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• Multiplex PCR: favors primer sets with reduced cross-dimers; use tighter Tm bounds.
• TaqMan / MGB probe assays: enable one option at a time; probes are designed within the amplicon under assay-specific constraints.
• RPA: shifts to longer primers and short amplicons suitable for ~37–42 °C isothermal workflows.
• Inverse PCR: assume circular template; ensure the marked region is consistent with circular amplification logic.
• C>>T bisulfite conversion: converts non-CpG cytosines for in silico evaluation on bisulfite-treated sequences.
• Overlapping primers: relaxes constraints on overlap where necessary (use cautiously).

Sequences use the standard IUB/IUPAC nucleic-acid codes. The full list of accepted letters (including U, I and LNA bases) is in Input formats → Allowed nucleotide codes.

The structure of the last nucleotides at the 3′-end of the primer can be specified to control primer specificity:

• N — any pattern (no constraint)
• One, two, or more characters using standard or mixed letters
• Multiple patterns of equal length separated by spaces: sws ssw sww wss www

For example, WSS corresponds to all 3′-end variants: acc acg agc agg tcc tcg tgc tgg.

The pre-designed primers/probes list is used for multiplexing with prior designed PCR primer/probe sets. This allows you to incorporate existing validated assays into new multiplex panels while ensuring compatibility and avoiding cross-reactivity.

Design of specific PCR primers for in silico bisulfite conversion for both strands. Only cytosines not followed by guanine (non-CpG context) will be replaced by thymines. CpG methylation sites are preserved.

Optimal primers should hybridize only to the target sequence. Common problems include annealing to repetitive sequences (retrotransposons, transposons, inverted tandem repeats), alternative product amplification, and multiple bands due to off-target binding.

Masks stretches that are poor targets for primer design because of their low sequence complexity — simple/microsatellite tandem repeats (SSRs) and telomere-like repetitive motifs, plus regions similar to them — the same low-complexity blocks detected by Repeat Finder. Such regions are prone to polymerase slippage, unstable annealing, and non-specific priming, so primers are not placed inside them.

This complements Non-specific priming control, which masks dispersed/interspersed repeats found elsewhere in the sequence: low complexity control targets local, in-place repetitiveness at the candidate site itself. Enabled by default.

Minor Groove Binders (MGBs) selectively bind non-covalently to the minor groove of the DNA helix, enabling shorter probe lengths and superior quenching for highly specific TaqMan-type assays.

Results in the PCR Primer Sets tab are not listed as simple one-to-one forward/reverse pairs. Instead, the output is organised by forward primer: for each forward primer candidate, all compatible reverse primers that pass the cross-dimer, Tm, and product-size filters are listed beneath it. When a probe mode is active (TaqMan, MGB, or Molecular beacon), every compatible probe found within the amplicon window of that F+R combination is listed under the reverse primer row.

This structure lets you:

• See at a glance how many workable reverse primers exist for a given forward primer — useful when one position has a narrow design window on one strand.
• Compare amplicon sizes and annealing Tm across all valid F+R combinations and pick the one that best fits your protocol.
• Choose, per F+R pair, the probe with the most favourable placement, Tm, or GC% without re-running the design.

Reading the output

• A forward primer row is marked F in the ID column and contains the primer sequence, length, Tm, CG%, LC%, and YR%.
• Each reverse primer row beneath it is marked R and additionally shows the amplicon size (bp) and the predicted annealing Tm (°C) for that specific F+R pair.
• If probes are designed, each probe row is indented below the reverse primer and marked with its probe type (P, MGB, or MB), along with its position within the amplicon.
• When no compatible reverse primer exists for a given forward primer, that forward primer is omitted from the output entirely.

• Paste or upload multi-FASTA DNA (no [brackets] markup is needed — the whole alignment is the target).
• Auto-correct orientation reverse-complements any read on the opposite strand before alignment, so mixed-orientation inputs still align.
• The alignment is computed in the browser; gap-open / gap-extend penalties tune how readily indels are introduced.

The Alignment & Conservation tab reports the alignment length, % conserved columns, the list of conserved blocks, and a consensus track (* = 100% conserved, lowercase IUPAC = variable, - = gap column).

Polymerase extension starts at the primer's 3′ terminus, so a mismatch there abolishes amplification. The tool therefore enforces:

• 3′-end window 100% conserved — the last N bases at the priming end (the “Conserved 3′-window”, default 8 nt) must be identical in every input sequence. This is the minimum guarantee for every primer reported.
• 5′-end may vary — with “Allow degenerate 5′-end”, upstream positions can sit on variable columns and are written as IUPAC codes (e.g., R=A/G, Y=C/T, N=any). With the option off, the whole primer is restricted to 100% conserved columns (plain A/C/G/T, no degeneracy).
• Gap columns are never spanned, so each primer has the same length in every template.

This mirrors the standard practice for consensus/universal primers, where the 3′ region is required to have zero variable sites while limited degeneracy is tolerated toward the 5′ end.

• Length range / Tm range — define the primer search window. Tm is computed on the majority-base (representative) sequence.
• Product size (bp) — filters pairs by amplicon length; reported as the range of ungapped lengths across all templates.
• Conserved 3′-window (nt) — number of fully conserved bases required at the 3′ end (≥3).
• Min. linguistic complexity (%) — rejects low-complexity / repetitive primers (see LC%).
• Max. pair Tm difference (°C) — keeps forward and reverse primers thermally matched.
• Allow degenerate 5′-end — toggles IUPAC encoding of variable 5′ positions (see the conservation rule above).
• Report overlapping primers — when off (default), clusters of overlapping primers are collapsed to the single best candidate by linguistic complexity (LC%), so the output is not flooded with near-duplicate primers whose 3′-anchors differ by only a nucleotide or two. Enable it to list every candidate, including overlapping ones.
• Max forward sets to report — caps the number of forward-primer blocks shown. Output is grouped: each forward primer is printed once with all its compatible reverse primers (each with product size and Ta) beneath it; blocks are separated by a blank line. The report shows totals vs. displayed.

• Alignment & Conservation — summary, conserved-block table and consensus/conservation track.
• Universal Primers — every candidate (Forward/Reverse) with sequence, length, Tm, CG%, LC%/LC_RY%, a 3′-conserved flag and a Degenerate flag.
• Primer Pairs — compatible F+R sets, each primer with its LC%/LC_RY%, plus product size (bp range) and predicted annealing Ta (°C).

Coordinates are 1-based alignment columns (e.g., F_1-20), so they map directly onto the consensus shown in the conservation track. Use the Copy / Download buttons to export each tab as TSV.

• No pairs found? Lower the conserved 3′-window, widen the Tm/length limits, or enable a degenerate 5′-end. Very divergent inputs may have no conserved block long enough for a primer.
• Keep degeneracy modest — highly degenerate primers lower effective concentration and specificity; prefer placing degenerate positions only at the 5′ end.
• Garbage in, garbage out — misaligned or unrepresentative input sequences yield misleading “conserved” blocks. Inspect the alignment first (the MSA tool helps).
• This is an in-browser consensus-primer designer; validate candidates by in silico PCR against your real templates before ordering.

Background reading on universal / consensus primer prediction from alignments:

• Sobhy H., Colson P. Gemi: PCR Primers Prediction from Multiple Alignments. Comp. Funct. Genomics 2012, 783138. doi:10.1155/2012/783138
• ConsensusPrime — A Bioinformatic Pipeline for Ideal Consensus Primer Design. BioMedInformatics 2022, 2(4), 637–642. doi:10.3390/biomedinformatics2040041

>1
gctctctgtgtctgatccaagaggcgaggccagtttcatttgagcattaa[A/G]tgtcaagttctgcacgctatcatcagggg

>2
tcatattccagtttgggcgagttttaagataggtccgg[S]acagtctttgcggcgccaacgcgtctttctccag

[S] represents G/C using the IUPAC ambiguity code.

Leave one side empty for deletions:

>1
tcatattccagtttgggcgagttttaagataggtccgg[AG/]acagtctttgcggcgccaac

>1
tgggcagcattagtagaagaaagtacaagaccgtgtgtagagg[GATATACTTGAG/CAGTCC]agcagatagcgttggatag

The [square brackets] should surround all SNPs that are part of the haplotype. Nearby SNPs not part of the haplotype should be outside brackets and identified using an IUPAC code.

ASP primers can carry standard KASP FAM and HEX tails (LGC Biosearch Technologies). Paste tails in FASTA-like format in the dedicated tab:

>FAM
GAAGGTGACCAAGTTCATGCT
>HEX
GAAGGTCGGAGTCAACGGATT

Custom tails can be used instead of the standard sequences.

