DNA Ligation Calculator — Vector and Insert Mass (T4 Ligase)
Why convert to molar ratio?
T4 DNA Ligase catalyzes formation of phosphodiester bonds between adjacent DNA ends, recognizing end count (i.e., moles) rather than mass. Ligation reactions therefore require converting mass (ng) to moles (fmol) and combining components at an appropriate insert : vector molar ratio. Mixing by mass ratio alone causes severe excess of small fragments or severe deficit of large ones, reducing cloning efficiency.
Core formula
The empirical average molecular weight per base pair of double-stranded DNA is 660 Da/bp (the four dNMPs average ~330 Da/nt; double-stranded × 2). The molar amount of any fragment is therefore:
n (fmol) = mass(ng) × 10⁶ / (size(bp) × 660)
Given vector mass, the required insert mass for a target molar ratio r (insert : vector) is:
insert mass (ng) = vector mass (ng) × insert length (bp) × r / vector length (bp)
This formula is consistent with the calculators used by NEB and Addgene.
Recommended molar ratios
| End type | Recommended insert : vector molar ratio |
|---|---|
| Sticky end (4-bp overhang) | 3 : 1 — 5 : 1 |
| Blunt end | 5 : 1 — 10 : 1 |
| TA cloning | 3 : 1 — 10 : 1 |
Blunt-end ligation efficiency is far lower than sticky-end ligation; it typically requires a higher molar ratio, longer reaction times (16 °C overnight), or addition of PEG 4000 to the reaction.
Applicability
- The formula applies only to linear double-stranded DNA; it does not apply to single-stranded DNA, RNA, or circular plasmids.
- Vector size should be the post-linearization size (i.e., the length after restriction enzyme digestion opens the circular plasmid), not the original total plasmid length.
- This tool is designed for classical T4 ligase reactions; it does not apply to Gibson Assembly, Golden Gate, or LR/BP recombination — these seamless cloning methods each have their own loading rules.
- Typical reaction: 10 µL total volume, 1 µL 10× T4 Ligase Buffer, incubate at 16 °C for 1–16 h, or 25 °C for 5–15 min (quick-ligation system).
- This calculator gives theoretical input amounts; actual cloning efficiency also depends on end-repair quality, degree of vector dephosphorylation (CIP/SAP treatment reduces ligation efficiency — increase the insert molar ratio accordingly), DNA purity, and competent cell efficiency.
Validation checks (verify by hand)
- Vector 50 ng, 3000 bp; insert 800 bp; molar ratio 3 → insert = 40.00 ng
- Vector fmol = 50 × 10⁶ / (3000 × 660) ≈ 25.25 fmol; insert fmol ≈ 75.76 fmol
- Vector 100 ng, 5000 bp; insert 500 bp; molar ratio 5 → insert = 50.00 ng
Common mistakes
- Wrong vector size: After restriction digestion, vector size = total plasmid length − replaced insert length, not the original plasmid size.
- Inverted molar ratio: This tool asks for the insert-to-vector fold excess; entering 3 means insert : vector = 3 : 1.
- Mixed units: Enter values directly in ng and bp; do not convert to µg or kb before entering.
- Ignoring volume constraints: In a 10 µL reaction, total DNA volume should generally not exceed 7 µL, leaving room for ligase and buffer; concentrate DNA first if it is too dilute.
FAQ
My plasmid is 5000 bp, and after digestion I removed a 200 bp insert. What size should I enter for the vector?
Enter the linearized backbone size: 5000 − 200 = 4800 bp. For insert size, enter the length of your new PCR-amplified insert (the few extra base pairs contributed by restriction sites in the primers have negligible effect).
The vector was dephosphorylated with CIP (alkaline phosphatase). Should I increase the insert molar ratio?
Yes. CIP treatment removes the 5′ phosphate from vector ends, preventing self-ligation (reducing empty-vector background), but it also reduces ligation efficiency. Increase the insert : vector molar ratio to 5 : 1 — 10 : 1 and extend ligation time to 16 °C overnight.
Can I use this tool for Gibson Assembly?
No. Gibson Assembly relies on exonuclease activity to expose 15–30 bp homology arms; optimal amounts and molar ratios are specified by the kit manufacturer (e.g., NEB HiFi Assembly), typically 0.02–0.5 pmol per fragment. The logic differs from T4 ligase — refer to the relevant product manual.
Transformation efficiency after ligation is very low. Is it a molar ratio problem?
Molar ratio is only one factor. Also check: ① whether ATP in the ligation buffer has degraded (ATP degrades with repeated freeze-thaw cycles; aliquot and store accordingly); ② whether vector linearization is complete; ③ whether DNA ends are flush (for PCR products, use a high-fidelity polymerase and add A-tails with Taq, or perform end-polishing); ④ competent cell efficiency.
The calculated insert amount exceeds what I have. What can I do?
Options: ① reduce the vector amount and scale down the entire reaction proportionally; ② concentrate the insert by ethanol precipitation; ③ lower the molar ratio to 2 : 1 (still acceptable for efficient sticky-end ligations); ④ scale up the total reaction volume to 20 µL, keeping molar amounts constant — use the entire volume for electroporation or 5 µL for chemical transformation.
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