PCR Master Mix Calculator
PCR Master Mix Calculator
Work out how much of one component a batch of PCR reactions needs, with the overage that stops the last tube coming up short — and see why the same overage has to be applied to every component alike.
PCR Master Mix
Per-reaction volume, count, overage → µL22 samples plus a no-template control and a positive control, 10% overage, and the 12.5 µL 2× master mix component of a 25 µL reaction
Formula
reactions = samples + controls
overage = 0.05 to 0.10 conventionally, more for small batches
- per-reaction volume
- how much of this one component a single reaction takes, in microlitres. The calculator handles one component at a time because that is how a mix is actually made up — run it once per line of the recipe
- reactions
- samples PLUS controls. A no-template control and a positive control consume master mix exactly as a sample does, and an extraction blank and an inter-run calibrator do too. Counting only patient samples is the commonest way to run out of mix on the last few wells
- overage
- the fraction of extra mix made to absorb pipetting losses. Conventionally 5–10%, and higher for small batches, because the volume lost to tip and tube surfaces is roughly fixed and therefore a larger proportion of a smaller mix
- why every component
- THE POINT OF THE PAGE: the overage must be applied to every component equally. Adding 10% extra master mix but the nominal volume of primer produces a mix that is diluted in primer — the composition has changed, not just the volume, and the assay is no longer the one that was validated
- what is not in the mix
- template. The sample DNA or cDNA is added to each well separately, which is why the master mix volume per reaction is the total reaction volume minus the template volume
Worked example
22 samples plus a no-template control and a positive control, 10% overage, and the 12.5 µL 2× master mix component of a 25 µL reaction
Reactions = 22 samples + 2 controls = 24. The controls are counted, because they consume mix
Reaction-equivalents with 10% overage = 24 × 1.10 = 26.4
2× master mix at 12.5 µL per reaction: 12.5 × 26.4 = 330.0 µL
Every other component takes the same 26.4 multiplier: forward primer 0.5 µL → 13.2 µL, reverse primer 0.5 µL → 13.2 µL, water 1.5 µL → 39.6 µL
Total mix per reaction = 12.5 + 0.5 + 0.5 + 1.5 = 15.0 µL, so the whole mix is 15.0 × 26.4 = 396.0 µL, and 10 µL of template per well brings each reaction to 25 µL
Had the overage been applied to the master mix alone, the batch would contain 330 µL of master mix with only 12.0 µL of each primer — a mix that is 10% short of primer and no longer the assay that was validated
A full 25 µL recipe at the default batch size
| Component | Per reaction (µL) | × 26.4 reaction-equivalents (µL) |
|---|---|---|
| 2× master mix | 12.5 | 330.0 |
| Forward primer | 0.5 | 13.2 |
| Reverse primer | 0.5 | 13.2 |
| Nuclease-free water | 1.5 | 39.6 |
| Master mix subtotal | 15.0 | 396.0 |
| Template (added per well) | 10.0 | not in the mix |
| Total reaction volume | 25.0 | — |
What the overage costs and covers
| Overage | 24 reactions become | 12.5 µL component (µL) | Suits |
|---|---|---|---|
| 5% | 25.2 equivalents | 315.0 | Large batches with careful, calibrated pipetting |
| 10% | 26.4 equivalents | 330.0 | The usual default for most runs |
| 15% | 27.6 equivalents | 345.0 | Small batches, viscous mixes, manual dispensing |
Why the overage goes on everything
Making a master mix is arithmetic with one wrinkle. The nominal calculation is simply the per-reaction volume of each component multiplied by the number of reactions, but a mix made to that figure never stretches to the last well. Some of it stays on the inside of the tube, some stays in the tip at every transfer, and each aliquot dispensed carries slightly more than the nominal volume because pipettes are calibrated with a positive bias in mind. The shortfall lands entirely on the final few reactions, which is why the classic failure mode is a run where the last column of a plate looks under-amplified.
The fix is to make extra — conventionally 5 to 10% more than the nominal requirement, and more than that for small batches, because the losses are roughly fixed in absolute terms and therefore a larger fraction of a smaller mix. Twenty-four reactions at 10% becomes 26.4 reaction-equivalents. The number is not sacred; what matters is that some margin exists and that it is applied consistently.
Consistency is the part that gets missed. The overage has to be applied to every component in the same proportion, because the mix is defined by its composition and not only by its volume. Adding 10% extra of the enzyme-containing master mix but the nominal volume of each primer produces 330 µL of something that is 10% dilute in primer. The reaction still amplifies, so nothing announces the error, but the primer concentration is now outside what was validated — and primer concentration is one of the parameters that sets both sensitivity and the tendency to form primer-dimer. Applying a single multiplier to the whole recipe, as the table above does, makes this impossible to get wrong.
The second thing that gets missed is the reaction count itself. A no-template control and a positive control are reactions: they take up wells, they consume mix, and they must be in the count. So must an extraction blank and an inter-run calibrator where the assay uses one. It is worth stating why they are not optional. Without a no-template control, a weak positive cannot be distinguished from contamination of the mix itself — and master mix is exactly where contamination does the most damage, because it reaches every well in the run. Without a positive control, a negative result cannot be distinguished from a failed reaction. Counting them into the batch from the start is both a technical and a practical necessity: it is how you avoid discovering, with the plate half-loaded, that there is no mix left for the controls.
Frequently asked questions
How do you calculate a PCR master mix?
Multiply the per-reaction volume of each component by the number of reactions, then by one plus the overage. For 22 samples plus 2 controls at 10% overage, the multiplier is 24 × 1.10 = 26.4, so a 12.5 µL component needs 12.5 × 26.4 = 330 µL. Use the same multiplier for every component.
How much overage should I add to a master mix?
Conventionally 5 to 10%, and more for small batches. The volume lost to tips and tube walls is roughly fixed, so it is a larger fraction of a small mix — a run of eight reactions may need 15 to 20% where a full plate is comfortable at 5%.
Why must the overage be applied to every component?
Because the mix is defined by its composition. Adding extra master mix but the nominal volume of primer gives a mix that is dilute in primer, which changes the assay's sensitivity and its tendency to form primer-dimer. Nothing in the result will reveal the error, so apply one multiplier to the whole recipe.
Do controls count as reactions in the master mix calculation?
Yes. A no-template control, a positive control, an extraction blank and an inter-run calibrator each consume master mix exactly as a sample does. Counting only patient samples is the usual reason a batch runs short on the last few wells.
Is the template included in the master mix volume?
No. Template is added to each well separately, so the master mix volume per reaction is the total reaction volume minus the template volume — 15 µL of mix plus 10 µL of template for a 25 µL reaction.
Related calculators
References
- Bustin SA, Benes V, Garson JA, et al. The MIQE guidelines: minimum information for publication of quantitative real-time PCR experiments. Clin Chem. 2009;55(4):611–622 — on reaction set-up reporting and the required controls.
- Kwok S, Higuchi R. Avoiding false positives with PCR. Nature. 1989;339(6221):237–238.
- Sambrook J, Russell DW. Molecular Cloning: A Laboratory Manual. 3rd ed. Cold Spring Harbor Laboratory Press; 2001 — chapter on the polymerase chain reaction.
Medical Disclaimer: The tools and content provided here are for educational and reference purposes only. They are not intended to substitute for professional medical advice, diagnosis, or treatment. Clinical decisions should always be based on the comprehensive assessment of a qualified healthcare professional.