Mx4000 – Standard Operating Procedure
Quantitative PCR (QPCR)– Standard Operating Procedure:
Stratagene Mx3005P
Purpose:
Real-time or quantitative PCR (QPCR) is an extremely sensitive and efficient method for quantifying target DNA or cDNA molecules (copies) in a sample. This technique is based on the detection of fluorescence, from a reporter molecule (e.g. SYBR Green), that increases as the PCR product accumulates during amplification. The use of template-specific primers in QPCR allows for the detection of only the targeted DNA at levels as low as one copy. An amplification plot is generated from the fluorescence data for standards and samples. Samples with the highest amount of target DNA, show the earliest increase in fluorescence. Therefore, a standard curve can be generated and the starting DNA copy number in unknown samples can be determined. The following protocol outlines the steps involved in running QPCR, including: 1) Template quantification; 2) Preparing a standard curve; 3) Preparing samples; 4) Set up of the Mx3005P instrument and software; and 5) Preparing and running the QPCR reactions.
I. Template Quantification for the Standard Curve:
In order to validate and calibrate the QPCR results, a standard curve must be created that correlates fluorescence with known template concentration. Any purified DNA (PCR product) or plasmid containing the sequence of interest may be used. Note that the following quantification method is non-specific and measures only total DNA.
1. Generate a PCR product (use the QPCR primers if it is more convenient).
2. Purify the PCR product using the Qiaquick PCR Purification kit (Qiagen #28104) or similar kit.
Note: It is important to purify the PCR product to remove any contaminants (such as DNA polymerase, dNTPs, primers, etc.) that may interfere with the QPCR reaction.
3. Determine the concentration of the purified PCR product by measuring absorbance at A260 (See protocol “Quantifying Nucleic Acids – UV/Vis Spectrophotometer”) or by using a dsDNA quantification kit such as Quant-iT DNA Assay Kits (Molecular Probes #Q33120 or #Q33130) or PicoGreen (Molecular Probes #P7589).
4. Calculate the number of copies (or molecules) of the purified PCR product using the online calculator at or the following formula:
DNA Amount (ng) ( 6.022(1023 (molecules/mole)
Number = _____________________________________________
of Copies
Length (bp) ( 1(109 (ng/g) ( 650 (g/mole of bp)
II. Preparing the Standard Curve:
The following steps outline the preparation of a standard curve from 1.0 x 108 to 1.0 x 101 copies/rxn using a purified PCR product. The standard curve can be prepared with nuclease-free water or with nuclease-free water containing 100 ng/(l yeast tRNA (Sigma #R8508). The yeast tRNA acts as a carrier molecule that will adhere to any binding sites available on the microcentrifuge tube. In our experience, preparation of the standard curve with yeast tRNA allows for multiple freeze-thaw cycles of the same standard curve, and greater sensitivity at the lower concentrations of the curve.
1. Prepare a sufficient amount of 100 ng/(l yeast tRNA in nuclease-free water for your standard curve.
2. Dilute the purified PCR product in nuclease-free water containing 100 ng/(l yeast tRNA to obtain 100 (l
of 2.0 x 108 copies/(l.
3. Prepare 10-fold serial dilutions (8 total) to obtain 8 standards from 2.0 x 107 to 2.0 x 100 copies/(l (e.g. 10.0 (l of sample + 90.0 (l of nuclease-free water containing 100 ng/(l yeast tRNA)
4. Gently mix standards by flicking tubes
5. Centrifuge for 15 seconds and keep on ice, or store at -20(C.
Notes:
• 5.0 (l of each standard is used per QPCR reaction (see below). Therefore, the 8-point standard curve generates a range from 1.0 x 108 to 1.0 x 101 copies/rxn.
• If samples continually fall in the same range on the standard curve, the number of points on the curve may be reduced, or the dilution series may be changed.
• Larger volumes of standards can be prepared, aliquoted and stored at –20oC to be used as needed.
• Dilutions can be prepared in 8 strip PCR tubes to facilitate using a multichannel pipette when transferring the standards to the reaction mixture.
• PCR reactions contain on the order of 1011 copies/(l. Therefore, contamination of the work area with your purified PCR product is a concern. If possible, set up standards and unknowns in different work areas, and use filter pipette tips to prevent contamination of pipettes.
III. Preparing Samples for a QPCR Experiment:
The user must determine the final concentration of template to use, as this is dependent on the abundance of the sequence of interest in the sample. Ideally, sample values should fall within the boundaries of the standard curve. For new users a serial dilution of the sample may be necessary. An example for creating 1, 10 and 100 ng/rxn follows:
1. Determine the DNA concentration in the sample by measuring absorbance at A260 or by using a dsDNA quantification kit such as Quant-iT DNA Assay Kits (Molecular Probes #Q33120 or #Q33130) or PicoGreen (Molecular Probes #P7589).
2. Use nuclease-free water (or nuclease-free water containing 100 ng/(l yeast tRNA) to prepare 50 (l of the sample at a concentration of 20 ng/(l.
3. Prepare two 10-fold serial dilutions to obtain 3 sample concentrations of 20, 2 and 0.2 ng/(l.
4. Gently mix samples by pipeting and keep on ice.
Notes:
• 5.0 (l of each sample will be added to a 20.0 (l PCR reaction to yield final concentrations of 1, 10 and 100 ng/rxn (see below).
• Nuclease-free water containing 100 ng/(l yeast tRNA should be used to dilute unknown samples if it was also used to prepare the standard curve.
