Molecular Tools

Oligo Dilution Calculator for PCR Primers and Probes

Compute exact stock and diluent volumes to prepare any working concentration from a concentrated oligonucleotide stock. Built on the C₁V₁ = C₂V₂ relationship with automatic µM, nM, and pM unit handling.

Calculate your dilution

Enter your stock concentration, target working concentration, and final volume. The recipe updates instantly — no Calculate button.

Quick presets

Stock (C₁)

Concentration

Concentration of the lyophilised oligo after resuspension.

Working (C₂)

Target concentration

Concentration needed for PCR, qPCR, sequencing, or transfection.

Final volume (V₂)

Total volume

Total volume of working solution to prepare.

Dilution recipe

Combine in a clean microfuge tube:

10.00 µL

of 100 µM stock oligo

90.00 µL

of nuclease-free water or TE buffer

= 100 µL

of 10 µM working solution

Stock to add

10.00

µL

Diluent to add

90.00

µL

Dilution factor

1:10

10.0× dilute

Moles delivered

1000.00

pmol total

Volume composition

10.0%

90.0%

Stock oligo

Nuclease-free diluent

Calculation

C₁V₁ = C₂V₂

V₁ = (C₂ × V₂) / C₁

V₁ = (10 µM × 100 µL) / 100 µM

V₁ = 10.000 µL

Diluent volume

V_diluent = V₂ − V₁

V_diluent = 90.000 µL

Pipetting check

✓Pipetting volumes are within the accurate range of standard P2/P20/P200 micropipettes.

Oligo dilution workflow diagram showing microfuge tube stock-to-working transfer alongside the C1V1 equals C2V2 formula — **Figure 1.** Transfer of a calculated stock volume (V₁) from a concentrated oligonucleotide solution (C₁ = 100 µM) into a destination tube containing nuclease-free diluent, producing a working dilution (C₂ = 10 µM) at the target final volume (V₂). The C₁V₁ = C₂V₂ relationship is a statement of mole conservation: dilution only adds solvent, so the number of moles of oligonucleotide transferred equals the number present in the final solution. This relationship underlies all single-step solute dilutions in molecular biology.

What is the C₁V₁ = C₂V₂ equation?

The dilution equation says that the moles of solute drawn from a stock equal the moles delivered to the final solution. Stock concentration C₁ multiplied by aliquot volume V₁ gives moles transferred. Final concentration C₂ multiplied by final volume V₂ gives moles in the diluted solution. Setting them equal is just stoichiometric bookkeeping — no oligo appears or disappears during dilution.

The equation works for any units, as long as C₁ and C₂ share units and V₁ and V₂ share units. Mixing µM with nM requires conversion (1 µM = 1000 nM); this calculator handles the conversion internally so you can pick any combination of units. The same equation applies to enzyme stocks, BSA solutions, sodium chloride buffers, and any other solute — it is not specific to oligonucleotides.

A non-obvious detail: at very low concentrations (below 10 nM), adsorption to polypropylene tube walls becomes proportionally significant. A nominal 1 nM solution prepared by serial dilution into standard microfuge tubes can lose 30–50% of the oligo to surface binding within hours. Low-binding tubes (silanised or coated) and carrier nucleic acid (10 ng/µL tRNA) mitigate this loss when working with sub-nM concentrations.

Standard oligo dilution workflow

Oligonucleotides ship lyophilised in known nmol amounts. A typical 25-nmol synthesis arrives in a 1.5 mL screw-cap tube with the exact yield printed on the label. The first dilution — resuspension — converts the dry pellet into a stock at a chosen concentration (usually 100 µM). The second dilution — the subject of this calculator — converts the concentrated stock into a working solution at 10 µM or below.

Most labs maintain three concentration tiers per oligo. The 100 µM concentrated stock sits at −80 °C in single-use aliquots and is never freeze-thawed. A 10 µM working stock sits at −20 °C in 50–200 µL aliquots and tolerates 10–20 freeze-thaw cycles. The reaction-ready dilution — typically 1 µM or lower — is prepared fresh weekly and kept at 4 °C. This tiered system limits exposure of the concentrated stock to handling and freeze-thaw stress.

