Why absorbance at 260 nm measures nucleic acid
The purine and pyrimidine rings in DNA and RNA absorb ultraviolet light with a maximum near 260 nm. Protein, by contrast, peaks near 280 nm because of tryptophan and tyrosine. That separation is the whole basis of nucleic acid quantitation by spectrophotometry: read at 260 nm and you are looking almost entirely at bases, and the amount of light absorbed is proportional to how many of them are in the beam.
The proportionality is the Beer–Lambert law, A = εcl: absorbance equals the extinction coefficient times concentration times pathlength. Rather than make you look up ε, the field long ago collapsed it into a single practical number per nucleic acid type. One absorbance unit at 260 nm through a 1 cm path corresponds to 50 µg/mL of double-stranded DNA, 33 µg/mL of single-stranded DNA, or 40 µg/mL of RNA — the conversion factors tabulated in Molecular Cloning and printed on the side of every spectrophotometer manual. Because 1 µg/mL is identical to 1 ng/µL, those same numbers are the ng/µL you want at the bench.
The three factors differ because of the hypochromic effect. Stacked bases in a duplex interact electronically and absorb less per base than the same bases would free in solution. Denature dsDNA and its A260 rises by roughly 30–40%; that is the same physics that makes a melting curve work, and the reason the calculator asks whether your sample is double- or single-stranded before it converts anything.
What this method cannot do is tell nucleic acid apart from anything else that absorbs at 260 nm. Free nucleotides, degraded RNA fragments, and phenol carry-over all read as signal. If you need to know that the mass you measured is intact, undegraded template, pair this with a purity check using the A260/A280 and A260/A230 ratio calculator and, for RNA, a fragment analyser trace.
The formula, term by term
Three multiplications and one division stand between the instrument reading and a usable concentration.
Divide by pathlength. Absorbance scales linearly with how far the beam travels through the sample. A 1 cm cuvette is the reference geometry the conversion factors assume. Microvolume pedestal instruments hold a 1 mm or 0.2 mm column of liquid and would therefore read ten or fifty times lower — so almost all of them rescale internally and display the 1 cm equivalent. Leave the pathlength at 1 unless your instrument explicitly reports raw absorbance for a short path.
Multiply by the dilution factor. The reading describes the cuvette, not the tube on ice. If you put 2 µL of sample into 98 µL of TE, the dilution factor is 50, and the stock is 50 times more concentrated than the reading implies. Work the dilution out with the solution dilution calculator if you built it in more than one step.
Multiply by the conversion factor. 50, 33 or 40 as above. Getting this wrong is a silent error of up to 50% — nothing about the reading itself will look suspicious.
Then convert mass to moles if you need to. A molar concentration requires a molecular weight, and a molecular weight requires a length. The standard approximations are 660 g/mol per base pair of dsDNA and about 330 g/mol per base of DNA (340 for RNA, whose ribose carries an extra oxygen). Multiply by the length in bases or base pairs and you have the MW of the whole molecule. Because 1 ng/µL is 1 mg/L, the molarity in µM works out as 1000 × (ng/µL) ÷ MW.
Worked example: a plasmid miniprep
You elute a 4,500 bp plasmid in 50 µL, dilute 2 µL into 98 µL of elution buffer, and read the dilution in a 1 cm cuvette against a buffer blank. The instrument shows A260 = 0.312.
- Pathlength. 1 cm, so the reading needs no rescaling: 0.312 ÷ 1 = 0.312 AU.
- Dilution factor. 2 µL into a final 100 µL is a 50-fold dilution. The undiluted stock would read 0.312 × 50 = 15.6 AU — well outside any instrument's range, which is exactly why you diluted it.
- Conversion factor. Plasmid DNA is double-stranded, so 50 ng/µL per AU.
- Concentration. 0.312 × 50 × 50 = 780 ng/µL.
- Total yield. 780 ng/µL × 50 µL = 39,000 ng = 39 µg.
- Molecular weight. 4,500 bp × 660 g/mol = 2,970,000 g/mol.
- Molar concentration. 1000 × 780 ÷ 2,970,000 = 0.263 µM, or 263 nM.
- Total picomoles. 0.263 µM × 50 µL = 13.1 pmol, since one µM in one µL is one pmol.
Sanity-check the yield independently: 39 µg from a 5 mL culture is a normal high-copy miniprep. And check the reading sat in range — 0.312 AU is comfortably inside the 0.1 to 1.0 window where the detector is linear, so the number is trustworthy. Had you read the neat eluate instead, the instrument would have been asked for 15.6 AU and would have returned something much lower and entirely fictional.
