If you are detecting bacterial sequences other than E.coli, the primers will have to exclude any part of the DNA that is shared between E.coli and the bacteria used (in other words, the primers have to be designed only in the part which is unique for the bacteria used).
If detecting E.coli sequences, There are some traces of bacterial DNA in the Taq polymerase and in the UNG (as it is produced in E.coli). You will have to determine the minimum level and subtract it from the positive signal. Everything above this signal should be considered as positive.
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When designing Molecular Beacons, pay attention to the folding of the probe. It might fold into alternate conformations, which are not well quenched. Change the stem or loop sequence, or both to avoid this.
If the salt concentration of the buffer is too low (below 1 mM MgCl2) the probe does also not fold correctly.
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Design assistance
In a one step RT qPCR reaction the RT reaction and the qPCR reaction are done in one and the same tube. The buffer is a combination of a RT and a PCR buffer, in which both enzymes do work. The one step RT qPCR reaction is a closed tube assay, so contamination can be avoided. It saves pipeting steps and time and is easy in handling.
In a two step RT qPCR the RT reaction and the qPCR reaction are done in separate tubes. It gives a more flexible way of working in that the cDNA can be used for more than one qPCR reaction and can be archived, so that it eliminates the need to continually isolate RNA.
Further more it allows the use of oligo d(T) and ramdom nonamer primers in the RT step and specific primers in the PCR step. This will increase the specificity and sensitivity of the assay.
Eurogentec offers both one and two step RT qPCR kits.
When performing a RT it is possible to use three different primer types:
Oligo d(T) primers, which bind to the poly A tail of the RNA and then only transcribe RNA. This will avoid contamination with genomic DNA. As the poly A tail is located at the beginning of the gene it will also lead to more full transcripts.
Random nonamers, which bind anywhere in the genome and allow the reverse transcriptase to fill up the gaps, will leads to high yields.
Specific primers, which bind to the gene of interest, and will therefore give specifics products.
The combination of oligo d(T) primers and random nonamers will give the highest yields and the longest transcripts, whereas specific primers transcribe only specific RNA but reduce the yield.
With an One step RT qPCR kit it is only possible to add specific primers, as it should be avoided that oligo d(T) primers and random nonamers participate in the PCR reaction, giving many aspecific products. As a RT reaction is performed at 40-50°C, the primers can bind with mismatches to the RNA and therefore transcribe unwanted sequences, which then also will be amplified in to consecutive PCR, leading to aspecific PCR products. This disadvantage is inherent to the method.
In a Two step kit the oligo d(T) primers and the random nonamers are included in the kit and will give ride to cDNA. Then two specific primers have to be selected and this will amplify the exact sequence.
When performing an assay with UNG, a first step at 50°C during 2min and a second step at 95°C during 10min has to be added before performing a normal qPCR:
50 °C during 2min
95 °C during 10min
94 °C during 15s
55 ºC-65 ºC during 30s
72 ºC during 60s
repeat during 35 cycles
Hold at 50 °C for ever
The fist step is needed to activate the UNG enzyme, to allow it to degrade U containing ds DNA.
The second step at 95 °C will deactivate the UNG and will activate the HotGoldStar.
For long term storage the Takyon™ qPCR MasterMixes and Core Kits should be stored at -15 °C to -25 °C in a constant temperature freezer. When stored under these conditions the reagents are stable during 12 months (MasterMixes) or 24 months (Core Kits).
For short term storage the Takyon™ qPCR MasterMixes and Core Kits can be stored at 4 °C for 6 months.
The Takyon™ Dry MasterMix dTTP can be stored at ambient temperature (15-35°C) up to 18-months. This is valid for unopened bottles (inert gas inside for long term stability)
!!! Do not expose the dried reagent to light as ROX normalisation dye is light sensitive
A jagged signal in an amplification plot is typically the result of poor probe signalling, which causes the instrument software to magnify the baseline noise to compensate for the low signal.
There are several potential causes for low signal. Check these three factors first:
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Concentration of the probe: low signal could be a sign that the concentration of your probe is too low.
Melting temperature of the probe: When the Tm of the probe is low, probe binding is inefficient under standard PCR conditions. When the Tm of the probe is close to that of the primers, probe hydrolysis may be compromised.
Probe length: If the distance between the fluorophore and the quencher dye is too far, then inefficient quenching, elevated background fluorescence and lower signal per cycle is expected.
