Hi Claire,

With TaqMan assays, Ct values >35 are not reliable. If you are making cDNA with at least ~200 ng total RNA (A260/280 values ~2.0 or better on NanoDrop) and diluting the cDNA 5X, I would be skeptical about the presence of your gene of interest.

Making cDNA with more total RNA may not help: Assuming near 100% qPCR efficiency, each doubling of concentration will only get you ~one Ct of detection. You'll still be stuck in the Ct 30+ range. As you get to the kit limit of total RNA input, your yield may actually go down due to the presence of inhibitors. We usually get nice TaqMan data with 200-400ng input and ABI's High Capacity cDNA kit.

As others have suggested, normalizing each sample to amplification of an endogenous control is a good idea. For TaqMan assays, I like human PPIA, mouse TBP, and rat RPLP2. Don't multiplex at this stage, and if possible run each assay in technical triplicate.

If you're using Sybr Green assays, good luck ... I've always had better luck with TaqMan.

Hi Claire

If you had ensured that there is no DNA or contamination of any other sort (that can give band)..Ct values aroud 35 are OK to make a point that your gene of interest is there...To further strengthen your point. you can additionally do the following things:

1) you can always keep a -RT contol along with your sample.....

2) you can increase the amount of cDNA so that your gene is detected early

3) Do a melting curve analysis for specificity of your product and

4) Also (initially) cross check the results by running the samples on a gel

5) Since you are getting High Ct ..use a reference gene such as Actin, GAPDH to further give an indication of relative levels of your target gene in the sample

bye

For RT-PCR, to avoid DNA contamination, you can choose primer on exon to exon jonction, so genomic DNA will not cross with your amplification.

To define that your sample is expressed but at low rate, you should compare it to a housekeeping gene of the cells of interest (as ABL, GUS, GAPDH...) and express your result as a ratio.

The best will to find a cell line that express your target, so you can make a calibration with logarithmic dilution of cell expressing your target in cells who did not express your target.

Hi Claire,

I totally agree with Ajay Saini

Hello Claire Morley,

Here already major issues are pointed out. There is no upper or lower limit for Ct value but from my experience Ct values in between 20-30 cycles give highly reliable/repeatable result. In addition to the above points, I wish to add some more

1. Confirm quality of your RNA (if it is total RNA, run in 1.5%-2.0% agarose gel and check for 28s:18s rRNA band).

2. Quantify your RNA and for reverse transcription, in most of the kits, can use at least 1 ug total RNA. Since your Ct value is very low, better to use max allowable (in your reverse transcription kit) RNA concentration.

3. Which primer you used for reverse transcription? oligo dT/random primer/mix of these 2/specific primer? Mix of random primer and oligo dT will give best result.

Hope your problem will be sorted out.

we do a lot of Taqman in our lab and we would classify a Ct of 36 as very low expression or basically non-existent.

Also you should always run a -RT sample along with a NTC on your Taqman.

Hi Claire,

With TaqMan assays, Ct values >35 are not reliable. If you are making cDNA with at least ~200 ng total RNA (A260/280 values ~2.0 or better on NanoDrop) and diluting the cDNA 5X, I would be skeptical about the presence of your gene of interest.

Making cDNA with more total RNA may not help: Assuming near 100% qPCR efficiency, each doubling of concentration will only get you ~one Ct of detection. You'll still be stuck in the Ct 30+ range. As you get to the kit limit of total RNA input, your yield may actually go down due to the presence of inhibitors. We usually get nice TaqMan data with 200-400ng input and ABI's High Capacity cDNA kit.

As others have suggested, normalizing each sample to amplification of an endogenous control is a good idea. For TaqMan assays, I like human PPIA, mouse TBP, and rat RPLP2. Don't multiplex at this stage, and if possible run each assay in technical triplicate.

If you're using Sybr Green assays, good luck ... I've always had better luck with TaqMan.

