SYBR Green Quantitative PCR

SYBR Green Quantitative PCR Protocol

Summary
Quantitative PCR is a method used to detect relative or absolute gene expression level. All qPCR involves the use of fluorescence to detect the threshold cycle (Ct) during PCR when the level of fluorescence gives signal over the background and is in the linear portion of the amplified curve. This Ct value is responsible for the accurate quantization of qPCR.
SYBR Green is a dye that intercalates with double-stranded DNA. This intercalation causes the SYBR to fluoresce. The qPCR machine detects
the fluorescence and software calculates Ct values from the intensity of the fluorescence.
This protocol will cover the SYBR Green quantitative PCR technique, and includes suggestions about which kits to use and an overview of
how to analyze your data.

Preparation
Quantitative PCR was initially developed to detect the copy number of transcribed mRNA, and that is basically what we still do when we
perform qPCR.

1. First, RNA must be isolated from your samples. Use a technique for isolating RNA that suits you best. TRIzol methods work well, and
   there are many good kits for it available.
   • Recommended: For isolating from cell culture and mouse liver, use Qiagen’s RNeasy Mini Kit (Cat. No. 74104)
   • This kit will work on many tissues and cell types. For aorta, use Qiagen’s RNeasy Fibrous Tissue Mini Kit (Cat. No. 74704)
2. In order to preserve RNA samples, which are very vulnerable to degradation at room temperature, I recommend using a first-strand DNA synthesis on your RNA in preparation for qPCR, as opposed to running RT-PCR simultaneously with qPCR.
   • Recommended: For first-strand DNA syntheses, I use Invitrogen’s SuperScript III First-Strand Synthesis System for RT-PCR (Cat. No. 18080-051)
   • This will create cDNA in a 1:1 ratio to the RNA in your sample.
3. For the qPCR itself, you will need your cDNA samples, standards made from the samples, primers specific for your genes of interest,
   and a SYBR Green mix (which will include SYBR Green dye, Taq Polymerase, ROX, and dNTP all in one).
   • Recommended: SYBR Green kits are available from many companies. I recommend Qiagen’s QuantiTect SYBR Green PCR Kit (Cat. No. 204143 for 500 rxns, Cat. No. 204145 for 2500 rxns)
   • You will also need an internal control as a point of comparison for your data. Choose a housekeeping gene that is endogenously expressed in your cell type (e.g. β-2 microglobulin, β-actin).
4. Finally, before making the plate, make sure to sign up to use a quantitative PCR machine ahead of time. Two available options:
   • You can sign up to use the one in the Genotyping Core in CHS 36-125 by calling x72461. The cost is $40 per plate, and you need to sign
     in when you submit the plate.
   • Or you can use the one in Dr. Steve Young’s laboratory. Sign up online at http://calendar.yahoo.com with login “younglabqpcr” and password “7500abi”.

Making the Standards
Every gene you run on qPCR will need to be run with a standard curve in order to relatively quantitate the Ct values of your samples.

The following protocol assumes that you have created cDNA from your RNA prior to qPCR. If you followed the Invitrogen SuperScript III kit's protocol, you should start with 21μl of cDNA per sample.

1. Dilute each cDNA sample ~4-fold. In this case, dilute your 21μl with 59μl H₂O for a final volume of 80μl. Vortex and spin down.
2. Pool an equal amount from each sample into a single tube. This will be Standard 1, your high standard. To determine how much to pull from each sample, calculate how much you will have left in each sample and what the final volume of your standards will be. (This is to have approximately the same final volume of standards and samples.)
   • Ex: Take 30μl from each of 12 samples and pool for Standard 1 with a final volume of 360μl.
   • Final sample volume will be sample volume * 5 after a five-fold dilution (see below). In this case, (80μl - 30μl) * 5 = 250μl of each sample, final volume.
   • Take 90μl of Standard 1 in a new tube labeled Standard 2. Dilute Standard 2 with 270μl H₂O for a final volume of 360μl. Repeat up to Standard 5.
   • Final standard volume will be initial standard volume - 90μl after making the next standard. In this case, 360μl Standard 1 - 90μl = 270μl of standard, final volume.
3. Create the rest of your standards by taking out 1/4 of the last standard and diluting it 4-fold. Each standard will have a value assigned to it, like the example below.

