Polymerase Chain Reaction And Gel Electrophoresis

Running the bench protocol

Polymerase chain reaction amplifies a target DNA sequence through repeated thermal cycling. You set up a reaction mix with template DNA, forward and reverse primers, dNTPs, buffer, and a thermostable polymerase. The cycler runs through denaturation at 94 to 98 degrees Celsius, annealing at a temperature calculated from your primer melting points, and extension typically at 72 degrees Celsius. Thirty to forty cycles usually gives you enough product for visualization. After the run finishes, you load the PCR product onto an agarose gel along with a DNA ladder. An electric current pulls the negatively charged DNA through the matrix. Smaller fragments move faster. You stain with something like ethidium bromide or SYBR Safe and image under UV or blue light. That is the basic workflow. The reality on the bench is messier.

Practical notes on Polymerase Chain Reaction And Gel Electrophoresis

The annealing temperature is where most people waste reagents. A rule of thumb is to set it three to five degrees below your primer Tm, but if your primer pair has mismatched Tms, use the lower one as your baseline. I run a gradient PCR once per new primer pair because the manufacturer Tm values are often inflated by salt correction methods that do not match my buffer conditions. It takes an extra tube but saves you from chasing a band that never appears. Agarose concentration matters more than most protocols mention. A 1 percent gel resolves fragments between 500 and 10,000 base pairs reasonably well. For smaller amplicons under 500 bp, bump up to 2 percent. Higher percentage gels run hotter though, so reduce voltage or run them shorter to avoid melting. I have ruined two gels by running a 2 percent at 150 volts for an hour. Twelve volts per centimeter is a safer ceiling. TAE versus TBE buffer is another thing people debate without reason. TBE gives sharper bands and better resolution for small fragments, but it buffers less effectively over long runs and borate can interfere with downstream enzyme digests. If you are doing a quick diagnostic gel, TAE is fine. If you need to cut out a clean band for cloning, use TBE and verify compatibility with your next enzyme. The phosphate in TAE can also chelate magnesium and inhibit restriction enzymes if you carry over too much buffer.

I ran into a problem last year where my PCR product appeared as a smear instead of a discrete band. The primers were fine, the cycling conditions matched the insert size, and the template quality was acceptable. I checked the polymerase storage history and realized the enzyme had been at room temperature during a brief power fluctuation in the lab. It had lost roughly half its activity. Low-fidelity polymerases compensate by non-specific binding when they are sluggish. I split a new aliquot from minus eighty, re-ran the reaction with an increased annealing temperature by two degrees, and got a clean single band on the first try. Thawing on ice instead of at room temperature would have prevented it, but the lesson was clear: enzyme activity drift is silent until your gel tells you something is wrong. Primer dimers are another classic issue that beginners blame on bad primers when the problem is usually the cycling program. If your primers have complementary sequences at their 3 ends, they will prime each other before they find the template, especially in early cycles when template concentration is low. The fix is not always redesigning primers. A hot-start polymerase or a 98 degree initial denaturation step for two to three minutes forces the polymerase to engage only after the primers have properly annealed to the template. I add a 15 second gap at the annealing step before ramping to extension so the primers settle into the correct position. This also reduces mis-priming on GC-rich templates. Loading dye is another overlooked variable. Some loading dyes contain SDS or high salt concentrations that alter migration speed. Xylene cyanol and bromophenol blue track at different rates depending on gel concentration, so do not assume the dye front position predicts fragment size accurately. Always run a ladder in a separate well, not mixed with your sample, because the ladder dyes can mask your bands if they overlap during imaging.

If you need quantitative data, gel electrophoresis is not the right tool. Band intensity on an agarose gel is only semi-quantitative at best, and intercalating dyes do not bind linearly across all fragment sizes. Use qPCR with SYBR Green or a standard curve if you need copy number. The gel is for confirmation and sizing, not measurement. Upstream of the PCR, the quality of your template DNA heavily influences everything downstream. Phenol-chloroform extractions leave residual organics that inhibit polymerase. If you are using a home-prepped extraction, a ethanol wash and a brief air dry before resuspending in TE or water makes a noticeable difference. Alternatively, column-based kits remove inhibitors faster, though they can shear genomic DNA if you vortex aggressively. Spin gently. A visible pellet means you have concentrated enough template for most applications. One microgram is overkill for plasmid templates. Nanogram amounts work fine. When you image the gel, exposure time matters more than people admit. Overexposing saturates the bright bands and compresses the dynamic range, making faint non-specific products look like artifacts. Underexposing hides weak but real bands. Take multiple images at different exposures if you are unsure. Most imaging software allows you to adjust brightness and contrast non-destructively, but starting with a properly exposed image saves time.

Post-electrophoresis, if you plan to excise a band for cloning or sequencing, use a clean scalpel under UV transillumination and minimize exposure time to prevent DNA damage. UV crosslinking can introduce thymine dimers that block subsequent PCR or ligation. Some labs use blue light transilluminators with SYBR Safe for this reason, which is gentler on the DNA. If you are sequencing the excised product, gel extraction kits recover yields around 50 to 70 percent of what is theoretically in the band, so factor that into your downstream reactions. Storage of prepared gels is straightforward but easy to mess up. A poured gel in TAE buffer stays usable for a few days if wrapped in plastic film and kept at four degrees. After that, the buffer leaches out, the gel dries, and migration patterns shift. TBE gels last longer but still degrade. I prepare gels the morning of use unless I am running a time-sensitive diagnostic, in which case I store them properly and verify migration with a ladder before loading samples. The whole process from PCR setup to gel result usually takes about four hours if you run it in one sitting. A typical 35-cycle PCR runs for roughly 45 minutes. Gel preparation, loading, and imaging adds another two hours depending on your gel thickness and voltage. If you batch multiple samples and use a multi-well comb, you can cut that imaging time significantly. Planning the layout on paper before pouring the gel prevents the common mistake of running out of wells for the ladder and your test samples at the same time.