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The 10 Most common mistakes

            with RT-PCR Tests


The 10 Most common mistakes with RT-PCR Tests .

1)  Poor primer and probe design .

For the most efficient design of PCR primer and probe sets for real-time qRT-PCR, we strongly recommend using primer design software. Most primer design programs include adjustable parameters for optimal primer and probe design. These parameters consider primer/probe Tm, complementarity, and secondary structure as well as amplicon size and other important factors. Restricting the number of identical nucleotide runs is also recommended. When designing amplicons in eukaryotic targets, choose PCR primers that span at least one exon-exon junction in the target mRNA to prevent amplification of the target from contaminating genomic DNA.


2)  Using poor quality RNA

Degraded or impure RNA can limit the efficiency of the RT reaction and reduce yield. RNA should either be prepared from fresh tissue, or from tissue treated with an RNA stabilization solution. The importance of using full length RNA for reverse transcription depends on the application. Amplicons for real-time qRT-PCR are typically short (70-250 bp). As a result, some degradation of the RNA can be tolerated. If it is not possible to use completely intact RNA, design primers to anneal to an internal region of the gene of interest. Note that for truly quantitative RT-PCR, partially degraded RNA may not give an accurate representation of gene expression.


3)  Not using “master mixes”

qRT-PCR is a highly sensitive tool for analyzing RNA. As the PCR amplifies the target, errors are simultaneously amplified. Therefore, variability should be kept to a minimum whenever possible. A “master mix”, or mixture of the reaction reagents, should be used when setting up multiple reactions to minimize sample-to-sample and well-to-well variation and improve reproducibility. To further reduce well-to-well variation, a reference dye such as ROX can be added to the master mix.


4)  Introducing cross-contamination

All surfaces in the PCR area should be routinely decontaminated to prevent cross contamination. Use of a DNA decontamination solution, such as DNAzap™, that destroys DNA, is recommended. A “No Template Control” (NTC) should be run to rule out cross contamination of reagents and surfaces. The NTC includes all of the RT-PCR reagents except the RNA template. Typically the RNA is simply substituted with nuclease-free water.

No product should be synthesized in the NTC; if a product is amplified, it indicates that one or more of the RT-PCR reagents is contaminated with the amplicon.


5)  Not using a “– RT” control

It is virtually impossible to completely eliminate genomic DNA from RNA preparations. Therefore, it is important to include a minus-reverse transcriptase control (“No Amplification Control” or NAC) in qRT-PCR experiments. Typically, the NAC is a mock reverse transcription containing all the RT-PCR reagents, except the reverse transcriptase. If a product is seen in the NAC, it probably indicates that contaminating DNA is present in the sample.


6)  Using an inappropriate normalization control

The reliability of any qRT-PCR experiment can be improved by including an invariant endogenous control in the assay to correct for sample to sample variations in qRT-PCR efficiency and errors in sample quantitation. The expression level of a good control should not vary across the samples being analyzed. 18S rRNA is often used as a control because it is less variant in expression level than other traditional internal controls such as ß-actin or GAPDH.


7)  Dissociation (melting) curves are not performed when using SYBR® Green

Ideally, the experimental samples should yield a sharp peak (first derivative plot) at the melting temperature of the amplicon, whereas the NAC and NTC will not generate significant fluorescent signal. This result indicates that the products are specific, and that SYBR Green I fluorescence is a direct measure of accumulation of the product of interest. If the dissociation curve reveals a series of peaks, it indicates that there is not enough discrimination between specific and non-specific reaction products. To obtain meaningful data, optimization of the qRT-PCR would be necessary.


8) Not setting the baseline and threshold properly

To obtain accurate Cq values the baseline needs to be set two cycles earlier than the Cq value for the most abundant sample. For real-time qRT-PCR data to be meaningful, the threshold should be set when the product is in exponential phase. Typically this is set at least 10 standard deviations from of the baseline.


9) The efficiency of the reaction is poor

The efficiency (Eff) of the reaction can be calculated by the following equation: Eff=10(-1/slope) –1. The efficiency of the PCR should be 90-110% (3.6 > slope > 3.1). A number of variables can affect the efficiency of the PCR. These factors can include length of the amplicon, secondary structure, and primer design, to name a few. Although valid data can be obtained that fall outside of the efficiency range, the qRT-PCR should be further optimized or alternative amplicons designed.