Standard Number	Dilution Factor	Dilutions	Value
Standard 1	Pool	360 μl Standard 1	25600
Standard 2	1:4	90 μl Stnd. 1 + 270 μl H2O	6400
Standard 3	1:16	90 μl Stnd. 2 + 270 μl H2O	1600
Standard 4	1:64	90 μl Stnd. 3 + 270 μl H2O	400
Standard 5	1:256	90 μl Stnd. 4 + 270 μl H2O	100

4. Remember to vortex and spin down after each dilution step!

Samples
Dilute your samples (cDNA) further in a 1:5 dilution with H₂O. For example, dilute your remaining 50μl of sample in 200μl H₂O for a final volume of 250μl.

Making the Plate
Before making the plate, draw a layout of how you will pipette it ahead of time so you know what is in each well.

Use plates appropriate to the machine you’ll be using. For the ABI7500 Fast PCR system, use Optical 96-Well Fast Thermal Cycling Plates from ABI (Part No.: 4346906). Use Optical Caps from ABI (Part No.: 4323032) with this plate.

Example: Two genes run on one plate. 18 samples total.

Your plate should look something like the above plate. It needs to include a standard curve for each gene being tested, and needs a water (no cDNA) control for each gene. Be sure to use the correct type of qPCR optical plates and caps for the specific machine you are using. A normal PCR plate will not work.

A typical plate will end up having 20μl in each well, final volume. Add samples and standards in the volumes listed below.

• For the No cDNA wells, pipette 8μl of H₂O.
• For standard wells, pipette 8μl of the corresponding standard.
• For sample wells, pipette 8μl of the corresponding sample.
• Note: Each sample, standard and no cDNA control will be in duplicate as above.

After pipetting the above, create 12μl of master mix for each well, as below, plus excess.

• 1 rxn: 12μl per well (+ 8μl cDNA = 20μl final volume)
  • 10μl SYBR Green
  • 0.4μl Forward Primer (10μM stock)
  • 0.4μl Reverse Primer (10μM stock)
  • 1.2μl H₂O
• 52 rxn: 624μl final volume
  • 520μl SYBR
  • 20.8μl F
  • 20.8μl R
  • 62.4μl H₂O

Pipette 12μl of your master mix into each well and mix gently by pipetting up and down a few times. Do not vortex, as shear
stress can damage your enzyme.

Running the qPCR
Quantitative PCR requires that a certain type of machine capable of detecting SYBR fluorescence while performing PCR be used. (The Young lab, for example, uses the ABI 7500 Fast Real-Time PCR system.)

Each system comes with different software to perform your PCR and analyze your data. Refer to the user’s guide for whichever machine you
are using to learn how to use it, or ask someone who knows to show you. At the end of this protocol, I will cover how to use the ABI7500 Fast Real-Time PCR system specifically.

These programs will invariably ask you to input the layout of your plate prior to running them. In addition, you will have to input your
PCR conditions. Please use conditions ideal for your own primers.

Note: I design all of my primers to ideally function at about 60°C. Therefore, my PCR conditions resemble the following:

• 94°C - 15 minutes (for the Qiagen mix mentioned earlier)
• 94°C - 15 sec. 40 cycles
• 60°C - 30 sec. 40 cycles
• 72°C - 30 sec. 40 cycles

Before running the plate, be sure to spin it down in an appropriate centrifuge to expel as many air bubbles as possible before running.

Analysis
On a basic level, all data from this type of quantitative PCR will be analyzed in a similar manner. However, depending on your experimental design, the way you analyze the data may vary. I’ll go over the basic analysis of data in a control vs. exposure type of experiment to give an example of how to analyze your data.

SYBR Green quantitative PCR machines take readings of the amount of double stranded DNA in each well at each cycle and give the critical threshold (Ct) value, which represents the quantization of your product.

The Ct is a relative value. This is why every experiment needs an internal control, usually a housekeeping gene.

Always check your standard curves for a good slope and R² value. The perfect slope would be -3.32, and R² would be 1. If your slope or R² deviate from these values too much (i.e. a slope of around -4.0, or an R² below .9), your primers are probably not very good.

