Polymerase Chain Reaction (PCR) of Cloned Genes
As part of the process to determine the validity of your clones, you will use polymerase chain reaction (PCR) to amplify all or just a portion of your ‘genes’ (‘inserts’), using the plasmid DNA as template. (Whether you amplify the entire gene or just a portion depends on which gene you are working with, because we have obtained different primer pairs for some of the plasmids.) Successful results will verify the inserts’ presence in the plasmids. In a subfolder in Bb, you will now find color-coded gene sequence Word documents for each gene (A, B6, and F1) as well as for the vector plasmid; please refer to the relevant document for your gene as you read the following.
For genes “A” and “F”: the upstream (forward) and downstream (reverse) PCR primers that we will use, will bind to sequences in the plasmid vector, as opposed to the insert itself (the shaded grey or blue area in the document), and so the entire insert (plus a little extra) should be amplified.
In the case of gene B6, previously untested in this respect, we have two sets of PCR primers available: the vector primers described above, plus an additional set of ‘internal’ primers that bind to sequences within the insert itself. Pending final decisions, it is likely that one B6 team will ‘test’ the PCR reaction with vector primers and the other B6 team(s) will ‘test’ the internal primers. Although the vector primers ‘should’ work on gene B6, the expected PCR product would be significantly larger than the PCR products from the other plasmids (~2059 bp as opposed to ~1000 bp). At the very least, the extension time would need to be lengthened to accommodate the larger PCR product, and we are still uncertain if it would properly amplify. On the other hand, the B6 PCR product generated from ‘internal’ primers is only ~900 bp, so would probably be amplified using the extension times of the original program (genes A and F). However, these primers may need a different annealing temperature from the vector primers, so a degree of experimentation in that respect is warranted. As you can see, the optimal PCR program for each new primer/template combination is usually empirically determined (trial and error!).
In the gene sequence documents for these 3 genes, the ‘vector’ primer binding sites are color-coded in turquoise blue. In addition, the ‘internal’ primer binding sites in gene B6 are color coded green (forward) and red (reverse).
Use the “Tools: word count” feature of Word to estimate the size (in bp) of the region to be amplified, between (and including) the two primer binding sites. To do this, highlight only the area of interest, then proceed to “word count”2.
Overview of PCR:
Polymerase Chain Reaction, or PCR, was invented in 1984 by Kary Mullis who was awarded a Nobel Prize for his work in 1994. The enormous utility of PCR is based on its ease of use and its ability to amplify DNA in relatively short time periods. The PCR process uses an enzyme known as Taq polymerase, or variants of this enzyme that are called for in specific circumstances. Taq polymerase is purified from a bacterium originally isolated from hot springs and is stable at the very high temperatures used for denaturation (separation) of DNA strands, typically 95-960C. Also included in the PCR reaction mixture are the DNA template- also known as the “target DNA” (genomic DNA, plasmid DNA, etc.), and two ~16-25-mer synthetic oligonucleotides known as “primers”. Single stranded DNA primers are designed to be complementary to the flanking (bordering) ends of the locus to be amplified, such that each target sequence to be amplified will have an ‘upstream’ (forward) and ‘downstream’ (reverse) primer. Also included in the PCR reaction mixture are dNTPs (the building blocks for DNA synthesis) and the appropriate reaction buffer with Mg+2 ions. The volume of a single reaction is usually in the range of 20- 50 µl, and the reactions are carried out in special thin-walled tubes that are designed to fit snugly into the PCR machine.
In the first step of a typical PCR reaction (inside the pre-programmed thermal cycler), the target complimentary DNA strands are melted, or denatured (separated from each other) at ~960C, while the Taq polymerase remains stable. In the second step, known as annealing, the sample is cooled to ~55-580C to allow hybridization of the two primers to their respective binding sites in the two strands of the target DNA. In the third step, the temperature is raised to 720C and the Taq polymerase then adds nucleotides to the 3’ ends of the primers to complete the synthesis of the new complementary strands.
These three steps – denaturation, annealing, and DNA synthesis (extension) – constitute one PCR “cycle”. This process is typically repeated for 30-40 cycles, amplifying the target sequence exponentially: after 30 cycles of amplification, in theory, the target DNA sequence is amplified by 230 or 109-fold in a period of ~2 hrs. (In reality, the efficiency of amplification of a given enzyme preparation is somewhat less than ideal.)
PCR is performed in a thermal cycler, which is programmed to rapidly heat, cool and maintain samples at designated temperatures for varying amounts of time, determined by the investigator.
For more information about PCR, please see:
- DNA engine (PCR machine)
- Template DNA: for each of your plasmid DNA samples, this will be an aliquot (small portion) that you have diluted to 10 ng/µl in dH2O (i.e, 0.010 µg/µl); so, if your original DNA was at 0.5 µg/µl (500 ng/µl), then this corresponds to a 1:50 dilution. You won’t need much for PCR; if you prepare, eg, 50 µl of this 1:50 dilution, you will only use 1 µl of your original DNA sample. Any excess diluted plasmid can be saved and used for future testing and/or controls.
