Gibson Assembly Fragment Amount Calculator (Multi-Fragment Molar Ratio and pmol Conversion)
The most common problem with Gibson assembly is not “what ratio should I use” but that when the number of fragments changes, the ratio has to change too — and the direction is counterintuitive.
Two Recommended Protocols
NEB’s publicly available NEBuilder HiFi reaction protocol divides this into two tiers:
| 2–3 fragments | 4–6 fragments | |
|---|---|---|
| Vector : insert (molar ratio) | 1 : 2 | 1 : 1 |
| Total fragment amount | 0.03–0.2 pmol | 0.2–0.5 pmol |
| Reaction conditions | 50 °C, 15 min | 50 °C, 60 min |
| Overlap length | 15–20 bp | ≥ 20 bp |
Note the direction of these two columns: with more fragments, the molar excess of insert actually drops from 2× to 1×, while the total fragment amount increases by nearly an order of magnitude. The reasoning is straightforward — more fragments means fewer correct ligation combinations and more chances for misassembly. In that case, equimolar amounts work better than having any one fragment in excess; the higher total amount compensates for the drop in efficiency as fragment count increases.
The protocol also gives an empirical guideline: 50–100 ng of vector with a 2-fold molar excess of insert gives the best cloning efficiency.
Conversion Formula
pmol = ng × 1000 ÷ (bp × 650)
650 daltons is the average molecular weight per base pair of double-stranded DNA.
Verification: the protocol itself provides two reference values — 50 ng of a 5000 bp dsDNA is approximately 0.015 pmol, and 50 ng of a 500 bp dsDNA is approximately 0.15 pmol. Plugging into the formula, this tool gives 0.0154 and 0.154 — consistent with the protocol values.
Two Rules That Are Easy to Miss
Fragments shorter than 200 bp need 5× the calculated amount. Short fragments are lost at a higher rate during the reaction and cleanup; the molar-ratio-based calculation typically underestimates how much you need. The protocol explicitly states this, and this tool flags it automatically.
Unpurified PCR products must not exceed 20% of the reaction volume. In a 20 µL reaction, that is 4 µL. Components in the PCR buffer inhibit assembly — this is unrelated to fragment amount, yet is often the real cause of failure.
Reaction Setup
Standard 20 µL reaction: 10 µL Master Mix + X µL fragments + water to 20 µL. When there are too many fragments to fit, the protocol says to scale up the reaction volume and increase Master Mix proportionally — not to force everything in. If concentration is limiting, concentrate the fragments first; do not compensate by reducing vector volume.
When These Numbers Do Not Apply
- These amounts are for seamless assembly (which depends on homology overlaps at fragment ends). Sticky-end or blunt-end T4 ligase cloning uses a completely different set of parameters — see DNA Ligation Molar Ratio.
- If you have more than 6 fragments, or a single fragment exceeding tens of kilobases, the table above no longer applies. Sequential assembly rounds are typically used rather than a single-pot reaction.
- Overlap design (length, Tm, avoiding secondary structure) is outside the scope of this tool; it only calculates amounts.
Related Tools
For sticky-end/blunt-end ligation molar ratios, see DNA Ligation Calculator; for converting OD₂₆₀ to fragment concentration, see Nucleic Acid Concentration Converter; for copy number and single-copy mass, see DNA Copy Number.
Original method: Gibson DG, Young L, Chuang RY, Venter JC, Hutchison CA 3rd, Smith HO. Enzymatic assembly of DNA molecules up to several hundred kilobases. Nat Methods 2009;6(5):343–345 (PMID 19363495). Amounts and reaction conditions from the publicly available NEB NEBuilder HiFi DNA Assembly reaction protocol.
FAQ
Why does increasing the number of fragments actually reduce the molar excess of insert?
Because more fragments means more possible misassembly combinations. With 2–3 fragments, a 2-fold excess of insert suppresses vector self-ligation; with 4–6 fragments, excess of any single fragment increases the chance of that fragment ligating end-to-end or inserting at the wrong site. **Equimolar** amounts work better in that case. So the molar ratio changes from 1:2 to 1:1 — but the total amount increases from 0.03–0.2 pmol to 0.2–0.5 pmol to compensate for the efficiency loss as fragment count grows. The two effects go in opposite directions, which makes this easy to confuse.
How do I convert between ng and pmol, and how can I verify the result?
pmol = ng × 1000 ÷ (bp × 650), where 650 daltons is the average molecular weight per base pair of double-stranded DNA. Verify against the protocol's own reference values: 50 ng of a 5000 bp dsDNA is approximately 0.015 pmol, and 50 ng of a 500 bp dsDNA is approximately 0.15 pmol. Plugging into the formula gives 0.01538 and 0.1538, which match.
Why do short fragments need extra amount?
Fragments shorter than 200 bp are lost at a higher rate during the reaction and subsequent cleanup; the molar-ratio-based calculation typically does not give you enough. The protocol's empirical guideline is 5× the calculated amount for these fragments. This tool flags them automatically — this is the most commonly missed rule, and the consequence is usually not complete failure but noticeably fewer colonies.
Why is there a volume limit on unpurified PCR products?
Components in the PCR buffer — particularly residual primers, dNTPs, and polymerase buffer salts — inhibit the assembly reaction. The protocol specifies that unpurified fragments must not exceed 20% of the total reaction volume, which is 4 µL in a 20 µL reaction. This has nothing to do with molar amounts, yet it is often the real cause of failure — even if the amounts are calculated correctly, exceeding this volume limit will still prevent colony formation.
Do these numbers apply to T4 ligase cloning as well?
No. These amounts are for **seamless assembly** (which relies on homology overlaps at fragment ends). Sticky-end and blunt-end T4 ligase cloning use a completely different system and different ratios. There is a separate [DNA Ligation Calculator](../ligation-calc/) on this site. Do not apply one protocol to the other.
What if I have more than 6 fragments?
The protocol's two tiers only cover up to 6 fragments; beyond that, you are outside the recommended range. The standard approach is sequential assembly: assemble a few fragments into an intermediate product, purify it, then proceed to the next round — rather than combining everything in a single pot. With too many fragments in one reaction, the fraction of correct assemblies drops rapidly.
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