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A SYBR Green real-time PCR protocol has four parts that must agree with each other: a reaction mix, a cycling program, a melt (dissociation) stage, and controls. This guide sets out the values from one manufacturer’s current user guide, the Thermo Fisher PowerUp SYBR Green Master Mix guide (MAN0013511 Rev G, 9 March 2026), and shows how to choose between its cycling programs. Those values belong to that master mix. If you use a different mix, follow the instructions for use that came with it.
Real-time PCR (also called quantitative PCR or qPCR) is ordinary PCR with fluorescence read at each cycle. SYBR Green is a dye that reports double-stranded DNA, so it needs no probe, but it also reports any unwanted double-stranded product. That is why the melt stage is part of the protocol rather than an optional extra. For the other half of the choice, see TaqMan qPCR; for the broader picture, see the qPCR and RT-qPCR guide.
What you need
- A SYBR Green master mix. The PowerUp mix is a 2X universal master mix containing SYBR Green dye, heat-labile UDG, ROX passive reference dye, and a dNTP blend with dUTP/dTTP. Other mixes differ; see master mix in PCR for what a mix contains.
- Forward and reverse primers designed for a short, specific product (see how to choose primers for PCR).
- Template: the guide specifies 1–10 ng cDNA or 10–100 ng genomic DNA per reaction.
- Nuclease-free water, an optical reaction plate or tubes, and an optical adhesive seal.
- A real-time instrument with its native analysis software (a thermal cycler with fluorescence detection).
- A no-template control (NTC): a well with every component except the sample.
Steps
1. Build the reaction mix
The user guide gives the following setup for a 10 µL or 20 µL reaction. Prepare the mix with 10% overage, and for other volumes scale every component in proportion.
| Component | 10 µL well | 20 µL well |
|---|---|---|
| PowerUp SYBR Green Master Mix (2X) | 5 µL | 10 µL |
| Forward and reverse primers | variable | variable |
| Template and nuclease-free water | variable | variable |
| Total volume | 10 µL | 20 µL |
The guide says to use 300–800 nM of each primer in both Fast and Standard modes.
2. Load the plate, including the NTC
Dispense the mix, add template to the sample wells, and add water in place of template in the NTC wells. Mix, centrifuge briefly to remove bubbles, load an optical plate, seal it with an optical adhesive seal, and centrifuge again. The guide states that the run can start up to 72 hours after setup if the plate is kept at room temperature, and that the plate should be protected from light.
3. Set the instrument
The guide lists these detection settings: reagent SYBR Green, reporter SYBR, quencher none, passive reference ROX. Applied Biosystems systems support ROX as a reporter or as an optional passive reference. The guide notes that Bio-Rad and Roche systems detect ROX but do not apply ROX-based passive reference normalization.
4. Choose the cycling program
The choice depends on the melting temperature (Tm) of your primers and on the template. The guide gives these programs.
| Program | UDG step | Polymerase activation | 40 cycles |
|---|---|---|---|
| Standard, primer Tm ≥ 60 °C | 50 °C, 2 min | 95 °C, 2 min | 95 °C 15 s, then 60 °C 1 min (anneal/extend) |
| Standard, primer Tm < 60 °C | 50 °C, 2 min | 95 °C, 2 min | 95 °C 15 s; anneal 55–60 °C 15 s (set to the melting point of your primers); extend 72 °C 1 min |
The guide also lists a Fast mode, with 40 cycles of 95 °C for 1 s (QuantStudio, ViiA 7, CFX Opus 96, LightCycler PRO) or 3 s (7500 Fast, 7900HT Fast, StepOnePlus, StepOne), then 60 °C for 30 s. Two points decide most choices. First, Fast mode is for primers with a Tm of 60 °C or higher. Second, the guide recommends standard cycling conditions for genomic DNA templates.
