Quantitative PCR (qPCR or also called Real-time PCR) is a very useful protocol for determining comparitive populations in a sample. This protocol is primarily for comparing bacterial populations in environmental samples (like silage), but can be used for other purposes as well. RT-qPCR can also be used to quantify and compare RNA in a sample including comparing relative mRNA levels. For more information on qPCR and its uses, please see the qPCR hub page or the PCR hub page. ==Equipment==
- The Genomics Core Facility at Penn State has two Applied Biosystems 7300 Real-Time PCR Systems that you can use. To access these systems you will need to in, and sign up on their calendar "CoreCal".
- To run your sample you'll need a 96 well optical plate with a half skirt (AB part #N801-0560) and an optical film to cover it. You can purchase these directly from the core facility.
- Your DNA can be extracted from any sample. We commonly use a kit for this purpose, but there are a variety of methods around to extract DNA from many materials.
- An important consideration is that the DNA is free of humic acids, which are common contaminants of dna extracted from soil or silage samples. Humic acids will screw up your qPCR.
- Typically 1-20ng of DNA will be added to each (20μL) reaction. This will generally involve diluting your DNA.
To design primers you should probably use a computer program. With the programs suggestions it is important to run a nBLAST to check for non specific binding. It is also important to make sure that their products are the same length if you're going to be comparing different populations using SYBR green. For considerations on how to choose which primers to use, see the analysis section. We use the following primers on a regular basis to identify specific populations:
- Quantification of total bacteria:
- Ftot - GCAGGCCTAACACATGCAAGTC
- Rtot - CTGCTGCCTCCCGTAGGAGT
- Quantification of Lactobacillus spp.
- Flac - GCAGCAGTAGGGAATCTTCCA
- Rlac - GCATTYCACCGCTACACATG
- Quantification of Ferulic acid esterase gene from Lactobacillus jonhsonii
- Ffae - TTAAAACAGATCCGCATGTACGTAA
- Rfae - AGCCCAGCTAACATTGAAGCA
While there are many commercial 2X PCR mixes to choose from, a cheap and easy homemade stock is really useful for most applications.
2X SYBR Mix
This recipe is to make 1ml of 2X qPCR master mix using Taq DNA polymerase (with thermopol buffer) available from New England Biolabs. This mix is enough to make an entire 96 well plate of 20μL qPCR reactions. After this mix is prepared it should be kept on ice until use.
- 690μL Water
- 200μL Thermopol Buffer (with Mg2+)
- 50μL dNTPs
- 40μL ROX 50x dye (this is optional but should be replaced with 40μL water if you're not using it)
- 10μL Taq DNA Polymerase (~15 units)
- 7μL each primer (100μM)
- 3μL SYBR Green II (100x)
You can also make a similar mix using a taqman probe instead of SYBR green. While the probes are expensive (over $100 each), they can lead to more reliable results, and eliminate the need for normalization based on the amplicon length when doing comparisons.
Running the sample
- 1. Keep the master mix and your sample apart until immediately before running.
- 2. Dilute a portion of your samples to have a DNA concentration of 0.1-2ng/μL.
- 3. Place 10μL of the diluted samples in each well in the plate.
- 4. Lightly put the optical film over the plate.
- 5. Take your prepped plate and your 2X master mix (in a tube on ice) down to the core facility.
- 6. Start up the machine.
- 7. Set up the following program.
- Detector = SYBR, Rox = NO
- 95°C for 10 min
- 95°C for 20sec
- 60°C for 60 sec
- Repeat the two-step cycle 45x
- If you want to, you can do a melt curve to be sure that your primers are only amplifying one thing.
- Ramp from 50.0°C to 95.0°C
- Read every 0.2°C h
- Hold 00:00:02 between reads
- 8. Add 10μL of 2X mix to each well (using a multi pipettor for this makes it much easier).
- 9. Quickly centrifuge the plate.
- 10. Place it in the machine and Run the program (you will need to save it first):
- Comparative analysis of qPCR data is usually based on CT (the cycle at which a given sample reaches some threshhold value of DNA fluorescence).
- The threshold for CT can be manually set by the user, but is typically close to the point when the sample fluroescence elevates above background levels.
- In general, the ratio between two CT's in one sample is not very interesting, and provides very little "real" information unless it's based on some kind of calibration. So the analysis methods are used primarily to compare two different CT's between different samples, or to compare many samples to one reference sample. This provides much more in the way of interesting data. An example would be thus In sample A, the CT of population 1 is 2.5 higher than the CT of population 2, but in the control sample the CT's are nearly equal. The 2-ΔΔCT method allows us to more precisely quantify this change.
As Percentage of Total DNA
Using this method the CT of a sample is compared to the CT values of a standard curve of DNA from a pure culture.
- By comparing the amount of DNA added to the sample to the amounts of DNA in the standard cuve, the "percentage of total DNA" can be calculated.
- Results typically feel like, "There were 10ng of DNA added to this well, but the CT is the same as the CT of the well containg 1ng of pure culture DNA; so our target DNA is 10% of the total DNA".
- The difficulty of this method lies in the replicability of DNA extraction from samples (especially plant samples like silage). Often extra plant DNA can be extracted very differently from one sample to another. Further complicating the matter is gleaning useful meaning from this "percentage of total DNA" output.
Using the 2-ΔΔCT Method
This method uses a second set of primers which anneal to a greater population of DNA as an reference. This eliminates the vagueness of the total DNA method and the error introduced by differential DNA extraction.
- The amount of the target DNA relative to the reference DNA is 2-ΔCT where ΔCT = ΔCT, target - ΔCT, reference.
- Typical output feels like "My population is 25% percent of the total bacterial population".
- You actually still have to make a standard curve (and sometimes more than one) for two reasons:
- 1. To be sure that your two primer pairs are amplifying at the same efficiency.
- The ΔCT values should be approximately the same for all dilutions.
- This standard curve can be done with DNA from any sample and need not necessarilly be from a pure culture.
- 2. To calibrate the relationship between the two amplicons.
- If your comparing a one copy gene (like an enzyme) to a many copy gene (like rRNA genes) then a numerical relationship needs to be established between them.
- If your amplicons are different lengths then this will also be taken care of in this calibration.
- Sometimes it helps to use a DNA from pure cultures of different genera to determine this relationship if that's important to you.
Using the Pfaffl Comparison Method
- This method corrects for differences in PCR efficiency as well as amplicon length.
- Should probably run a dilution just to see if your values jive.
- Understand the following:
- Target - The population you're trying to quantify
- Reference - The population you're comparing it to
- Control - A treatment for which you are measuring both target and reference populations
- Sample - A treatment you are comparing to the control treatment and measuring both target and reference populations
- Define an efficiency for each reaction using the slope during the exponential phase of the reaction:
- Efficiency = E = 10-1/slope.
- This is the major difference with the previous method.
- The slope of different dlutions should be the same.
- Define the ΔCT for target and reference:
- ΔCT = Control Cttarget - Sample Cttarget.
- Compare the sample to the control using the CT's (the Pfaffl paper calls these values "CP").
- Ratio = (Etarget)ΔCttarget(control-sample) / (Ereference)ΔCtreference(control-sample)