• Non-specific priming control (default on) — keeps both the allele-specific and common primer off dispersed/interspersed repeats elsewhere in the flanking sequence.
• Low complexity priming control (default on) — keeps primers off other SSR/microsatellite and telomere-like stretches in the flanks, the same blocks detected by Repeat Finder. Disable either control only if it prevents a primer from being found and the flanks cannot be replaced.

Enter one or more rsIDs (space/comma separated), select a species, and set the desired flank size. The tool retrieves the flanking sequence from Ensembl and populates the input area automatically.

• Biallelic SNP: flank [A/G] flank or single IUPAC code flank [R] flank
• Tri- / tetra-allelic SNP: [A/C/G], [A/C/G/T] or [N]
• Insertion / deletion: use - for the empty side, e.g. [ATCG/-] (4-bp deletion) or [-/T] (single-base insertion)
• Excluded region — wrap with /.../ to keep primers out; see Targeting markup.

>rs1801133
tggggggaaaattagaggtaaccaaaatgggg...gatgaaatcg[Y]ctcccgcagacaccttctcc...

>rs3918290
gccacatacagtgaaaaccaactcaataaa...aatcacactta[ATCG/-]gttgtctggaaagtcagcc...

All standard IUPAC ambiguity codes are accepted both inside and outside the brackets — see the Input formats section.

Exactly one probe format may be active at a time — checking one option automatically clears the others.

• Length range (nt) — default 20–22. Keep the window narrow for uniform amplification across loci.
• Tm range (°C) — default 61–63. Match it to your master-mix recommendation. Probe Tm is internally targeted ~5–10 °C above primer Tm. Computed under the site-wide standard-PCR conditions (see Common parameters → Tm calculation conditions).
• PCR product size (bp) — default 200–500. For optimal qPCR efficiency keep amplicons ≤200 bp where possible.
• 3′-end pattern (default sws ssw sww wss www) — restricts the last bases of each primer; use S=G/C, W=A/T, N=any. Multiple space-separated patterns are tried and the best result is kept.
• Forward / Reverse tail (5′-3′) — optional universal tails appended to the 5′ end of each primer (e.g. M13 sequences) for downstream universal-primer detection.
• Mask repeats & low-complexity regions (on by default) — see Common parameters → Mask repeats.
• Allow overlapping primer positions — relaxes positional constraints when the search space is tight (use cautiously).
• Multiplex PCR — designs assay sets for all input sequences to run together in one reaction. Each candidate primer/probe is checked against every primer/probe already picked for other sequences, so only mutually dimer-compatible sets are kept; sequences with few candidates are prioritized first so they aren't crowded out. Requires ≥2 input sequences (ignored for a single sequence). See Output tabs and columns for how the assay-set table changes when this is on.
• C → T bisulfite conversion — switches the design into bisulfite-treated mode for methylation analysis (non-CpG cytosines are converted in silico; CpG sites preserved).

• Input Sequences — your FASTA / retrieved sequences with live formatting and a per-tab counter.
• My Primers — paste pre-existing primers/probes (optional) to be checked for compatibility with newly designed sets, e.g. for adding the new variant to an existing multiplex panel.
• Report (Primer list) — flat list of every primer and probe candidate with their physico-chemical properties.
• qPCR Assay Sets — final ready-to-order assays grouped by target: forward primer + reverse primer + allele-discriminating probe. With Multiplex PCR checked, this tab instead lists complete panels: each block groups one compatible assay per input sequence, and successive blocks are alternative panels — the number of blocks is capped by whichever sequence yielded the fewest compatible sets.

Per-oligo and per-pair columns follow the Common parameters → Standard output columns (ID · Sequence · Length · Tm · CG% · LC% · YR%, plus Fragment size and Annealing Tm on each pair).

1. Load the variant — paste FASTA with [REF/ALT], upload a file, or retrieve by rsID from Ensembl. Use Load Example to see the expected formatting.
2. Pick a probe format (TaqMan by default; switch to MGB for short / AT-rich targets, or Molecular beacon for difficult discrimination).
3. Adjust Tm range, product size and 3′-end pattern if your master-mix or chemistry requires it.
4. (Optional) Paste universal forward/reverse tails or pre-existing primers in My Primers.
5. Click Generate. Review the assay sets in the qPCR Assay Sets tab.
6. Copy the result area (Ctrl+A → Ctrl+C) and paste into Excel for ordering — see Exporting results.

• No assay set returned — widen the Tm window / product size or uncheck Mask repeats; full checklist in Troubleshooting → Results are empty.
• Probe not designed — for AT-rich variants switch to MGB; for very low-complexity flanks try Molecular beacon; check that the polymorphic position has at least 30 bp of unique flanking sequence on each side.
• Ensembl retrieval fails — confirm the rsID and species id are correct, see Sequence retrieval fails.
• Adding the assay to an existing panel — paste the existing primer/probe set into My Primers; new candidates incompatible with the panel will be filtered out.

Strengths

• Sensitivity. Detects mutant fractions ≥0.1%, compared with the ~5–20% floor of Sanger sequencing.
• Speed. Sample-to-result in under three hours on a standard real-time instrument.
• Sample tolerance. Compatible with fresh-frozen tissue, cell lines, cfDNA, and FFPE material.
• Robustness. Less affected by common PCR inhibitors (e.g. melanin in melanoma extracts) than several alternative allele-specific methods.

Typical applications

• Targeted oncology. Genotyping actionable hotspots in EGFR, BRAF, KRAS and similar genes to inform therapy selection.
• Resistance monitoring. Tracking minor resistant sub-clones (e.g. FLT3 in AML) that drive relapse.
• Liquid biopsy. Detection of circulating tumor DNA and rare tumor cells from peripheral blood.

• Paste FASTA in the Input Sequences tab with the polymorphism enclosed in square brackets (e.g. [A/G], [ATCG/-], [R]).
• Upload a FASTA file — plain FASTA, multi-FASTA or sequences already containing [REF/ALT] brackets.
• Retrieve from Ensembl by rsID — choose a species from the drop-down (Human, Mouse, Rat, Cow, Pig, Chicken, Arabidopsis), enter rsIDs separated by spaces, set the flank size (≥100 bp) and click Retrieve from Ensembl. The forward-strand context with the allele bracket is fetched automatically.
• Custom species — type any Ensembl species id (lowercase, underscores, e.g. canis_lupus_familiaris) into the "Other species" field and click Add. Custom species are stored in your browser (localStorage) for later use.

Provide ≥250 bp of flanking sequence on each side of the variant so the primer search has enough room.

• Biallelic SNP: flank [A/G] flank — alleles separated by /
• Shorthand (IUPAC): flank [R] flank — any IUPAC ambiguity code
• Tri- / tetra-allelic SNP: [A/C/G], [A/C/G/T] or [N]
• Insertion / deletion: [ATCG/-] (deletion) or [-/T] (insertion)
• Excluded region — wrap with /.../ to keep primers and probes out; see Targeting markup.

All IUPAC ambiguity codes are accepted inside and outside the brackets — see Input formats → Allowed nucleotide codes.

Select the probe chemistry — one format at a time; checking one option clears the other. The probe is placed in the conserved amplicon region shared by all alleles (between the ASP and the common reverse primer) and detects all amplicons regardless of allele. Allele discrimination comes entirely from the ASP.

• Length range (nt) — default 20–22. Defines the search window for the common counter-direction primer.
• Tm range (°C) — default 61–63. Match it to your master-mix recommendation. Probe Tm is internally targeted ~5–10 °C above primer Tm (TaqMan: +8 °C, MGB: −2 °C of the primer Tm window).
• PCR product size (bp) — default 200–500. For optimal qPCR efficiency keep amplicons short.
• SNP at 3′-end of ASP — number of bases at the 3′-end of the allele-specific primer that must differ between alleles (default 2; use 1 for simple SNPs and increase for tri-/tetra-allelic or short InDels). A larger value adds extra mismatches on non-target templates and suppresses leak amplification.
• 3′-end pattern (default sws ssw sww wss www) — restricts the last bases of the common primer; S=G/C, W=A/T, N=any. Multiple space-separated patterns are tried and the best primer is kept.
• Forward / Reverse tail (5′-3′) — optional universal tags appended to the 5′ ends of the common primers (e.g. M13 sequences) for downstream universal-primer detection.
• Mask repeats & low-complexity regions (on by default) — see Common parameters → Mask repeats.
• Allow overlapping primer positions — relaxes positional constraints when the search space is tight (use cautiously).
• Multiplex castPCR — designs one dimer-compatible castPCR set per locus so that all input targets can be amplified together in a single reaction. Each candidate set for a locus is screened for primer-dimers against the primers and probes already committed for the other loci, so only mutually compatible sets are kept. Requires ≥2 input loci (ignored for a single sequence). See Output tabs and columns for how the assay-set tab changes when this is on.
• C → T bisulfite conversion — switches the design into bisulfite-treated mode for methylation analysis (non-CpG cytosines are converted in silico; CpG sites preserved).

