The qPCR assay worked well previously with amplification efficiency of 1.901. Thing goes wrong in recent. I made a standard curve with 10-fold serial dilution gDNA as template. Late Ct value was obtained in the amplification curve (attached).
I suspect either something is wrong with my dilution skill or the primer is already too old and lost the efficiency.
Could anyone share with me the method to dilute the template? Please let me know if you have any thought. I will be greatly appreciate.
Hi,
first of all you should know what is your sample concentration it shouldn't less than 0.5-1 ng/ul .
Second: for serial dilution you simply need to bring your extract in a tube no1 "undiluted" then tube no2 put 10 ul from your sample + 90 ul PCR grad water and mix well this become 1:10 dilution. take 10 ul from tube no2 put in tube no3 +90 ul PCR grad water this dilution will be 1:100 ......etc,
Small tips:
just make sure you always using PCR grade consumables it mean it is DNAse RNAse free filter tips ,unpowdered gloves. with clean area of work.
- make sure of your master mix sometimes it lose activity cause of freezing and thawing or being forget on bench top also activity varies from company to another.
Are you preparing your working primer fresh each time from a concentrated stock? At dilute concentrations, the effective primer concentrations can go off quickly.
In addition to the suggestions above:
Prepare all triplicates in a single tube, then pipet into plate from there.
Don't freeze-thaw your template gDNA too much.
Use a carrier nucleic acid (e.g., 10165948001 Roche RNA, MS2
from bacteriophage MS2) in your gDNA template during storage and afterwards: this helps the gDNA not stick to the plastic used. (The carrier will offset the sequestering of gDNA onto plastic surfaces).
Negatively charged nucleic acid backbone is attracted to the +charged plastic surfaces, and during serial dilutions, this can result in uneven final replicate Cq values.
Use a diluent spiked with carrier (e.g., 800 ng/uL) for the standard serial dilutions.
Use also for sample dilutions of course.
Check the delivery of your pipets by weighing water with every adjustment (e.g., 900 uL of water, should weigh 0.897 g at 25C (so ~0.9 g). At 4C, 900 uL should weigh almost exactly 0.9 g (but most of us don't work in cold rooms).
And, never adjust the pipet during serial dilutions (one pipet for loading e.g. 0.9 mL water/carrier solution, and another to carry 100 uL from one tube to the next).
Due to the sensitivity of the technique, too low amplification is usually "inefficient". Subsequently, too low Cq/Ct are often outside the linearity range of the curve. that's why you allways need to do diiferent dilutions when doing a QPCR = to check that you are in the linearity range of your curve. else, it is not OK. Don't worry, just don't use the results outside the linearity of your curve.
Your main problem is why you had a larger linearity range before than now, especially if you do exactly the same dilution than before, for this, see the answers before.
But you should certainly check the freshness of your products (particularly PCR mixes and co....) and your DNA concentration.
Another thing that helps is to prepare the standards fresh each time - (e.g., avoid using already freeze-thawed standards). Normally, this is especially true of RNA samples used in one-step qPCR. As Mélanie Beraud has stated above, your curious loss of efficiency indicates some parameter that has changed. Primer concentration may be too low, and/or primers have been freeze-thawed too much. But, often, primers can withstand up to 10 freeze-thaws or more without compromise. In any case, with absolute quantification, you are still left with the dilemma of how to reconcile the different efficiency of the 'absolute' template curve with the different efficiency you will often experience for the same target in your experimental samples. This can be done using the attached transformation.
- Badges
- Science topic