Hot Start Amplification of DNA with SYBR^® Green JumpStart™ Taq ReadyMix™

Preparation Instructions

DNA Preparation

The single most important step in assuring success with PCR is high quality DNA preparation. Integrity and purity of DNA template is essential. Quantitative PCR involves multiple rounds of enzymatic reactions and is therefore more sensitive to impurities such as proteins, phenol/chloroform, salts, EDTA, and other chemical solvents. Contaminants can also interfere with fluorescence detection. The ratio of absorbance values at 260 nm and 280 nm gives an estimate of DNA purity. Pure DNA has an A₂₆₀/A₂₈₀ ratio of 1.8-2.0. Lower ratios indicate the presence of contaminants such as proteins.

Primer Design

Specific primers for PCR should be designed with the aid of primer design software to eliminate the complications introduced with primer-dimers and secondary structures. Lower primer concentrations decrease the accumulation of primer-dimer formation and nonspecific product formation, which is critical in using SYBR Green I dye in quantitative PCR.

Magnesium Concentration

Lower magnesium chloride concentrations usually result in the formation of fewer nonspecific products. The ReadyMix solution is provided at a 2x concentration of 7 mM magnesium chloride (final concentration 3.5 mM). A vial of a 25 mM magnesium chloride solution is provided for further optimization of the final magnesium chloride concentration if necessary.

Internal Reference Dye

A vial of internal reference dye is included for reaction normalization. Maximum excitation of this dye is 586 nm and maximum emission is 605 nm. Standard instrument settings for ROX reference dye are satisfactory for the measurement of the internal reference dye. This internal reference dye is necessary for ABI Sequence Detection Systems.

Controls

A positive control is always helpful to make sure all of the kit components are working properly. A negative control is necessary to determine if contamination is present. A signal in the no template control demonstrates the presence of DNA contamination or primer dimer formation. See Lovatt, et al., for a thorough discussion of qPCR controls.3

Data Analysis

Follow the recommendations of the real time instrument manufacturer to perform quantitative PCR using SYBR Green I dye. Generally, the log of relative fluorescence is plotted against the number of cycles to determine the threshold cycle (Ct) or crossing point. The Ct value is used to determine the amount of template in each sample.

Consider the following points when determining the Ct:

• Ct is the first detectable increase in fluorescence due to PCR product formation
• Cycles before the Ct are the baseline cycles
• The threshold can be adjusted manually
• Threshold should always be set using a logarithmic amplification plot
• Threshold should be set in the most exponential phase of the reaction, not after reaching the plateau.

Melting Curves

Performing a melting curve analysis at the end of the run will help analyze only the PCR product. Follow the real time instrument manufacturer’s instructions for melting curve analysis. After running a melting curve, any additional runs involving the same PCR product can be done with data collected in an additional detection step to eliminate primer-dimer and other misprimed product signal.

Methods of Quantification

Standard Curves

Standard curves are necessary for both absolute and relative quantification. When generating standard curves, different concentrations of DNA (typically five) should be used to generate a standard curve that will bracket the concentration of the unknown. Each concentration should be run in duplicate.

Absolute and Relative Quantification

This product may be used to quantify target DNA using either absolute or relative quantification. Absolute quantification techniques are used to determine the amount of target DNA in the initial sample, while relative quantification determines the ratio between the amount of target DNA and a reference amplicon. The ideal reference amplicon would have invariant, constitutive expression. In practice, a housekeeping gene is chosen for this function, but there are other reference choices which better adhere to the above requirements.⁴

Absolute quantification uses external standards to determine the absolute amount of target nucleic acid. These external standards contain sequences that are the same as the target sequence or which vary only slightly from the target sequence. The primer binding sites of the external standards are always identical to the target sequence. The similarity between the external standard sequence and the target sequence is necessary for amplification efficiencies between the two to be essentially equivalent. Equivalent amplification efficiencies between the target and external standard are necessary for absolute quantification. A standard curve of external standard dilutions is generated and used to determine the concentrations of unknown target samples.

Relative quantification calculates the ratio between the amount of target template and a reference template in a sample. The relative amount of gene expression is a common application for relative quantification. The reference gene, usually a housekeeping gene, must not vary in concentration in different experimental conditions or tissue states for relative quantification to be possible. Amplification of the target and reference template dilutions in the sample should be performed in separate tubes. SYBR Green PCR quantification does not allow for multiplexing. If the reference template and the target template have different amplification efficiencies, then two standard curves need to be generated. The ratio of the resulting amounts of target and reference in the sample of interest can then be determined from these two standard curves. If the reference template and the target template have very similar amplification efficiencies, then only one standard curve for the reference template needs to be generated to determine the ratio of the amounts of target and reference in the sample.