• Input Sequences — your FASTA / retrieved sequences with live formatting and a per-tab counter.
• My Primers — paste pre-existing primers/probes (optional) to be checked for cross-dimers against newly designed candidates, e.g. when adding a new variant to an existing panel.
• Report (Primer list) — flat list of every candidate ASP, common detection probe and common primer with their physico-chemical properties.
• castPCR Assay Sets — final ready-to-order assays grouped by target. For each locus two sets are reported: the forward set (ASP_F per allele + one shared Probe_F + R common primer) and the reverse set (ASP_R per allele + one shared Probe_R + F common primer). The probe is the same for all alleles within each orientation; pick the orientation that gives the better probe placement and allele discrimination. With Multiplex castPCR checked, this tab instead reports a single combined panel — one dimer-compatible castPCR set per locus (forward or reverse, whichever fits), chosen so the whole panel can be amplified together in one reaction. A locus for which no set compatible with the rest of the panel can be found is omitted.

Per-oligo and per-set columns follow the Common parameters → Standard output columns (with Fragment size and Annealing Tm on the common-primer row).

Cross-locus combinatorial use

All listed candidates are pre-checked for cross-dimers between sequences, so a chosen ASP + probe + common primer set from one locus can be combined with any other locus in a low-multiplex run without re-validation. To have the tool assemble those cross-compatible panels for you automatically, enable Multiplex castPCR (see above).

1. Load the variant — paste FASTA with [REF/ALT], upload a file, or retrieve by rsID from Ensembl. Use Load Example to see the expected formatting.
2. Pick a probe format (MGB by default; switch to TaqMan when the SNP flanks are GC-balanced and longer probes are acceptable).
3. Adjust Tm range, product size and the SNP at 3′-end of ASP value for your variant type (1 for simple SNPs, 2+ for tri-/tetra-allelic or short InDels).
4. (Optional) Paste universal forward/reverse tails or pre-existing oligos in My Primers.
5. Click Generate. Review the assay sets in the castPCR Assay Sets tab — both forward and reverse triplets are reported; choose the orientation with the cleaner discrimination.
6. Copy the result area (Ctrl+A → Ctrl+C) and paste into Excel for ordering — see Exporting results.

• No assay set returned — widen the Tm window / product size or uncheck Mask repeats; full checklist in Troubleshooting → Results are empty.
• Probe not designed — for AT-rich variants switch to MGB; check that the polymorphic position has at least 30 bp of unique flanking sequence on each side.
• Poor allele discrimination at qPCR — increase SNP at 3′-end of ASP from 1 to 2 so the last two bases of the ASP differ between alleles; the additional mismatch on the non-target template suppresses leak amplification.
• Ensembl retrieval fails — confirm the rsID and species id are correct; see Sequence retrieval fails.
• Adding the assay to an existing panel — paste the existing primer/probe set into My Primers; new candidates incompatible with the panel are filtered out.

Paste a multi-FASTA where each sequence contains one SNP marked in square brackets. Provide ~150–300 nt of flanking context on each side so the tool has room to choose PCR primers and probes.

• Biallelic SNP: ...flank[C/T]flank...
• Multi-allelic: ...flank[G/A/C/T]flank...
• IUPAC shorthand: ...flank[R]flank... — any IUPAC ambiguity code
• InDels: ...flank[ATCG/-]flank... (one allele empty for a pure deletion)
• Excluded region — wrap with /.../ to keep PCR primers and SBE probes out; see Targeting markup.

>rs3918290
gccacatacagtgaaaaccaactcaataa…ttagatgttaaatcacactta[C/A]gttgtctggaaagtcagcc…aatatacat
>rs1801133
tggggggaaaattagaggtaaccaaaatgg…aagaatgtgtcagcctcaaagaaaagctgcgtgatgatgaaatcg[Y]ctcccgcagacaccttctcc…ggaggtggctcagag

Sequences can also be retrieved by rsID via Ensembl directly from the input tab.

• Length range (default 20–22 nt) and Tm range (default 61–63 °C). Tight Tm windows are essential for multiplex — all loci must amplify under one cycling profile.
• PCR product size (default 200–500 bp). For SNaPshot it is the SBE probe — not the amplicon — that resolves the genotype, so the amplicon can be small.
• 3′-end pattern (default sws ssw sww wss www): a space-separated list of 3-letter composition codes (S=G/C, W=A/T, N=any) describing the last three bases. The tool tries every pattern and keeps the best primer; the default favours one G/C clamp without G/C runs.
• Forward / Reverse primer 5′-tails (e.g. M13 universal sequences): added to the 5′ end of every primer for compatibility with downstream sequencing or universal-tail PCR strategies. Leave empty if no tail is required.
• Mask repeats & low-complexity regions — see Common parameters → Mask repeats.
• Allow overlapping primer positions: returns multiple alternative primers covering the same window. Useful for finding workable candidates in difficult templates.

A pre-existing primer list (My Primers tab) is checked against every newly designed primer — any new primer that would form a heterodimer with one of yours is rejected.

An SBE probe has its 3′-end fixed one nucleotide before the SNP on the PCR amplicon. After the SAP/ExoI clean-up, the probe is extended by a single fluorescent ddNTP whose colour identifies the allele. The tool generates the probe in both orientations (forward and reverse) and you choose the one with the better local context (no nearby secondary SNPs, balanced Tm, no homopolymer at the 3′-end).

• Probe core length 16–35 nt with the same Tm window as the PCR primers, so the SBE step works under the standard SNaPshot cycling profile.
• Length staggering for CE: probes are sorted by core length and a 5′-tail is appended so adjacent probes differ by ≥4 nt in total length (the typical CE peak resolution limit). The shortest probe gets no tail; the next gets just enough tail to be 4 nt longer; and so on.
• Tail composition: by default the tail is poly-T. If you fill in Forward primer tail / Reverse primer tail, those sequences are also reused as the SBE-probe tail (cycled if shorter than required). Tails are printed in UPPERCASE to visually separate them from the lowercase probe core, e.g. TTTCACacgtacgtacgtacgtac.
• Mobility caveat: tail composition affects CE mobility almost as much as length. Pure poly-dT is the safest baseline; mixed-composition tails (e.g. dGACT) shift mobility differently and may compress two adjacent probes into one peak. If using a non-poly-T tail, increase the minimum length gap or run a CE pre-test on the panel.
• The tool does not account for the 1-nt extension and dye-induced mobility shift after SBE (typically 1–2 nt depending on dye/ddNTP). For very dense panels, add a 1–2 nt safety buffer to the staggering.

The full panel-level SBE probe table appears in the Multiplex Assay Sets tab under the heading === SBE probe panel ===.

Multiplex compatibility is enforced at design time, not as a post-hoc filter. As primers are designed locus-by-locus, every new candidate is checked against the cumulative pool of all previously accepted primers from every other locus. Any primer that would form a heterodimer with an already-accepted primer is rejected and an alternative is chosen.

• For each locus you may see one or several PCR primer pairs in the Multiplex Assay Sets tab.
• Any forward + reverse pair from one locus can be freely combined with any pair from any other locus in a single mPCR. All listed pairs are pre-validated to be cross-locus dimer-free.
• This lets you choose, per locus, the pair with the best Tm match, smallest amplicon, or highest specificity, without re-checking compatibility against the rest of the panel.

Note: cross-locus compatibility is enforced for the PCR primers. The SBE step uses different chemistry and is run after SAP/ExoI removes excess PCR primers and dNTPs, so probe-vs-PCR-primer interactions are not the limiting factor in practice.

Two output tabs serve different purposes:

• Report (Primer list) — every candidate primer and probe found per locus, with full quality metrics. Use this to inspect alternatives or to debug a locus that produced no multiplex set.
• Multiplex Assay Sets — only the validated PCR primer pairs (cross-locus dimer-free) plus the panel-level SBE probe table with length-staggering tails. This is the orderable output.

Column headers in both tabs are tab-separated and Excel-friendly:

ID    Sequence(5'-3')    Length    Tm(°C)    CG(%)    LC(%)    YR(%)    Fragment_Size(bp)/Tm(°C)

• LC% (linguistic complexity) penalises repeats and homopolymers — values ≥70% are recommended for primers.
• YR% reports purine/pyrimidine balance — extremes can predispose to secondary structure.
• Fragment size / Annealing Tm is shown on the reverse-primer row of each PCR pair: the predicted amplicon size in bp and a Tm estimate of the full amplicon (informational, not the cycling Tm).
• Probe ID encodes locus and orientation, e.g. 1F_… = locus 1, forward probe.

