Ligation Calculator - Insert:Vector Molar Ratio to Microlitres
Turn a 3:1 insert-to-vector molar ratio into the microlitres you actually pipette, for one insert or a whole multi-fragment assembly.
🌐 Nothing you paste is logged or stored — every tool is also callable via REST & MCP, and in bulk from the batch tools
Give each part its length and stock concentration and get the volumes to pipette to hit the molar ratio you want. The reason this needs a calculator at all is that a molar ratio counts molecules while a bench works in nanograms per microlitre, and the two parts have different lengths - so 3 molar equivalents of a 900 bp insert against a 3 kb backbone is about 0.9 times the backbone's mass, not three times it. Doing that conversion by eye is the classic way to end up an order of magnitude out on a ligation, and it fails silently: the reaction just gives you background. Multi-fragment mode gives every fragment its own equivalents, which is what Golden Gate and multi-part Gibson need. The plan also checks the two things that go wrong at the bench rather than on paper - a volume below what a single-channel pipette measures reliably, and a DNA volume that leaves no room for 10x buffer and enzyme - and tells you how far to dilute rather than just flagging it.
The backbone mass is the quantity you fix. Every insert amount below follows from it and the molar ratio — 50–100 ng of vector is the usual range for a 20 µL reaction.
3 : 1 is the usual starting point for a sticky-end ligation; 1–2 : 1 is more common for Gibson and Golden Gate, where excess fragment promotes concatemers.
How to use the Ligation Calculator tool
- 1Enter the backbone's length, stock concentration, and how much of it to put in the reaction. 50-100 ng is the usual range for a 20 uL ligation; this is the quantity you fix and everything else follows from it.
- 2Enter the insert's length and stock, and the molar ratio you want. 3:1 is the usual starting point for a sticky-end ligation; 1-2:1 is more common for Gibson and Golden Gate, where excess fragment promotes concatemers.
- 3For a multi-fragment assembly, switch to multi-fragment mode and give each piece its own length, stock and molar equivalents. Equimolar is the standard for Golden Gate.
- 4Pipette the volumes in the table, then make up to the reaction volume with the water, buffer and ligase shown on the last row. Copy the table if you are setting several up.
Frequently asked questions
Why is 3:1 not three times the mass of insert?
Because a molar ratio counts molecules, and a shorter fragment has fewer nanograms per molecule. Three molar equivalents of a 900 bp insert against a 3000 bp backbone works out to 3 x (900/3000) = 0.9 times the backbone's mass. Use three times the mass instead and you have put in roughly ten molar equivalents, which for a sticky-end ligation mostly produces multiple-insert clones and for Gibson or Golden Gate produces concatemers. This conversion is the whole reason the tool exists.
What ratio should I use?
3:1 insert to vector is the standard starting point for a sticky-end or blunt ligation, and 1:1 to 2:1 is more usual for Gibson, NEBuilder and Golden Gate, where an excess of fragment drives concatemer formation rather than helping. For a multi-fragment one-pot assembly, equimolar across all fragments is the normal choice. These are starting points, not optima: if you are getting empty vector, more insert helps; if you are getting multi-insert clones, less does.
How much vector should I put in?
50-100 ng in a 20 uL reaction covers most cases. Much less and the reaction becomes sensitive to pipetting error and to how good your concentration measurement is; much more and you are wasting prep and pushing the DNA volume into the space the buffer needs. The tool takes the vector mass as the quantity you fix because that is how the decision is actually made at a bench.
It says a volume is too small to pipette. What now?
Dilute that stock and use the same plan - the tool tells you roughly how far to dilute. Below about 0.5 uL a single-channel pipette is not reliable, and a 0.2 uL error on a 0.3 uL aliquot is a 60% error in the ratio, which is larger than the difference between the ratios you are choosing between. Diluting is nearly free; guessing at a sub-microlitre volume quietly invalidates the calculation.
Why does the plan warn about room for buffer and enzyme?
Because ligase buffer is normally supplied at 10x, so a 20 uL reaction needs 2 uL of buffer plus the enzyme before any DNA goes in. If the DNA alone fills 18 uL, the plan does not physically work and the usual fix is a more concentrated stock or a larger reaction rather than shaving the buffer. The gate on the results checks this explicitly rather than leaving you to notice it while pipetting.
Do the pmol figures agree with the DNA molarity tool?
Yes, exactly - both use the same molar-mass function, and a test asserts they agree for the same input. That is worth stating because two tools on one site disagreeing about how many pmol are in 50 ng of a 3 kb plasmid is the kind of defect nobody notices until it has already cost someone a cloning round. Supply a sequence for the backbone and the molar mass is computed from its actual base composition instead of the 650 g/mol per base pair average.
What does this not tell me?
Whether the ligation works. This is stoichiometry - it says nothing about whether your ends are compatible, whether the vector was dephosphorylated, whether salt or enzyme carried over from a gel extraction, or whether the insert is intact. It also cannot check your stock concentrations: every volume here is only as good as the Nanodrop or Qubit reading behind it, and a 20% concentration error moves the ratio by 20% no matter how exact the arithmetic is.
Is my data stored, and can I run this from code?
The inputs are lengths and concentrations, and nothing is persisted. ligation_setup is also callable from the REST API and the MCP server, which is how you would plan a plate of assemblies at once, and it returns pmol, ng and volumes per part so a downstream script or liquid handler can consume the plan directly.
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