5. Add the melt stage
Because the dye detects any double-stranded DNA, the guide says it is important to check for nonspecific amplification, using dissociation (melt) curves. Its melt stage is: 95 °C for 15 s at 1.6 °C/s; 60 °C for 1 min at 1.6 °C/s; then a ramp to 95 °C at 0.15 °C/s, ending with 15 s at 95 °C. The melt can be run up to 72 hours after the amplification run if the plate is stored in the dark, or up to 24 hours if it is exposed to light.
6. Run, then analyze
Analyze with the native software of your instrument. Check that the sample wells amplify, that the NTC returns no Ct value, and what the melt stage shows. For reading the peaks, see qPCR melt curve analysis; for turning Ct values into quantities, see building and validating a qPCR standard curve.
What can go wrong
Nonspecific amplification and primer dimers
SYBR Green binds double-stranded DNA regardless of sequence, so primer dimers and off-target products produce signal just as the wanted product does. Karsai et al. (BioTechniques 2002) describe the principle: the dye reports any double-stranded product, accuracy depends on minimizing nonspecific products including primer dimers, and probes give sequence specificity that the dye lacks. In their Arabidopsis example, the specific UBQ5 product gave a melt peak at 92 °C, while nonspecific products showed a broad melt starting at about 71 °C, and identity was confirmed by melt analysis and an agarose gel. Those temperatures belong to that assay; they are not a general rule, and your own assay has its own expected peak.
Signal in the no-template control
The guide states that the NTC contains all components except the sample and therefore should not return a Ct value. A Ct value in the NTC means the run does not meet that expectation and needs investigating before the sample results are interpreted.
Late or no amplification, and baseline and threshold settings
The user guide has a troubleshooting appendix that covers baseline and threshold settings, primer-dimer formation, and no or late amplification. Use the appendix for the master mix you are running. The PCR protocol basics guide covers the same failure modes for end-point PCR.
Product length
In the Karsai et al. home-made-mix study, fragments of 200–400 bp performed similarly, while efficiency fell for a 709 bp fragment. That result comes from one home-made reaction mixture and one gene, so treat it as an illustration of why short products are usual in SYBR Green assays, not as a limit that applies to every mix.
When to get professional help
Ask the instrument vendor’s support team or your institution’s core facility when the instrument itself misbehaves, or when software settings cannot be reconciled with the user guide. Ask the master mix manufacturer’s technical support when a run follows its guide but still fails. Reporting standards are a separate matter: see the MIQE reporting checklist.
FAQ
What is the difference between PCR and real-time PCR?
End-point PCR amplifies a target and you look at the product afterwards, for example on a gel. Real-time PCR (qPCR) reads fluorescence in every cycle, so the run itself produces data. The variants in searches, such as “pcr real time”, “real time pcr and pcr” and “real-time pcr”, all refer to this technique.
Why does a SYBR Green protocol need a melt curve?
Because the dye detects any double-stranded DNA. The melt stage separates the wanted product from primer dimers and other products by how they dissociate on heating.
Should I use standard or fast cycling?
For the PowerUp mix, use Fast mode only with primers whose Tm is 60 °C or higher, and use standard cycling for genomic DNA templates, as the guide recommends.
How much template and how much primer?
The PowerUp guide specifies 1–10 ng cDNA or 10–100 ng genomic DNA per reaction and 300–800 nM of each primer. Other mixes have their own ranges.
SYBR Green or TaqMan?
SYBR Green needs no probe but cannot tell products apart without a melt analysis; probe chemistries add sequence specificity. See TaqMan qPCR.
Sources
- Thermo Fisher Scientific. PowerUp SYBR Green Master Mix User Guide, MAN0013511 Rev G, 9 March 2026. https://documents.thermofisher.com/TFS-Assets/LSG/manuals/MAN0013511_PowerUp_mastermix_UG.pdf. Date checked: 29 October 2026.
- Karsai A, et al. Evaluation of a homemade SYBR Green I reaction mixture for real-time PCR quantification of gene expression. BioTechniques 2002;32(4):790–796. https://pmc.ncbi.nlm.nih.gov/articles/PMC4353838. Date checked: 29 October 2026.
