Answers on plasmid editing, PCR primer design and specificity, colony PCR, Sanger sequencing, restriction digestion, codon optimization, molarity, buffers and antibody dilution.
Open GenBank, ApE, EMBL, SnapGene .dna, FASTA, plain DNA or a saved editor project. Sequence files can be up to 25 MiB, projects up to 80 MiB, and individual records up to 5 million bases. Multiple records remain separate. FASTA and plain DNA have no feature table, and the editor does not automatically identify missing annotations.
Yes. Choose Edit sequence, enter the position or inclusive base range, and select Review edit. The review shows the exact coordinates, bases added or removed, resulting length, and annotations that will disappear or change. Confirmation applies the edit; Cancel keeps the record. Manual edits use a contiguous range in the current coordinate system. For a range crossing a circular origin, choose a suitable new origin first or make two edits.
Right-click an annotation and choose Delete annotation to remove its label and feature entry while keeping the DNA. Delete annotated DNA removes its exact base segments and shows the effects on other annotations before confirmation. DNA between the segments of a joined feature is kept. Both applied actions support Undo and Redo.
Map rotation changes the display only. Record settings can change a circular sequence’s base-1 position, which rotates the DNA and remaps annotations in one Undo step. Supported structured coordinate qualifiers are remapped; free-text references are unchanged. Unsupported or uncertain locations can prevent an origin change. Save a topology change before choosing a new circular origin.
Use Save project to reopen the records and continue editing. It retains annotations, display settings and original import/edit metadata, but not the Undo history for the open session. Click Export, choose Format, then Download for GenBank, SnapGene .dna, FASTA, SVG, PNG or JPEG. The formats preserve different information: see the Help guide's format limits. Original SnapGene primers, alignments, history and other uneditable data stay in the project and are not included in a new .dna export.
Zoom enlarges the complete map. Scroll horizontally or vertically within it to inspect a region, and use Reset view to return to the beginning at 100%. These display controls do not change DNA, feature coordinates or the record’s topology.
Open Export, choose PNG, JPEG or SVG under Format, then click Download. PNG and JPEG save a raster image; SVG saves a scalable vector map. Each includes the current filters, rotation and selection. Scrolling and zoom do not crop the exported figure. Explicit font settings help preserve labels during import, while font substitution and label editing depend on the receiving application. Desktop PowerPoint supports inserting and resizing SVG. Inspect the saved figure at its intended size and check your journal’s accepted formats. Dense maps retain the editor’s display limits, so select important annotations before export when needed.
The editor draws up to 1,000 features and bounds the displayed labels so dense maps remain usable. Selecting a feature can add its label. The feature table and sequence view use bounded batches, while the record retains the supplied annotations. Map exports retain the current map appearance rather than guaranteeing a label for every feature.
Sequence processing runs in your browser without uploading the sequence. Local recovery starts off and stores applied edits only when enabled. Up to ten workspace copies can be kept, with an 80 MiB limit per workspace. Browser cleanup or storage restrictions can remove access to them. Download a project for a lasting backup; turning recovery off deletes editor recovery copies in that browser.
Copy the required DNA or export FASTA, then supply it to the receiving tool. Confirm its topology and range because edits and origin changes can alter coordinates. The editor has no direct send buttons and does not generate primers. Other tools have their own input limits and may omit annotations. Checker and Primer Scan accept ten primer pairs per checking batch; supply the appropriate reference and run the receiving calculation.
Copying a feature joins its segments in biological 5′ to 3′ orientation, including reverse-strand features. Copy reverse complement reverses and complements the chosen DNA. Selection copying follows the selected range; check which feature or bases are selected before using the sequence in another tool.
Select the bases you want to annotate, right-click and choose Add feature / note. You can also use + Add feature and enter the location yourself. Enter the name, type, location and any notes, then click Save feature.
To edit an existing annotation, select it on the map or in the Features table, change its fields and click Save feature. Locations use 1-based, inclusive coordinates. Saving an annotation changes its description or location; it does not rewrite the DNA sequence.
Select annotations with the checkboxes in the Features table, then open Edit multiple features. Choose a colour or map symbol and apply it to the selected annotations. Locations, strands and properties you do not choose remain as they were.
Select all matching includes matching annotations beyond the currently visible table rows. An annotation you selected remains in the batch even if a later search hides its row, so review the selection count before applying changes. Undo reverses the whole batch.
If browser recovery was enabled and a copy is available, choose it under Saved workspaces and click Restore selected copy. Check the restored records before continuing. Recovery restores saved workspace content, but not the previous session’s Undo history or unsaved form entries.
Delete selected copy removes only the chosen recovery copy. Turning recovery off deletes all of the Editor’s browser recovery copies. Use Save project for a downloaded backup before deleting a copy you may need. Browser recovery is off by default and depends on browser storage.
The .dna writer cannot safely represent uncertain, remote, between-base, ordered or mixed-strand feature locations, so it disables that export for affected records. Use GenBank or Save project to retain those annotations, or resolve their locations before exporting .dna.
A new .dna export includes the current DNA, name, topology and supported feature annotations. Original SnapGene primer objects, alignments, history and other imported data remain only in the saved project. Other applications may draw annotation symbols differently.
