Convert between plasmid DNA mass (ng) and copy number. Calculate molar equivalents for cloning, transfection, and qPCR standard preparation.
For Research Use Only. This calculator uses the average molecular weight of 660 g/mol per base pair for dsDNA. Actual values may vary based on sequence composition and DNA form (supercoiled, nicked, linear).
Enter parameters and click Calculate to see results
You miniprepped a 6-kb expression plasmid and the NanoDrop reads 250 ng/µL. The transfection protocol calls for 500 ng total, but the cloning protocol next door wants 50 fmol of vector for the ligation. Same tube of DNA, two completely different units. A plasmid copy number estimator converts between mass (ng), moles (fmol, pmol), and absolute copy number for any plasmid size so you can switch between protocols without pulling out a calculator app.
The mistake that derails most ligations: treating all plasmids as if they weigh the same per molecule. 50 ng of a 3-kb plasmid contains twice as many molecules as 50 ng of a 6-kb plasmid because each molecule weighs half as much. If you set up a 3:1 insert-to-vector ligation by mass alone, the actual molar ratio depends entirely on the size of each fragment. You need fmol, not ng, to get the ratio right.
The molecular weight of a plasmid is the total length (backbone + insert) multiplied by 660 Da per base pair. A pUC19 backbone is ~2.7 kb. Drop in a 3.3-kb insert and the total plasmid is 6 kb, with MW = 6,000 × 660 = 3.96 × 10⁶ g/mol. If you use only the backbone size and ignore the insert, the calculated MW is 40% too low and every molar quantity you derive from it is 40% too high.
For empty vectors (no insert), use the published backbone length from the vendor’s map. For constructs with inserts, add the insert length. If you cloned by restriction digest and the vector lost a stuffer fragment, subtract the stuffer and add the insert. The calculator needs the total circular molecule size as it exists in the tube — not the backbone catalog size.
Supercoiled, relaxed, and linear forms of the same plasmid all have the same mass per molecule. Topology changes shape, not molecular weight. However, supercoiled DNA migrates differently on gels and absorbs UV slightly differently on a NanoDrop, so quantification can vary by ~10% depending on form. For most applications, this is within pipetting error.
Co-transfection experiments often require two or three plasmids mixed at defined molar ratios — for example, 1:1 transfer vector to packaging plasmid, or 4:2:1 for a three-plasmid lentiviral system. Mixing by equal mass gives unequal molar ratios whenever the plasmids differ in size, which they almost always do.
The workflow: convert each plasmid from ng/µL to fmol/µL using its specific size, pick the target fmol for each component based on the molar ratio, then calculate the volume of each stock to pipette. If plasmid A is 8 kb at 200 ng/µL (38 fmol/µL) and plasmid B is 4 kb at 150 ng/µL (57 fmol/µL), a 1:1 molar mix is not equal volumes — you need more volume of A (lower fmol/µL) to match the moles of B.
For ligation reactions, the same logic applies but the ratio is typically 3:1 or 5:1 insert to vector (molar). Calculate fmol of vector first, multiply by the desired ratio to get fmol of insert, then convert back to ng using the insert’s MW to find the mass to add.
Linearized plasmid DNA is the gold standard template for absolute-quantification qPCR. You digest the plasmid at a single site outside your amplicon, purify, quantify, and then dilute to a series of known copy numbers. The calculator tells you how many copies are in your stock so you can build the dilution series.
A typical standard curve spans 10⁷ to 10² copies per reaction in 10-fold steps (six points). Make each dilution fresh in low-bind tubes with TE + 0.1% Tween-20 or carrier DNA (like yeast tRNA at 10 ng/µL) to prevent template adsorption to plastic at low concentrations. Below 10³ copies, adsorption losses can eat 50% or more of your template if you dilute in plain water.
One pitfall: using supercoiled plasmid instead of linearized. Supercoiled DNA amplifies less efficiently than linear DNA in the first few PCR cycles because the helicase activity of Taq has to fight the superhelical tension. The result is a standard curve with a slightly different slope than the unknowns (which are linear genomic fragments), introducing systematic error. Always linearize.
My ligation always fails even though I use the “right” insert-to-vector ratio. What is going on?
