DNA Ligation Calculator — Vector and Insert Amount (T4 Ligase)
Why convert to molar ratio?
T4 DNA Ligase catalyzes phosphodiester bond formation between adjacent DNA ends, recognizing end count (i.e., moles) rather than mass. Ligation reactions therefore require converting mass (ng) to moles (fmol) before mixing at the appropriate insert : vector molar ratio. Mixing by mass ratio alone causes severe excess of small fragments or severe deficiency of large fragments, reducing cloning efficiency.
Core formula
The empirical average molecular weight per base pair for 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 the vector mass, the required insert mass at 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 those used by the NEB and Addgene official calculators.
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 is far less efficient than sticky-end ligation and typically requires a higher molar ratio, longer reaction time (16 °C overnight), or addition of PEG 4000 to the reaction.
Applicable conditions
- 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 linearized size (i.e., the length after restriction enzyme digestion opens the circular plasmid), not the original plasmid total length.
- This tool is designed for classical T4 ligase reactions; it does not apply to Gibson Assembly, Golden Gate, or LR/BP recombination and other seamless cloning methods, which have their own input rules.
- Typical reaction setup: 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 is also affected by the quality of end repair, the degree of vector dephosphorylation (CIP/SAP treatment reduces ligation efficiency — increase insert molar ratio accordingly), DNA purity, and competent cell efficiency.
Verification assertions (can be checked 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 digestion, vector size = total plasmid length − length of the removed old insert, not the original plasmid size.
- Molar ratio direction reversed: This tool takes the fold-excess of insert over vector — entering 3 means insert : vector = 3 : 1.
- Mixed units: Enter mass in ng and size in bp directly; do not convert to µg or kb before entering.
- Ignoring volume constraints: In a 10 µL reaction, total DNA volume is typically no more than 7 µL, leaving room for ligase and buffer; concentrate DNA if it is too dilute.
After transformation of the ligation product, see the Transformation Efficiency Calculator for how to convert colony counts to efficiency — note that ligation products naturally give far lower efficiency than supercoiled plasmid standards, so the two should not be compared directly.
FAQ
My vector is a 5000 bp plasmid. After digestion I removed a 200 bp old insert and am cloning in a new fragment. 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 bases introduced by restriction sites in the primers are generally negligible).
My vector was CIP-dephosphorylated. Should I increase the insert molar ratio?
Yes. CIP treatment removes the 5′ phosphate from vector ends, preventing self-ligation (which reduces empty-vector background), but also lowers overall ligation efficiency. Raise the insert : vector molar ratio to 5 : 1 — 10 : 1 and extend ligation to 16 °C overnight.
Can I use this tool for Gibson Assembly?
No. Gibson Assembly relies on exonuclease exposure of 15–30 bp homology arms; recommended molar ratios and optimal DNA amounts are specified by the kit manufacturer (e.g., NEB HiFi Assembly), typically 0.02–0.5 pmol per fragment. The underlying logic differs from T4 ligase — consult the relevant product manual.
Transformation efficiency after ligation is very low. Could the molar ratio be the issue?
Molar ratio is only one factor. Also check: ① whether ATP in the ligation buffer has degraded (ATP is easily inactivated after repeated freeze-thaw cycles — aliquot into single-use portions); ② whether the vector is fully linearized; ③ whether DNA ends are properly prepared (for PCR products, use a high-fidelity polymerase then add A-tails with Taq, or perform end-polishing); ④ competent cell efficiency.
The calculator shows I need more insert than I currently have. What are my options?
Options include: ① Reduce vector amount (scale down the entire reaction proportionally); ② Increase insert concentration (concentrate by ethanol precipitation); ③ Lower the molar ratio to 2 : 1 (still acceptable for efficient sticky-end ligations); ④ Scale up total reaction volume to 20 µL while keeping moles constant — then use the full volume for electroporation or 5 µL for heat-shock transformation.
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