Why you make a master mix at all
A master mix exists to remove variation you cannot see. Every component of a PCR except the template is identical across all the wells on a plate, so pipetting each of them separately into every tube adds one pipetting error per component per well. Combine them once in a single tube, dispense that mix, and each well inherits exactly the same buffer, dNTP, primer and enzyme concentration. The only thing that differs between wells is the sample — which is the only thing you wanted to differ.
The second reason is volume. A 25 µL reaction needs 0.125 µL of a 5 U/µL polymerase. No air-displacement pipette delivers that accurately. Multiply it by 26 reactions and you are pipetting 3.25 µL once, well inside a P10's working range. Small volumes become tractable purely by batching them.
The arithmetic is a single relationship applied component by component: the dilution equation C₁V₁ = C₂V₂. Rearranged for what you actually need, the volume of a component is the reaction volume times the concentration you want, divided by the concentration you have. Water is not calculated from a concentration at all — it is simply whatever volume is left over once every component has claimed its share, which is why it has to be worked out last.
The three calculations, and the one trap in them
Component volumes. For each component, V = Vrxn × Cfinal ÷ Cstock. The units cancel, so it does not matter whether you are working in mM, µM or fold-strength as long as the stock and final figures use the same one. A 10× buffer is the same calculation with Cfinal = 1 and Cstock = 10: one tenth of the reaction volume.
Water. Vwater = Vrxn − ΣVcomponents. If the sum exceeds the reaction volume, the recipe is simply impossible and no amount of arithmetic rescues it; you need a larger reaction, more concentrated stocks, or less template. This calculator returns no water figure at all in that case rather than a negative volume.
Scale-up. Multiply every component by n × (1 + excess). Ten reactions with 10% excess means eleven reactions' worth of every component. The excess is not waste for its own sake: liquid clings to tip walls, the mixing tube has a dead volume, and the last well of a plate is the one that comes up short.
The trap is the template. It belongs in the reaction but not in the master mix, because it is the one thing that differs between wells. So the mix is made to the reaction volume minus the template volume, and dispensed at that reduced volume per tube. Forget this and every reaction ends up over-volume and under-concentrated in everything. This calculator keeps template out of the mix column automatically and dispenses Vrxn − Vtemplate per tube.
Worked example: 24 reactions of 25 µL
You are screening 22 colonies plus a positive and a no-template control — 24 reactions at 25 µL each, with 10% excess. Your stocks are a 10× Taq buffer that already supplies magnesium, a 10 mM dNTP mix, 10 µM primer working stocks, and a 5 U/µL Taq. You will add 1 µL of template per tube.
- Buffer. 25 µL ÷ 10 = 2.50 µL per reaction.
- dNTPs. 25 × 0.2 ÷ 10 = 0.50 µL, giving 0.2 mM of each dNTP.
- Each primer. 25 × 0.4 ÷ 10 = 1.00 µL. Two primers, so 2.00 µL in total.
- Polymerase. 0.125 µL, which is 0.625 units — inside the 0.5–1.25 U most suppliers recommend for a 25 µL reaction.
- Template. 1.00 µL, added per tube, not into the mix.
- Components subtotal. 2.50 + 0.50 + 2.00 + 0.125 + 1.00 = 6.125 µL.
- Water. 25 − 6.125 = 18.875 µL per reaction.
- Scale-up factor. 24 × 1.10 = 26.4 reaction-equivalents.
- Mix per tube. 25 − 1 = 24.00 µL of master mix into each well.
- Master mix total. 24.00 × 26.4 = 633.6 µL.
Inside that 633.6 µL: water 18.875 × 26.4 = 498.3 µL, buffer 2.50 × 26.4 = 66.0 µL, dNTPs 0.50 × 26.4 = 13.2 µL, each primer 1.00 × 26.4 = 26.4 µL, and Taq 0.125 × 26.4 = 3.3 µL. Those five figures sum to 633.6 µL, which is the check worth doing before you touch a pipette.
Note the order of assembly: water first, then buffer, then dNTPs and primers, and the polymerase last, immediately before dispensing. Adding enzyme to concentrated buffer or to neat dNTPs exposes it to conditions it was not stabilised for.
Sanity checks on the numbers you get back
Is any single volume below 0.5 µL? That is the practical floor for a standard air-displacement pipette used well. If a primer comes out at 0.2 µL per reaction, dilute the stock so the volume lands between 0.5 and 2 µL, or pre-combine the forward and reverse primers into one tube and pipette them together. The master mix column often rescues you here: 0.2 µL per reaction is 5.3 µL across 26.4 reactions, which is fine.