Fluorophore selection is another significant factor. Some pairings of fluorophores and quencher dyes form intramolecular dimers, resulting in a pseudo-beacon structure that is very efficiently quenched. In a robust assay, switching dyes is usually seamless, with only a slight change in quantification cycle (Cq
) value. Such variation is typically in the order of one to two cycles, and relates to the differences in dye intensity and the variation in instrument optics across the different channels. Fundamentally, all real-time thermal cyclers are engineered to detect fluorescein (FAM) most effectively. For some instruments, dyes with longer wavelength emission may be detected less sensitively.False positives or negatives may result from several factors:
A false positive can be recorded if contamination occurred during the PCR setup. This would be evident if negative controls were also returning a positive result. In this case, the source of the contamination needs to be addressed. Be sure to change pipette tips after each use to prevent cross-contamination between samples. Cleaning the reaction setup area and pipettes with a 10% bleach solution is an effective way to remove any potential contaminants. Ensure that pre-PCR and post-PCR processes are physically separated.
False negatives may result from a component missing from the reaction. In many cases, this could mean that no template was added to the sample well or that the sample concentration is too low. However, if the master mix is not appropriately mixed, a reaction component (primer, probe, etc.) could be missing from that sample. Therefore, for the most consistent and accurate results, thoroughly mix and briefly centrifuge all solutions before pipetting.
One source of trouble can be the preparation of the dilutions. A drop-out effect can be observed when performing dilutions with limited content of target DNA. It could be that the template did not make it into the reaction at specified concentrations because DNA may have been absorbed into the plastic surface. Plastics are porous surfaces and are well known to absorb some portion of their contents, including proteins, DNA or other biomolecules.
When manipulating only one or a few molecules of the target sequence, the drop-out becomes much more apparent. We recommend performing all dilutions with a buffer containing nucleic acid unrelated to the target for that situation. Frequently used carriers include yeast tRNA, PolyA, PolyC and linearised acrylamide. This carrier molecule is used in vast excess, e.g. 100 ng/uL, and preconditions the plastic so that the target molecules are less likely to become absorbed.
When using plasmid DNA as a template, amplification is aided by ensuring that supercoiled preparations are nicked or linearised before use.
For qPCR, only the end-users can determine their own experimentally derived threshold setting so they can discriminate between true low-level RFU and signal noise. Ensure the qPCR instrument is calibrated for the dyes being used and check the manufacturer's specifications for their recommended peak height minimum RFU. Judge anything below these levels very cautiously.
It is essential to set the threshold correctly so that the relative cycles between samples remain constant. The qPCR software may initially select an incorrect baseline setting. These baseline fluorescent values are used for threshold setting and so could result in a suboptimal threshold. The threshold should lie above the background noise, evident as the baseline, and within the phase of exponential amplification.
If the threshold is set incorrectly, Cq values would be affected for all assays and may result in lack of detection of samples with lower amounts of template or differences between Cq values between different samples and controls, thus affecting relative quantification.
DNAse contamination (for example, due to bacterial contamination of reagents) or reducing agents carried over into the reaction result in progressively increasing baseline fluorescence. Trace amounts of microbial contamination in reagents can be enough to introduce the DNAse. A microbial cell that makes it into the reaction would be lysed at the high temperature for PCR, releasing DNAse, which degrades the probe and releases the fluorophore from the quencher.
Reducing agents such as DTT are sometimes present in master mixes and reverse transcription reagents (and different batches of polymerase will have different levels of DTT). This can be carried over from reverse transcription reactions, causing reaction degradation. All dark quenchers contain an azo bond (N=N) that represents a site of potential reduction. Once the azo bond is reduced, the quencher becomes non-functional.
Dithiothreitol (DTT is a strong reducing agent capable of reducing the azo linkages present in BHQ™ (Black Hole Quencher™) dyes. Reduction of the azo bond in the BHQ dye results in a loss of quencher activity. Loss of quenching appears as increasing fluorescence and the background is then observed to increase, evident as baseline drift.
Inhibitors in the PCR reaction may cause the efficiency to change. These inhibitors may be carryover contaminants from the RNA or DNA isolation steps. To determine if an inhibitor is present in the template, a careful review of the Cq values of a serial dilution will reveal differences in delta Cq. For a sample without inhibitors present, the delta Cq should remain constant through the dilution series. When a sample does contain inhibitors, differences between Cq for the lower concentrations will be closer to expectation than for those at higher concentrations. When a sample that contains an inhibitor is diluted, the inhibitor is also diluted, therefore the inhibition is evident in high concentrations but less apparent at low concentrations. Diluting away the contaminant is not always an option, and in these situations, an ethanol precipitation or other DNA purification method may be beneficial.
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