10)  Using an inappropriate range for standard curves

Standard curves should be prepared for each gene under study for RNA quantitation (absolute or relative quantitation), or for verification of the efficiencies of the reactions for comparative quantitation (delta-delta-Ct). The standard curve should extend above and below the expected abundance of your target. Additional input quantities can be included such as the minimum and maximum RNA amounts above and below the limit of detection to help differentiate between specific and non-specific products.

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What does "most common" mean? It means they happen. Right?

Ten Things That Can Kill Your PCR

(are any of these repeated with RT-PCR ?)

A blank PCR gel has got to be one of the most aggravating things about molecular biology. We've all had PCRs work one day, then fail inexplicably the next. And we've all banged our heads against a wall trying to figure out what went wrong. If your gels are turning up blank, check out this list of things that can kill your PCR -- and get your experiments working! Nowhere is the old adage "an ounce of prevention is worth a pound of cure" more applicable than to PCR. Depending on your level of PCR expertise, some of these hints may seem obvious. But how many times have you kicked yourself because you overlooked the obvious? Whether you're an expert or a novice, any of these may be just what you're searching for to get your PCR working.


1) Too much dNTP, or degraded dNTP.

Too much dNTP can actually inhibit your PCR reaction. Between 40 - 200 uM is the optimal range. Also, dNTPs are sensitive to repeated freeze-thaw cycles. Make small aliquots when you get a fresh batch, and turn over your stock frequently since dNTPs frozen at -20 C will eventually go bad.


2).  Not mixing MgCl2.

Magnesium chloride solutions form a concentration gradient when frozen and need to be vortexed prior to use (1).


3)  Wrong MgCl2 concentration. Every PCR reaction has an optimal MgCl2 concentration range, usually between 1 - 4 mM. Mg2+ ions form complexes with dNTPs and can also act as a co-factor for polymerases, so you'll need to try several conditions to optimize your concentration.


4)  Inhibitors in your reaction.

Make sure you know how you got your source DNA. Chloroform, phenol, EDTA, ionic detergents (SDS and Sarkosyl), xylene cyanol, bromophenol blue and ethanol -- among many other things -- can inhibit PCR. An extra clean-up step on your template may do the trick. Also, certain polymerases can be more susceptible to certain substances, so be sure to check your polymerase for possible inhibitors.


5)  Poor quality mineral oil.

Lower-grade preparations may contain nucleases that can kill your PCR. Also, avoid autoclaving your mineral oil if possible. Exposure to high heat may cause reaction-inhibiting hydrocarbons to form. Similarly, do not irradiate mineral oil with UV for long periods (2).


6)  Too much enzyme.

Excess enzyme in your PCR can lead to smearing of PCR products. Most people seem to use 0.5 ul of their stock Taq per reaction, but that may contain way more than necessary for your particular reaction.


7)  Wrong primer concentration.

If you have too little primer you won't see any product. Too much primer and you may get primer dimerization and not enough amplification. Stay within 0.1 - 1.0 uM of primer.


8)  Wrong PCR program.

Make sure the program you selected on your PCR machine is actually the one you want! It only takes a slip of a finger, or some klutz, to alter your personal program on a common PCR machine. Check your program while it's cycling to make sure it's what you wanted.


9)  Excess or insufficient template. Too much template can inhibit PCR by binding all the primers. Too little template, and amplification may not be detectable. For 25 - 30 cycles, 104 copies of the target sequence are sufficient.


10)  Poor primer design.

While primer design can seem like a black art, avoid obvious errors like self-complementarity, complementarity between paired primers, or excessively long oligos (>30 bp). Often, making a new primer next to a suspect one can solve the problem and can be faster and cheaper than trying numerous variations in reaction conditions. Of course, this list is by no means comprehensive. Like any experiment, many things can go wrong.


If you have anything you'd like to add, we'd love to know about it.

Drop us an email at We'll compile another set of PCR Do's and Don'ts and publish them in the future. In the meantime, may all your PCRs work the very first time!

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the list of things that can go wrong with these tests on this website are just the beginning of possible errors. And with these stats they wish to starve to death millions of babies and children in the Third World.

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