DNA/RNA Molarity Calculator: ng/µL, nM, Copies

Your NanoDrop reads 120 ng/µL for a 500-bp PCR product and the qPCR standard curve protocol asks for a 10 nM stock. Those two numbers live in different universes — mass concentration vs. molar concentration — and the bridge between them is molecular weight. A DNA/RNA molarity calculator takes the mass reading from your spectrophotometer, the sequence length (or explicit MW), and returns nanomolar concentration plus absolute copy number so you can set up standards, normalize ligation inserts, or dilute template to a specific molecule count.

The mistake that wrecks the most standard curves: treating ng/µL as if it were already nM. A 100 ng/µL solution of a 500-bp dsDNA fragment is about 303 nM, but the same mass concentration of a 5-kb plasmid is only about 30 nM. Length matters because longer molecules weigh more per molecule. If you skip the MW conversion, your 10-fold dilution series will be off by whatever factor the assumed and real molarities differ.

Every qPCR standard curve and every digital PCR absolute-quantification experiment needs a known number of template copies. The path from mass to copies runs through Avogadro’s number: Copies = (mass in g × 6.022 × 10²³) / (length × MW per bp). For dsDNA, the average molecular weight per base pair is ~660 Da. For ssDNA it is ~330 Da per nucleotide, and for ssRNA ~340 Da per nucleotide.

At the bench, you rarely need more precision than the ~660 average. GC-rich sequences are slightly heavier than AT-rich ones, but the difference is under 2% and is swamped by pipetting error and NanoDrop measurement noise. Use 660 for dsDNA, 330 for ssDNA, and 340 for RNA unless you are doing something that demands nucleotide-level accuracy (like mass spec of synthetic oligos).

One practical note: copy numbers for qPCR standards are typically expressed per µL after dilution, not per total tube volume. Make sure you track which volume basis you are reporting, because confusing “copies per µL” with “copies per reaction” throws off the standard curve by whatever your template volume is (usually 1–5 µL).

Once you know your stock molarity, the next question is how to dilute it to the working concentration. For a qPCR standard curve you typically want a 10-fold series spanning 10⁷ down to 10² copies per µL. Start by calculating the copy number in your neat stock, then figure out the first dilution that lands at your top standard.

If the stock is 3 × 10¹ copies/µL and your top standard is 10⁷, the first dilution is 1:300. Make that in two steps (1:30 then 1:10) rather than trying to pipette 0.33 µL into 100 µL — sub-microliter volumes from a standard pipette are unreliable. From the 10⁷ stock, serial 1:10 dilutions give you 10⁶, 10⁵, 10⁴, 10³, 10².

For ligation reactions, the target is usually a molar ratio (e.g., 3:1 insert:vector). Convert both insert and vector from ng/µL to nM, then calculate the volume of each that gives the desired molar ratio at your total reaction mass. The calculator handles this conversion so you do not have to estimate MW by hand each time.

The average MW per nucleotide differs by strand type because of chemistry. dsDNA has two strands, so 1 bp = ~660 Da (two nucleotides of ~330 Da each). ssDNA (primers, synthetic oligos, M13 phage genomes) uses ~330 Da per nucleotide. ssRNA uses ~340 Da per nucleotide because the ribose sugar is one oxygen heavier than deoxyribose.

Picking the wrong strand type is a 2-fold error for ds vs. ss and a smaller but real error for DNA vs. RNA. If you enter a 20-nt primer as dsDNA, the calculator doubles the MW and halves the molarity — your “10 µM stock” is actually 20 µM, and every downstream dilution is 2x off.

dsRNA (like siRNA duplexes) uses ~660 Da per bp, same as dsDNA. The extra oxygen per ribose adds only ~1% to the total MW of a 21-bp duplex, which is negligible. For short synthetic RNA duplexes, the MW printed on the synthesis report is more accurate than any per-nucleotide estimate — use the manufacturer’s value when available.

My qPCR standard curve is shifted by exactly 2-fold. Where did I go wrong?
You probably entered a ssDNA amplicon length as dsDNA, or vice versa. The MW doubles when you switch from ss to ds, cutting the calculated molarity in half. Check the strand type dropdown and make sure it matches your actual template.

The NanoDrop reading is 200 ng/µL but the Qubit says 85 ng/µL. Which do I use?
Qubit is fluorescence-based and specific to dsDNA (or RNA, depending on the kit). NanoDrop measures A₂₆₀ absorbance, which picks up free nucleotides, ssDNA, RNA, and phenol contamination. For molarity calculations, use the Qubit value because it reflects intact target molecules. If you only have NanoDrop, check the 260/280 and 260/230 ratios to assess contamination before trusting the number.

I converted ng to copies but the number seems impossibly large.
It probably is not wrong — 1 ng of a 500-bp dsDNA fragment contains about 1.8 × 10⁹ copies. Molecular biology works with enormous molecule counts. The issue usually surfaces when people forget to dilute to a working range: 10⁹ copies/µL is far above any qPCR standard curve.

Can I use this calculator for circular plasmids?
Yes. A circular plasmid has the same MW per bp as linear dsDNA (~660 Da/bp). Topology does not change mass. Enter the total plasmid size (backbone + insert) as the length.

Three equations cover the full conversion chain from spectrophotometer to pipette:

Units note: the 10⁶ in the nM formula converts ng to g and µL to L simultaneously. The 10⁹ in the copies formula converts ng to g (10⁻⁹) and cancels with Avogadro’s number. If your stock is in µg/mL instead of ng/µL, note that 1 µg/mL = 1 ng/µL — same number, different label.

Scenario: You gel-extracted a 500-bp PCR amplicon and the NanoDrop reads 100 ng/µL (260/280 = 1.85, clean prep). You need the molar concentration in nM and the copy number per µL to set up a qPCR standard curve.

Step 1 — Molecular weight.
MW = 500 bp × 660 Da/bp = 330,000 g/mol = 3.3 × 10⁵ g/mol.

Step 2 — Molar concentration.
nM = (100 × 10⁶) / 330,000 = 100,000,000 / 330,000 = 303 nM.

Step 3 — Copy number.
Copies/µL = 303 × 6.022 × 10¹⁴ = 1.82 × 10¹⁷ copies/µL.
Alternatively: (100 × 10⁻⁹ × 6.022 × 10²³) / (3.3 × 10⁵) = 1.82 × 10¹⁷. Same answer.

Step 4 — Dilute to top standard.
Target: 10⁷ copies/µL. Dilution factor = 1.82 × 10¹⁷ / 10⁷ = 1,820-fold.
Two-step dilution: 1:100 (1 µL into 99 µL TE), then 1:18.2 (5 µL into 86 µL TE). That gives 10⁷ copies/µL. Serial 1:10 dilutions from there build the rest of the curve.