After extracting RNA, we can get cDNA by reverse transcriptase reaction with a conventional PCR. Then we can amplify the target gene and compare the PCR products with the house keeping gene products. We can also go for the qPCR/RTPCR. But in absence of qPCR, we can compare the gene expression by the above mentioned process. Is it a correct process?
Dear Mohammed
as mentioned in the above very good answer it is possible to look at gene expression by RT PCR but it is semi quantitative at best
- part of the issue is sensitivity: agarose gels can reliably detect products down to about 10ng. In contrast qPCR can reliably detect products down to at least 1ng: to put this in some sort of context, products that yield Ct values of 25-30 can be reliably quantified by qPCR but such products would amount to less than 20ng and therefore could not be reliably quantified on an agarose gel; assuming they could be detected at all
- the fact that small qPCR products - usually 70-150bp - are so small and therefore bind minimal sybr dye or ethdium bromide does not help with sensitivity:
- in other words high copy number genes can be semi quantified on agarose gels whereas low copy number genes cannot
- the most fundamental problem with conventional PCR however is the fact that it is an end point assay: in other words you detect your product at the end and with 25x cycles or more your signal might be saturated precluding reliable distinction between a gene under 2 different conditions
- In contrast, real time PCR by definition measures the product from the outset as it is amplified and so you can set the measurement point within dynamic range; in other words during exponential amplification alllowing you to discern maximal difference between 2 samples with maximal sensitivity
- Nevertheless for high copy number genes if you titrate the cycle number in 3 different PCRs at x20 cycles; x25 cycles; and x30 cycles it is likely that one of the first 2 conditions will put you in suitable dynamic (log) range allowing you to semi estimate gene expression differences. After all this was the accepted procedure before 2000 when real time qPCR started to be used universally
Dear Mohammed
as mentioned in the above very good answer it is possible to look at gene expression by RT PCR but it is semi quantitative at best
- part of the issue is sensitivity: agarose gels can reliably detect products down to about 10ng. In contrast qPCR can reliably detect products down to at least 1ng: to put this in some sort of context, products that yield Ct values of 25-30 can be reliably quantified by qPCR but such products would amount to less than 20ng and therefore could not be reliably quantified on an agarose gel; assuming they could be detected at all
- the fact that small qPCR products - usually 70-150bp - are so small and therefore bind minimal sybr dye or ethdium bromide does not help with sensitivity:
- in other words high copy number genes can be semi quantified on agarose gels whereas low copy number genes cannot
- the most fundamental problem with conventional PCR however is the fact that it is an end point assay: in other words you detect your product at the end and with 25x cycles or more your signal might be saturated precluding reliable distinction between a gene under 2 different conditions
- In contrast, real time PCR by definition measures the product from the outset as it is amplified and so you can set the measurement point within dynamic range; in other words during exponential amplification alllowing you to discern maximal difference between 2 samples with maximal sensitivity
- Nevertheless for high copy number genes if you titrate the cycle number in 3 different PCRs at x20 cycles; x25 cycles; and x30 cycles it is likely that one of the first 2 conditions will put you in suitable dynamic (log) range allowing you to semi estimate gene expression differences. After all this was the accepted procedure before 2000 when real time qPCR started to be used universally
Dear Mohamed,
Though qPCR/qRT-PCR is the 'most' reliable and widely accepted method which uses standard curve to quantify the absolute/relative quantity of mRNA levels. But the RT-PCR-based semi quantitative method could also address this issue, IF the suitability of the method for the purpose and the quality with what the method is applied are well documented.
Kindly refer to: https://www.ncbi.nlm.nih.gov/pubmed/12734582
Good luck !
qRT PCR is always the best method for accurate quantification because it guarantees selection of data points within dynamic range
RT PCR as an end point assay does not
Dear Mohammad,
You can do sqRT-PCR yes, however, it depends on the type of study you are doing. If you only need to confirm expression, or at most show changes in expression you can use this approach. I have done so previously, but designing primers that amplify the same size (500bp or 1kb) is important. So you can still use a reference gene (actin/ubiquitin) to establish a baseline and ensure that all your samples contain equal amount of RNA/cDNA and also to determine the amount of cycles where expression of your reference gene is still in the linear phase. This is usually around 24-26 cycles when you used 1µg of RNA in your cDNA synthesis reaction (20µL) and 3µL of cDNA in a 50µL PCR reaction with Promega GoTaq. However, if your gene of interest is not expressed at high levels, it might not be visible after 25cycles, and therefor I would advise you to use a amplicon size of 1kb when designing your primers, as smaller amplicons require more copies to be visualized on agarose gels.
In conclusion, you can use this method to determine expression levels, however nowadays when you want to publish reviewers will request qPCR instead.
Yes the procedure you mentioned is correct. It can only tell you Whether expression is there or not.
Just one suggestion, kindly treat your RNA with DNase to remove any DNA contamination and set up a PCR reaction for housekeeping gene with RNA and DNase treated RNA before making cDNA.
I think it is hardly possible and if you will fulfill to the MIQE criteria (Bustin 2009) for gene expression studies you will find a number of criteria that cannot be met when using conventional PCR.
FIRST you must figure out 3-5 reference genes that express to the same level as your gene at interest and do not change with respect to experiment-control situations.
SECOND, you must determine the efficiency of the individual PCR reactions (see Radonic 2004) (control/experiment) by dilution series up to 10^7 for at least in 5-fold. Measure bands by densidometry and construct a calibration curve for each of the genes. Exclude data points that show a variation above 5% (> 2SD) from that calibration curve.
THIRD: Detemine that your reference genes are constant during the experiment
FOURTH: find a dilution that for all genes show least variation and calculate the dilution factor.
FIFTH: calculate the starting concentration (only possible if you know which concentration counts for the chosen dilution) and that is not possible with gene expressions as you don’t know what is lost during post-mortem processing and which recovery pertains to your individual genes.
SIXTH: With the efficiency in mind try to use the formula of Pfaffl as publishe in A-Z of PCR Chaper 3 pages 87 - 112
in:
A-Z of quantitative PCR (Editor: S.A. Bustin)
International University Line (IUL)
La Jolla, CA, USA
publication year 2004
Apart from the formula of Pfaffl that corrects for different efficiencies, it is even better to calclate the N0 with LinRegPCR; freeware that can be downloaded from http://www.hartfaalcentrum.nl/index.php?main=files&sub=0
Next to the download extensive description and a tool to combine results from different 96 or 364 wells plates.
Mark that the program works with non-baseline corrected fluorescence data.
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