Is the water volume comfortable? Water should normally be the largest single component of a conventional PCR. If it has shrunk to under half a microlitre, you have no headroom — any additive, any volume adjustment, any second template will overflow the reaction.
Does the enzyme amount make sense? Most Taq suppliers specify 1.25 units per 50 µL reaction as a starting point; more enzyme buys little and costs specificity. Check what your own supplier states for the exact formulation, because concentrated and hot-start preparations differ.
Does the total reconcile? Add the master-mix column and compare it with the reported total. They must agree exactly. A mismatch means a component was left at zero when it should not have been.
One thing this calculator will not do is decide your final concentrations for you. Those come from your polymerase supplier's protocol and from optimisation. What it guarantees is that whatever concentrations you choose are delivered accurately once you have chosen them.
Typical final concentrations in a conventional PCR
| Component | Common stock | Usual final | µL per 25 µL reaction |
|---|---|---|---|
| Reaction buffer | 10× | 1× | 2.50 |
| dNTP mix | 10 mM each | 0.2 mM each | 0.50 |
| Forward primer | 10 µM | 0.2–0.5 µM | 0.50–1.25 |
| Reverse primer | 10 µM | 0.2–0.5 µM | 0.50–1.25 |
| MgCl₂ (if not in buffer) | 25 mM | 1.5–2.0 mM | 1.50–2.00 |
| Taq polymerase | 5 U/µL | 0.5–1.25 U | 0.10–0.25 |
| Template (genomic) | varies | 1–100 ng | 1–5 |
| Nuclease-free water | — | to volume | remainder |
The final-concentration ranges are the conventional starting points given in standard PCR protocol manuals; always follow the figures your own polymerase supplier specifies for that formulation. The µL column is each range converted for a 25 µL reaction using the stock in the second column.
Magnesium is the one to titrate first
Free magnesium concentration is the parameter that most often decides whether a stubborn PCR works. It matters because dNTPs chelate Mg²⁺ roughly mole for mole, so raising dNTPs without raising magnesium lowers the free Mg²⁺ that the polymerase actually sees. Most 10× buffers supply 15 mM MgCl₂, giving 1.5 mM final, and that is where to start. When a reaction fails, a magnesium titration in 0.5 mM steps from 1.0 to 3.0 mM is usually more productive than changing anything else — and it is easy to set up here by making one mix without magnesium and adding a different supplemental volume to each tube.
Master mix mistakes worth avoiding
- Putting template in the mix. It is the one component that must differ per well. Keep it out and dispense reaction volume minus template volume.
- Forgetting the excess. Without it the last two wells of a plate come up short, and they are usually your controls.
- Calculating water from a concentration. Water has none. It is the remainder, and it must be recalculated whenever any other component changes.
- Confusing dNTP mix concentration with total dNTP. A “10 mM dNTP mix” normally means 10 mM of each nucleotide, 40 mM in total. Using the total figure gives you a quarter of the dNTPs you intended.
- Adding the polymerase first. Enzyme goes in last, into a mix already at working strength, and the mix goes back on ice or in a cold block immediately.
- Vortexing after the enzyme is in. Shearing and foaming denature polymerase. Flick or pipette-mix gently, then spin briefly.
- Ignoring what a ready-made 2× mix already contains. It supplies buffer, dNTPs, magnesium and enzyme. Set those components to zero here or you will double them.
Adapting this to qPCR, multiplex and high-fidelity setups
qPCR with a 2× ready mix. Set the buffer stock to 2×, which claims half the reaction volume, and set dNTPs, magnesium and polymerase to zero because the mix already contains them. Primer final concentrations for SYBR-based assays typically sit between 0.1 and 0.5 µM; probe-based assays add a probe, which you can enter in place of the supplemental magnesium row by giving it the probe's stock and final concentration. Analyse the resulting Ct values with the ΔΔCt fold change calculator.
Multiplex PCR. Each additional primer pair claims more volume and more of the free magnesium and dNTP pool. Work out each pair's volume separately with this calculator at the same reaction volume, add them up, and check the total still leaves water in the reaction. Balancing primer concentrations pair by pair — often lowering the efficient pairs rather than raising the weak ones — matters more than the total.
High-fidelity polymerases. Proofreading enzymes usually come with a 5× buffer, run at a higher primer concentration (commonly 0.5 µM), and are supplied at a lower unit concentration, so the volumes shift. They also degrade single-stranded DNA, which means primers can be chewed back if the enzyme sits with them at room temperature — assemble on ice and start the cycler promptly.
Whatever the chemistry, the two upstream numbers that decide whether the reaction works are the template amount and the primer melting temperature. Quantify the template with the A260 concentration calculator, convert mass to molecules with the DNA copy number calculator, and set the annealing temperature from the primer Tm calculator.