Often, yes. The designer groups nearby changes when the primer-length limit and unchanged 3′ binding region permit it. Overlap PCR can combine edits in one linear product. Inverse PCR may require sequential rounds for distant sites. All positions refer to the original uploaded DNA.
Choose Overlap PCR for an edited linear product. Choose Overlap assembly to join separately amplified pieces into a circular construct. Choose Inverse PCR for a circular template opened by a back-to-back primer pair, with end joining afterward. Supply the complete plasmid for either circular method and follow the reaction plan returned for that method.
Inverse PCR groups edits only when the primer-length budget and exact binding regions permit it. Distant or difficult groups can require sequential rounds. Use the verified product of each round as the template for the next, while keeping edit positions tied to the original input. Do not combine round-specific primer pairs in one PCR.
Inverse PCR produces back-to-back primers rather than a QuikChange overlapping pair. The tool reports Tm for the exact 3′ binding region; it does not reproduce a kit-specific annealing-temperature calculation. Use the design with a matching inverse-PCR/end-joining protocol and consult your kit guidance. The overlap modes have a different fragment-and-joining workflow.
The current Mutagenesis designer accepts nucleotide substitutions only. For an amino-acid change, identify the coding-strand DNA and desired codon first, then enter the required base substitutions at their DNA coordinates. Protein sequence alone does not identify a unique DNA template. Insertions, deletions and combinatorial libraries are outside this tool’s current scope.
60 nt is the default total-primer design budget, including rewritten sequence and the binding region. You can set 40–120 nt, but a higher limit can require additional synthesis review. The engine still needs separate exact binding regions. If a design cannot fit, inspect the changes and method before increasing the limit or planning separate rounds.
Positions are 1-based in the exact uploaded DNA, so position 1 is its first base. Check the sequence version and strand, and ensure you have not entered a protein coordinate or a coordinate relative to an extracted subregion. Every change refers to the same original input. Correct the coordinate or template rather than selecting an arbitrary original base.
The transfer sends the final edited construct and the named primers. Primer Binding Checker checks where those primers map on that final DNA; it does not check each round against its actual starting template.
For sequential Inverse PCR, check each round’s pair separately against the intermediate template reported in the reaction plan. The first round uses the original DNA, and each later round uses the verified product of the preceding round. Do not combine the round-specific pairs in one PCR. Enter all mutation positions using the original uploaded DNA’s coordinates.
Yes. Assembly PCR Primer Designer and Gibson Assembly Primer Designer count one vector plus the inserts. A vector with two inserts is three total fragments; a vector with three inserts is four. Overlap PCR has no vector and counts only the fragments being fused. Kit-based overlap defaults use this total.
Paste or upload your exact vector sequence in Gibson Assembly Primer Designer. For circular DNA, Linearize after base opens between that base and the following base. Enter the sequence length to open immediately before base 1. Leave the position blank only for an already-linear, blunt-ended backbone; its first and last bases are the physical ends.
Enter the vector and each insert separately, then arrange the inserts in the final order. Total fragments includes the vector. Each junction needs overlap with its actual neighbor: vector to first insert, insert to insert, and last insert back to vector. The tool adds the primer tails. Review every junction and the final sequence before exporting.
Use the profile matching your kit and total fragment count. This site defaults to 20 bp for two or three total fragments in Gibson/HiFi; with four or more, Gibson defaults to 30 bp and HiFi to 25 bp. GeneArt starts at 15 bp. In-Fusion starts at 15 bp for vector plus one insert and 20 bp for more pieces. These are tool-profile defaults; review reported warnings and your kit instructions.
The 3′ region binds the original PCR template; the added 5′ tail carries sequence for joining to the next fragment. Order the full primer shown in the result. The reported binding Tm excludes the added tail. Total primer length, binding-region length and shared-overlap length describe different parts of the design.
Enter the sequence you want to retain from each source fragment, without manually adding the assembly primer tails. The designer creates those tails. If two supplied fragments already contain the same terminal homology, the tool does not automatically trim it; prepare the retained fragments so the intended construct does not duplicate that region.
An overlap is the shared sequence that joins adjacent PCR products and is retained once during assembly. A linker is new DNA you deliberately add between source fragments. Enter the linker once in the field for that junction. It contributes to the shared overlap; a linker longer than the requested overlap expands the reported overlap and can make the primers longer.
Add the retained vector length, all retained insert lengths and one copy of each added linker. Do not add primer-introduced homology again. A 3,000 bp vector plus 600 bp and 900 bp inserts gives 4,500 bp without linkers. Individual PCR products can be longer because they contain joining tails. Compare the final sequence as well as its length.
Assembly and Gibson exports begin at the first retained backbone base after the opening. For an opening after source base 250, the displayed construct starts at source base 251. Rotating a circular sequence changes its numbering without changing the molecule. Use the reported opening and fragment order to verify the insertion, and avoid treating exported coordinates as unchanged source coordinates.
A blank Linearize after base field tells the tool that your vector is already linear and blunt-ended, so it designs insert primers without a vector PCR pair. For a circular vector that you want opened by PCR, supply its full sequence and the opening position. Confirm that the selected preparation matches the DNA you actually have.
