Predict how two restriction enzymes cut linear DNA or a circular plasmid. The tool reports recognition sites, cut positions, fragment sizes, compatible sticky ends, and reaction setup warnings. Use it before diagnostic digestion, insert excision, or vector preparation.
Paste a DNA sequence, select two enzymes, and compare the double digest with both single digests. Basic mode answers the fragment-size question. Advanced mode adds reaction-volume checks for the wet-lab setup.
Start with a realistic plasmid or insert example, then paste your own sequence and enzyme pair.
Paste the full linear sequence or circular plasmid sequence. The tool ignores spaces, numbers, and FASTA headers.
Sequence length
233 bp
Unique cuts
2
EcoRI sites
1
BamHI sites
1
Ends are not directly compatible
EcoRI leaves AATT (5-overhang) and BamHI leaves GATC (5-overhang). Use ligation only after compatible end design, blunting, adapters, or a different enzyme pair.
EcoRI: Produces a 5′ AATT cohesive end.
BamHI: Produces a 5′ GATC cohesive end.
Compare each single digest with the combined digest before you interpret gel lanes.
A double digest cuts one DNA molecule with two restriction endonucleases. Most users want one practical answer: “What band sizes should I see on my gel?” This calculator answers that by scanning the sequence for both recognition sites and measuring the distance between cut positions.
Circular plasmids need different math than linear PCR products. A plasmid has no true first base during digestion, so the final fragment wraps from the last cut site back to the first cut site. Linear DNA has physical ends, so fragments also run from each end to the nearest cut.
Molecular cloning uses this logic every day. A diagnostic digest can confirm an insert, check orientation, or distinguish similar plasmid clones. Addgene describes diagnostic digests as a way to cut plasmids into predicted fragments and analyze those bands by gel electrophoresis. Read Addgene’s diagnostic digest guide.
Each input controls a specific biological assumption. Use the table before you trust a fragment pattern from any digest calculator.
| Tool part | What you enter | Why it matters |
|---|---|---|
| DNA sequence | A plasmid, insert, PCR product, or synthetic fragment | The sequence determines the exact recognition sites and fragment sizes. |
| Template type | Circular plasmid or linear DNA | Topology changes the fragment calculation at the sequence ends. |
| Enzyme pair | Two restriction enzymes such as EcoRI and BamHI | Each enzyme contributes a recognition site, cut offset, and DNA end type. |
| Cut map | No manual input | The visual map shows where cuts fall before you interpret gel bands. |
| Advanced reaction setup | DNA mass, concentration, enzyme volume, buffer stock | The volume check catches negative water and high glycerol before the tube is prepared. |
Use Basic mode when you need cut sites, fragment sizes, and sticky-end compatibility. This fits classroom problems, plasmid map checks, and quick insert-excision planning. It keeps the workflow focused on sequence logic.
Use Advanced mode when you also need DNA volume, buffer volume, water volume, and enzyme-volume warnings. It does not replace the supplier protocol, but it helps you catch pipetting problems early.
The calculator treats the cleaned sequence as a coordinate system. It records each cut position, sorts unique cuts, then calculates distances between neighboring cuts.
| DNA type | How fragments are measured | Common result |
|---|---|---|
| Linear DNA | From the left end to the first cut, between cuts, and from the last cut to the right end. | Two cuts usually create three fragments. |
| Circular plasmid | From each cut to the next cut around the circle, including the wraparound segment. | Two unique cuts usually create two fragments. |
| One-cut plasmid | One cut opens the circle at a single position. | The gel often shows one linear band near the full plasmid size. |
A 5,200 bp circular plasmid has one EcoRI site at 800 bp and one BamHI site at 2,100 bp. The distance from EcoRI to BamHI equals 1,300 bp. The wraparound fragment equals 5,200 − 1,300 = 3,900 bp.
A correct gel should show two main bands near 3.9 kb and 1.3 kb after complete digestion. If the 5.2 kb band remains strong, one or both enzymes may have cut incompletely.
BamHI recognizes GGATCC and BglII recognizes AGATCT. Both leave a 5′ GATC overhang. The fragments can ligate together because the cohesive ends match.
The final ligation junction usually does not regenerate the original BamHI or BglII recognition site. That matters when you plan a scar, a diagnostic digest, or a second cloning step.
Fragment prediction solves only half of the problem. The physical reaction also needs compatible conditions. NEB notes that double-digest planning should account for buffer selection and provides tools for choosing reaction conditions. Check NEB’s double-digest guidance.
| Check | Why users miss it | Practical action |
|---|---|---|
| Buffer compatibility | Different enzymes can lose activity in the wrong buffer. | Use the supplier chart for the exact enzyme versions. |
| Incubation temperature | Most enzymes use 37°C, but not all do. | Avoid combining enzymes with conflicting temperature needs without a protocol. |
| Glycerol level | Large enzyme volumes add glycerol from storage buffer. | Keep enzyme volume modest or increase total reaction volume. |
| Methylation sensitivity | Some sites cut poorly when methylated. | Check Dam, Dcm, or CpG sensitivity when bacterial plasmids fail. |
| Partial digestion | Too little time, wrong buffer, or too much DNA can leave uncut template. | Run single-digest controls and include an uncut lane. |
Map single-enzyme and multi-site digestion patterns before you design a cloning check.
Compare cohesive ends from enzymes such as BamHI, BglII, XbaI, and SpeI.
Calculate insert mass after you choose compatible vector and insert ends.
It tells you where two restriction enzymes cut a DNA sequence and what fragment sizes those cuts create. The result helps you plan a diagnostic digest, confirm a plasmid map, or predict whether two enzymes release an insert. The calculator also reports sticky-end compatibility and a basic reaction-volume check. You should still confirm buffer activity, incubation time, and enzyme units with the supplier protocol before running the experiment.
For circular DNA, each fragment equals the distance from one cut site to the next around the plasmid. One cut linearizes the plasmid and produces one band near the full plasmid size. Two unique cuts usually produce two fragments. More cuts create more bands, although similar-size fragments may merge on an agarose gel.
EcoRI and BamHI cut common multiple-cloning sites and produce different 5′ sticky ends. Different ends can support directional cloning because the insert has a preferred orientation. EcoRI leaves AATT, while BamHI leaves GATC. Those overhangs do not ligate directly to each other, but each ligates to a matching compatible end from the same or a compatible enzyme.
Yes, BamHI and BglII both leave a 5′ GATC overhang. The ends can anneal and ligate, but the final junction often destroys the original recognition sites. That can help cloning because the joined scar may resist recutting. Always check the final sequence if you need to preserve a diagnostic site.
Unexpected bands can come from partial digestion, star activity, wrong plasmid topology, methylation sensitivity, or an incorrect sequence map. Supercoiled, nicked, and linear plasmids also migrate differently before digestion. Run single-digest controls when a double digest fails to match the predicted pattern. The single lanes show whether one enzyme failed or whether the input plasmid differs from the map.
Many restriction enzymes ship in about 50% glycerol. A high enzyme volume can push final glycerol above 5%, which may increase star activity for some enzymes. The Advanced mode estimates glycerol from the total enzyme volume and reaction volume. Increase total reaction volume or use less enzyme when the warning appears.
No. The tool helps you model cut sites, fragments, ends, and pipetting feasibility. Buffer compatibility depends on enzyme version, supplier, buffer formulation, methylation sensitivity, and incubation temperature. Use the supplier double-digest chart or online protocol tool before you prepare the tube.