Gibson Assembly Fragment Amount Calculator (Multi-Fragment Molar Ratio and pmol Conversion)
The most common problem in Gibson assembly is not “what ratio should I use” — it’s that when the number of fragments changes, the ratio has to change too, and in the opposite direction from what you might expect.
Two Protocol Tiers
NEB’s published NEBuilder HiFi reaction protocol divides recommendations 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 |
| Homology arm length | 15–20 bp | ≥ 20 bp |
Note the direction of these two columns: as fragment count increases, the insert excess drops from 2-fold to 1-fold, while total fragment amount rises by nearly an order of magnitude. The reason is straightforward — more fragments means fewer correctly assembled combinations and more opportunities for mis-assembly, so keeping all fragments equimolar outperforms having any single one in excess; the higher total amount compensates for the efficiency drop.
The protocol also gives an empirical optimum: 50–100 ng of vector with a 2-fold insert excess gives the best cloning efficiency.
Conversion Formula
pmol = ng × 1000 ÷ (bp × 650)
650 daltons is the average molecular weight per base pair of dsDNA.
You can verify this: 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 those into the formula gives 0.01538 and 0.1538, which match. This calculator uses exactly this equation.
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, so the molar-ratio-based amount is typically insufficient. The protocol explicitly states this, and this calculator will flag such fragments automatically.
Unpurified PCR product must not exceed 20% of the total reaction volume. In a 20 µL reaction that is 4 µL. Buffer components in the PCR mix inhibit assembly — this is unrelated to molar amounts but is frequently the actual cause of failure.
Reaction Setup
Standard 20 µL reaction: 10 µL Master Mix + X µL fragments + water to 20 µL. If the fragment volume does not fit, the protocol calls for scaling up the reaction volume and Master Mix proportionally, not squeezing everything in — if concentration is the bottleneck, concentrate the fragments first; do not reduce vector volume to make room.
When These Values Do Not Apply
- This is the protocol for seamless assembly (which depends on homology arms at fragment ends). Cohesive-end or blunt-end T4 ligase cloning uses a different system with different ratios; see DNA Ligation Calculator.
- For more than 6 fragments, or single fragments larger than tens of kilobases, the table above no longer applies — multi-round assembly is usually required rather than a one-pot reaction.
- Homology arm design (length, Tm, avoiding secondary structure) is outside the scope of this tool; it only calculates amounts.
Related Tools
For cohesive/blunt-end ligation molar ratios, see DNA Ligation Calculator; for converting fragment concentration from OD₂₆₀, see Nucleic Acid Concentration Converter; for copy number and single-copy mass, see DNA Copy Number Calculator.
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). Fragment amounts and reaction conditions from NEB’s published NEBuilder HiFi DNA Assembly reaction protocol.
FAQ
With more fragments, why does the insert excess ratio actually decrease?
With more fragments, there are more possible mismatch combinations. For 2–3 fragments, a 2-fold insert excess suppresses vector self-ligation; for 4–6 fragments, excess of any single fragment increases the chance of head-to-tail circularization or incorrect insertion, so **equimolar** amounts work better. The molar ratio therefore shifts from 1:2 to 1:1 — but total fragment amount rises from 0.03–0.2 pmol to 0.2–0.5 pmol to compensate for the efficiency drop with more fragments. These two changes go in opposite directions, which makes them easy to confuse.
How do you convert between ng and pmol, and how can you verify the result?
pmol = ng × 1000 ÷ (bp × 650), where 650 daltons is the average molecular weight per base pair of dsDNA. To verify, use the two reference examples from the protocol: 50 ng of 5000 bp dsDNA ≈ 0.015 pmol, and 50 ng of 500 bp dsDNA ≈ 0.15 pmol. Plugging into the formula gives 0.01538 and 0.1538, which match.
Why do very short fragments need to be added in larger amounts?
Fragments shorter than 200 bp are lost at a higher rate during the reaction and subsequent cleanup, so the molar-ratio-calculated amount is often insufficient. The protocol recommends using 5× the normal amount for these fragments. This calculator will automatically flag them — this is the easiest rule to overlook, and when missed the result is usually not complete failure but a noticeably low colony count.
Why is there a volume limit on unpurified PCR product?
Components in the PCR buffer — especially residual primers, dNTPs, and polymerase buffer salts — inhibit the assembly reaction. The protocol specifies that the total volume of unpurified fragments must not exceed 20% of the reaction volume, which is 4 µL in a 20 µL reaction. This has nothing to do with molar amounts, yet it is often the actual cause of failure — even perfectly calculated amounts won't give you colonies if this volume limit is exceeded.
Do these amounts apply to T4 ligase cloning as well?
No. These are the recommended amounts for **seamless assembly** (which relies on homology arms at fragment ends). Cohesive-end and blunt-end T4 ligase cloning use a different system with different ratios — there is a separate [DNA Ligation Calculator](../ligation-calc/) for that. Do not use these values interchangeably.
What do you do with 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 multi-round assembly: join several fragments into an intermediate product, purify it, then proceed to the next round, rather than putting everything in one pot. With too many fragments in a single reaction, the fraction of correctly assembled products drops sharply.
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