When C→T bisulfite conversion is enabled, the input sequence is first virtually converted (every unmethylated C becomes T on each strand independently), and primers/probes are designed against the converted sequence. This is the standard preparation for bisulfite-sequencing assays where the methylation status at a CpG site is read out as a C/T polymorphism after conversion.

• Mark the CpG of interest with [C/T] in the original (unconverted) sequence — this represents methylated vs unmethylated allele after conversion.
• Avoid placing PCR primers over additional CpG sites whenever possible (the tool does not do this automatically — use /.../ markup if needed).
• The two strands diverge after conversion, so the forward and reverse SBE probes are designed independently on each converted strand.

1. Enter sequence(s) — paste a multi-FASTA with each SNP marked as [REF/ALT] (e.g. …flank[C/T]flank…), upload a local FASTA, or retrieve flanking context by rsID from Ensembl.
2. Set parameters — PCR primer length / Tm / amplicon size, optional 3′-end composition pattern, and 5′ mobility-shift tails for the forward and reverse SBE primers (poly-dT or poly-dGACT). Tick Mask repeats to avoid low-complexity flanks; tick C→T conversion for bisulfite-methylation assays.
3. Press Generate — the Report tab lists every candidate PCR primer and SBE probe per locus; the Multiplex Assay Sets tab returns validated, dimer-free PCR + SBE combinations ready for ordering. Pairs from different sequences are pre-validated to be cross-locus compatible, so any F+R pair from one locus can be mixed with any pair from any other locus in the same mPCR.
4. Plan the multiplex — stagger SBE primer lengths by ≥6 nt (using the 5′ tails) so all loci resolve as distinct peaks on capillary electrophoresis.

1. Multiplex PCR — pool all forward and reverse primers from the chosen pairs at equal final concentration (typically 0.05–0.2 µM each). Use a hot-start polymerase and the Tm window reported by the tool as the annealing temperature.
2. SAP/ExoI clean-up — remove unincorporated dNTPs and excess primers (Shrimp Alkaline Phosphatase + Exonuclease I). This is mandatory before the SBE step.
3. SBE reaction — combine the cleaned PCR product with the pooled SBE probes and the SNaPshot reaction mix (contains the four fluorescent ddNTPs and a thermostable polymerase). Cycle per the manufacturer's protocol.
4. Post-extension SAP — dephosphorylate unincorporated fluorescent ddNTPs (otherwise they appear as dye blobs on the trace).
5. Capillary electrophoresis — run with a size standard (e.g. GeneScan 120 LIZ) and analyse with GeneMapper or comparable software. Each locus appears as a coloured peak at the expected probe length.

If two adjacent loci share a peak position in the trace, increase the length-stagger gap or swap one probe to the opposite orientation.

• Paste one target sequence (5′→3′), with or without a FASTA header.
• Mark the variant as [A/G] (slash-separated alleles) or a single IUPAC code such as [R], [Y], [S].
• Single-base substitutions only; indel brackets are reported and skipped.
• Provide enough flanking sequence for the footprint (default 24 nt) so the SNP can sit centrally in the branch domain.

• Probe (C) / Target (X) concentration (µM): set the equilibrium yields and the reported Q.
• Operating temperature (°C): ΔG is computed at this temperature from ΔH/ΔS.
• Total salt / Mg²⁺: entropy salt correction (monovalent-equivalent).
• Probe footprint (nt) and toehold length range (nt): the search space for F and M.
• Target ΔG°rxn (kcal/mol): 0 = thermoneutral = maximal discrimination.

• Probe design: the F/M/R segments, the final C and P strands (5′→3′), a schematic, and the discrimination factor Q.
• Thermodynamics: per-allele ΔG(X·C), ΔG(C·P), ΔG°rxn, ΔΔG and equilibrium yield χ; the forward/reverse toehold ΔG; and the forward-toehold length scan.
• Warnings: SNP placement, toehold-window flags, low Q, self-structure/leak, and the modelling caveats below.

• Equilibrium, not kinetics. Yields and Q are equilibrium values; forward/reverse toehold ΔG are reported as rate proxies (Zhang & Winfree 2009), not a simulation.
• Mismatch thermodynamics are measured. Off-target alleles use internal single-mismatch nearest-neighbour parameters (Allawi & SantaLucia 1997/1998; Peyret et al. 1999; SantaLucia & Hicks 2004), not step-zeroing.
• Salt is approximate. A monovalent-equivalent entropy correction (SantaLucia 1998; von Ahsen 2001 Mg term) is applied; ΔG°rxn is a difference of similar duplexes, so salt largely cancels.
• The clamp is synthetic sequence. Confirm it is free of secondary structure and cross-hybridisation in your assay context.

A synthetic MLPA probe is two oligos that, once ligated, read 5′→3′ as a single PCR template:

LPO (5'->3')  =  [forward tag] + LHS
RPO (5'->3')  =  5'P + RHS + [stuffer] + [reverse tag]

amplicon      =  [forward tag] + LHS + RHS + [stuffer] + [reverse tag]

• LHS / RHS — the target-specific left and right hybridising sequences. Concatenated, they equal a contiguous window of the input strand, so the ligation junction is shown directly in the input, between LHS and RHS; the half-probes anneal to the complementary strand. The LHS provides the 3′-OH and the RHS the 5′-phosphate at the nick.
• Forward tag (LPO 5′-end, default GGGTTCCCTAAGGGTTGGA) and reverse tag (RPO 3′-end, default TCTAGATTGGATCTTGCTGGCAC) — the universal primer sequences. The reverse PCR primer is the reverse complement of the reverse tag (GTGCCAGCAAGATCCAATCTAGA). Both are editable; the defaults are the standard SALSA MLPA primers.
• Stuffer — filler added to the RPO so each amplicon reaches a unique length. Printed in UPPERCASE to separate it from the lowercase target arms. The stuffer is tiled from the source pattern and is not automatically screened for secondary structure or complexity, so inspect a custom or long stuffer (and avoid homopolymers/strong hairpins).

The RPO must be ordered 5′-phosphorylated; without it the ligase cannot seal the nick.

Paste a multi-FASTA with one target region per record — typically an exon or dosage-sensitive segment of ~120–300 nt. The optional [REF/ALT] markup decides where the ligation junction goes.

• Allele-specific (1–4 alleles) — mark the interrogated site and the junction is placed at it, with the allele as the 3′-end of the LHS: a separate LPO is emitted per allele while the RPO is common. Works for SNPs ...gtcaccat[C/T]tatggg... (or the IUPAC ambiguity code [Y] = C/T, [R] = A/G, [N] = all four, …), InDels [CCC/T], and deletions [CCC/] or [CCC/-] (the empty side is the del allele, whose LHS is just the upstream flank). Only the perfectly matched allele ligates. Because alleles can differ in length, their amplicons can differ in size — so InDel alleles resolve by size on CE, while equal-length SNP alleles share one size (tell them apart by ligation/sequence).
• Position marker — a single-token or empty bracket ([G], []) just fixes the junction location; one probe is designed there.
• No bracket — for plain copy-number targets the best repeat-free, G/C-clamped ligation site is chosen automatically.
• Targets can also be retrieved by rsID via Ensembl directly from the input tab — the returned variant is bracketed automatically and becomes the ligation site.

The optional Existing Oligos tab lets you supply oligos that the new probe arms should not dimerise with.

• Hybridising arm length (default 21–30 nt). Each arm grows to the shortest length that reaches the minimum Tm, within this window.
• Min Tm per arm (default 64 °C, nearest-neighbour). Hybridisation/ligation runs ~60 °C overnight, so each arm must melt comfortably above it; MRC-Holland recommends arms several °C above the hybridisation temperature.
• Complexity — a linguistic-complexity floor (LC% ≥ 70) is applied internally to reject low-complexity arms. GC% is reported for each arm but is not constrained; Tm is the governing criterion.
• G/C base pair at the ligation junction (default on) — the 3′ base of the LHS and the 5′ base of the RHS are constrained to G/C to stabilise the nicked junction. The LHS 3′ base is the key ligation-fidelity position; the RHS 5′ base is clamped as well as a conservative default. If no compliant junction exists, the tool relaxes this constraint and flags it.
• G/C clamp at both outer arm ends (off by default) — additionally constrains the 5′-end of the LHS and the 3′-end of the RHS.
• Mask repeats — exclude repeat-masked positions from the arms. A marked [REF/ALT] site is the design target (not avoided); any other brackets in the same record are kept off the arms.
• Universal tags & stuffer source — editable; defaults are the standard SALSA primers and a balanced filler.

If a target is too short, too AT-rich to reach the Tm, or fully repeat-masked, that target is reported as skipped / no-probe rather than forcing a poor design.