Determination of PCR Reaction Efficiencies

The PCR efficiency between a reference sample and a target sample is determined by preparing a dilution series for each target. The Ct values from either the reference or target is then subtracted from the other. The difference in Ct values is then plotted against the log of the template amount. If the resulting slope of the straight line is less than 0.1, the amplification efficiencies are similar.

Procedure

Note: Because SYBR Green I binds to all double-stranded DNA, it is important to test primers and cycling conditions to ensure that the PCR product is a single band, or the results will be uninterpretable. It is best to ensure PCR specificity by checking the reaction on a normal (non-quantitative) thermocycler and analyzing the result using agarose gel separation.²

For best results, optimal concentrations of primers, MgCl₂, KCl and PCR adjuncts need to be determined. Testing various combinations of primer concentrations (50-1000 nM) is most efficient for primer optimization. If maximum sensitivity is not required and your PCR target is abundant, satisfactory results for SYBR Green based qPCR are often obtained with final concentrations of both primers 200-400 nM.

The following procedure serves as a guideline to establish optimal primer concentrations. Further optimization may be necessary due to primer specificity.

Note: The use of up to 5% (v/v) dimethyl sulfoxide (DMSO) will not disturb the enzyme-antibody complex. Other co-solvents, solutes (salts) and extremes in pH or other reaction conditions may reduce the affinity of the JumpStart Taq antibody for the Taq polymerase and thereby compromise its effectiveness.

A. Optimizing Primer Concentrations

1. Prepare and dispense diluted primers (Fig 1).
   1. Prepare 60 µL of 8 µM working solutions of both forward (fwd) and reverse (rev) primers in the first tubes of 2 separate 8-tube strips.
   2. Dispense 30 µL of water into tubes 2-5.
   3. Transfer 30 µL of the 8 µM primer solution from tube 1 into tube 2. Mix thoroughly by pipetting up and down at least 5 times.
   4. Repeat transfer and mixing from tube 2 to 3, 3 to 4, and 4 to 5.
   5. Using a multichannel pipettor, transfer 5 µL from the strip-tubes containing diluted fwd primer into the first 5 wells down columns 1-5 of a 96-well PCR plate. After adding fwd primer, PCR mix and template, final concentrations of fwd primer will be 1000, 500, 250, 125, 62.5 nM.
   6. Similarly transfer 5 µL from the strip-tubes containing diluted rev primer into the first 5 wells across rows A-E. After adding PCR mix and template, final concentrations of rev primer will be 1000, 500, 250, 125 and 62.5 nM
2. Prepare qPCR master mix:
   Add reagents below in an appropriate sized DNase-free tube. Mix gently by vortexing and briefly centrifuge to collect all components at the bottom of the tube.

3. Aliquot 26 µL master mix into all wells in the PCR plate that contain primers (A1-E5)
4. Mix Thoroughly and transfer 18 µL from each of wells A1 through E5 to wells A8 through E12.
5. Add 2 µL template DNA (10-50 ng genomic DNA or 0.1-1 ng plasmid) to one set of reactions (columns 1-5) and 2 µL of water to the other columns (8-12).
6. Mix gently by vortexing and briefly centrifuge to collect all components at the bottom of the tube.
7. Perform Thermal cycling:
   
   Optimal cycling parameters vary with primer design and thermal cycler. Consult your thermal cycler manual. It may be necessary to optimize the cycling parameters to achieve maximum product yield and/or quality.
   
   Typical cycling parameters for 100 bp – 600 bp fragments:
   
   This protocol has been successfully tested on the following thermal cyclers: Stratagene MX 3000P, BioRad iCycler, MJ Opticon

8. Evaluate fluorescence plots (DRn) for reactions containing target nucleic acid (columns 1-5). Primer combinations with the lowest C_t and the highest fluorescence will give the most sensitive and reproducible assays.

B. Procedure for Routine Analysis

1. Preparation of a reaction master mix is highly recommended to give best reproducibility. Mix all reagents but template in a common mix, using ~10% more than needed. Once template is diluted into the reaction vessel, master mix is aliquoted into the proper tube or plate for thermocycling.

* Volume for 50 mL reaction, however component volumes may be scaled to give the desired reaction volumes.
** Use 0.1x for ABI 7500 and Stratagene instruments; replace with FITC for BioRad iCycler.

2. Mix gently by vortexing and briefly centrifuge to collect all components at the bottom of the tube.
3. Perform Thermal cycling

Troubleshooting Guide