Here's already a long list of suggestions- one more from me is that QPCR machines (from my experience on ABI) have an option to increase cycles from 40 to 45. It can not increase the Ct value in your templates but it will definitely help you determine the confidence level in target amplification by makingcomparison between -Rt and RT reactions. Beside, even at Ct 35 the beauty of peak itself tells the real story.

Good Luck

Hi everyone, I am blown away by all the answers, there is a lot for me to consider now.

Just wish to clarify a few things on a few details which might help with answers; ...RNA extraction was via TriSure (Bioline) protocol, nanodrop assessment confirmed little gDNA or protein in the samples- all ratios indicated ideal RNA standards. Samples were diluted to 100ng/uL. I used 1uL of each RNA sample in the High Capacity RNA-to-cDNA kit (Applied Biosytems) for reverse transcription and did not dilute my cDNA. I then used 1uL of my cDNA sample within a 50uL TaqMan Gene Expression assay for standard qPCR.

For my target gene Taqman assay; each sample was in triplicate (with ct values between 32-37) and had a -RT sample for which the ct value was 'not determined'.

My endogenous/housekeeper control was b-actin as my samples are from smooth muscle cells, this assay produced ct values between 18-22. The b-actin Taqman assay was primer limited (for use in multiplex) - I used it in a simplex reaction- could this make any difference when I compare the ct values to my target gene?

In light of this extra knowledge; would it be worth me adding more cDNA to my Taqman assay?

Furhtermore, with b-actin being so abundant and my target gene being much less so - is it necessary to compare their expression levels when all I want to do is state its presence/absence?

Thank you again for all those who have already participated in this conversiation, I greatly value your opinions and expertise.

Hi Claire

In my opinion..you have taken care of most of the things ..in view of the fact that you only want to highlight the presence or absence of the gene of your interest..and if your controls (-RT) does not show any indication of involvement of the genomic DNA in your Ct values...you can confortably conclude that the gene is present...However still adjusting the cDNA so that you get Ct values below 30 would be fine (if you can spare that much RNA)...

bye

Hi Claire Morley,

In my view, you have followed all most all the required steps, but I have some more doubts or issues to share.

1. How about your primer? Did you check its efficiency? If beta actin is highly expressed but not your target gene, then there are two possibilities (a) the target gene is not expressed (b) primer may be not good.

So if you have cloned the target gene, check the primer's PCR efficiency

2. I do not favour on using more template, in that case you have to increase concentration 10-100 times to get 3-7 CT value lower to your present CTs (if your primer is 100% efficient).

3. There are some genes which are not expressed all the time. So if your target gene falls in that category, may be use of inducers will give better CTs and hence can confirm the sample has no or undetectable level of expression of the target gene.

Good luck

Thank you for your further answers Ajay and Satheesha, they have been very insightful. I shall look into using an inducer as this target is known to alter its expression levels, a would feel more content achieveing a lower Ct value for my results. Many thanks again.

Ct changes are an effect of using diluent for DNA/RNA. For example: using TE buffer for DNA makes Ct higher than in case of pure H2O. Generally conent of salt can inhibit reaction and affect Ct. So your first step should be standarising all samples.

I would run the product on a gel, isolate the band and send it for sequencing (you may need to clone it). That way nobody can question the specificity of your reaction. Even if you Ct value is above 35 cycles, it may be a gene that is expressed at extreme low levels (it happens sometimes). Also, are your primers in a region highly specific for the gene you are looking to amplify (e.g. look for homology regions among genes, etc…).

Also, because amplification starts so late, make sure your controls are completely negative. Also, run a PCR reaction on your isolated RNA to check if you are carrying any genomic DNA contamination; or to be safe it is best if your primers are designed not to amplify genomic DNA (or pseudogenes).