The transfer sends the final assembled or edited construct and the named primers. It checks their mapping on that final construct. To check an initial fragment PCR, open the corresponding original template separately and enter that fragment’s primer pair. Imported GenBank gene annotations are not carried over by the designers.
Choose Custom for a circular overlap assembly using your own vector and fragments, without a named kit profile. Choose Overlap PCR to fuse fragments into linear DNA; this mode has no vector or closing junction.
For Overlap PCR, amplify the separate fragments with their listed pairs, then use the reported outer primer pair for the fusion PCR. Turning the linear product into a plasmid requires a separate joining or cloning step.
Clear that junction’s Overlap (bp) field, then click or Tab out of it. The field takes the main overlap length again and follows later changes to that setting.
Use default resets the main overlap to the selected profile’s default. It leaves individually entered junction lengths in place. Clear each override that you want to reset. An optional DNA linker is a separate sequence setting; resetting an overlap length does not clear the linker.
Primer Scan (Beta) checks existing primers against one selected host genome or your custom reference. It searches both strands and predicts possible PCR products, including sites with substitutions within your settings. Unlike NCBI Primer-BLAST, it does not design primers, search NCBI databases or assess gapped alignments. Equivalent sensitivity has not been established.
Yes, host DNA may produce a background amplicon if your primers bind in suitable positions and orientations. Select the cloning or expression host in Primer Scan and enter the intended plasmid PCR product size, not the total plasmid length. Genomic candidates near that size are flagged even if the intended plasmid sequence is absent from the reference. The size entry does not verify amplification from the plasmid.
Product lengths help identify potentially confusing bands, but do not guarantee separation. Enter an intended plasmid amplicon size in Primer Scan to flag genomic products within plus or minus 10% of it. The fixed window is a screening aid, not a validated gel-resolution cutoff. Agarose concentration, fragment size, loading and electrophoresis conditions also matter; products outside the window are not guaranteed to be removable by gel purification.
The library includes E. coli K-12 MG1655, DH5alpha, TOP10 and BL21(DE3); B. subtilis 168, WB800N and PY79; S. cerevisiae S288C, CEN.PK113-7D and BY4742; and A. niger ATCC 1015 and NRRL3. Bundled references contain chromosomal DNA only, excluding plasmids and mitochondrial DNA. NRRL3 scaffolds remain separate. Choose the closest relevant strain, or supply your own assembly when its sequence differs.
Enter up to ten forward/reverse pairs. Each primer is searched on both strands, and each pair is checked independently, including possible products formed by one primer at two sites. Enable Check products between different primer pairs when the pairs will be mixed in one reaction. Cross-pair products are listed separately. This does not predict primer dimers or multiplex PCR efficiency.
No. Primer Scan's default minimum binding length is 15 bp, excluding non-binding 5' tails. It allows shorter binding regions to be considered, but they must still pass the selected screen. Consecutive-match mode separately defaults to 18 exact bases at the 3' end; set that threshold to 15 to check 15 consecutive matches. Neither threshold is a recommended primer design length or a guarantee of PCR success.
Choose annealing temperature to compare estimated binding-site Tm values with your planned PCR annealing temperature (Ta), using the selected salt and primer concentrations. Choose consecutive matches for an uninterrupted exact match ending at each primer's 3' end; this mode does not apply a temperature filter. Use full primer length requires an exact match across the entire primer, including any added tail.
Results show binding coordinates, strands and final product lengths, including inferred non-binding primer tails. Gene context is added where sequence-matched annotations are available. NRRL3 and custom references currently have no gene labels; this is not evidence that a product is intergenic. A product covering an annotated gene or CDS does not prove that the gene is intact, expressed or functional.
No. Custom references are read and scanned in your browser, not uploaded to our server or saved in an online account. Keep your original file for later scans. Supply assembled DNA, FASTA or GenBank sequence, not raw reads or protein sequences. Files must be uncompressed, no larger than 60 MB, and contain at most 50,000,000 bases across all records. Each chromosome, plasmid or contig should remain a separate record.
No preprocessing or indexing software is required. Choose a bundled reference or provide custom DNA, FASTA or GenBank, then scan in the browser. Bundled files use compact encoding internally; custom files need no conversion. Keep records separate, choose the correct topology and check size limits. Large or repetitive inputs can take longer or reach a resource limit.
No. It means no product passed the selected criteria in the supplied reference. Missing plasmids, strain differences, assembly gaps, short binding regions or restrictive settings can hide possible off-targets. Binding sites requiring insertions or deletions are not searched. Review near-threshold and unscored candidates, check your reference and size limits, and verify experimentally. An incomplete-search error is not a negative result.
Open Custom genome, paste or upload the complete plasmid, and select All circular for a FASTA record. Enter both primers 5′ to 3′ as ordered; do not reverse-complement the reverse primer again. Set the size limits to include the intended product and run the scan. Circular mode checks products across base 1. Use linear mode for an isolated fragment.