How to judge the number you get
Start with the absorbance, not the concentration. The single most useful check is whether the 1 cm equivalent reading fell between about 0.1 and 1.0 AU. Below 0.1 the signal competes with blank drift and with any contaminant that absorbs at 260 nm, so a small absolute error becomes a large relative one. Above 1.0 most detectors have run out of linear range and quietly under-report. The calculator flags both cases.
Then ask whether the yield is plausible for the prep. A high-copy plasmid miniprep from 1–5 mL of culture typically lands in the tens of micrograms; a genomic DNA prep from a few million cultured cells lands in the same range; a total RNA prep from 106 mammalian cells commonly yields around 10 µg. These are rules of thumb from routine practice, not specifications — use them to notice when a number is off by an order of magnitude, not to grade a prep.
Finally, remember what the number is not. A260 measures every 260-absorbing species in the cuvette. For a genomic DNA prep contaminated with RNA, the reading counts both, and the DNA concentration you write on the tube is too high. This is the single biggest reason that quantitative applications — library preparation for sequencing, digital PCR, transfection of defined mass — use a fluorescent intercalating dye assay instead, which responds only to double-stranded DNA. Use the spectrophotometer for a fast yield check and for purity ratios; use the dye assay when the number has to be right.
Conversion factors and molecular weights
| Nucleic acid | ng/µL per A260 unit | MW per residue (g/mol) | ng/µL at A260 = 0.25 | Typical A260/A280 when pure |
|---|---|---|---|---|
| Double-stranded DNA | 50 | 660 per bp | 12.50 | ≈ 1.8 |
| Single-stranded DNA | 33 | 330 per base | 8.25 | ≈ 1.8 |
| RNA | 40 | 340 per base | 10.00 | ≈ 2.0 |
| Short oligonucleotide | 33 | 330 per base | 8.25 | ≈ 1.8 |
The per-residue molecular weights are averages over the four bases; a sequence with skewed base composition departs from them by a few percent. The A260/A280 figures are the conventional purity targets, not a measure of quantity.
Blank with the same buffer you eluted into
The blank defines zero. Elution buffers, TE, and especially anything containing EDTA, guanidinium salts or residual phenol absorb in the ultraviolet, and a water blank against a guanidinium-carrying sample can add tenths of an absorbance unit that the calculator will faithfully convert into nucleic acid that does not exist. Blank with the identical buffer, at the identical pH, and re-blank whenever you change buffer. On microvolume instruments, wipe and re-read the blank two or three times: a drifting blank is the commonest cause of a concentration that will not reproduce.
Where A260 quantitation goes wrong
- Using 50 for RNA or for an oligo. The factor is 40 for RNA and 33 for single strands; using 50 overstates them by 25% and 52% respectively.
- Forgetting the dilution factor. A 50-fold dilution ignored is a result fifty times too low, and nothing about the reading looks wrong.
- Reading outside the linear range. Above roughly 1.0 AU the detector saturates; below 0.1 AU the blank dominates.
- Counting RNA as DNA. An RNase-free genomic prep reads its co-purified RNA at 260 nm too. If the ratio matters, digest with RNase A and re-read.
- Trusting A260 on a sample with visible turbidity. Scattering raises absorbance across the whole spectrum. Check A320: it should be near zero, and whatever it reads should be subtracted from A260 before conversion.
- Assuming 330 g/mol per base for a short oligo. Base composition matters most when the molecule is short. For anything under about 30 bases where exact molarity matters, use the sequence-specific extinction coefficient from the synthesis report.
- Quoting molarity without a length. Mass concentration is length-independent; molarity is not. A 100 ng/µL solution of a 200 bp amplicon is 20 times more concentrated in molecules than 100 ng/µL of a 4,000 bp plasmid.
Where this fits among the other quantitation methods
Spectrophotometry is fast, non-destructive of anything but 1–2 µL, needs no standard curve, and reports purity ratios in the same read. That combination is why it survives despite being the least specific of the common methods.
Fluorometric dye assays (PicoGreen, Qubit and equivalents) bind selectively to double-stranded DNA and are two to three orders of magnitude more sensitive. They require a standard curve and consume sample, but they ignore free nucleotides, RNA and protein — so when a spectrophotometer and a dye assay disagree, the dye assay is usually the one to believe for dsDNA.
Agarose gel densitometry against a mass ladder is cruder but tells you something neither instrument does: whether the mass is where you think it is, in one band or smeared across the lane.
qPCR quantifies only amplifiable template, which is the relevant quantity when the DNA is degraded or when only a small fraction of a mixed sample is your target. Convert between mass and molecules with the DNA copy number calculator, and set up the reactions themselves with the PCR master mix calculator.
In routine practice you use them together: the spectrophotometer for a yield and purity check straight off the column, and a dye assay or qPCR immediately before any step where the input mass is written into a protocol.