Check whether you set the ratio by mass or by moles. A 3:1 ratio by mass for a 1-kb insert into a 5-kb vector gives only a 0.6:1 molar ratio — less insert than vector. Convert both to fmol and recalculate. You probably need 5x more insert by mass than you think.
I used the backbone size from the vendor map but my insert is already cloned in. Does that matter?
Yes. If the vendor lists pUC19 as 2,686 bp and your construct is pUC19 + 3,000-bp insert, the total is 5,686 bp (minus any stuffer fragment removed during cloning). Using 2,686 instead of 5,686 overestimates your fmol by ~2.1-fold.
Can I use this for BACs or fosmids?
The formula works for any dsDNA regardless of size. A 150-kb BAC at 660 Da/bp has MW = 9.9 × 10⁷ g/mol. The numbers get large, but the math is the same. Just be aware that BAC yields from minipreps are much lower than standard plasmids, so your stock concentration may only be 10–50 ng/µL.
Why does my copy number disagree with the qPCR result?
The calculated copy number assumes 100% of the measured DNA is intact, amplifiable template. Nicked, degraded, or denatured plasmid contributes to A₂₆₀ absorbance but may not amplify. Run the plasmid on a gel — if you see a degradation smear, the effective copy number is lower than the calculated value.
Four equations handle every plasmid stoichiometry question:
Units note: 1 fmol = 6.022 × 10⁸ molecules. The 10⁶ in the fmol formula converts ng to µg then to pmol then to fmol in one step (ng / g/mol × 10⁶ = fmol). If your mass is in µg, multiply by 1,000 to convert to ng first.
Scenario: You have a 6-kb expression plasmid (4-kb backbone + 2-kb insert) at 250 ng/µL from a miniprep. You need to know how many copies are in 50 ng for transfection math and how many fmol that represents for a ligation side-experiment.
Step 1 — Molecular weight.
MW = 6,000 bp × 660 Da/bp = 3,960,000 g/mol = 3.96 × 10⁶ g/mol.
Step 2 — fmol in 50 ng.
fmol = (50 × 10⁶) / 3,960,000 = 50,000,000 / 3,960,000 = 12.6 fmol.
Step 3 — Copy number.
Copies = 12.6 × 10⁻¹⁵ × 6.022 × 10²³ = 7.6 × 10⁹ copies.
Or directly: (50 × 6.022 × 10¹⁴) / 3.96 × 10⁶ = 7.6 × 10⁹. Same answer.
Step 4 — Volume to pipette.
50 ng from a 250 ng/µL stock = 50 / 250 = 0.2 µL. That is too small to pipette reliably. Dilute the stock 1:10 first (25 ng/µL), then pipette 2 µL for 50 ng.
Step 5 — Ligation check.
If your insert is a 1-kb fragment at 30 ng/µL, its fmol/µL = (30 × 10⁶) / (1,000 × 660) = 45.5 fmol/µL. For a 3:1 insert:vector molar ratio with 12.6 fmol vector, you need 37.9 fmol insert = 37.9 / 45.5 = 0.83 µL of insert stock.
NEB — Ligation Calculator: Insert-to-vector ratio calculator with molar and mass inputs.
Addgene — Plasmid Quantification: Guide to measuring plasmid concentration and assessing quality.
Thermo Fisher — DNA/RNA Molecular Weights: Reference for MW per bp and per nucleotide conversions.
IDT — Nanograms to Copy Number: Practical walkthrough of the mass-to-copies conversion for molecular cloning.
This calculator converts between plasmid DNA mass (in nanograms) and copy number (number of molecules). Given the plasmid size in base pairs, it uses Avogadro's number (6.022 × 10²³ molecules/mol) and the average molecular weight of double-stranded DNA (660 g/mol per bp) to compute the conversion. It also provides molar equivalents in mol, fmol, and pmol. The calculations are: Molecular Weight = Size (bp) × 660 g/mol, Moles = Mass (g) / MW, Copies = Moles × Avogadro's Number. Understanding these calculations helps you convert between mass and copy number for ligation reactions, qPCR standards, and transfection optimization.