Use full primer length requires every primer base to match the reference, including a cloning tail. If the original template lacks that tail, leave this option off and select Allow non-binding 5' tails in Specificity scan settings. Review the binding thresholds and check that inferred tails match your design rather than a sequence error.
In temperature mode, Needs review can indicate a near-threshold Tm, a site outside the selected 3′ limits, or a mismatch pattern with no supported Tm estimate. A blank Tm is an unscored duplex, not zero degrees or proof of no binding. Inspect the alignment and reason; qualifying review candidates remain in the report and CSV.
A useful starting design has a template-binding region of roughly 18-30 bases, similar forward/reverse melting temperatures, and no strong 3' complementarity or long single-base runs. Primer Designer defaults to 25 bases. Check the actual template and your polymerase protocol before ordering.
Leave Target Tm blank and set Primer length for a fixed binding length (25 bases by default). Enter Target Tm to adjust each binding region separately toward a melting-temperature target. Added 5' tails do not count toward binding length or binding-region Tm; the target is not the PCR annealing temperature.
Primer melting temperature (Tm) is the temperature at which half the primer molecules are bound to their perfect complements under the specified conditions. The tools estimate it for the template-binding region, excluding added 5' tails. Tm is not the temperature you necessarily set for PCR annealing.
Check for strong self-complementarity and complementarity between primers, especially near their 3' ends. Primer Designer provides screening estimates for hairpins and dimers, not full folding predictions. Primer Binding Checker does not perform these thermodynamic calculations.
Yes. Primer Designer is free and needs no login. Enter your template and target region, choose a fixed primer length or target Tm, and check the resulting pair with Primer Binding Checker. These tools do not search genome databases for specificity.
Use Primer Binding Checker for an exact map and predicted products on one supplied template. Use Primer Scan to search both strands of a selected host reference or custom reference, allowing substitutions within your settings. Neither searches every public sequence database. Choose the relevant reference and check the intended product and any additional candidates.
DNA polymerase extends from the 3' end, so a suboptimal 3' end can give inefficient or nonspecific amplification. Check the last few bases for off-target binding, excessive complementarity, or too many G/C bases in a row.
Use the enzyme supplier's end-cleavage guidance. Primer Designer's Auto option adds a stored enzyme-specific clamp before the restriction site; you can override it. There is no single flanking-base count that guarantees efficient cutting for every enzyme.
Try a longer template-binding region or a nearby region with higher GC content, then recalculate Tm under the same reaction conditions. Added non-binding 5' tails do not raise the binding-region Tm during the first PCR cycles.
The primer sequence may be repeated or partly repeated in the template, especially at the 3' end. The primer pair summary identifies the chosen Forward/Reverse combination. In Full-template context, choose the pair and enable Show additional binding sites to inspect its other matches. Pair details and the TSV retain all compatible products. Check the exact binding ranges and product sizes before ordering or using the primers; additional sites alone do not establish extra PCR products or PCR success.
They do not need to be identical, but they should be close enough to work under the same annealing condition. If one primer is much lower, redesign that primer or adjust the target region.
Yes. Enter Target Tm to adjust forward and reverse binding lengths independently toward similar melting temperatures. Leave it blank to use fixed Primer length. The selected target is not a recommendation for PCR annealing temperature.
To translate into a downstream C-terminal tag, omit the insert's stop codon and keep the insert, junction and tag in the same reading frame. Confirm that a stop codon follows the tag in the final construct. Include an in-frame stop before the tag if you do not want it translated. Primer Designer does not automatically validate fusion-protein reading frames.
Primer Designer and Sanger Primer Designer use SantaLucia 1998 nearest-neighbor parameters with Owczarzy 2008 salt correction. Enter total dNTP across all four nucleotides: 0.2 mM each means 0.8 mM total. These estimates do not reproduce every manufacturer's proprietary buffer model.
It is the minimum consecutive exact match at the primer's 3' end, 15 bases by default. The tool reports each primer's longest matching suffix and treats unmatched 5' bases as a tail. It does not accept internal mismatches or exhaustively report every shorter suffix.
Primer Binding Checker accepts up to 10 forward/reverse pairs against one template and analyzes each pair separately. It does not predict cross-pair products. Primer Scan accepts up to ten pairs and has an optional Check products between different primer pairs setting for primers that will be mixed in one reaction. This sequence screen does not predict multiplex efficiency.
The template amplicon spans the selected binding regions. The final PCR product length also includes detected 5' tails from both primers. Check both lengths when cloning or comparing the expected product with a gel band.
Enter the sequence of the DNA you will actually submit for sequencing, then the target region and mutation positions using 1-based coordinates. Do not include flanks absent from a purified PCR product. Sanger Primer Designer predicts forward and reverse read coverage; it does not introduce mutations or analyze chromatograms.
A long insert may require overlapping Sanger reads from several primer positions. Forward and reverse reads can also help confirm a variant. Each standard Sanger sequencing reaction uses one primer; run the planned forward and reverse primers in separate reactions.
It is the part of a predicted read left after trimming the selected low-quality edge from both ends. With a 650 bp read and 50 bp edges, the planning window is 550 bp. Actual quality depends on your template, reaction and sequencing provider.