For high-throughput work, robotic liquid handlers prepare 96-well plates of pre-diluted primer pairs at 5 µM each, ready to be added in a single transfer step to qPCR master mixes. The dilution equation determines the volumes per well; the calculator output here is identical to what those systems compute.

Worked examples

Example 1: 100 µM stock to 10 µM working

Goal: prepare 100 µL of 10 µM working stock from a 100 µM resuspended primer.

V₁ = (C₂ × V₂) / C₁ = (10 µM × 100 µL) / 100 µM = 10 µL stock

Diluent = V₂ − V₁ = 100 − 10 = 90 µL TE buffer

Result: 1:10 dilution. P20 pipette handles both transfers comfortably.

Example 2: 100 µM to 250 nM (high dilution factor)

Goal: prepare 50 µL of 250 nM TaqMan probe from 100 µM stock.

Convert: 250 nM = 0.25 µM. V₁ = (0.25 µM × 50 µL) / 100 µM = 0.125 µL stock

Pipetting 0.125 µL is below the accurate range of any pipette. Use a two-step dilution: first dilute 100 µM to 10 µM (5 µL into 45 µL), then dilute 10 µM to 250 nM (1.25 µL into 48.75 µL).

Final dilution factor is 400×, distributed across two 20× and 40× steps.

Practical applications

Standard PCR setup. A 50 µL reaction typically uses 200 nM each of forward and reverse primer, delivered as 1 µL of a 10 µM working stock. The 10 µM dilution prepared with this calculator gives you enough primer for 50–100 reactions per 50 µL of working stock.

qPCR primer-probe mixes. Multiplex qPCR assays combine forward primer, reverse primer, and probe in a single tube. Each component is diluted separately to a working stock (10 µM primers, 5 µM probe), then mixed in defined ratios for the assay format. The dilution calculator runs once per oligo.

siRNA transfection. Knockdown experiments use siRNA at 10–100 nM final concentration in the cell culture well. A standard workflow dilutes 50 µM siRNA stock to 10 µM working concentration, then further dilutes into Opti-MEM for the transfection complex with Lipofectamine.

Hybridisation probes. Northern blots, in situ hybridisation, and FISH all use labelled oligo probes at defined concentrations (typically 100 ng/mL for FISH, 1–10 ng/µL for Northern). Mass-based concentrations require knowing the molecular weight — use the Oligo Concentration Calculator alongside this tool for mass-to-molar conversions.

Limitations and caveats

The C₁V₁ = C₂V₂ equation assumes ideal mixing and no loss of solute. In practice, oligonucleotides adsorb to plastic tube walls — especially at concentrations below 10 nM in standard polypropylene. Dilutions made into low-binding tubes or with carrier nucleic acid (10 ng/µL tRNA or salmon sperm DNA) retain closer to nominal concentration.

Single-step dilutions above 100× compound pipetting error and amplify any concentration drift in the stock. For dilutions exceeding 50×, prefer a two-step serial approach. The calculator flags problematic dilution factors but does not enforce a redesign — the choice depends on your accuracy requirements.

This calculator is educational. For GMP, GLP, or clinical assay preparation, follow the SOP for your facility and validate dilutions empirically by spectrophotometric or fluorometric measurement of the final solution. Reference: IDT oligonucleotide handling guidelines.

Frequently asked questions

What is the standard working concentration for PCR primers?

Most PCR and qPCR protocols use 10 µM working stocks of each primer, then add 0.5–2 µL per 20–50 µL reaction to give a final concentration of 200–500 nM. Probe-based qPCR assays typically use 5 µM probe stocks delivering 100–250 nM final. Working at 10 µM hits the sweet spot: low enough to avoid primer-dimer formation, high enough that small pipetting errors do not translate into large concentration errors. Oligos arrive lyophilised, and the manufacturer ships the absolute nmol amount — you choose the resuspension volume to set the stock concentration, then dilute again to reach the working concentration.