Because all probes share one primer pair, they are resolved only by amplicon length. Probes are sorted by their stuffer-free length and a per-probe stuffer is added so each successive amplicon is at least the minimum size spacing longer than the previous one, starting at the minimum amplicon size.

• Amplicon size range (default 130–500 nt) and min size spacing (default 8 nt). Wider spacing is safer; CE peak resolution and length-dependent mobility both degrade as peaks crowd. The 130 nt floor keeps the panel clear of the <120 nt control-fragment band (see note below).
• Amplicon length = forward tag + LHS + RHS + stuffer + reverse tag. With the default tags that is 42 nt + arms + stuffer.
• If the staggered sizes exceed the ceiling, the tool warns; reduce the probe count, shorten arms, or raise the maximum.

Commercial MLPA kits add separate denaturation/quantity control fragments below ~120 nt; leave headroom for them when choosing the minimum size.

Two tab-separated, Excel-friendly outputs:

• Probe Report — per target: the ligation junction position, the LHS and RHS with length/Tm/GC/LC% and coordinates, then the full LPO and RPO sequences to synthesise (tags UPPERCASE, target arms lowercase). For an allele-specific site, one LHS/LPO line is listed per allele above the common RHS/RPO; allele probe IDs are suffixed with the allele — [C], [T], [CCC], [del], …
• Probe Panel — the universal primer pair, a size-sorted table (ProbeID, Locus, Amplicon, LHS, RHS, Stuffer, Ligation@, TmL/TmR), the full ligated probe sequences, and the cross-dimer summary.

Ligation@ is written as a^b — the junction sits between input nucleotides a (LHS 3′-end) and b (RHS 5′-end); for an allele-specific site it reads p(C), the variant position and allele. Equal-length alleles (SNPs) share one amplicon size — distinguish them by allele-specific ligation/sequence (digitalMLPA); length-varying alleles (InDels) resolve by size on CE.

1. Enter targets — paste one region per FASTA record (exon / dosage segment), upload a FASTA, or fetch by rsID from Ensembl. Mark any known variant to avoid with [REF/ALT].
2. Set parameters — arm length / Tm / GC, the amplicon size range and spacing, and (if needed) custom universal tags or stuffer. Keep the junction G/C clamp on for ligase fidelity.
3. Press Design Probes — review the Probe Report per target, then order the LPO (unmodified) and RPO (5′-phosphorylated) oligos from the Probe Panel.
4. Run MLPA — denature DNA, hybridise the probe mix overnight at ~60 °C, ligate, then PCR with the single universal primer pair (label the forward primer, e.g. 6-FAM) and size the products on a CE instrument. Normalise each peak area to reference probes to call copy number.

Always include reference (copy-number-stable) probes and run several normal controls; MLPA reports relative dosage, so calls depend on the reference set and on consistent DNA quality.

• Jan P. Schouten, Cathal J. McElgunn, Raymond Waaijer, Danny Zwijnenburg, Filip Diepvens, Gerard Pals Relative quantification of 40 nucleic acid sequences by multiplex ligation-dependent probe amplification. Nucleic Acids Research 2002, 30(12): e57. doi:10.1093/nar/gnf056
• Principle of MLPA

Paste a multi-FASTA where each record contains one microsatellite with ≥ ~25 nt of unique flank on each side (e.g. paste the region reported by Repeats).

• Auto-detect — the strongest tandem repeat (motif 2–6 nt, ≥ the minimum repeat length) is found, extended to whole copies and normalised to its primitive motif, so (CACACA) is reported as (CA); homopolymer (1 nt) runs are down-weighted as poor markers.
• Marked — wrap the repeat in [..] (e.g. ...acctg[ca]n... or the literal repeat [cacacacaca…]) to fix it explicitly; everything inside the brackets is taken as the repeat.
• Mononucleotide runs and very short flanks may yield no primer pair — widen the size range or supply longer flanks.

Direct dye-labelling of a primer per locus is expensive. The Schuelke (2000) method adds a universal 5′-tail (default M13(-21) TGTAAAACGACGGCCAGT) to the forward primer; a single dye-labelled universal primer equal to the tail is added to the PCR and labels every locus.

• Run the tailed forward primer at ~¼ the reverse concentration plus the labelled universal primer; a two-step anneal (high then low) lets the tail prime in later cycles.
• The reverse primer gets a 5′-PIG-tail (default GTTTCTT) that promotes full non-template +A addition by Taq, sharpening called peaks and avoiding split (−A/+A) alleles.
• Both the M13 tail and the reverse PIG-tail add their length to every product — the reported sizes already include them (PIG-driven +A adds ~1 nt more).
• For multi-colour panels, give each dye channel its own universal tail (e.g. the four tails of Blacket et al. 2012) by editing the tail field.

• Primer length / Tm (default 18–27 nt, 57–63 °C) — a tight Tm window is important for multiplex co-amplification.
• Product size (core, default 90–400 bp) — every Tm-valid combination in range is ranked by amplicon size and Tm match, and the top 3 are reported as alternative pairs; the multiplex panel picks whichever one best avoids a same-dye size collision (not always the best-ranked one). The lengths of both 5′-tails (M13 forward + PIG reverse) are added on top.
• Buffer from repeat — keep the primer 3′-end this many nt away from the repeat edge (avoids slippage).
• Allele range (± repeats) — defines each marker's expected size window (= ± repeats × motif length) for multiplex packing.
• Dye channels — different dyes may share a size range freely (colour tells them apart); markers are packed greedily by size within each channel. Set this to 1 for a single-colour panel, where every product must instead be told apart purely by length.
• Min size gap, one dye (nt) (default 15) — the minimum separation required between product sizes within the same dye channel (10–20 nt is typical for reliable CE resolution). The packer tries each locus's alternative primer pairs to fit; if none clear the gap on any channel, the locus is placed at its least-bad size and flagged ⚠.
• Motif length (nt) (default 2–6, di- to hexanucleotide; the minimum cannot go below 2) — the window scanned by auto-detect; homopolymer (1 nt) runs are down-weighted, as they make poor STR markers.
• Min repeat length (nt) (default 12) — the minimum tandem-repeat tract length to call a microsatellite (also enforced as ≥ 2× the motif length and ≥ 2 copies).
• Non-specific priming control (default on) — keeps both primer arms off dispersed/interspersed repeats elsewhere in the flanking sequence (the microsatellite itself is already excluded by the buffer setting).
• Low complexity priming control (default on) — keeps both primer arms off other SSR/microsatellite and telomere-like stretches in the flanks, the same blocks detected by Repeat Finder. Disable either control only if it prevents a primer from being found and the flanks cannot be replaced.

Always co-load an internal size standard (e.g. GeneScan LIZ) and bin alleles against a reference ladder; report repeat-unit allele names rather than raw bp where a nomenclature exists.

1. Paste/upload target sequences in FASTA.
2. Provide the primer/probe list in the dedicated tab.
3. Run analysis and review both Results and Log output tabs.
4. If no output: confirm hits exist within the allowed product size range, and verify primer orientation (forward/reverse).

These Tm values are computed under the same fixed, standard-PCR-like conditions used by every design tool on this site (see Common parameters → Tm calculation conditions), not LAMP's typically higher Mg²⁺ and primer concentrations. Keeping one condition set makes Tm comparable across tools, but it will read a few degrees lower than a LAMP-specific calculator would report for the same primers — treat it as a relative design target, not the literal isothermal reaction temperature.

• Use [ ... ] to constrain primer design to a specific region.
• Exclude regions with /.../ (e.g. repeats); see Targeting markup.

Enabling C→T bisulfite conversion automatically adjusts the default length (17–28 nt), Tm (57–59°C), and overlapping primer settings — suitable for designing LAMP primers on bisulfite-converted templates for methylation analysis.

• Sequences must be in FASTA format, listed in the desired assembly order.
• Adjacent fragments must share an overlap of ≥20 bp with a Tm ≥50°C to ensure efficient exonuclease digestion and annealing.
• For circular constructs, include vector (backbone) sequence at both the first and last positions. The tool will automatically generate primers spanning the junction.
• Fragment names (FASTA headers) are used in the output to label each primer pair — use meaningful names.

• Min. length (nt): minimum length of the annealing part of the primer (default 18; the overlap tail is added on top of it). There is no maximum length — the primer is extended as far as needed to reach the minimum Tm. If reaching the minimum Tm would push the Tm above the maximum, the primer is instead shortened (even below the minimum length) so that it stays under the Tm ceiling; these cases are flagged in the report Notes.
• Tm range (°C): the maximum Tm is an enforced ceiling — no primer ever exceeds it. The minimum Tm is the target, and the shortest primer reaching it is selected. Default 60–62°C.
• Min. linguistic complexity (%): rejects low-complexity / repetitive primers (≥70% recommended). If no primer can be found at the requested value, the tool automatically relaxes it step-by-step (down to 10%) and notes the value actually used.
• 3′-end pattern: use N for any nucleotide, or specify patterns like WSS to enforce a particular 3′-terminus composition.

