Having difficulties with your experiment?

We are dedicated to your success. Get back on track. View our expert recommendations for commonly encountered problem scenarios.

View the relevant questions below:

Having problems with your experiment? 
Visit our

Troubleshooting page

Browse our FAQ database for
more information ›

Reverse Transcription

The High Capacity RNA-to-cDNA™ Kit includes a 2X RT buffer mix, which is composed of dNTPs, random octamers, and oligo(dT)16.

The main differences are in the enzyme and type of reverse transcription primer used. The High-Capacity kits use MultiScribe™ Reverse Transcriptase, while the SuperScript® VILO™ Master Mix uses SuperScript® III Reverse Transcriptase. The High-Capacity RNA-to-cDNA™ Kit uses a blend of random primers and oligo(dT), while the High-Capacity cDNA Reverse Transcription Kit and the SuperScript® VILO™ Master Mix use only random primers.

Please review this selection table to help choose the best reverse transcription kit for your needs.

With the High Capacity Reverse Transcription Kit, you can use from 20 ng up to 2 µg of total RNA in a 20 µL reaction.

Yes. You can order the RNase Inhibitor separately as Cat. No. N8080119.

No. The 10X random primers are only available as part of the High-Capacity Reverse Transcription Kit.

You can store your cDNA at 2–6°C for up to 24 hours. For long-term storage, store the cDNA at –15 to –25°C and add EDTA to a final concentration of 1 mM to prevent degradation.

DNase treatment would depend on how your assay or primers were designed. If the probe or primer sits on an exon–exon junction, then the design is such that it will not amplify gDNA. In these cases you do not have to DNase-treat your sample to remove contaminating gDNA. However if, for example, your assay or primers are designed within a single exon, then you will want to treat your RNA with DNase. Our TURBO DNA-free™ Kit (Cat. No. AM1907) is a good option for this.

It is most common to purify the RNA from your sample before proceeding with the next steps in a qPCR reaction. However, depending on your sample type, you may be able to skip the purification process and go directly from a lysate to the reverse transcription step. When used with compatible cell lines, Cells-to-CT™ kits allow you to streamline your workflow. You can find more information on how this works here.

You can use the GeneAmp® 10X PCR Buffer II, which you can order separately as Cat. No. N8080010.

qPCR General

The volume will depend on the starting amount of RNA used for first-strand synthesis, and the abundance of the target gene. We recommend starting with 5 µL of the first-strand reaction in a 50 µL PCR reaction (i.e., 1/10 volume). More than 10% may inhibit downstream reactions. In general, for gene expression you will want to start with 1–100 ng of cDNA per well.

Please see this selection table to compare options for a master mix.

ROX™ is a passive reference dye that is included in many qPCR master mixes. For more details on what it is and its use in qPCR, please check out our short video, “What is ROX™?”

One-step RT-PCR is convenient, and less prone to contamination as there is less opportunity for pipetting error. This method is also faster than two-step. However, the cDNA cannot be archived, and fewer genes can be analyzed. Two-step RT-PCR gives you the ability to archive cDNA, analyze multiple genes, and gives greater flexibility. This table also provides a comparison.

Check out this short video to understand the different phases of the PCR reaction and why they are important.

ROX™ dye is a passive reference that is used to help with data analysis and troubleshooting, such as by reducing the deviation among replicates. ROX™ dye can help, but it is not required for the qPCR reaction or the instrument. For use with an Applied Biosystems® real-time PCR instrument, make sure to change the passive reference to “None”, as it will be set to “ROX” by default in all software.

Please view this short video, which explains some best practices for replicates and plate setup.

Chemistry and Analysis Options

TaqMan® and SYBR® Green chemistries are two different methods of detection for qPCR. Please see this detailed comparison of these two approaches. You can also watch this short video on how TaqMan® assays work.

The optimal primer concentration needs to be determined empirically, but a good starting point is 200–300 nM (each primer). If you are using a master mix, check the manual for specific recommendations.

Multiplexing in qPCR experiments means that you use multiple probes in the same reaction well but with different reporter dyes. This allows you to interrogate more than one gene at a time per reaction. For more details on the pros and cons of multiplexing, please watch our short video, “Advantages and Disadvantages of Singleplex and Duplex qPCR”.

No. A TaqMan® probe, once cleaved, cannot be re-quenched. Therefore a melt curve does not apply when using a TaqMan® assay.

Absolute quantification will quantitate unknowns based on a known quantity. It involves the creation of a standard curve from a target of known quantity (i.e., copy number). Unknowns can then be compared to the standard curve and a value can be extrapolated. Absolute quantification is useful for quantitating copy number of a certain target in DNA or RNA samples. The result usually is a number followed by a unit, such as copy number and ng, etc.

Relative quantification can quantitate a fold difference between samples. It involves the comparison of one sample to another sample (calibrator) of significance. For example, in a drug treatment study you could compare a treated to an untreated sample. The quantity of the calibrator is not known and cannot be measured absolutely. Therefore the calibrator (untreated sample) and samples (treated samples) are normalized to an endogenous control (a gene that is consistently expressed among the samples) and then compared to each other to get a fold difference. Relative quantification is useful for quantitating messenger RNA levels. Since the result is a fold change or ratio, it is not followed by a unit.

The method that you choose will depend on the type of data you need from your experiment. You can find more information here as well.

In a standard curve experiment, you must generate a standard curve for each target gene. The standards should closely represent the sample (i.e., RNA for RNA input, plasmid or gDNA for DNA input). This reference is a good review of standard curves and the experimental setup. You can also review this short video on standard curve experiments.

In a relative quantification experiment, you will need to identify an endogenous control and a reference (or calibrator) sample. An endogenous control is a gene that does not change in expression across all the samples in your study. A reference sample is the sample that you are comparing all others to. This is often the untreated, or control, sample. Please see our Relative Gene Expression Workflow bulletin for more step-by-step guidelines on how to design your experiment.

Comparative Ct experiments use an endogenous control gene to normalize the cDNA input. Please watch this short video for more details on how this works. For a protocol workflow, please refer to our Guide to Performing Relative Quantitation of Gene Expression.

Need more information? Contact us ›