Hi, the question "is the transcript of this gene present or not?" is as such not a biologically very sensible question - on two grounds:

PCR methods are able to amplify a single molecule up to detectable amounts. When it comes to realy tiny target concentrations, some PCRs will accidently have several target molecules, some may have one or two, others may have none. If the PCR is well optimized, you will get ct values whenever there is at least a single molecule in the particular reaction mix. The distribution of the number of expected molecules per reaction is the Poisson distribution. Consider this example: your source contains, say, 1 transcript per 100 cells. After purification, RT, dilution steps of the cDNA and so on you may have cDNA refering to about 100 cell equivalents per PCR (just as example). In this setting you can expect an AVERAGE number of one target molecule per reaction. According to the Poisson distribution, you can expect that 37% of all PCRs will be negative just because they missed to contain the target molecule. The remaining 63% will be positive (again 37% containing a single molecule, 18% with 2 molecules, 8% with 3 or more molecules). If you perform triplicates, there is almost no chance to reliably estimate the proportion of positive/negative reactions. Three negative reactions are not reliably indicating the systematic absence of target molecules. Three negative PCRs would be expected with a 5% chance if the true average concentration is 1 molecule per PCR. For 1 molecule per 10 PCRs, three negative PCRs are expected with a 74% chance. Thus, one quite likely will come to the false conclusion "there are no target molecules".

This leads to the second ground: What concentration can be considered "absent"? Would 1 molecule in 100 cells have a biological effect? Is 1 in 1000 already enough? What about leaky genes? Is is reasonable to think that an "off-gene" does not not produce any mRNA at all?

So you should first define a critical concentration of molecules per cell or sample (less is considered "absent"). Then you can set a level of confidence with what you like to detect the "absence". Then you can employ the Poisson distribution to find out how many reactions you need to analyse to reach this confidence.

Concerning the ct values: If your PCR and/or the detection is specific (I recommend here to use sequence specific probes like hydrolysis or hybridization probes), you can use a dilution series of a standard with known concentration of target molecules (e.g. a purified PCR product quantified by OD260; this PCR must be done with "normal" dNTPs, no dUTP) to find out if samples with an average concentration of 1 molecule do give ~63% positive reactions (i.e., with a detectable ct). If you did this, you know that each PCR with a detectable ct containes at least one target molecule. It is not relevant what exactely the ct value is (whether it is 30, 35, 40 or anything else). IMPORTANT: Beware of contaminations when working with PCR products (standard)! Do this in a different lab, best in a different building. Purify, measure and dilute the standard there and take only the highest dilutions back into your real-time-PCR lab. Change the lab coat and gloves. Once you have contamination, too many (probably all) PCRs will be positive and it's VERY difficult to get rid of them (best burn down the lab...)! If you work with highly diluted standards (say, 1-10 molecules per µl) contaminations are no problem (the purified PCR product contains in the order of 10^11 molecules per µl!).

Hi Claire, the best way to present your results will certainly be the number of copy of RNA of interest per µg of total RNA used in the reverse transcription step. The easiest way to make this calculations and the assay will be the use of the methematical model from Pfaffl (2001) in respect of the MIQE guideline from Bustin et al 2010. To perform this calculations you will need first to validate your primer on a positve DNA control diluted 10 by 10 until the limit of detection. If this standard reference reveal a limit of detection of 33ct you have to consider your sample (at 35ct) as negative. If your standard reference limit is over 35ct ,your sample is postive. if your standard limit is about 34 ct then you will have to decide. finally it is extremely important to use housekeeping genes in the calculation and as a reference of quality of your samples.

Bye, good luck

Cyrille, fail to get a signal above the limit of detection does *not* indicate that the sample is negative. The PCR might not be sensitive enough. Also you seem to confuse LOD and LOQ (limit of quantification). If the LOD is at 33 cycles, you just won't get higher ct values (unless unspecific products are amplified and detected, e.g. with SYBR Green). To determine the LOD you must know the average number of target molecules in the standards and make use of the Poisson distribution. If the LOD is as low as a single molecule, 37% of the PCRs are expected to be negative anyway, just because of random distributional effects. Taken the other way around, a negative PCR does not rule out that the original sample did not contain any target molecules.

