PCR Master Mix Calculator
Tell the calculator the reaction volume, how many reactions you are running, and each reagent's stock and final concentration. It returns the volume of every reagent per reaction and in the master mix, with overage and water, as a table you can paste into your notebook. Start from an NEB Taq or Q5 recipe or build your own. Runs entirely in your browser. Free, no sign-up.
Count your controls too.
Extra mix to cover pipetting loss; 10% is a common choice.
Added to each tube separately, not to the master mix.
Reagents
Use the same unit for a reagent's stock and final concentration; the volume per reaction is final ÷ stock × reaction volume.
Results
211.2
Master mix to make (µL)
24
Per tube (µL)
8.8
Reactions made for
Master mix recipe
| Reagent | Stock | Final | Per reaction (µL) | Master mix (µL) |
|---|---|---|---|---|
| 10X Standard Taq reaction buffer | 10 X | 1 X | 2.5 | 22 |
| dNTP mix (each) | 10 mM | 0.2 mM | 0.5 | 4.4 |
| Forward primer | 10 µM | 0.2 µM | 0.5 | 4.4 |
| Reverse primer | 10 µM | 0.2 µM | 0.5 | 4.4 |
| Taq DNA polymerase | 5 U/µL | 0.025 U/µL | 0.125 | 1.1 |
| Nuclease-free water | 19.875 | 174.9 | ||
| Total (template not included) | 24 | 211.2 |
At the bench
- Combine the reagents and water in the table to make 211.2 µL of master mix, adding the polymerase last. Mix gently and keep it on ice.
- Dispense 24 µL into each of the 8 tubes.
- Add 1 µL of template to each tube.
How to Use the PCR Master Mix Calculator
- 1. Pick a starting recipe. The Taq and Q5 options load the stock and final concentrations from NEB's protocol tables. Choose the blank option to enter your own enzyme and buffer.
- 2. Set the reaction volume, number of reactions and overage. Count every tube, including your positive and no-template controls. 10% overage is a common allowance for pipetting loss.
- 3. Check the reagent list. Each row is a reagent with its stock concentration, the final concentration you want, and one shared unit. Edit, add or remove rows to match your kit.
- 4. Mix and dispense. Make the master mix from the Master mix column, dispense the volume shown into each tube, and add template to each tube separately. Copy the table into your notebook.
The Arithmetic
Each reagent is a dilution from its stock to its final concentration in the finished reaction, so its volume follows from C₁V₁ = C₂V₂:
Per reaction
volume (µL) = final concentration ÷ stock concentration × reaction volume (µL)
Master mix
mix volume = per-reaction volume × N × (1 + overage)
Water
water = reaction volume − template − sum of reagent volumes
Stock and final must be in the same unit — a 10 mM stock to a 0.2 mM final, or 10 µM to 0.2 µM — and only the ratio matters. A 10X buffer to a 1X final means one tenth of the reaction volume. To dilute a concentrated stock into a working stock first, use the Dilution Calculator.
Starting Points From NEB
| Reagent | Taq (M0273), 25 µL | Q5 (M0491), 25 µL |
|---|---|---|
| Buffer | 2.5 µL of 10X Standard Taq (1X) | 5 µL of 5X Q5 (1X) |
| 10 mM dNTPs | 0.5 µL (200 µM each) | 0.5 µL (200 µM each) |
| 10 µM primers | 0.5 µL each (0.2 µM) | 1.25 µL each (0.5 µM) |
| Polymerase | 0.125 µL of 5 U/µL (1.25 units per 50 µL) | 0.25 µL of 2 U/µL (0.02 U/µL) |
| Template | variable | variable |
| Water | to 25 µL | to 25 µL |
NEB's 50 µL Taq reaction doubles every volume except that the enzyme is 0.25 µL, and NEB gives the usable primer range as 0.05–1 µM. Its notes say to assemble reactions on ice, add the polymerase last, and move the tubes to a thermocycler already at the denaturation temperature (95 °C for Taq, 98 °C for Q5). The Standard Taq Buffer contains 1.5 mM MgCl₂, and NEB suggests raising magnesium in 0.5 mM steps, up to 4 mM, if an amplification needs optimising. For Q5, the optional 5X High GC Enhancer is added at 1X for difficult, GC-rich templates; add it as an extra row (stock 5, final 1, unit X). Check the current datasheet for your lot, since kit formulations change.
Worked Example: Eight Reactions of Taq PCR
Eight 25 µL Taq reactions with 10% overage (the calculator's defaults) are made for 8 × 1.1 = 8.8 reactions. With 1 µL of template per tube, each tube receives 24 µL of master mix, so the batch is 24 × 8.8 = 211.2 µL:
- 10X buffer: 2.5 × 8.8 = 22 µL
- dNTPs: 0.5 × 8.8 = 4.4 µL; forward primer 4.4 µL; reverse primer 4.4 µL
- Taq: 0.125 × 8.8 = 1.1 µL
- Water: 19.875 × 8.8 = 174.9 µL
The six volumes add up to 211.2 µL. Without the master mix you would pipette Taq at 0.125 µL, which no pipette can do accurately; at 1.1 µL it is well within range. This is the main reason to make a master mix: fewer pipetting steps, less error between tubes, and volumes that are actually measurable.
Controls and Common Mistakes
- Forgetting the controls. Run a no-template control (water in place of template) to catch contamination and a positive control to show the reaction can work. Include them in the reaction count.
- Mixing up stock units. A primer stock in µM and a final in nM gives a thousand-fold error. Keep each row's two concentrations in one unit.
- Putting the template in the master mix. Template varies between tubes, so it goes into each tube individually; the calculator subtracts it before computing water.
- Too little overage. With only 2 or 3 reactions, 10% is less than a single pipetting loss. Make enough for one extra reaction instead.
- Vortexing the enzyme. Mix the finished master mix gently by pipetting or a brief flick and spin, and keep it cold.
Frequently Asked Questions
- How do you calculate a PCR master mix?
- For each reagent, divide the final concentration by the stock concentration and multiply by the reaction volume to get the volume per reaction. Multiply by the number of reactions plus overage to get the master mix volume. Water is the reaction volume minus the template and all reagents.
- How much overage should I add to a master mix?
- 10% is typical, and 5–20% covers most practice. For very small batches, round up to one extra reaction instead, because a percentage of two or three reactions is smaller than the volume lost to the pipette tip and tube walls.
- Does template go in the master mix?
- Normally no. The template differs between samples, so it is added to each tube after the master mix is dispensed. The exception is a set of identical reactions, such as replicates of one sample, where you can include it in the mix and scale it like any other reagent.
- How do I make a 1X buffer from 10X?
- Use one tenth of the reaction volume. For a 25 µL reaction that is 2.5 µL of 10X buffer, and for a 5X buffer such as Q5's it is 5 µL. The calculator does this with stock 10 and final 1 in the X unit.
- What final primer concentration should I use?
- NEB lists 0.2 µM each for its Standard Taq reactions, with a working range of 0.05–1 µM, and 0.5 µM each for Q5. Start with the polymerase manufacturer's recommendation and optimise from there; too much primer promotes primer dimers and non-specific products.
- Should the polymerase go in the master mix?
- Yes, for most protocols. NEB's guidance is to add the polymerase last, mix gently, and keep everything on ice until the tubes go into a pre-heated thermocycler. The enzyme volume is tiny per tube, which is exactly where a master mix improves accuracy.