Double-stranded DNA consists of two complementary strands. Each nucleotide averages about 330 Da (Daltons), so a base pair (one nucleotide on each strand) totals approximately 660 Da. This is a well-accepted average for dsDNA calculations. The exact value varies slightly based on GC content (A, T, G, and C have slightly different molecular weights), but 660 g/mol per bp is accurate enough for most applications. Understanding this constant helps you see why molecular weight calculations are straightforward and why this value is used consistently in molecular biology.
The calculation is accurate for pure, intact plasmid DNA of known size. However, real-world accuracy depends on: (1) accurate concentration measurement (NanoDrop readings can vary with purity, Qubit is often more accurate for dsDNA), (2) correct plasmid size (verify from sequence or restriction map), and (3) DNA quality (degraded DNA or contaminants give less accurate results). The calculation assumes pure double-stranded DNA—contaminants, different plasmid forms (supercoiled vs. linear), or degraded DNA can affect accuracy. Understanding these factors helps you interpret calculated copy numbers correctly and know when to verify with additional methods.
Copy number (molecules) is useful for qPCR standard curves, viral particle calculations, and thinking about absolute quantities. Molar units (fmol, pmol) are more practical for enzymatic reactions like ligations and Gibson Assembly, where you mix fragments in specific molar ratios (e.g., 3:1 insert:vector). For example, in ligation reactions, you typically use 3-5× more insert than vector (molar), not mass. Understanding when to use each unit helps you work with the appropriate format for your application and achieve proper stoichiometry in enzymatic reactions.
First, calculate fmol or pmol for both your insert and vector using this tool. Then, to achieve a 3:1 molar ratio, use 3× as many pmol of insert as vector. For example, if you have 50 fmol of vector, use 150 fmol of insert. Convert back to ng if needed for pipetting. The key is using molar ratios (not mass ratios) because equal masses of different-sized fragments contain different numbers of molecules. Understanding this helps you achieve optimal ligation efficiency and avoid incorrect ratios that reduce cloning success.
The molecular weight calculation is the same regardless of DNA form (supercoiled, nicked, or linear all have the same molecular weight). However, supercoiled, nicked, and linear DNA can behave differently in applications like transfection or ligation. Supercoiled DNA often gives higher transfection efficiency, while linear DNA is preferred for in vitro transcription. For copy number calculations, form doesn't matter—the calculation assumes the same molecular weight. Understanding this helps you see that copy number calculations are independent of form, but form affects downstream applications.
Yes! The same formula applies to any double-stranded DNA. Just enter the fragment length in base pairs. For PCR products, use the expected amplicon size. The calculation assumes pure dsDNA. For example, a 1 kb PCR product uses the same calculation as a 1 kb plasmid fragment. Understanding this helps you use the calculator for any dsDNA fragment, not just plasmids, making it useful for cloning, qPCR, and other molecular biology applications.
This calculator is designed for double-stranded DNA (660 g/mol per bp). For single-stranded DNA, use ~330 g/mol per nucleotide. For RNA, use ~340 g/mol per nucleotide. These require a different calculator or manual adjustment. The key difference is that single-stranded molecules have half the molecular weight per base/nucleotide compared to double-stranded DNA. Understanding this helps you know when this calculator is appropriate and when you need different constants for single-stranded molecules.
NanoDrop (A260) measures all nucleic acids and can be affected by contaminants that absorb at 260 nm (RNA, proteins, salts). Qubit uses fluorescent dyes that bind specifically to dsDNA, giving more accurate measurements for pure dsDNA quantification. For plasmid calculations, Qubit values are often more reliable because they're specific to dsDNA and less affected by contaminants. Understanding this helps you choose the appropriate measurement method and interpret concentration values correctly for accurate copy number calculations.
qPCR standard curves often span 10¹ to 10⁸ copies per reaction. A common approach is to prepare a stock at 10¹⁰ copies/µL and make 10-fold serial dilutions. The specific range depends on your expected sample concentrations. For example, if your samples are expected to have 10⁵-10⁷ copies, prepare standards from 10¹ to 10⁸ copies to cover the range. Understanding typical copy number ranges helps you design qPCR standard curves that accurately quantify your samples.
From copy number to molar ratios, get accurate calculations for your cloning, transfection, and qPCR experiments
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