Compute per-reaction and scaled master-mix volumes for Taq PCR from stock/final concentrations. Flags sub-0.5 µL pipetting precision issues and exports a bench-ready protocol with component addition order.
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Load example PCR master mix scaler data to see the full workflow
Typically 10× stock → 1× final.
Stock mM
Final mM
Stock µM / Final µM
Stock µM / Final µM
Stock (U/µL)
U per reaction
Added per tube (NOT in master mix).
| Component | Stock | Final | Per rxn (µL) | Master mix (µL) | Flag |
|---|---|---|---|---|---|
| 10× PCR buffer | 10 × | 1 × | 2.500 | 132.0 | |
| dNTP mix | 10 mM | 0.2 mM | 0.500 | 26.4 | |
| MgCl₂ | 25 mM | 1.5 mM | 1.500 | 79.2 | |
| Forward primer | 10 µM | 0.5 µM | 1.250 | 66.0 | |
| Reverse primer | 10 µM | 0.5 µM | 1.250 | 66.0 | |
| Taq polymerase | 5 U/µL | 0.04 U/µL | 0.200 | 10.6 | < 0.5 µL |
| Water | — | — | 16.800 | 887.0 | |
| Template (per tube) | — | — | 1.000 | (per tube) |
PCR master mix for 48 reactions × 25 µL (10% overage included).
Master mix (combine in order):
• 887.0 µL nuclease-free water
• 132.0 µL 10× PCR buffer (10 × stock)
• 26.4 µL dNTP mix (10 mM stock)
• 79.2 µL MgCl₂ (25 mM stock)
• 66.0 µL Forward primer (10 µM stock)
• 66.0 µL Reverse primer (10 µM stock)
• 10.6 µL Taq polymerase (5 U/µL stock)
Aliquot 24.0 µL master mix into each tube/well.
Add 1.0 µL template DNA per tube (final volume: 25 µL).
Per-reaction breakdown:
10× PCR buffer 2.50 µL (1 × final)
dNTP mix 0.50 µL (0.2 mM final)
MgCl₂ 1.50 µL (1.5 mM final)
Forward primer 1.25 µL (0.5 µM final)
Reverse primer 1.25 µL (0.5 µM final)
Taq polymerase 0.20 µL (0.04 U/µL final)
Water 16.80 µL
Template 1.00 µL (added per tube)
──────
Total 25.00 µLWhen to use
Do not use for
Add water, then buffer (establishes ionic strength), then small-molecule components (dNTPs, MgCl₂), then primers, then enzyme. Enzyme in a low-salt, room-temperature environment loses activity fast. This order is not a suggestion — it is in every enzyme spec sheet for a reason.
If your Taq is 5 U/µL and you need 1 U per 25 µL reaction, that is 0.2 µL per tube — below the P2 precision floor. Make a 1:5 working dilution in the enzyme's storage buffer (1 U/µL) and pipette 1 µL instead. The working dilution is good for a day at 4 °C; discard after.
P200 tips retain 1-3 µL of dead volume per aspiration, and a 48-reaction master mix has 48 opportunities to lose a little. 10% overage covers this. If you are pipetting with a multichannel into strip tubes, bump to 15% because the dead volume per strip is higher. Only drop to 5% for calibrated robotic dispense.
Check your buffer spec sheet. If the buffer includes 1.5 mM MgCl₂ at 1× and you add another 1.5 mM, you are running at 3.0 mM — enough to generate non-specific bands in most primer sets. If the buffer is MgCl₂-free and you forget to add it, you get no amplification. The checkbox in this calculator ("include MgCl₂") must match your buffer's spec sheet.
Computes per-reaction volume for each component as (final_concentration / stock_concentration) reaction_volume. Polymerase is computed as units_per_reaction / stock_U_per_µL. Water is the balance: reaction_volume − sum(components) − template_volume. Master-mix total scales each component by reaction_count (1 + overage_fraction). Template is excluded from the master mix. Pipetting precision warnings fire when any per-reaction volume falls below 0.5 µL — the practical floor for manual P2 pipettes (CV > 10%). Plate layout reuses the planPlateLayout helper from the tissue digestion calculator.
Last validated 2026-04-07. Calculations are designed for planning and documentation support; verify procurement decisions against manufacturer specifications or institutional SOPs.
ConductScience PCR Master Mix Scaler (v1.7.0). ConductScience, Inc. 2026. Available at: https://conductscience.com/tools/pcr-master-mix-scaler
Saiki RK, Gelfand DH, Stoffel S, et al. Primer-directed enzymatic amplification of DNA with a thermostable DNA polymerase. Science. 1988;239(4839):487-491.
Lorenz TC. Polymerase chain reaction: basic protocol plus troubleshooting and optimization strategies. J Vis Exp. 2012;(63):e3998.
Pipetting 6 components into 48 individual tubes is 288 pipetting steps. Pipetting 6 components into one master-mix tube and aliquoting once is 54 steps — a 5× reduction in opportunities for error. Every extra pipetting step adds 1-3% CV on top of whatever concentration you are targeting, so the master-mix approach is not just faster but also more precise.
Template DNA (the crude tail lysate or purified genomic DNA) varies per sample by definition. It gets added after aliquoting, one tube at a time. The calculator excludes it from the master-mix math and tells you the per-tube aliquot + per-tube template volume as a pair.
Add nuclease-free water to the tube first, then buffer, then dNTPs, then MgCl₂, then primers, then enzyme last. The enzyme goes last because it is the most expensive component and the most vulnerable to denaturation if it sits in a low-salt, high-temperature environment. The bench protocol in this calculator lists components in recommended addition order.
1. dNTP concentration too low — 0.2 mM (each) is standard; dropping to 0.05 mM starves the polymerase. 2. Template volume too high — more than 10% of the reaction volume introduces too many lysate inhibitors (SDS, proteinase K, salt). 3. MgCl₂ omitted when the buffer is MgCl₂-free — uncheck the box only if your buffer truly includes it.
1. MgCl₂ too high — try 1.0-1.5 mM instead of 2.5 mM. 2. Primer concentration too high — drop from 0.5 µM to 0.2-0.3 µM final. 3. Annealing temperature too low — not a master-mix issue but frequently misdiagnosed as one.
1. Template too little — check tissue lysis completeness. 2. Old proteinase K in the lysis step upstream (see the Tissue Digestion Volume Calculator). 3. Pipetting precision on the Taq aliquot — if you are pipetting 0.2 µL, a 20% error means ±0.04 µL, which can halve the enzyme in some tubes. Pre-dilute to a working stock.
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