PCR master mix calculator
Maintained by the Talindrew team · Last updated
A PCR master mix scales each component by the number of reactions plus an overage for pipetting loss: volume = per-reaction volume × n × (1 + overage), with 10% the standard overage. Premixing everything except template cuts pipetting steps from n × 7 to n + 7 and removes per-tube variation.
Result
Each component: per-reaction volume × reactions × (1 + overage). 10% overage compensates pipetting loss. Add template separately when it differs per reaction, zero its row here and dispense the mix first.
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What it looks like
Formula
component volume = per-reaction volume × reactions × (1 + overage)
Source: Vendor protocol convention (e.g. NEB Taq/Q5 protocols, 10% overage)
Worked example
Given: 8 reactions of a 25 µl recipe with 1.25 µl of each 10 µM primer, 10% overage
- 1.Scale factor: 8 × 1.10 = 8.8.
- 2.Each primer: 1.25 × 8.8 = 11 µl.
- 3.Repeat per component; dispense 23.5 µl mix per tube, then add template individually.
11 µl of each primer in the master mix
How the calculation flows
Units & constants
| Standard overage | 10% (round up to 1 extra reaction for n < 10) |
|---|---|
| Template | Added per reaction, NOT in the mix (set its row to 0 µl here) |
| Default recipe | 25 µl Taq-style reaction (editable per component) |
| Typical final concs | 0.2 mM each dNTP · 0.5 µM each primer · 1× buffer |
| Price | Free |
Why use a master mix at all?
Pipetting seven components into twenty tubes means 140 chances to vary; premixing collapses that to seven, and every reaction sees exactly the same reagent ratios. The enzyme also survives better in a buffered mix than dosed in 0.125 µl increments: sub-microlitre transfers of glycerol stocks are the least accurate pipetting a bench does.
How much overage do you actually need?
10% covers wetting losses and meniscus error for 8+ reactions. For very small batches (2–4 reactions) make one full extra reaction instead, the fixed dead volume of the tube dominates. For 96-well plates dispensed by multichannel, 10–15% plus reservoir dead volume is realistic.
What goes in last, and why?
Template, always: it is the one component that differs between reactions, and adding it to the shared mix is the classic cross-contamination route. Polymerase goes into the mix second-to-last, on ice, and the mix gets vortexed gently and spun before aliquoting so the glycerol-dense enzyme is evenly distributed.
Frequently asked questions
They mirror a generic 25 µl Taq protocol (NEB-style: 5 µl 5× buffer, 0.5 µl 10 mM dNTPs, 1.25 µl each 10 µM primer, 0.125 µl enzyme). Every row is editable: enter your kit's protocol volumes and the scaling math is identical.
Treat it as one component at 12.5 µl per 25 µl reaction, zero the buffer/dNTP/enzyme rows, and keep primers, water and template as separate rows. The calculator only cares about per-reaction volumes.
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