I’ve already validated the use of a good housekeeping gene in my samples with Sybr Green.
However, for one of my genes of interest, I need to use Taqman assay.
So, I’m wondering if I can use Taqman for my gene of interest and Sybr Green for the housekeeping gene to normalize gene expression.
I'm a little bit surprised by the many responses that advice against the combined use of a DNA binding dye and a sequence specific probe. If the aim of your study is to find relative differences between samples or sample groups (which is the case for >95% of the studies), then this is perfectly possible. At Biogazelle, we sometimes boldly state in our qPCR courses (see link) that there is no mathematical nor technological reason why you could not normalize your gene of interested measured by a hydrolysis probe on a 96-well instrument from manufacturer X with a reference genes (or preferably at least 2 validated reference genes) measured by SYBR Green I on a 384-well instrument from manufacturer Y. Importantly, every sample should be measured with the same method for the same gene. However, it is no problem that different genes are measured with different methods. My answer implicitly also states that a gene of interest and reference gene do not need to be measured in the same plate.
http://www.biogazelle.com/training
Does your qPCR software allows running SYBR and TaqMan simultaneously, and accepts that for ddCt? Try. Otherwise you will need to run the assays separately and calculate manually. Anyway I have never noticed anything like that in a paper.
I think, you should order TaqMan for the same housekeeping gene as you use with SYBR and run it for you gene that requires TaqMan. You may try to design a probe for the same primers you use in SYBR assay for housekeeping gene.
No, you can`t compare samples that have amplified and detected differently. Keep in mind that in order to compare gene expression, the efficiency of the amplification must be the same within the same detection method.
The SYBR dye detects all double stranded DNA, so if you are multiplexing, SYBR signal would reflect both genes. If you are not multiplexing, the problem is that dye-based and probe-based chemistries are different. The optimal formulations for master mixes are different and probe-based chemistries use significantly higher primer concentrations. If you want to compare expression of 2 genes, you need to have reaction conditions as similar as possible, which is one reason why multiplexing is ideal.
I second Kent's comment- literature is showing more and more that one reference gene is not enough to be confident in conclusions.
Make sure you review the MIQE guidelines when designing your experiments as well. Here is a website that lists the original paper and many other useful references: http://miqe.gene-quantification.info/
practically, You can. First, find the value of delta ct of gene expression taqman prob and secondly find the value of delta ct of the housekeeping gene. lastly give fold change by DDCt methot mathematically. but This is not possible in theory and unacceptable. because, Syber green probe less specific than Taqman prob. house keeping gene and the reference gene should be read in the same PCR protocol and in the same plate. Cp value of the housekeeping gene will be required entry into before the reference gene. standardization and normalization studies will not give accurate resultsof different plates and probes.
Nope. Spend the extra money and get a TaqMan probe for your housekeeping gene. Also, several companies routinely sell housekeeping genes primer/probes for gene expression.
I'm a little bit surprised by the many responses that advice against the combined use of a DNA binding dye and a sequence specific probe. If the aim of your study is to find relative differences between samples or sample groups (which is the case for >95% of the studies), then this is perfectly possible. At Biogazelle, we sometimes boldly state in our qPCR courses (see link) that there is no mathematical nor technological reason why you could not normalize your gene of interested measured by a hydrolysis probe on a 96-well instrument from manufacturer X with a reference genes (or preferably at least 2 validated reference genes) measured by SYBR Green I on a 384-well instrument from manufacturer Y. Importantly, every sample should be measured with the same method for the same gene. However, it is no problem that different genes are measured with different methods. My answer implicitly also states that a gene of interest and reference gene do not need to be measured in the same plate.
http://www.biogazelle.com/training
@Jo
I also agree with Jo....There should not be any problem if two different genes are analyzed by different methods....finally we are measuring levels of transcripts....if gene 1 is measured in all samples by method X and gene 2 is estimated by method Y ....as far as relative quantification is concerned....it should be possible.....only the sensitivity of the methods will differ....it should not have significant effect on the results......the problem arise if your plan to analyze some samples of gene 1 with one method and remaining with another and then try to compare....
Because of difference in specificity of SYBER green (lower specificity) and Taq man probe, it would be difficult to compare the results, therefore in my opinion its not advised to use both together in same set of experiment.
Ample evidence here that lots of biologists don't believe in math! If you have a reliable TaqMan assay, then it will give you the relative amounts of your target of interest in your set of samples. And if you have a reliable SYBR green assay, it will do the same. (The information from equally efficient SYBR green and TaqMan assays for the same target should be essentially identical for one set of samples. The Ct values will be different, but the relative Ct values should not be.) So calculations of relative levels--which depend only on the differences between Ct values--should not be affected. The idea is similar to choosing different threshold values for determining Ct. You can move the threshold up or down within the exponential range of the amplification and the Ct values for each sample will change, but the differences between the Ct values will not.
The real problem here may be that reviewers of your manuscript won't believe in math either--so you may need to follow the majority to facilitate acceptance of your manuscript! It's also interesting that using two kinds of assays would be perceived as a "red flag", but most reviewers would never question whether the amplification efficiency of your GOI assay is the same as that for your endogenous reference assay, or even demand to see evidence that your GOI assay is fully specific.
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