I also agree with Jochen .......absence of signal in any technique.....for example in qPCR at a aprticular cycle....doesnot mean that the sample in negative...it is basically the issue with sensitivty and detection limit....of a technique.......Like a DNA band may not be visible on a gel stained with Ethidium bromide (and we conclude that band is absent)....however same sample will showup band on Silver staining which is more sensitive than EtBR.......whether a gene is present (especially whether.absent) should also be supported by other evidences...

bye

In answer to Jochen- what you have elaborately explained is a very insightful philosophy of Science but the original question still stands on its place. As for the presence or absence of any target RNA molecule, it is where "relative" expression comes to its actual meaning. Rightly said the control sample may even have not one but even 100 copies of RNA, but you also know that the effect you study relevant to the transcript is not or say minimally being affected in presence of those much copies. So what your purpose is i.e. to find ou t the relative no of copies ( fold change) that are present there in order to cause the altered/ observed / desired effect. For Claire- given you have taken care so well, your purpose is to determine fold change in transcript in a condition w.r.t. control condition. The purpose of internal control is to ensure equal loading of template in two PCR reaction tubes. So these above articles are good in philosophy but dont be taken away from your original query.

Mala, I don't see what "fold-changes" do have to do with question if a target sequence is present in a sample. To prove presence, a single positive PCR is enough. Strictly proving absence is not possible, but one can set up an experiment to conclude absence with a given confidence. There is no task to compare "treated" and "controls" or something like this. Such a question might arise when you have three groups, for example say healthy, diseased, and diseased&treated. Now it may be known that th etarget is not expressed in healthy but in diseased specimens. "Not expressed" may be seen more precisely as "if expressed at all, then at a very low level". The upper bound of this level is to be determined as "biologically irrelevant" (for the disease). The diseased specimens do express the target at some higher (i.e. relevant) levels. The treatment is considered effective when the expression of diseaseds specimens is suppressed below the level of relevance. Here, the problem might be formulated in terms of relative expression.

Very rightly said and there's a quantitative term for " some higher level and that becomes "x-fold higher than in control condition" simply in case of QPCR- Quantitative PCR that's why it is known as...........By the way fold change comes in question only when you have some thing present , may be in traces. You can't have a fold change to something which is absent. "0" how many folds changed would remain zero. Thats only what i meant without offending anybody.

Just a short question. How would you recommend to present qPCR data in the article in case when there's no Ct?? i.e. fluorescence does not cross threshold.

In my case I'm comparing the transcipt level in a pair of clinical samples: tumor vs. normal tissue. The gene is expressed in tumor, and I get Ct values after qPCR, but not expressed in normal tissue.

Mikhail, you dont have a quantitative information about an "absent transcript", so there is nothing to show anyway. The principle of real-time PCR does not allow to measure "no transcript", the ct-value would be infinitive, and this in principle can't be measured. What you could do: determine your limit of detection (or the limit of quantification) and define reactions with ct-values above this limit as "negative" (no transcript). You can show this limit as a horizontal line in your graph and indicate the number of reactions having greater ct-values. For dct values this is a littel more complicated - you would have to determine a reasonable "minimal dct" where you can draw the horizontal line.

Jochen, thanks a lot!

I´m very confortable when people use the qPCR graphic given by the computer as a figure in the paper/presentation. To my is the easiest way to represent if a sample is positive or negative

The higher Ct values perhaps depends on relative abundace of target transcript. The low abundant transcripts may require more template to amplify and to give reasonable Ct. First thing needs to be verified is integrity of RNA. Generally if you run your RNA samples on agarose gel you can see the double intensity of upper band than lower band. This indicates RNA is intact. Secondly, you alter the amount of template in your reaction ranging from 10 - 200ng. But, higher concentration of template sometimes having inhibitory effect on amplification. Hence, make different dilutions of your template and test it. Good luck.