• Report (Primer list): all designed primers with name, sequence, length, Tm, GC%, and LC%. A # Notes block at the end flags every fragment where the minimum complexity was relaxed, where the primer was shortened below the minimum length to respect the Tm ceiling, or where no primer could be found — together with the likely reason.
• PCR primer pairs: the forward and reverse primer for each amplification reaction. The annealing portion is in lowercase and the 5′ overlap tail (reverse-complement of the neighbouring fragment end) is in UPPERCASE.
• Assembled-construct amplification pair: a separate, clearly labelled block at the bottom of the PCR primer pairs tab — see the next panel.

At the bottom of the PCR primer pairs tab the tool reports one extra vector-anchored primer pair for amplifying or verifying the whole joined cassette after assembly:

• A forward primer placed anywhere on the first (left) vector arm and a reverse primer anywhere on the last (right) vector arm, both pointing inward across the inserts.
• The position is free — it slides along the arm to find a good primer — so the primer is not forced onto the palindromic cloning site at the fragment end. Where only a short primer is dimer-free, it is reported with a note (it may be below the minimum length).
• An approximate amplicon size is given (left-arm reach + inserts + right-arm reach), with a warning if the two primers might form a primer-dimer.

Tip: provide a sufficiently long vector sequence as the first/last entries. If a vector arm is only a short polylinker of palindromic restriction sites, dimer-free primers may be short or unavailable — extend the vector sequence or move the fragment boundary.

• Target sequence(s) in FASTA; mark the region with […] and exclude problem areas with /.../ (see Targeting markup).
• Amplicon size range: choose by platform (e.g., shorter for Illumina; longer for ONT).
• Gap between amplicons: set to 0 for full coverage; increase for fewer primers.
• Optional pre-designed primers: provide a list for compatibility with existing assays.
• Non-specific priming control (default on) — keeps primers off dispersed/interspersed repeats elsewhere in the flanking sequence.
• Low complexity priming control (default on) — keeps primers off other SSR/microsatellite and telomere-like stretches in the flanks, the same blocks detected by Repeat Finder. Disable either control only if it prevents a primer from being found and the flanks cannot be replaced.

Each target's block in the PCR Primer Pairs tab opens with a Target coverage line, reporting what fraction of the marked target region (not the flanking primer-search window) is spanned by amplicons:

• Panel A / Panel B — each panel's own coverage, on its own. Neither panel is designed to reach 100% alone — that is what the other panel's amplicons are for.
• Combined A+B — the union of both panels' amplicons; this is the number that matters for "did we cover the whole target". 100% combined coverage with two genuinely gapped individual panels is the ideal outcome.
• If combined coverage falls short, widen the amplicon size range, lower Gap between amplicons, or relax the masking/complexity controls above — the usual reasons a stretch has no valid primer on either panel.

• Pick any region within the insert except the Inverted Terminal Repeats (ITRs).
• For genome integrity assessment, target the ends of the insert, e.g., the CMV enhancer at one end and SV-40 at the other.
• Use [ … ] directly inside the pasted sequence to pin each probe's location individually.
• Use /…/ to exclude problematic regions (repetitive elements, secondary structure hotspots) from probe placement; can be repeated multiple times.

• Probe length (35–40 nt): length of each individual oligo probe.
• Probes distance (>5 nt): minimum gap (in nucleotides) separating the two probes along the target strand. The tool also reports pairs at the configured separation.
• Calculated energy: probe self-folding energy should be above −10 kcal/mol to minimize homo- and hetero-dimer formation.
• Linguistic Complexity (LC%): measures sequence vocabulary richness; 100% = maximum diversity. Higher values reduce non-specific hybridization risk.

• Paste or upload sequences in FASTA format; name is optional but recommended (e.g., >EGFP).
• Retrieve a sequence directly from NCBI by entering a nucleotide accession ID (e.g., A02710) and clicking Retrieve Sequence.
• The name and sequence string can be separated by either a space or a tab; style must be consistent for all entries.
• Input is case-insensitive; only standard IUPAC nucleic acid characters are accepted.

• Results are reported as FWD (forward strand, positive strand) and RVS (reverse strand, negative strand) probe pairs in separate tabs.
• Pairs are presented in ranked order; test several top-ranked pairs to determine the most optimal one experimentally.
• Submit the final probe sequences to an oligo manufacturer (e.g., Eurogentec, LGC Biosearch Technologies, Eurofins) with the appropriate fluorescein and biotin modifications specified.

• Paste a single oligo sequence (5′→3′) with or without a FASTA name header.
• Standard IUB/IUPAC degenerate codes are supported — see Input formats → Allowed nucleotide codes.
• U = Uracil (RNA), I = Inosine.
• LNA codes: E=LNA-dA, F=LNA-dC, J=LNA-dG, L=LNA-dT.
• Tm calculations use nearest-neighbor thermodynamic parameters, accounting for primer concentration, salt, and Mg²⁺.

Three calculators are available:

• Dilution: calculate stock volume needed to reach a target working concentration and volume.
• Resuspension: calculate how much TE/water to add to a lyophilised oligo to reach a desired stock concentration.
• Amount from OD/mass/nmol: inter-convert between OD₂₆₀, mass (µg), and nanomoles using the oligo's extinction coefficient and molecular weight.

• Primer concentration (0.01–5 µM): concentration of the oligo used in your reaction.
• Total salt concentration (Na⁺, K⁺, NH₄⁺, Tris⁺; 1–1000 mM): matches your buffer conditions.
• Mg²⁺ concentration (0–10 mM): free magnesium concentration in your reaction.
• Dimer sensitivity (1–10): controls the detection threshold for self-dimer reporting.

• Features: tabular summary — Tm, GC%, LC%, length, base composition, Mw, extinction coefficient, OD values — one row per primer.
• Dimers: self-dimer and cross-dimer analysis for all primer combinations, ranked by binding strength.

PrimersList handles specialized oligonucleotide types:

• Molecular beacons — stem-loop probes; the stem sequence lowers the apparent Tm and affects dimer analysis.
• Scorpion primers — self-probing amplification primers with an internal probe region.
• Standard degenerate primers, RNA oligonucleotides (U), and LNA-modified sequences (E/F/J/L).

• Reverse Complement — the sequence of the opposite strand, read 5′→3′ (what you'd order as a reverse primer).
• Complement (no reverse) — complements each base in place without reversing the order.
• Reverse (no complement) — reverses the letter order only; rarely needed on its own, included for completeness alongside the two above.
• UPPERCASE / lowercase — case conversion only, letters are otherwise untouched.
• C→T Bisulfite (sense / reverse strand) — in silico bisulfite conversion for both strands, same rule as the PCR/qPCR/KASP tools: only cytosines not followed by guanine (non-CpG context) become thymine; CpG sites are preserved.
• FASTA → Column — converts to one name<TAB>sequence row per record (paste straight into Excel).
• → FASTA — converts column or bare input back into standard >name / sequence FASTA records.
• Sort by Length — reorders multi-record input, longest sequence first.
• Length & CG% report — a quick per-record table, no sequence transform.
• Cluster Sequences — groups the input records into homology clusters; needs at least 2 sequences. See Cluster Sequences by Homology below.

All input is cleaned to valid IUPAC nucleotide letters first (see Input formats → Allowed nucleotide codes); a record that has no valid bases after cleaning is dropped.

Fills the Input tab with a fresh random sequence (uniform A/C/G/T composition, no biological meaning) of the given length — default 1000 nt, up to 1,000,000 nt. Useful as quick scratch/test input, e.g. to sanity-check another operation or a downstream tool. This replaces the current Input tab contents, like Load Example.

Groups the input records into homology clusters by the fraction of shared 12-mers — the overlap coefficient of each pair's distinct 12-mer sets, taken relative to the shorter sequence so a length difference doesn't deflate it. Shared long k-mers stay high for genuinely homologous sequences even across substantial divergence and indels, and are ~0 for unrelated sequences, so this reliably separates homologs from noise. Both strands are compared. Clustering is single-linkage (A~B and B~C group A, B and C together). Needs at least 2 sequences. Each cluster is reported as a labelled FASTA block, one record per member, ready to copy straight into another tool or an alignment.