What is the C1V1 = C2V2 formula?

The dilution equation C₁V₁ = C₂V₂ states that the moles of solute in the aliquot drawn from the stock (concentration C₁, volume V₁) equal the moles in the final diluted solution (concentration C₂, volume V₂). Rearranged for the volume of stock needed: V₁ = (C₂ × V₂) / C₁. Diluent volume is simply V₂ − V₁. The relationship holds because dilution only adds solvent — no oligo is created or destroyed. Both concentrations must use the same units; this calculator converts internally so you can mix µM, nM, and pM freely.

Should I dilute oligos in water or TE buffer?

TE buffer (10 mM Tris-HCl, 0.1–1 mM EDTA, pH 8.0) is the standard for long-term oligo storage because EDTA chelates divalent cations that activate trace nucleases, and the buffered pH minimises depurination. For working dilutions used within a week, nuclease-free water is acceptable and avoids introducing EDTA into downstream PCR mixes (high EDTA can sequester Mg2+ and inhibit Taq polymerase). For probes carrying fluorophores like FAM or HEX, follow the manufacturer recommendation — some dyes hydrolyse faster at the slightly alkaline pH of TE.

How accurate are micropipette dilutions?

Calibrated air-displacement micropipettes achieve ±1–3% accuracy in the middle of their range and ±5–10% at the lower limit. A P2 (0.1–2 µL) pipette dispensing 0.5 µL carries roughly twice the relative error of a P20 dispensing 10 µL. For dilutions below 2 µL, prepare an intermediate stock to keep transferred volumes in the accurate range of a P20. For viscous solutions or repeated transfers, positive-displacement pipettes outperform air-displacement designs because they eliminate the air cushion that causes volume variation.

Why does the calculator warn about high dilution factors?

Single-step dilutions above 50× compound pipetting error. Drawing 1 µL of stock into 99 µL of diluent (100× dilution) means a 5% pipetting error on the 1 µL transfer becomes a 5% error on the working concentration. A two-step serial dilution — first 10×, then 10× again — distributes the error across two larger transfers and typically achieves better final accuracy. The calculator flags single-step dilutions above 50× so you can decide whether the precision matters for your downstream assay.

How long do diluted oligos last in storage?

Concentrated stocks at 100 µM in TE buffer remain stable for at least two years at −20 °C with minimal degradation if freeze-thaw cycles are limited to 10 or fewer. Working dilutions at 10 µM or below degrade faster because adsorption to plastic tube walls becomes proportionally more significant at low concentrations. Use low-binding tubes (Eppendorf LoBind or equivalent) for working dilutions below 1 µM. Aliquot stocks into single-use volumes to avoid repeated freeze-thaw, which causes phosphodiester bond cleavage and loss of full-length oligo.

Can I dilute multiple oligos in the same tube to make a primer mix?

Yes, and pre-mixed forward and reverse primers reduce per-reaction pipetting steps in high-throughput PCR. A common format is to mix forward and reverse primers each at 5 µM in the same tube, then add a fixed volume per reaction. The calculator gives the volume of each individual primer to add; the diluent volume then accounts for the combined oligo volume removed. Avoid mixing TaqMan probes with primers in the same tube long-term, as the fluorophore can degrade slowly and the probe will be exhausted before the primers.

What is the difference between this calculator and the oligo resuspension calculator?

The oligo resuspension calculator takes the manufacturer-reported oligo amount in nmol and computes the diluent volume needed to reach a chosen stock concentration — typically 100 µM. The oligo dilution calculator starts from an already-resuspended stock at a known concentration and calculates further dilution to a lower working concentration. The two tools chain together: resuspend the lyophilised pellet first to 100 µM, then use this dilution calculator to reach 10 µM working stocks or 500 nM PCR-ready dilutions.

Educational note. This calculator is built for teaching, coursework, and routine lab planning. For regulated diagnostic or pharmaceutical work, follow facility SOPs and validate empirically. References: IDT oligo handling guidelines, NCBI.