I got the ct values for Experimental target gene(35.2ct), control target gene(6.7ct), control House Keeping gene(7.0ct) and experimental housekeeping gene(7.1ct).
now i got the fold change value about 1.7. now i am a little bit confused that how to interpret the data in graphical representation. and there is also one thing which i want to know that if some one explain me the qPCR Ampplot attached below. the later two line are experimental target gene.
A few basic guidelines: assuming a robust PCR reaction with good (95-105%) efficiency, a Cq value of 35 corresponds to roughly 1-2 transcripts per well, i.e. "not really expressed". Values greater than 35 correspond to 'mispriming/primer dimers', usually (and melt curves help identify these, assuming SYBR green qPCR).
You may also find that your replicate Cq values are highly variable (32, 35, 38 etc) because you're measuring stochastic noise: at such low levels of template molecules you could easily have five molecules in one well, one in another, and none in the third.
I would not trust a Cq of 35.
Conversely, a Cq of 6 or 7 represents a MASSIVE excess of template: mathematically speaking, a Cq of 6 vs a Cq of 35 is a 29-cycle difference, or 200000000000000000000000000000-fold higher.
You should never need Cq values that low, and similarly, you should never trust Cq values that low: when there is that much template, you cannot really ensure your amplification is 100% efficient.
And you should certainly not be normalising data with a Cq of 35 to reference genes with Cqs of 6 or 7. That would be like calibrating a microgram balance with 'the moon'.
What I suspect you have done here (and I only know this because I have made this mistake myself) is not dilute your cDNA. cDNA synthesis involves various buffer components that are not good for PCR, and moreover a 2ug cDNA prep contains untold gazillions of template molecules. If you use concentrated cDNA (i.e. you don't dilute it), you will usually find that it significantly interferes with your PCR efficiency: moderately expressed genes become hugely inefficient and appear very late or not at all, and highly expressed genes (18S etc) give very low Cq values because even though the reaction is massively inhibited, there is simply bucket tonnes of template there.
You'll also burn through all your cDNA quickly.
I typically dilute my cDNA 1/20, and then use 1-2ul of this in a 10ul reaction.
You can test this by doing a serial 10-fold dilution of your cDNA: if all is well, Cq values should increase by ~3.32 with each 10-fold dilution (of by 1 if you do a 2-fold dilution series). If you don't see this, then you have inhibition issues: avoid concentration, avoid over-dilution, and pick a dilution somewhere in the middle where the data behaves in a linear fashion.
Hi,
could you please send a picture of the multicomponent plot and also check your baseline settings? There seems to be a lot of background but sometimes this might also come in part from the ROX normalization step
Sven
it seems that your qPCR is not optimal...I suppose that first of all, you should improve your PCR conditions...maybe...
A few basic guidelines: assuming a robust PCR reaction with good (95-105%) efficiency, a Cq value of 35 corresponds to roughly 1-2 transcripts per well, i.e. "not really expressed". Values greater than 35 correspond to 'mispriming/primer dimers', usually (and melt curves help identify these, assuming SYBR green qPCR).
You may also find that your replicate Cq values are highly variable (32, 35, 38 etc) because you're measuring stochastic noise: at such low levels of template molecules you could easily have five molecules in one well, one in another, and none in the third.
I would not trust a Cq of 35.
Conversely, a Cq of 6 or 7 represents a MASSIVE excess of template: mathematically speaking, a Cq of 6 vs a Cq of 35 is a 29-cycle difference, or 200000000000000000000000000000-fold higher.
You should never need Cq values that low, and similarly, you should never trust Cq values that low: when there is that much template, you cannot really ensure your amplification is 100% efficient.
And you should certainly not be normalising data with a Cq of 35 to reference genes with Cqs of 6 or 7. That would be like calibrating a microgram balance with 'the moon'.
What I suspect you have done here (and I only know this because I have made this mistake myself) is not dilute your cDNA. cDNA synthesis involves various buffer components that are not good for PCR, and moreover a 2ug cDNA prep contains untold gazillions of template molecules. If you use concentrated cDNA (i.e. you don't dilute it), you will usually find that it significantly interferes with your PCR efficiency: moderately expressed genes become hugely inefficient and appear very late or not at all, and highly expressed genes (18S etc) give very low Cq values because even though the reaction is massively inhibited, there is simply bucket tonnes of template there.
You'll also burn through all your cDNA quickly.
I typically dilute my cDNA 1/20, and then use 1-2ul of this in a 10ul reaction.
You can test this by doing a serial 10-fold dilution of your cDNA: if all is well, Cq values should increase by ~3.32 with each 10-fold dilution (of by 1 if you do a 2-fold dilution series). If you don't see this, then you have inhibition issues: avoid concentration, avoid over-dilution, and pick a dilution somewhere in the middle where the data behaves in a linear fashion.
John Hildyard I have similar thoughts, i should reconsidered the quantification of cDNA and i think i should also need to check for non specific amplification.
Hi,
you can see a Dip of your Rox fluorescence from cycle 1-17. As the insturment is doing a Rox nomralization this might lead to the effect observed on your end. In general the FLU values look a bit low for me but I have mostly worked on ABI7500 so it might be due to the instrument. Still wondering why you see such a dip in Fluorescence (and BTW your target FLU curve looks good). So would try to reanaylze with adapted settings for Baseline
Sven
I saw your amplification curves and saw some important items:
1) A Cq of about 7 indicates a too high amount of target in your sample. About 3 log more dilution would be helpful at first. I suggest you to make a 10X dilution series to begin with a Cq of about 15. Then look if the plateau remains horizontal (an obigation for a correct amplification curve)
2) there is no plateau, which indicates a wrong baseline correction
3) As your Cq is about 7, there are not enough cycles for the software to determine the baselinecorrection. From about 15 cycles the software is able to do a correct baseline correction
4) You use an outdated terminology. Ct is replaced by Cq in 2009. Please read the paper of MIQE https://www.gene-quantification.de/miqe-bustin-et-al-clin-chem-2009.pdf -
5) the MIQE criteria will help you to make a correct setup for your experiments.
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