• Shared 12-mers % (default 30, range 5-95) — the minimum percentage of shared 12-mers (relative to the shorter sequence) required to place two sequences in the same cluster. Homologous genes typically share 30–60%; unrelated sequences ~0%. Lower it to catch more diverged homologs, raise it to be stricter. Note this is a shared-k-mer fraction, not a literal alignment percent-identity — a pair at ~85% identity with indels may share only ~20–30% of its 12-mers.
• Sequences that don't match anything closely enough are listed separately under Unclustered, not forced into a cluster.
• A member flagged reverse-strand match (noted on its header line) matched the rest of its cluster on the opposite strand — reverse-complement it before aligning by eye against its cluster-mates; the sequence itself is still reported exactly as given, not auto-flipped.
• Normalising by the shorter sequence means a length difference on its own does not weaken the match (a short homolog still clusters with a much longer one). It compares whole records as given without aligning or trimming, so extra unrelated flanking sequence on the shorter member — which adds non-shared 12-mers — can still lower the score.

Enter a NCBI nucleotide accession (e.g. NC_045512, NM_000546) and choose what to retrieve:

• Fetch FASTA → Input — loads just the sequence into the Input tab, ready for any operation above.
• View GenBank → Result — loads the full GenBank flat-file record (features, annotations, references) into the Result tab to inspect or copy.
• Download GenBank (.gb) — saves the same record straight to a .gb file, without going through the Result tab.

The tool can automatically detect and correct sequence orientation (forward vs. reverse-complement) before alignment. This is especially useful when working with Sanger sequencing reads or assembly contigs that may be reported in either direction.

• Auto-correct orientation (Alignment Parameters) is on by default and applies to DNA/RNA only; protein input is never reverse-complemented.
• Each non-reference sequence is scored against the first sequence in both orientations; if the reverse-complement scores higher, that strand is used.
• Corrected sequences are flagged in the Log/summary tab and tagged above the visual alignment, so the displayed strand is unambiguous.
• Turn the switch off to keep every sequence exactly as entered — useful when the strand is already known and you do not want it changed.

• Visual: colour-coded alignment shown directly in the browser for quick inspection.
• FASTA: aligned sequences with gap characters (-), suitable for downstream phylogenetic or variant analysis.
• CLUSTAL: standard interleaved CLUSTAL format compatible with most alignment viewers.
• The visual view shows a conservation line beneath each block: * marks fully conserved non-gap columns and . marks the rest.
• Output block width controls how many columns are shown per line in the visual output (line wrapping).
• The Log/summary tab reports identity and, for pairwise runs, the alignment score; for local (Smith–Waterman) it also lists the matched region coordinates in each sequence.
• Use Copy to copy the alignment to the clipboard, or Save view to export it exactly as shown (colours preserved) as a standalone HTML file.

• FASTA headers start with >; each header begins a new sequence record.
• Use standard one-letter alphabets (DNA/RNA or protein). Both upper- and lower-case are accepted.
• Automatic cleaning: position numbers and whitespace from GenBank/NCBI-style listings are stripped, case is normalised, and gap symbols (.) are converted to -, so you can paste blocks straight from a record viewer.
• Validation notices: if a sequence contains characters that are not valid for its detected type (DNA/RNA or protein), the offending characters and their positions are listed under Notices in the Log/summary tab.
• Auto mode selects pairwise alignment for 1–2 sequences and multiple sequence alignment for 3+ sequences.
• The sequence-count badge on the input tab updates as records are detected.

• Gap open penalizes starting a gap; gap extend penalizes extending an existing gap.
• Penalties are typically negative (e.g., -10 open, -1 extend). More negative values generally produce fewer/shorter gaps.
• Global aligns end-to-end (Needleman–Wunsch); Local finds best-matching subregions (Smith–Waterman).

• The alignment engine uses dense typed-array dynamic-programming matrices with cached substitution-score lookups, so pairwise alignments of longer sequences run substantially faster than a naive implementation.
• Orientation checks score each strand without building a full traceback matrix, keeping memory low even on long reads.
• The progressive guide tree is built with the nearest-neighbor-chain UPGMA algorithm (O(n²) instead of O(n³)), so adding more sequences scales much better.
• The Time field in the Log/summary tab reports how long the last run took.
• It remains an in-browser tool: very large inputs (many long sequences) are still bounded by browser memory, since the multiple-alignment step holds full matrices for the longest profile.

• Privacy: alignments run locally in JavaScript; sequences are not sent to a server by this page.
• Limitations: a compact, educational implementation inspired by MUSCLE; results may differ from full MUSCLE builds and other aligners.
• If the output looks wrong, try: (i) switching output format, (ii) simplifying input characters, or (iii) adjusting gap penalties.

Non-parametric bootstrap (Felsenstein 1985): alignment columns are resampled with replacement, the tree is rebuilt for each replicate with the same method and model, and each internal branch is labelled with the percentage of replicates that recovered it.

• 100 replicates is a quick default; 500-1000 is usual for publication.
• Values above ~70% are normally treated as reasonable support; low values mean the data do not distinguish that grouping — not that the tool failed.
• Set the count to 0 to skip bootstrapping (much faster on large sets).

Click any node (leaf or internal) to select it, then use the buttons:

• Re-root here — place the root on the branch above the selected node (outgroup rooting). Branch lengths and topology are preserved.
• Rotate — swap the order of a clade's children; cosmetic only.
• Collapse — fold a clade into a triangle; click again to expand.
• Rename — relabel a leaf or annotate an internal node.
• Delete — prune a taxon or a whole clade; the remaining branch lengths are merged so distances stay correct.

Layouts: phylogram (x-axis is evolutionary distance, with a scale bar), cladogram (topology only) and circular. Support values, branch lengths, row spacing and width are toggled in the toolbar.

• Input: FASTA (aligned or unaligned, DNA or protein), or a Newick string to load and edit a tree computed elsewhere.
• SVG — vector, exactly as drawn; opens in Illustrator or Inkscape for figure preparation.
• PNG — raster at 2× for slides and quick sharing.
• Newick / NEXUS — standard tree files (with support values) for MEGA, FigTree, iTOL and similar.

Exports reflect your edits: what you see is what is written out.

• Distance-based inference only. Maximum likelihood and Bayesian methods are not included — they need run times and numerical machinery that do not fit a browser tab. For those, export the alignment and use IQ-TREE, RAxML or MrBayes.
• Saturated pairs are flagged. When sequences are too divergent for the chosen correction (for example p ≥ 0.75 under Jukes-Cantor), the distance cannot be computed. Such pairs are placed beyond the largest measurable distance so a tree can still be drawn, and the run is reported as containing saturated pairs — treat those branch lengths as lower bounds, or switch to p-distance or a protein alignment.
• Alignment quality dominates. A tree can only be as good as the alignment it was built from; check the Alignment tab before trusting the topology.

The defaults suit Sanger-quality reads of a few hundred to a few thousand bases. Three parameters do almost all the work:

• Min. overlap (40 nt) — the shortest alignment accepted as evidence that two fragments belong together. Raise it if unrelated fragments are being joined; lower it only if you know your fragments barely overlap.
• Min. identity (90 %) — lower it for noisy reads; raise it to keep similar-but-distinct sequences (paralogues, gene family members, related strains) in separate contigs.
• Max. unmatched overhang (30 nt) — the repeat filter. In a genuine join, one fragment runs off each end of the alignment; two fragments that merely share an interspersed repeat both continue past the alignment on both sides. Raising this admits more joins and more mistakes.

• Seed word length k (12) — how overlaps are found before they are aligned. A smaller k finds more overlaps in error-rich data at the cost of speed; a larger one is faster and more specific. Fragments shorter than k cannot be seeded at all.
• Min. fragment length (40 nt) — shorter records are reported as dropped rather than silently used.
• Trim from both ends / Quality clip — fixed or quality-driven clipping of unreliable read ends before assembly. Quality clipping needs the Quality tab.
• Search both strands — leave on unless every fragment is already known to be on the same strand.
• Worker threads — see “Speed and threads” below.

When reads disagree at a column, there are two possible reasons and the tool has to tell them apart. An isolated wrong base is sequencing error and should be out-voted. The same minority base seen in several independent reads is polymorphism — a heterozygous site, two alleles of a gene family, two strains in a mixture — and should be preserved.

A column is called polymorphic only when the minority base clears all of:

• at least Min. reads carrying it (2 by default),
• at least Min. minor-allele fraction of the column (25 % by default),
• at least Min. column coverage (3 reads by default), and
• a statistical test: given the sequencing error rate measured from your own data (it is reported in the Log tab), the chance of that many reads carrying that base by error alone must be below roughly one expected false call per assembly.

The last test is what makes the calls trustworthy at higher coverage. Without it, two coincident errors among eight reads reach 25 % of the column and pass the plain fraction rule — on a 30 kb assembly that produced 23 “polymorphic” sites in sequence that had none.

