Making a PCR master mix

Setting up more than a couple of PCR reactions one tube at a time is slow and inconsistent. A master mix — one batch of everything the reactions share, made once and aliquoted — is faster and far more reproducible.

Why batch it

Pipetting 1 µL of primer into twenty tubes twenty times invites variation: each small volume carries its own error. Mixing enough primer for all twenty at once, then dispensing a larger combined volume per tube, spreads that error out and keeps every reaction identical in composition.

Scaling the recipe

For each shared component, the total you need is:

total = per-reaction volume × reactions × (1 + overage ÷ 100)

A 20 µL reaction across 20 samples with 10% overage behaves like22 reactions. A component at 10 µL/reaction needs 10 × 22 = 220 µL; a 1 µL/reaction primer needs 22 µL; the whole mix totals 20 × 22 = 440 µL, which you then split 20 µL per tube.

Why the overage

Liquid clings to tips and tube walls, so if you make exactly 20 reactions’ worth you run short before the last tube. Adding 5–10% extra — a couple of reactions’ worth — covers that loss. Use the higher end for viscous mixes or very few reactions, where the fixed loss is a bigger fraction.

What stays out of the mix

Only the shared reagents go in: buffer or 2× mix, dNTPs, primers (if common to all wells), polymerase and water. Template usually stays out — each sample is different — and is added to individual wells after the mix is dispensed. Keep the polymerase cold and add it last.