- Taq polymerase (5U/µl stock) and 10X PCR buffer with Mg+2
- Deoxynucleotides (‘dNTPs’, a mixture of 10 mM of each)
- Thin-walled PCR tubes (0.5 ml capacity, NOT TO BE CONFUSED with our typical “microfuge” tubes of 1.5 ml capacity)
- Forward and reverse PCR primers:
- ‘Vector primers’ (genes A, B6, and F):
called “ABC-f3 ”: 5’-GATCCGGTACCGAGGAGAT-3’
Stock 50 µM in dH2O
called “BC-r”: 5’-CCACCAGCTCGAACTCCA-3’
Stock 50 µM in dH2O
‘Internal primers’ (gene B6 only):
called “ITGB6 F” 5’-GAGAGAAGAAGCAGGCACATTA-3’
Stock 50 µM in dH2O
called “ITGB6 R” 5’-GTAGCTTCCAGATGCACAGTAG-3’ Stock 50 µM in dH2O
Master Mixes: (Prepare master mix on ice, and keep it there!)
Teams with Genes A and F will prepare 4 samples for PCR: DNA #1, DNA #2 (your two plasmid samples), a negative control (consisting of dH2O instead of DNA), and a positive control (provided: a previously isolated, purified clone of your gene, already at 10 ng/ul). (Obviously we don’t yet have a positive control for gene B6, so those teams will prepare 3 samples.) Think about why we should include these controls. For our protocol, a typical 50 µl PCR reaction would have the following components (per tube):
10X PCR Buffer w/Mg+2 5 µl
dNTPs (@ 10 mM each) 1 µl
Forward Primer (50 µM concentration) 1 µl
Reverse Primer (50 µM concentration) 1 µl
(for Gene A4 ONLY: -5M Betaine 10 ul)
dH2O 39.8 µl (29.8 µ l for Gene A)
Taq DNA Polymerase (5U/µl) 0.2 µl (1U per reaction)
-Plasmid template DNA*, (10 ng/µl) 2 µ l (20 ng)
50 µl total
* or dH2O for neg ctl, or positive ctl DNA
Typically, a “Master Mix” (bracketed items) consisting of everything except the template DNA is initially prepared in a single 1.5 ml eppendorf tube, then allocated into each sample PCR tube (on ice); template DNA is then added last, right before the tubes are placed into the PCR machine. For genes A and F, you would need Master Mix for four samples minimum (including the provided positive control DNA), so you should prepare enough MM for FIVE samples (always figuring that there will be some loss of reagent to the sides of the tube and the inside of pipette tips). Gene B6 teams, who don’t have a positive control, will only be making 3 samples, so therefore their master mix should be made x4.
- Example: To prepare your Master Mix for five samples, label an eppendorf tube ‘MM’ and keep it on ice (as well as all other PCR reagents); multiply the above volumes of each reagent within the bracket by five, then add that amount to the MM tube. For example, you would add 5 µl x 5 = 25 µl of the 10X PCR buffer to the tube, 1 µl x 5 = 5 µl of dNTPs, etc. Add the Taq DNA polymerase last, and take care with pipetting, as it will still be a very small volume (1 µl) and it is stored in glycerol, so it will be viscous. After the Master Mix is complete, gently vortex briefly, a couple of times, and/or ‘flick’ the tube with your fingers several times to mix the reagents, then pulse-spin in the microfuge to bring all components to the bottom of the tube.
- Pipette 48 µ l of this Master Mix into each of your labeled PCR tubes (on ice) (the thin-walled 0.5 ml tubes, NOT standard eppendorf tubes). There should be some Master Mix left over, if you have pipetted properly (which can then be discarded).
- Finally, add 2 µ l of either your diluted plasmid DNA, the positive control plasmid DNA (provided), or dH2O (for negative control) to each appropriately labeled PCR tube, for 50 µl total per tube. Briefly vortex or ‘flick’ the tubes to mix reagents; pulse-spin in microfuge (using tube adaptors) to bring all contents to the bottom of tubes. (Prior to adding your DNA, please check with your classmates to verify that they are at or near this step, so that you don’t have to keep your samples ‘waiting’ very long before placement in the PCR machine.)
Note to Teams with Gene “A”:
Some DNA templates present problems of secondary structure (if high G-C content), which require an ‘additive’ such as 1M betaine (N,N,N-trimethylglycine) to facilitate strand separation6. Apparently “Gene A” (RASSF1A) is one of those templates.
The amplification reactions take place in the black ‘DNA engine’ from MJ Research. It has two independently controlled 30-well heating blocks.
Teams with Genes A and F: Place your tubes together in one block. When everyone’s samples are loaded, we will hand-tighten the lid with the thumbscrew and begin the program. We will use the program ‘OLINBIO’ to amplify:
Step #1 95˚C 1 minute
Step #2 94˚C 15 seconds (denaturation)
Step #3 56˚C 30 seconds (annealing)
Step #4 72˚C 1 minute (polymerization)
Step #5 Go to Step #2 (32x)
Step #6 4˚C forever*
DMSO (dimethylsulfoxide) added to the Master Mix (to 5%), works just as well as 1M Betaine; our choice to use 1M Betaine in today’s reactions is entirely arbitrary. Either would work.
Teams with Gene B6: Place your tubes in the other block (separate from genes A and F samples). We will use the program called ‘B6’, which is a slightly modified version of ‘OLINBIO’; the only difference is that step #4 (extension, 72 deg C) is for 2 min 30 seconds in ‘B6’, to accommodate the longer PCR product.
When the reactions are complete, the machine is programmed to store the tubes at 40C until they are removed, which for our purposes can wait till the next morning. Instructor will remove samples and store them in the -20°C freezer until further analysis, by agarose gel electrophoresis the following week.