Sometimes yes, but PCR primers may sit too close to the region you need to read. For important checks, design sequencing primers so the mutation or insert junction falls in the reliable read window.
The signal close to the sequencing primer can be noisy, so the first stretch of bases may not be the best place to confirm a mutation. Plan coverage to account for low-quality read ends.
When sequencing a plasmid, use separate reactions with primers that read from the vector into each insert junction. When sequencing a purified PCR product, primer-binding sites must be present in that product. Add internal primers if the region extends beyond the usable read lengths.
Yes. Leave Read zones / primer pairs blank in Sanger Primer Designer for automatic planning of up to 10 pairs. Expected read length, low-quality edges and mutation positions guide the plan. If the suggested count exceeds 10, choose smaller overlapping regions. You can enter 1–10 manually, but review any coverage gaps or boundary warnings.
Listed mutation or SNP positions influence the planned read zones so those bases are less likely to fall in trimmed, low-quality edges. The result reports predicted forward and reverse coverage; it does not measure read quality or call a mutation.
A restriction site is a DNA sequence recognized by a restriction enzyme. Cleavage may occur within the site or outside it, as with Type IIS enzymes. Finding the sequence does not prove the enzyme will cut under your reaction conditions.
Paste or upload raw DNA, FASTA or GenBank into Restriction Site Analyzer. Choose linear or circular topology, search enzyme names or select a group, then analyze. Recognition positions and counts are separate from the resolved cleavage positions used for digest fragments.
Analyze the sequence, then select one or more enzyme badges or map labels for a combined digest. Review strand-specific cuts, fragment sizes and the virtual gel. One cut linearizes a circular template into a full-length fragment; no separate dashed reference is needed. The gel is a schematic, not a prediction of enzyme activity or the migration of uncut plasmid forms.
Possible causes include incomplete digestion, unexpected restriction sites, methylation-sensitive sites or star activity. Uncut supercoiled and nicked plasmids migrate differently from linear DNA. Confirm the sequence and topology; the virtual gel does not model these reaction or migration effects.
Sticky-end enzymes make staggered cuts that leave single-stranded overhangs. Complementary overhangs can base-pair before ligation. Blunt-end enzymes cut both strands at the same position and leave no overhang; their ligation conditions often need more optimization.
Check for unwanted internal sites in the vector and insert. For directional restriction cloning, choose two ends that cannot ligate to each other, while giving each insert end a compatible vector end. Different enzyme names alone do not ensure this. Confirm digestion conditions and methylation sensitivity with the supplier.
Some enzymes cut poorly when their recognition site is too close to the end of a DNA fragment. Add extra bases before the site in the primer, then recheck the designed sequence.
Star activity is cutting at sites that are similar to, but not exactly, the normal recognition site. It can happen under non-ideal digest conditions, too much enzyme, high glycerol, or long incubation.
Often yes, but both enzymes need compatible buffer and temperature conditions. If the activities do not match well, a sequential digest may be safer.
Circular DNA can produce fragments across the coordinate origin, meaning base 1 / the start of the entered sequence. If a plasmid is treated as linear, fragment sizes and map positions near base 1 can be misleading.
In circular mode, origin means the coordinate origin: base 1 / the start of the entered sequence. It does not mean the biological ori or origin of replication unless that feature happens to be located at base 1 in your sequence map.
Yes. SalI and XhoI recognize different sequences but both leave compatible 5' TCGA overhangs. Their ends can ligate, so using them at opposite ends does not enforce directional cloning. A SalI-XhoI hybrid junction does not regenerate either original recognition site.
Type IIS enzymes cut outside their recognition sequence. The complete cleavage region must be present in the supplied DNA. If a cut extends beyond a linear end, it is excluded with a warning. Overlapping cleavage regions also suppress fragment and gel predictions.
Enter vector mass, vector length, insert length and the insert:vector molar ratio in Ligation Calculator to find the insert mass. Enter either or both stock concentrations in ng/µL to see pipetting volumes. An optional reaction volume gives final DNA concentrations; leave space for buffer, ligase and other reagents.
A 3:1 insert:vector molar ratio is a common starting point, not a universal optimum. Follow the ligase and cloning protocol. The calculator offers preset ratios and a Custom multiplier; enter 6 for a 6:1 ratio.
The calculation only handles DNA amount. Ligation can still fail because of bad vector prep, incomplete digestion, missing phosphorylation, inactive ligase, poor insert purity, or difficult blunt-end ligation.
Ligation depends on the number of DNA molecule ends, not just the mass in the tube. A short insert has more molecules per ng than a long insert, so length must be included.
Use the amount recommended by your ligase or cloning protocol. Enter that mass in ng, together with vector length, insert length and the insert:vector molar ratio. The calculator determines insert mass; it does not choose an experimentally optimal DNA amount.
Sticky ends can base-pair with matching overhangs before ligase seals them. Blunt ends do not have that help, so they often need cleaner DNA, higher concentration, or optimized ligation conditions.
Transform uncut plasmid to check competent cells and selection. Transform digested vector without ligase to assess residual uncut-vector background, and a vector-only ligation to assess additional background from re-ligation. Compare these with the insert-plus-vector reaction, following your cloning protocol.
