Honestly, "no signal at 40 cycles" is pretty much concrete evidence that either your primers do not work, or your target is not there.

PCR is a very, very sensitive technique, and can detect even a single target molecule: by convention, 40 cycles is considered sufficient to amplify anything specific up to reaction saturation, with the corollary that a Cq of 35 or so generally corresponds to amplification of single template (Cq is measured in the log phase, several cycles before reaction plateau).

So, we use 40 cycle protocols, and ignore anything with Cq > 35.

Obviously there is some wiggle room here: an inefficient PCR might produce Cqs of ~38 for a single molecule, but as a general rule of thumb it suffices.

I work with low abundance genes, and I detect them just fine. Sometimes I need to use greater numbers of replicate wells to account for stochastic partitioning (see below), but if a target is there, PCR can detect it.

Also note that "JUST BUNG IN MORE CDNA" is usually counterproductive: cDNA synthesis uses various buffer components that may not play nice with PCR, so you usually should dilute your cDNA before using it in qPCR (I dilute mine 1/20, then use ~2ul in a 10ul reaction, so effectively 0.1ul of cDNA stock).

So. Things we need here.

Positive controls. You absolutely need a sample that HAS your gene of interest (GOI), or failing that, a purified PCR product of your GOI.

You need to know that your PCR works, basically. This is really key. You can also use this to determine reaction efficiency (how much does your signal increase, per cycle: it should double -100%- but anything from 95-105% is considered acceptable -if it's lower, consider redesigning your primers).

You also need negative controls: you need to know that your PCR doesn't detect anything when there's nothing there. I think we can safely say that this is probably the case here, but still.

Once we have these, we can go further: create a standard curve.

Using a purified PCR product of known concentration (and thus, known molecules per ul), ideally padded out with non-specific, confirmed GOI-negative cDNA to replicate crowding effects, prepare a 10-fold dilution series from say, 1x10^9 molecules per ul all the way down to 1x10^-1 molecules. Then run qPCR on this.

What you should see is a clear, linear increase in Cq with greater dilution: about 3.32 cycles for each 10-fold dilution (since 2^3.32 = 10), and at some point it will just go really scatty and start producing wildly different values for replicate wells. This is the stochastic partitioning range: when you only have

Hi,

Technically maximum volume is limited due to other reagents (mastermix, primers). But before this, do optimisation with serial dilution to see your optimum Ct range. For example Try 100, 50, 25ng in the same reaction. This also shows your reaction success.

You can use as much cDNA as technically possible for you (limittations are cDNA concentration and PCR volume).

Another option is to use a pre-amplification (IVT, WTA, PCR).

Honestly, "no signal at 40 cycles" is pretty much concrete evidence that either your primers do not work, or your target is not there.

PCR is a very, very sensitive technique, and can detect even a single target molecule: by convention, 40 cycles is considered sufficient to amplify anything specific up to reaction saturation, with the corollary that a Cq of 35 or so generally corresponds to amplification of single template (Cq is measured in the log phase, several cycles before reaction plateau).

So, we use 40 cycle protocols, and ignore anything with Cq > 35.

Obviously there is some wiggle room here: an inefficient PCR might produce Cqs of ~38 for a single molecule, but as a general rule of thumb it suffices.

I work with low abundance genes, and I detect them just fine. Sometimes I need to use greater numbers of replicate wells to account for stochastic partitioning (see below), but if a target is there, PCR can detect it.

Also note that "JUST BUNG IN MORE CDNA" is usually counterproductive: cDNA synthesis uses various buffer components that may not play nice with PCR, so you usually should dilute your cDNA before using it in qPCR (I dilute mine 1/20, then use ~2ul in a 10ul reaction, so effectively 0.1ul of cDNA stock).

So. Things we need here.

Positive controls. You absolutely need a sample that HAS your gene of interest (GOI), or failing that, a purified PCR product of your GOI.

You need to know that your PCR works, basically. This is really key. You can also use this to determine reaction efficiency (how much does your signal increase, per cycle: it should double -100%- but anything from 95-105% is considered acceptable -if it's lower, consider redesigning your primers).

You also need negative controls: you need to know that your PCR doesn't detect anything when there's nothing there. I think we can safely say that this is probably the case here, but still.

Once we have these, we can go further: create a standard curve.

Using a purified PCR product of known concentration (and thus, known molecules per ul), ideally padded out with non-specific, confirmed GOI-negative cDNA to replicate crowding effects, prepare a 10-fold dilution series from say, 1x10^9 molecules per ul all the way down to 1x10^-1 molecules. Then run qPCR on this.

What you should see is a clear, linear increase in Cq with greater dilution: about 3.32 cycles for each 10-fold dilution (since 2^3.32 = 10), and at some point it will just go really scatty and start producing wildly different values for replicate wells. This is the stochastic partitioning range: when you only have

Hi, i completely agree with John and his recommendations. I never use more than 10% PCR reaction volume of cDNA (so max 2uL for 20 uL reaction).

I assume 10ng is your total RNA, in which your mRNA would be about 1% and within that 1% is your gene of interest. Unless your primers are very efficient your chances of picking up signals with 10 ng of total RNA seem dim.

Jochen Wilhelm suggested pre-amplication , but I not so sure if that's the right way to go ahead. But in case you do not have that option, you can do pre-amplification. Buti would advise you to sequence and confirm that PCR done using pre-amplification is the right product.

I would suggest better to obtain RNA for doing your qRT PCRs. This is a vague suggestion as I do not know, why you have low yield of RNA (10 ng), is it due to limited tissue/cell availability?

All the best.

Thanks.

Prasad.

10 ng is not a lot of RNA to start with and it is possible that you might not detect a lowly expressed gene. If RNA is not a limitation use 500 ng or 1µg RNA from your control sample (if you hve more than one sample to test) to make cDNA and take an aliquot of the cDNA reaction and prepare 5-fold or 10-fold serial dilutions. Use these serially diluted cDNAs to first test the efficiency of you primers (standard curve) to see whether thay can amplify your gene and what is the best dilution of your cDNA.

As per my knowledge 10ng is a low concentration. I would suggest using 100-200 ng concentration of your cDNA and try to use 2-4 ul of your cDNA. Because when we use 1-2ul cDNA, there may be the possibility of pipette loss too.

Rest it also depends upon your master mix which you are using.

All the best.