence in the case of the control tube) of PCR product by running 15 ml of tube contents in a 2% agarose gel with DNA marker, looking for an 821 bp band. Our website has sample protocol . Our website protocol uses slightly different volumes but the final concentrations are the same. Our rate of success with this protocol is very high, and when students perform the hand-cycling for PCR, it makes for much more meaningful learning than just adding some ingredients into a tube and putting it into a thermal cycler to do the work. I make all our biotechnology program students do hand cycling before they get to use our thermal cyclers for more advanced experiments. As for the teacher, once you know how to order primers and use the PCR beads, you can do many other PCR experiments with the beads, changing only the template DNA and the primers.
References:
Bloom, Freyer, Micklos. (1996). Laboratory DNA Science. Menlo Park, CA. Benjamin Cummings Publishing.
Devor, E.J., and Behlke, M.A. (2005). Oligonucleotide Yield, Resuspension and Storage. Available online at the website of Integrated DNA Technologies: www.idtdna.com/support/technical/TechnicalBulletinPDF/Oligonucleotide_Yield_Resuspension_and_Storage.pdf .Mulvihill, C. (2007). Water Bath PCR. Available online at: http://www.occc.edu/BBDiscovery/documents/Modules/water_bath_pcr.htm
Seidman, L.A.(2008), Basic Laboratory Calculations for Biotechnology. San Francisco, Pearson Benjamin Cummings.
Appendix
GE Healthcare: go to www5.gelifesciences.com, put PCR beads into the search window to reach the ordering information for illustra PuRe Taq Ready-to-Go PCR beads (100 reactions for $140; select 0.2 mL or 0.5 mL size tubes as needed if using a thermal cycler.)
Carolina Biological: www.carolina.com cat.# 21-1408 for Lambda DNA at a concentration of 0.1 mg/ml, $13.75.
Primers: there are many companies that manufacture primers – here are a few:
MWG biotech: www.MWGbiotech.com
Integrated DNA Technologies: www.idtdna.com
Operon: www.operon.com
FIGURE ONE.
Suppose you are looking at a piece of DNA for PCR amplification. Typically the sequence shown is just one strand of the two, and by convention, it shows the strand in the 5’>3’ direction left to right. Of course, if you know one strand, the base-pairing rules give you the sequence of its complementary strand. The primer sequences are shorter than what would normally be used, just to simplify the illustration.
DNA:
5’ A T C G C C …………………………. C C G G A T 3’
What would the primers be?
For the downstream (Reverse) primer, you fill in the complementary strand
<3’G G C C T A 5’ R primer
5’ A T C G C C …………………………. C C G G A T 3’
But when you order this downstream primer, you write it in the 5’>3’ direction as
5’A T C C G G 3’
The upstream/Forward primer would be complementary to the opposite DNA strand:
5’ A T C G C C …………………………. C C G G A T 3’
3’ T A G C G G …………………………..G G C C T A 5’
5’ A T C G C C 3’> F primer
5’ A T C G C C 3’ – this primer is written as it appears: note it is the same sequence as the original 5’ strand of the DNA template.
Copyright: C.Mulvihill, 2008.