For acid/base pairs and pH-adjusted components, the requested stock volumes must fit within the final volume. Use more concentrated stocks or lower the target concentration when they do not fit. McIlvaine recipes show a warning for an oversized stock mixture. Always follow the selected recipe’s pH-adjustment instructions.
The transfer preserves liquid reagent volumes calculated from the selected % w/w assay and density, as well as volumes from the component-stock concentrations. Molarity Calculator receives these volumes as % v/v, with the preparation assumptions in each component name; dry solids retain their molar targets. Changing an assay, density or stock concentration requires recalculating and sending the buffer recipe again. The receiving calculator scales quantities but does not recalculate buffer pH.
Enter the molecular weight, target concentration, and final volume. The molarity calculator gives the mass to weigh and can also show stock-multiplier recipes.
For a dilution, stock concentration times stock volume equals final concentration times final volume. This conserves the amount of solute. Use consistent units for C1 and C2 and for V1 and V2; the calculator handles supported unit conversions.
A dilution factor tells how many fold lower the final concentration is. A 1:N antibody dilution means one volume of antibody stock in N total volumes. Always check the intended final concentration and protocol wording.
Only when converting between molar concentration and mass concentration, including % w/v. Molar-to-molar and mass-to-mass conversions do not need it. X units work only with X units. The calculator does not convert % v/v or % w/w.
Select Multi-solute in Molarity Calculator, enter final recipe volume, and add each component with its own concentration and units. Enter 1X targets before choosing a stock multiplier. This calculates amounts; it does not determine pH, solubility or biological suitability.
Very small stock volumes are easier to read in µL, while the final mixture may be easier to read in mL. The dilution tool keeps the quantities correct and formats each value in a practical unit.
Enter the intended 1X targets and select the 10X stock multiplier. The calculator multiplies each component amount for the entered final stock volume. Check solubility and pH; not every buffer or medium can be prepared at 10X.
Dissolve the solute in less than the final volume, then add solvent to reach the final solution volume specified by the recipe. Adding solute to the full solvent volume can give a larger final volume and a lower concentration than intended.
Percent w/v means grams per 100 mL. To convert it to molarity, you need the molecular weight. Without molecular weight, percent w/v and molar units cannot be converted exactly.
Compare it with your pipette's reliable range. Dilution Calculator flags stock or antibody volumes below 1 microlitre and suggests an intermediate dilution when possible. Larger volumes are not automatically accurate; consider pipette calibration and handling technique.
Final volume is the total volume after stock plus diluent are combined. Diluent volume is only what you add after measuring the stock volume.
Yes, but only if you know the molecular weight. µM is a molar unit, while mg/mL is mass per volume, so molecular weight is required to convert between molar and mass concentration.
Enter 1X working targets and choose the stock multiplier in Molarity Calculator. Above 1X, results show the working amount alongside the stock amount. The multiplier changes concentration, not the entered final preparation volume.
Yes. Percent w/v means grams per 100 mL of final solution. Multiply molarity in mol/L by molecular weight in g/mol, then divide by 10. For example, 1 M NaCl at 58.44 g/mol is 5.844% w/v.
Yes. Buffer Calculator supports named systems such as phosphate, citrate, Tris and HEPES, plus recipes such as PBS, TBS, TAE, TBE, SSC and TE. Set strength, final volume and target pH, then review the calculation type and suitability warnings.
Use Buffer Calculator for named buffer recipes and pH guidance. Use Molarity Calculator when you already know each component's target concentration. Buffer Calculator uses acid/base equations, mixing tables or fixed recipes depending on the buffer; changing target pH does not change the composition of a fixed recipe.
No. The stock multiplier changes the concentration or amount used to prepare a concentrated stock recipe. The final volume is still the volume you bring the prepared stock or multi-solute recipe to, not extra diluent added on top.
Yes. Protein molarity mode converts mass concentration to molarity, molarity to mass concentration, total mass to mole amount, and mole amount to mass. Enter protein molecular weight in kDa. For a 50 kDa protein, 1 mg/mL equals 20 micromolar.
Use amount (pmol) = mass (µg) × 1000 / molecular weight (kDa). For example, 25 µg of a 50 kDa protein is 500 pmol. In Protein molarity mode, select protein mass to mole amount, enter the molecular weight and mass, and choose pmol output. No sample volume is needed for this conversion.
Use the molecular weight of the species you want to count. A 50 kDa subunit at 1 mg/mL corresponds to 20 µM monomer equivalents. If all protein forms intact 100 kDa dimers, the dimer concentration is 10 µM. The calculator cannot determine the oligomeric state or the proportions in a mixture.
A hydrate contains water of crystallization, which contributes to its formula mass. For the same number of moles of the salt, a hydrate therefore requires more mass than the anhydrous form. Select the exact salt and hydrate shown on the bottle; do not substitute gram-for-gram. Check the supplier's molecular weight and assay.
For a dilution, maximum final volume = stock concentration × available stock volume / target concentration, using consistent units. For example, 100 µL of 10 mM stock makes 1 mL at 1 mM. Enter Available stock volume in Dilution Calculator to see this limit and check whether your requested preparation is possible. The estimate does not include pipetting losses.