Example:
In this experiment, we are comparing cells exposed to a certain oxidized phospholipid with cells that are not to see how this exposure effects their expression of Gene X.
Using a six-well cell culture plate, we exposed two of our wells to media (the controls) and two to the oxidized phospholipid (the exposure). We lysed the cells, collected the lysate and isolated RNA, then ran a first-strand synthesis of DNA.
We then created standards from our samples just like above and diluted our samples as above.
Sample 1 + 2 - Control (no oxidized phospholipid)
Sample 3 + 4 - Exposure (+ oxidized phospholipid)
We’ll use the gene β-2 microglobulin (β2M) as our internal control.
Our plate layout looks something like this:

After running in the qPCR machine, we obtain Ct values and relative quantities calculated by the program after data analysis. It calculates these quantities by comparing the Ct of your samples with the standard curve’s arbitrary values.
Since we ran duplicates of each sample, the program should average them and give you a quantity mean for each sample.
β2M is a housekeeping gene, so its expression should not change between control and exposure. Therefore, we use it as a point of comparison for Gene X's expression.

Say we get the following data for each sample:

Start by dividing Gene X's values by β2M's values. This will establish the relative values for Gene X's induction.

Now divide this value by the average of your control values for each one. This normalizes your control's value to 1 and allows you to see the relative induction or reduction of your exposure samples. Average your replicates to get your final value.

In this example, Gene X in the experimental samples (3 and 4) is induced about 70% in response to exposure to the oxidized phospholipid.

Your method of analysis depends on the type of experiment you are performing. However, you will always need to normalize your data at some
point, as above, in order to obtain the relative values for your gene of interest's change in expression.

SYBR qPCR Quick Protocol
Standards

• Dilute the cDNA ~4-fold (e.g. 21μl sample + 59μl H₂O = 80μl)
• Pool an appropriate amount from each sample to create standard 1 (i.e. 30μl x 12 samples = 360μ standard 1; Calculate out the final volume of your standards and try to make it as close to the final volume of your samples as possible.)
• Create standards as follows
  • Pool - 360μl standard 1 - (25600)
  • 1:4 - 90μl standard 1 + 270μl H₂O - (6400)
  • 1:16 - 90μl standard 2 + 270μl H₂O - (1600)
  • 1:64 - 90μl standard 3 + 270μl H₂O - (400)
  • 1:256 - 90μl standard 4 + 270μl H₂O - (100)
• Mix well at each step.

Samples

• Dilute the remaining samples (cDNA) further in a 1:5 dilution with H₂O. For example, dilute your remaining 50μl sample in 200μl H₂O.
• Your samples will end up being about 1/5^th of the high standard.

Plate

• For the No cDNA wells, pipette 8μl of H2O.
• For the standard wells, pipette 8μl of standard.
• For the sample wells, pipette 8μl of sample.
• Run each sample and standard in duplicate.

Assay

• 1 rxn: 12μl per well (+ 8μl cDNA = 20μl final volume)
  • 10μl SYBR Green
  • 0.4μl Forward Primer (10μM stock)
  • 0.4μl Reverse Primer (10μM stock)
  • 1.2μl H₂O
• 52 rxn: 624μl final volume
  • 520μl SYBR
  • 20.8μl F
  • 20.8μl R
  • 62.4μl H₂O

Analysis (on the ABI7500 Fast Real-Time PCR System in Dr. Steve Young’s Lab)

1. Sign up online at http://calendar.yahoo.com with login "younglabqpcr" and password "7500abi".
2. Bring your plate up to MRL 4629. The ABI machine is in a room at the back right of 4629.
3. Start up the computer and ABI7500 machine to allow it to heat up before use.
4. Spin down your plate in the Eppendorf centrifuge for large tubes and plates just outside the ABI room. Make sure the wells are free of bubbles.
5. Open the ABI7500 software and prepare your template.
6. Set your standard values accordingly (25600 for the high standard down to 100 for the low standard).
7. Set up the temperatures and times for your runs accordingly. Be sure to set it to run "standard", not "fast". Set the volume to 20μl and have it take data during the third step.
8. Start the run.
9. After the run, use the analysis tools to analyze your data. Set it to "Auto Ct" and have the program analyze. Check your standard curves for a good slope and R² value (the perfect slope would be -3.32, and R² would be 1).
10. Analyze your data in Excel. Your method of analysis depends on your experimental design.