DNA Concentration Calculator
Estimate double-stranded DNA concentration from absorbance at 260 nm using the standard 50 μg/ml per A260 conversion. The default UV-spectroscopy method used in every molecular-biology lab for quick nucleic-acid quantification.
Last updated: May 2026
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About this calculator
Nucleic acids absorb UV light strongly at 260 nm because of the conjugated π-electron systems in their nitrogenous bases. For double-stranded DNA, the empirically determined relationship is: one absorbance unit at 260 nm (1 cm path length) corresponds to approximately 50 μg/ml of dsDNA. This calculator multiplies the measured A260 by 50 (μg/ml per A260 unit) and by the dilution factor to back-calculate the original undiluted concentration: concentration = A260 × 50 × dilution_factor. Variables: absorbance260 is the measured A260 of your diluted sample; dilutionFactor is how many times the original sample was diluted before measurement (1 if you measured neat). Edge cases: the 50 μg/ml conversion is specific to dsDNA — for single-stranded DNA use 33 μg/ml per A260; for RNA use 40 μg/ml per A260; for short oligonucleotides use the molar extinction coefficient calculated from the specific sequence. A260 readings below ~0.1 are unreliable due to instrument noise; readings above 1.0 may be in the non-linear range of older spectrophotometers (most modern instruments handle up to ~2.0). The A260/A280 ratio measures purity: pure DNA gives ~1.8, RNA gives ~2.0, values below 1.6 indicate protein contamination, and values above 2.0 may indicate RNA contamination. A260/A230 measures other contaminants (phenol, ethanol, salts) and should be > 1.8 for clean samples. Common interferences: residual phenol from extraction (peak at 270 nm), ethanol carryover (peak at 230 nm), guanidine salts (peak at 230 nm), and heme-pigment contamination from blood (peak at 414 nm).
How to use
Example 1 — Standard DNA prep. You diluted your DNA stock 10-fold (1 μl into 9 μl water for a NanoDrop reading) and measured A260 = 0.8. Enter Absorbance at 260nm = 0.8, Dilution Factor = 10. Concentration = 0.8 × 50 × 10 = 400 μg/ml = 400 ng/μl. ✓ This is a typical concentration for a standard DNA prep — well above the working range needed for most downstream applications like PCR (1–10 ng/μl) or restriction digestion (200 ng total). Example 2 — Dilute genomic DNA. After ethanol precipitation you suspect low yield. A260 reads 0.12 on a 1:1 measurement (no dilution). Enter Absorbance = 0.12, Dilution Factor = 1. Concentration = 0.12 × 50 × 1 = 6 μg/ml = 6 ng/μl. ✓ Low but usable for most PCR applications; for cloning or library prep where you need 100+ ng total, you'll need to concentrate (vacuum centrifugation, alcohol precipitation, or column concentration). Note: A260 = 0.12 is approaching the lower limit of instrument reliability — a fluorescence-based method (Qubit, PicoGreen) is more accurate at this concentration.
Frequently asked questions
Why is the conversion factor 50 μg/ml per A260 for dsDNA?
It's an empirical average derived from the fact that dsDNA has a per-base molar extinction coefficient of about 6,600 M⁻¹cm⁻¹ at 260 nm. Combined with the average molecular weight of a base pair (~660 Da), this gives an absorbance of 1.0 per cm path length per 50 μg/ml. The factor varies slightly with sequence composition (GC content affects extinction modestly), buffer, pH, and temperature, but 50 is a good universal approximation for typical genomic or plasmid DNA. The corresponding factors: 33 μg/ml for ssDNA (less base stacking gives higher per-base extinction, so same absorbance comes from less material); 40 μg/ml for RNA; and for synthetic oligos the per-base extinction coefficient should be calculated from the specific sequence (online calculators provide this). Using 50 for the wrong sample type can over- or under-estimate by 20–50%.
What do the A260/A280 and A260/A230 ratios tell me?
A260/A280 is the purity ratio for nucleic-acid samples. Pure dsDNA gives ~1.8, pure ssDNA ~1.7, pure RNA ~2.0. Protein contamination drops the ratio (proteins absorb strongly at 280 nm via tryptophan and tyrosine); a ratio below 1.6 typically indicates significant protein impurity. A ratio above 2.0 in a sample expected to be DNA suggests RNA contamination, which is common in plasmid preps and requires RNase treatment. A260/A230 detects organic-solvent and salt contamination (phenol, ethanol, guanidine, chaotropes all absorb at 230 nm); a clean sample should give A260/A230 > 1.8. A low A260/A230 means impurities will probably interfere with downstream enzymatic steps (PCR, cloning, sequencing) and you should re-precipitate or purify before use.
When is UV absorbance inadequate and you should use a fluorescence-based method instead?
Use fluorescence-based methods (Qubit, PicoGreen, SYBR Safe) when (1) you need quantification of very low concentrations (below ~5 ng/μl, where UV is noise-limited); (2) you need to distinguish dsDNA from ssDNA, RNA, or free nucleotides (UV can't tell them apart, fluorescence dyes are selective); (3) your sample has potentially contaminating absorbing impurities (proteins, organic solvents, salts) that would inflate the UV reading; (4) you need accurate quantification for sensitive downstream applications (NGS library preparation is the classic case — you need precise dsDNA quantification, and Qubit is dramatically better than UV). UV is fine for routine plasmid preps, PCR products, and rapid checks; for anything requiring accuracy or low quantities, the fluorescence-based methods are now standard in well-equipped labs.
What are the most common mistakes people make with UV DNA quantification?
The first is using the wrong conversion factor — 50 is for dsDNA only; ssDNA uses 33 and RNA uses 40, so applying 50 to all samples produces 20–50% errors. The second is ignoring the dilution factor — forgetting to multiply by the dilution back to original concentration is a routine mistake that distorts everything downstream. The third is not checking purity ratios; a sample with A260/A280 = 1.3 is more protein than DNA, and the A260 reading is essentially meaningless for DNA quantification. The fourth is reading samples with A260 outside the linear range (below 0.05 or above 2.0 on older instruments); use a different dilution or a more sensitive method. The fifth is sample contamination by RNA (very common in plasmid preps) inflating the DNA reading by 20–50%; treat with RNase A and re-measure if the prep wasn't clean. Finally, single readings have ~5% noise — duplicate or triplicate measurements give more confidence, especially near the detection limit.
When should I not use this calculator?
Skip it for single-stranded DNA, RNA, or oligonucleotides — use the appropriate per-base conversion factor (33 for ssDNA, 40 for RNA, or sequence-specific extinction for short oligos). Don't use it when sample contains absorbing contaminants (proteins from inadequate purification, phenol from extraction, dyes from gel extraction); the A260 reading is inflated and the DNA concentration overstated. It's the wrong tool for samples below ~5 ng/μl (UV noise dominates); use Qubit, PicoGreen, or similar fluorescence-based methods. Avoid it for next-generation sequencing library quantification where accuracy matters more than convenience; qPCR or fluorescence methods are now standard. Don't use it for distinguishing dsDNA from ssDNA, RNA, or degraded DNA — UV can't tell them apart, only fluorescence dyes can. Finally, don't use it without checking A260/A280 and A260/A230 purity ratios; bad ratios mean the reading isn't valid regardless of the math.