The pKa of Tris changes with temperature, shifting the acid/base equilibrium. For a fixed Tris buffer composition, pH generally decreases as temperature rises and increases on cooling. Measure or adjust pH at the temperature specified by your protocol, and record that temperature. Buffer Calculator does not predict temperature-dependent pH changes.
Select Antibody dilution and Dilution ratio (1:N), then enter 1000 and the final volume. One stock volume is included in 1000 total volumes, not added to 1000 volumes of diluent. For 10 mL final volume, use 10 microlitres antibody and 9.99 mL diluent.
Yes. In Antibody dilution, select Stock and target concentration, choose the units and enter final volume. The result uses C1V1 = C2V2. It does not account for antibody activity, carrier proteins or assay performance; titrate for your application.
First prepare a lower-concentration intermediate stock using the listed stock and diluent volumes. Then take the specified amount of that intermediate to make the final working solution. These are two separate mixtures; do not add both recipes' diluent volumes to the final tube.
For supported acid entries, Molarity Calculator and Buffer Calculator use editable % w/w assay and density to estimate stock volume. Check these values against the product certificate or SDS. They are not the same as the final % w/v or molarity target.
Yes. The citrate-only recipe uses a published 100 mM citric-acid/trisodium-citrate table from pH 3.0 to 6.2, with interpolation between entries and no extrapolation. At other concentrations, the calculated amounts do not ensure the same pH. Measure and adjust the finished buffer at its working temperature.
The reverse complement is the complementary DNA strand written 5' to 3'. Use it to design a reverse primer's binding region, check insert orientation or inspect an ORF on the opposite strand.
DNA sequence is normally written from the 5' end to the 3' end. Primer sequences and coding sequences should be checked in this direction to avoid orientation mistakes.
In ORF and Protein Translator, select Quick translate with DNA input. Choose a single frame or All 6 frames. Use ORF finder instead to list candidate coding regions with positions and amino-acid lengths. Translation uses the standard genetic code.
An open reading frame is a stretch without an in-frame stop codon. This ORF finder starts candidates at exact ATG codons and ends them at TAA, TAG or TGA. Include open-ended ORFs also reports candidates with no downstream stop in the selected sequence.
Codon optimization means changing synonymous codons so the DNA sequence is better matched to an expression host while encoding the same protein. It should keep the amino acid sequence the same.
No. Codon choice is only one factor. Expression also depends on the promoter, host strain, mRNA structure and stability, induction conditions, protein folding and toxicity. Purification affects recovered yield; it is not a measure of expression alone.
For a target strand written 5' to 3', the reverse primer's binding sequence is the reverse complement of the target's right-hand end. It anneals to that displayed strand and extends toward the forward primer. Both primer sequences are entered and ordered 5' to 3'.
Translate the strand that corresponds to the coding sequence in the correct 5' to 3' direction. If you are not sure, check all reading frames and look for the expected ORF.
Each DNA strand can be read in three frames, depending on where codon grouping starts. Because DNA has two strands, there are six possible reading frames to check.
Very high or very low GC content can affect synthesis, PCR, sequence stability, and expression. Codon optimization should balance host codon preference with practical sequence constraints.
Choose the organism in which the protein will be expressed, not its source organism. The 20 available host tables use species-level genomic codon counts. Cell-line labels such as CHO, HEK-293 and Sf9 are host-species approximations, not cell-line-specific expression data.
Yes. The Reverse Complement Tool accepts raw DNA sequence, FASTA, and plain-text GenBank input. For GenBank, it reads bases from the ORIGIN sequence section and ignores feature annotations, including ori/origin-of-replication annotations.
Check the strand, reading frame, selected region and minimum ORF length. This tool requires an exact ATG start and uses the standard genetic code, so alternative starts or genetic codes can be missed. Incomplete sequence, introns, sequencing errors or internal stops can interrupt the expected ORF. Try all six frames; for missing terminal stops, enable Include open-ended ORFs.
For DNA entered 5' to 3', Complement returns the paired bases in 3' to 5' order. Reverse only reverses the input. Reverse complement both reverses and complements it, giving the paired strand 5' to 3'. Cleaned output keeps the input orientation. The tool supports IUPAC ambiguity codes.
Usually not: most amino acids have more than one codon, so a protein sequence does not generally identify its original DNA sequence. Quick translate returns one possible coding sequence and a degenerate IUPAC library. The library can encode additional amino acids or stops. Use Codon Optimizer for host-specific codon choices.
In Optimize DNA CDS mode, enter a codon-aligned, 1-based range such as 4-300. Valid starts are 1, 4, 7 and so on; valid ends are 3, 6, 9 and so on. Bases outside the range stay unchanged. Leave the range blank for the full CDS.
The 20 host tables use FDA HIVE-CoCoPUTs genomic CDS counts from its public September 2021 release, based on RefSeq 99 and GenBank 238.0. Our snapshot was retrieved on 14 September 2026. The newer retrieval date does not change the source dataset's age.
No. The optimizer tries to meet the selected GC range and reduce unwanted restriction sites or DNA motifs while preserving translation. Synonymous choices and a restricted editing region can make constraints incompatible. Check final GC%, remaining motif counts and warnings.