IV. Preparing the Mx3005P Instrument and Software
Following are the steps involved in preparing the Mx3005P instrument and software for running a QPCR experiment. However, the user should review the appropriate Instruction Manual prior to use.
Note: The Mx3005P should be left ON at all times (unless shutdown by the GSC manager).
1. Open the MxPro software on the desktop.
2. Select the appropriate method in the New Experiment Options window (eg. SYBR Green – with dissociation curve).
3. Complete the Plate Setup (See user manual)
• Import the plate setup from a previous experiment, OR
• Select wells to be used in the plate
• Select well type and optical path (dyes) to be used for each well
• Identify replicate samples
• Specify standard quantities and select standard units
5. Select the Thermal Profile Setup tab to complete setup (See user manual).
• Import a previously used Thermal Profile, OR
• Setup a new Thermal Profile using the default profile as a template
o Add Segments or Plateaus as needed
o Set temperatures, times and number of cycles for each segment
o Drag and drop a data collection marker over the appropriate plateau (this is the point where fluorescence data is collected by the instrument)
• Add a “Hold” segment if you will not return when the run is finished
V. Preparing Reactions for QPCR
The Brilliant SYBR Green QPCR Master Mix Kit (Stratagene #600548) is recommended for use with the Stratagene Mx3005P. If you are using a dual-labeled (TaqMan-like) probe instead of SYBR Green, you will need to include the probe in the Master Cocktail and use the Brilliant Quantitative PCR Master Mix Kit (Stratagene #600549). Store all reagents at < -20o C until use.
1. Thaw the Brilliant SYBR Green QPCR Master Mix. Keep on ice during set-up procedure and store unused portion at 4o C for up to 6 months (Avoid multiple freeze-thaw cycles).
Note: The SYBR Green I dye present in the master mix is light-sensitive. Therefore, the master mix should be protected from light whenever possible.
2. Determine the number of reactions in the experiment - including standards, samples and controls - and label the well map accordingly.
Note: A No Template Control (NTC) reaction should be run with each experiment to screen for contamination of reagents or false amplification.
3. Dilute the reference dye (ROX) 1:500 using nuclease-free water and keep on ice. For example, add 1.0 (l of ROX to 499 (l of nuclease-free water in a 1.5 ml microcentrifuge tube.
Note: Keep all solutions containing the reference dye protected from light
4. Using the table below determine the volume of each reaction component to add to the Master Cocktail:
|Component |Volume for One |# of Rxns |Component Volume |
| |Rxn | |in Master Cocktail|
| |6.875 (l | | (l |
|PCR H2O | | | |
|2x Brilliant SYBR |12.5 (l | | (l |
|Green Master Mix | | | |
| |0.125 (l | | (l |
|Primer 1 | | | |
|(10 (M stock) | | | |
| |0.125 (l | | (l |
|Primer 2 | | | |
|(10 (M stock) | | | |
|Reference Dye |0.375 (l | | (l |
| |20.0 (l | | (l |
|Total Volume | | | |
Notes:
• Calculate volumes assuming one additional reaction to compensate for pipeting error
• Add the components to an appropriate size tube in the order described in the table
• The user must optimize the final primer concentration (50 nM to 150 nM final concentration is recommended). The table at the right assumes an optimal primer concentration of 50 nM.
5. Gently mix the Master Cocktail with a pipette, being careful not to create bubbles (Do NOT vortex).
6. Keep the Master Cocktail on ice.
7. Assemble the appropriate number of Mx3005P reaction tube strips in a 96-well cold block and keep covered (e.g. lid from pipette tip box).
8. Transfer 20.0 (l of Master Cocktail to each reaction tube.
9. Add 5.0 (l of standard, sample or control to the appropriate tubes.
Notes:
• Standards: 5.0 (l of standards prepared in Section I will provide a standard curve from 1.0 x 108 to 1.0 x 101 copies/rxn.
• Samples: 5.0 (l of sample prepared in Section II will provide final DNA concentrations of 1, 10 and 100 ng/rxn.
• Controls: 5.0 (l of nuclease-free water is added to the NTC tubes.
10. Place and secure a cap strip on top of each tube strip
Notes:
• Gloves should be worn at all times to avoid contamination, but it is especially important to keep fingerprints off of the optical plastic surface of the cap.
• Check that caps are secure. If caps are not seated properly on the tubes, fluorescence readings will be affected.
• After caps are secured label the side of the tube with the appropriate row number.
11. Gently mix the reactions by flicking the bottom of the tubes, being careful not to create bubbles.
Note: Bubbles will interfere with fluorescent detection
12. Centrifuge the reactions briefly.
13. Lift the outer door on the instrument, and then lift the lid on the thermal block.
14. Place the strip tubes in the thermal block in the proper orientation to coincide with your Plate Setup
Note: Wipe tops of strip tubes with a Kimwipe as you are placing them into the thermal block.
15. Confirm that ALL tubes and caps are level and seated completely in the thermal block.
16. Close the lid on the thermal block, and then close the outer door.
17. Run the Real-Time Experiment by selecting the Start Run button on the Thermal Profile Setup page (See user manual).
Note: Be sure the “Turn off lamp at end of run” box is selected.
18. When the run has been completed, remove the tubes from the thermal block.
19. Leave the instrument TURNED ON.
20. Please record your use of the Mx3005P in the logbook, noting the number of hours the lamp was used.
21. For data analysis refer to the MxPro software user manual.
[Created by Ryan Rhodes; 6/2004; Last revision, 9/27/2010]
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