DNA Molarity (nM) Calculator
DNA Molarity (nM) Calculator
Convert a double-stranded DNA concentration in ng/µL into nanomolar, using 660 g/mol per base pair — the calculation that normalises sequencing libraries, where molarity and not mass is what gets loaded.
DNA Molarity (nM)
ng/µL, bp → nM10 ng/µL of a library with a mean fragment length of 350 bp
Formula
- 660 g/mol
- the average molar mass of one base pair of double-stranded DNA, backbone and counter-ion included. It is an average over base composition, so an unusually GC-rich or AT-rich fragment departs from it by a few per cent. For single-stranded DNA the corresponding figure is about 330 g/mol per base
- 1e6
- the unit conversion, and worth verifying rather than trusting. 1 ng/µL is 1e-9 g in 1e-6 L, which is 1e-3 g/L. Dividing by the molar mass 660 × bp gives mol/L, and multiplying by 1e9 gives nmol/L — net effect, multiply the ng/µL figure by 1e6 and divide by 660 × bp
- length
- the length of the whole double-stranded molecule. For a sequencing library that means the insert plus both adapters, roughly 120 bp of extra length, taken as a mean from an electrophoresis trace. Using the insert length alone overstates molarity
- why molarity
- because clustering and loading are molecular events. A sequencer’s loading concentration is specified in picomolar or nanomolar, and two libraries at identical ng/µL but different mean lengths contain different numbers of molecules — so pooling by mass gives unequal read shares
- what it shares with copy number
- the same 660 g/mol constant. Copy number answers how many molecules are in a stated MASS; molarity answers how many are in a stated VOLUME. One is an absolute count for a standard curve, the other a concentration for loading or pooling
Worked example
10 ng/µL of a library with a mean fragment length of 350 bp
Molar mass of the fragment = 660 × 350 = 231,000 g/mol
Numerator = 10 × 1e6 = 1e7
nM = 1e7 ÷ 231,000 = 43.29 nM
To load at 4 nM, the dilution factor is 43.29 ÷ 4 ≈ 10.8 — so roughly 1 part library to 9.8 parts buffer
Now double the length and hold the mass concentration: 10 ng/µL of a 700 bp library is 21.65 nM, exactly half. Same mass, half the molecules, because each molecule is twice as heavy
That halving is the argument for pooling by molarity. Two libraries both at 10 ng/µL, one at 350 bp and one at 700 bp, differ twofold in molecule number, and pooling them equally by volume would give the shorter one twice the reads
Molarity of 1 ng/µL at different fragment lengths
| Fragment | Length (bp) | nM at 1 ng/µL |
|---|---|---|
| Short amplicon or adapter-dimer | 100 | 15.15 |
| Typical short-read library | 350 | 4.33 |
| Long-insert library | 500 | 3.03 |
| Long amplicon | 1,000 | 1.52 |
| Small plasmid | 3,000 | 0.51 |
| Large construct | 10,000 | 0.15 |
Molarity against mass at 350 bp
| Concentration (ng/µL) | Length (bp) | nM |
|---|---|---|
| 2 | 350 | 8.66 |
| 10 | 350 | 43.29 |
| 10 | 700 | 21.65 |
| 20 | 450 | 67.34 |
Why sequencing works in nanomolar and not nanograms
Almost every measurement of DNA is a mass measurement. A fluorometer reports nanograms per microlitre, an absorbance reading reports micrograms per millilitre, and both answer the question of how much material is present. But the things that happen to a library on a sequencer are molecular events: a molecule binds to the flow cell, a molecule seeds a cluster, a molecule is read. Loading concentrations are therefore specified in picomolar or nanomolar, and the conversion from mass to molarity is the step that has to happen in between.
The conversion needs only the average molar mass of a base pair of double-stranded DNA, taken as 660 g/mol. Multiplying the length in base pairs by 660 gives the molar mass of the fragment; dividing the mass concentration by that gives a molar concentration. Folding the unit conversions in leaves the compact form used here: multiply the ng/µL figure by a million and divide by 660 times the length. The constant is an average across base composition, so an extremely GC-rich fragment departs from it slightly, but over a library of a few hundred base pairs the departure is far smaller than the uncertainty in the mass measurement.
The length is where the errors are. Molarity is inversely proportional to it, so a twofold error in length is a twofold error in molarity. Two habits cause most of the trouble. The first is using the designed insert size rather than the actual molecule: adapters add roughly 120 base pairs to each fragment, and ignoring them overstates the molarity by that proportion. The second is using a nominal figure instead of the mean from an electrophoresis trace — real libraries have a distribution of lengths, and where that distribution sits is an empirical fact about the preparation, not a design parameter. A trace also reveals adapter-dimer, a short all-adapter species that quantifies well, clusters efficiently and produces reads containing nothing.
The practical payoff is in pooling. When several libraries are sequenced together, what determines each one’s share of the reads is its share of the molecules, not of the mass. Two libraries at the same ng/µL with mean lengths of 350 and 700 base pairs differ twofold in molarity, so pooling them equally by volume gives the shorter one twice the coverage of the longer. Converting each to nanomolar first, then pooling by molarity, is what makes the read allocation match the intention. This page and the copy number calculator use the same 660 g/mol constant to answer two different questions: copy number gives an absolute molecule count in a stated mass, which is what a standard curve needs, while molarity gives a concentration per unit volume, which is what loading and pooling need.
Frequently asked questions
How do you convert ng/µL to nM for DNA?
nM = (ng/µL × 1,000,000) ÷ (660 × length in bp). The 660 is the average molar mass of a base pair of double-stranded DNA. At 10 ng/µL and 350 bp the result is 10,000,000 ÷ 231,000 = 43.29 nM.
Should adapters be included in the fragment length?
Yes. The molecule being counted is the insert plus both adapters, which adds roughly 120 bp. Using the insert length alone overstates the molarity in proportion, and the length should be a mean taken from an electrophoresis trace rather than the size you designed for.
Why pool sequencing libraries by molarity rather than by mass?
Because read share follows molecule number. Two libraries at the same ng/µL but mean lengths of 350 and 700 bp differ twofold in molarity, so pooling them equally by volume gives the shorter library twice the reads. Equalising molarity equalises the molecules loaded.
What is the difference between DNA molarity and DNA copy number?
They use the same 660 g/mol constant to answer different questions. Copy number converts a stated mass into an absolute number of molecules, which is what a qPCR standard curve needs. Molarity converts a concentration into molecules per unit volume, which is what loading and pooling need.
Why does my library molarity look unexpectedly high?
Usually because the length used is too short. Check for adapter-dimer — a 120 to 130 bp all-adapter species that quantifies and clusters well but yields no usable reads. It shortens the mean fragment length and inflates the calculated molarity at the same time.
Related calculators
References
- Bustin SA, Benes V, Garson JA, et al. The MIQE guidelines: minimum information for publication of quantitative real-time PCR experiments. Clin Chem. 2009;55(4):611–622 — on nucleic acid quantification and reporting.
- Sambrook J, Russell DW. Molecular Cloning: A Laboratory Manual. 3rd ed. Cold Spring Harbor Laboratory Press; 2001 — appendix on nucleic acid mass and molar conversions.
- Head SR, Komori HK, LaMere SA, et al. Library construction for next-generation sequencing: overviews and challenges. BioTechniques. 2014;56(2):61–77.
Medical Disclaimer: The tools and content provided here are for educational and reference purposes only. They are not intended to substitute for professional medical advice, diagnosis, or treatment. Clinical decisions should always be based on the comprehensive assessment of a qualified healthcare professional.