Yes. The current Mol Biology Tools pages are free to use in the browser, with no login needed.
For quick checks, use the separate tools for primer design, primer binding, restriction sites, reverse complement, ORF translation, ligation, molarity, and dilution. Plasmid Editor can edit supplied DNA and annotations, review deletions, and export sequence files or SVG, PNG and JPEG maps. It does not provide a full laboratory platform, automatic annotation, alignment editing or every proprietary desktop feature.
It supports quick PCR primer design from a supplied template, with target Tm, fixed length, overhangs and quality screening. Its code is not the Primer3 engine. Use NCBI Primer-BLAST or another database search when genome-wide primer specificity is needed.
It provides an exact-match in-silico PCR-style check for your supplied linear or circular template. It predicts products for each forward/reverse pair, but does not model mismatches or cross-pair products, exhaustively screen every orientation, or search a genome database.
Yes, for site mapping and predicted digests of supplied DNA. It finds recognition sites, calculates strand-specific cuts and fragments, and displays linear or circular maps. Compatible-end warnings concern ligation of DNA ends, not whether the enzymes work together in one buffer. Check reaction conditions with the supplier.
It supports browser-based ORF finding, six-frame DNA translation and protein-to-DNA reverse translation. It currently uses only the standard genetic code and exact ATG starts, so it does not reproduce every option of NCBI ORFfinder or ExPASy Translate.
Sequence analysis and calculator computations run locally in the browser. Tool-to-tool transfers use browser-local storage. The website still makes normal requests to load pages and services; see the Privacy page for analytics and hosting details.
Choose Primer Designer for ordinary PCR, Assembly PCR or Gibson for fragment joining, Mutagenesis for nucleotide substitutions, or Sanger Primer Designer for sequencing coverage. Review the full oligos and the method-specific reaction plan, then check binding on the appropriate template and use Primer Scan for the relevant host or custom reference.
For restriction-and-ligation cloning, inspect sites with Restriction Site Analyzer and calculate insert mass with Ligation Calculator. For overlap joining, use Assembly PCR or Gibson to set vector ends, fragment order and overlaps. Review the resulting sequence and primer plan, then check the relevant template or host with the binding tools.
Use calculators to avoid arithmetic mistakes, then still check the biology: enzyme conditions, primer specificity, sequence orientation, solubility, and the actual wet-lab protocol.
Yes. Supported transfer buttons pass the current template, primers or calculated components through browser-local storage. Open the tools on the same site in the same browser, then check the received inputs before calculating. Manual copying remains available if browser storage is blocked.
Yes. After generating PCR primers, open the designed primer pair directly in Primer Binding Checker to check binding sites, orientation, and expected amplicon size against the same template sequence.
Yes. Sanger generates up to 10 pairs per design and transfers every pair with the template for an exact binding check. Primer Binding Checker accepts all ten together. Original names and full primers are retained.
For many routine planning steps, yes. Use Restriction Site Analyzer to check enzyme sites, Primer Designer to create primers or overhangs, Primer Binding Checker to validate binding and amplicon size, and Ligation Calculator to calculate insert-to-vector molar ratio amounts.
Use raw sequence, FASTA or GenBank ORIGIN text where the selected tool supports them. Plasmid Editor also opens annotated GenBank, ApE, EMBL and SnapGene .dna files, up to 25 MiB per sequence file and 5 million bases per record; its saved projects can be up to 80 MiB. Other sequence tools do not accept SnapGene binary files and have their own smaller limits. Restriction and reverse complement allow 100,000 bases; ORF, codon and Sanger allow 250,000; ordinary primer design/binding allow 500,000. Assembly/Gibson allow 100,000 per sequence and 200,000 per final construct; Mutagenesis allows 100,000. Their uploads must be under 1 MB. Primer Scan allows 50,000,000 bases across all records and uploads up to 60 MB. Check the individual guide for alphabet, record and protein limits.
Primer Scan searches each reference record independently and does not form products between records. Assembly PCR, Gibson and Site-Directed Mutagenesis require one record per sequence input. The older sequence tools concatenate multiple FASTA records into one sequence; process unrelated records separately there to avoid artificial junctions, ORFs or binding sites. Plasmid Editor keeps records separate and lets you switch with its Record selector.
Each tool has a References section, and each help guide links to it. These identify implemented calculations, database snapshots and their limits. Classic methods papers remain relevant; a database citation year is not necessarily the date of every stored value.
The tool has rejected characters outside the alphabet allowed for that field or mode. Read the existing error message, correct the highlighted symbols and run the tool again; the highlights update while editing. A malformed FASTA or GenBank record can also fail its structure checks without a single highlighted character. Export binary sequence files as plain-text FASTA or GenBank.
It depends on the button. Assembly, Gibson and Mutagenesis send the final construct, topology and named primers to Binding Checker. Primer Designer’s input buttons send the template; its result button also sends the designed pair. Transfers to Primer Scan send primer pairs, so choose the reference there. Buffer Calculator sends recipe components and volume to Molarity Calculator. Each tool guide lists its direct destinations and manual follow-up steps.