The consensus then carries the standard IUPAC code: R = A/G, Y = C/T, S = G/C, W = A/T, K = G/T, M = A/C, and B, D, H, V, N for three or more alleles. Every such column is also listed in the report with its coverage, the count of each base, and the code.

Insertions and deletions have no IUPAC letter. A minority indel is therefore reported in the same table as an indel variant, with the number of reads on each side, and the consensus takes the majority.

Repeated sequence is the hardest part of any assembly. If two different places in your target contain the same sequence, a fragment that ends inside one copy looks exactly like a fragment that ends inside the other, and joining them splices two unrelated regions into one contig that looks perfectly confident and is wrong.

Three independent checks guard against this:

• Overhang — an alignment that leaves a long unmatched tail on both fragments at both ends is a shared repeat, not a join.
• Repeat content — the tool measures how many times each word occurs across all your fragments, which tells it directly which stretches are single-copy and which are repeated. An overlap that lies almost entirely inside repeated sequence is refused, because nothing distinguishes it from the same repeat elsewhere.
• Branching — if a fragment's end overlaps several fragments that do not overlap each other, the continuation is ambiguous and the contig stops there.

The consequence is deliberate: the assembly breaks at unresolvable repeats instead of guessing, and the report tells you where. Look at the “Rejected overlaps” and “Layout conflicts” tables — a cluster of rejections mentioning repeat content is exactly the boundary of a repeat. Longer reads that span the repeat with unique sequence on both sides are what resolves it.

If you are assembling something you know has no repeats and the tool is splitting it too readily, lower Min. overlap or raise Max. unmatched overhang, and check the rejection reasons to see which filter is firing.

• Assembly report — a run summary (reads in, reads placed, singletons, contigs, N50, mean coverage, elapsed time and threads), the parameters actually used, then tab-separated tables: one row per contig, one per read placement, one per polymorphic site, one per overlap used, one per rejected overlap with its reason, and one per layout conflict. Downloadable as text; the tables paste straight into a spreadsheet.
• Consensus (FASTA) — the contigs, upper-case, with length, reads, cov and variant counts in each header. Unassembled fragments (“singletons”) are appended and marked, unless you turn that off.
• Alignment view — every read shown under its contig's consensus. A dot means the read agrees with the consensus, so disagreements stand out; - is a gap, +/- after the read name is the strand, and polymorphic columns are marked with * and shown in colour. Downloadable as plain text or as a standalone colour HTML page.
• Read placements (TSV) — contig, read, strand, start, end and clipping for every fragment, for downstream scripting.
• Log / summary — the short version, plus the seeding statistics, the measured error rate, and any notices.

Almost all the time goes into aligning candidate fragment pairs, and those alignments are completely independent of one another, so that stage — and only that stage — is spread across Web Workers. Seeding, layout and consensus stay on the page but run in short slices, so the tab keeps responding either way.

Measured on 300 reads of 600–900 nt covering 30 kb: 3.6 s on one thread, 2.4 s on two, 1.8 s on four, 1.5 s on eight. The speed-up flattens because the stages that stay serial take about 1 s of that, however many threads you add. The output is byte-identical whatever you choose.

Small jobs stay single-threaded on purpose — handing the work to other threads would cost more than the alignments it saves. Choosing an explicit thread count overrides that. A page opened straight from disk (file://) cannot start Workers at all and falls back to one thread automatically; the Log tab always reports how many were really used.

• Scale. This is intended for tens to a few hundred fragments, not for whole-genome shotgun data. Large inputs are accepted but the pairwise stage grows with the number of candidate pairs.
• No paired-end or mate-pair information. Repeats longer than your fragments cannot be resolved by any method without long-range links, and the tool stops rather than guessing.
• Contig orientation is arbitrary. With no reference, a contig may come out reverse-complemented relative to what you expect; that is not an error. An R in one orientation is a Y in the other.
• Nothing is uploaded. The whole assembly runs in your browser.
• Everything is one contig and it should not be — raise Min. identity, raise Min. overlap, or lower Max. unmatched overhang.
• Nothing joins — check the Rejected overlaps table for the reason. Usually the fragments overlap by less than Min. overlap, or are more divergent than Min. identity allows. Fragments shorter than the seed word length k can never be joined.
• Expected polymorphism is missing — check the coverage at that column in the Alignment view. Two reads cannot establish a minority allele; the thresholds and the statistical test both need enough depth.

1. Paste or upload one or more sequences in FASTA format.
2. Select the repeat types to detect and set minimum repeat unit / copy number thresholds.
3. Click Analyze. Results are reported per sequence with coordinates and consensus repeat units.
4. Use the masking output to generate a repeat-masked FASTA for primer design tools (paste it into PCR / LAMP / KASP tool inputs).

Repeat-rich regions are a major source of primer design failures. The recommended workflow is:

1. Run TotalRepeats on your target sequence to identify repeat coordinates.
2. Export the masked sequence (repeat regions in lower case) or manually add /…/ exclusion markup around identified repeat coordinates.
3. Paste the masked/marked sequence into the PCR, KASP, LAMP, or Assembly tool for repeat-aware primer design.

The Picture tab draws a map of the detected repeat blocks and clusters along the sequence on a zoomable canvas.

• Zoom: scroll the mouse wheel over the map (zoom centres on the cursor).
• Pan: click and drag left or right.
• Double-click: recentres and resets the view to the default zoom, showing the whole sequence from the start — use it whenever the map has drifted off-centre.

Two buttons below the map save the picture:

• Save Image — exports the current view as a PNG raster image (map.png).
• Save SVG — exports the current view as a scalable SVG vector file (map.svg) that stays sharp at any zoom and can be edited in vector-graphics software.

Both buttons capture the current zoom and pan, so double-click first to reset the view if you want the entire map in the saved file.

The browser-based tool is suitable for sequences up to 100 Mb. For whole-genome analyses, use the command-line TotalRepeats Java application, which supports unlimited input size.

All reagent labels and concentrations are editable. Available slots include:

• PCR buffer (label editable)
• MgCl₂ or MgSO₄ (mM)
• dNTP mix (mM)
• Forward and reverse primers (µM) — or FIP/BIP for LAMP
• Up to 4 additional probes or primers (e.g., F3/B3, loop primers for LAMP)
• An extra component slot with a custom unit label (e.g., Betaine, enzyme enhancer)
• DMSO (%), ROX reference dye (×), SYBR Green II (×)
• Primary polymerase (U/µl) and optional secondary Pfu polymerase (U/µl)
• DNA template volume (µl)

• Set a final concentration of 0 for any component you do not need — it will be excluded from the output.
• Include a 5–10% overage by entering a slightly higher number of reactions (e.g., 11 instead of 10) to account for pipetting losses.
• For LAMP, the preset loads the full 6-primer set (FIP, BIP, F3, B3, FLOOP, BLOOP) with recommended concentrations and isothermal buffer.
• Export the result table with Ctrl+A → Ctrl+C → paste into Excel or your lab notebook.

• Units must be consistent across all concentration fields (all in mM, all in %, etc.) and across all volume fields (all in mL, µl, etc.).
• pH mixing uses a linear approximation (C₁·V₁ + C₂·V₂ = C_f·V_f), valid only for rough educational estimates; true pH requires accounting for buffer capacity and activity coefficients.
• The tool assumes ideal mixing (volumes are additive). In practice, some solutions (e.g., concentrated sulfuric acid) show significant volume changes on mixing.
• The formula applied is the standard mixing equation: C₁V₁ + C₂V₂ = CfVf.

• Reduce the probe distance parameter if the target region is short — a very large required gap can exhaust available candidates.
• Confirm the input sequence is long enough: two non-overlapping 35–40 nt probes plus the required distance gap requires at least 75–80 nt of usable target sequence.
• Remove or narrow any /.../ exclusion regions — they may be blocking too much of the target.
• Verify you are not targeting the ITR region, which is excluded by design.
• Try slightly expanding the allowed probe length range (e.g., 33–42 nt) if your probe design criteria are flexible.

• Confirm all final concentrations and the template volume are filled in correctly — any missing entry is treated as 0 and excluded.
• The calculator uses the reaction volume field as the total. Water (or buffer) is added automatically to reach that total; if the components already exceed the reaction volume, a negative water volume will be shown.
• Check that stock concentrations are in the correct units — stock and final concentrations must use the same unit system (both in µM, or both in mM, etc.).

• Confirm exactly 4 fields are filled and 2 are left empty. Providing 5 or 6 values over-constrains the system.
• For pure dilution (adding water/buffer), set C₂ = 0, not blank — blank means "unknown".
• If a calculated volume is negative, the combination of concentrations and volumes is physically impossible (e.g., the desired final concentration is higher than both stock concentrations).
• Ensure all concentration values use the same units and all volume values use the same units.

If the page suddenly stops working after a server update (buttons do nothing, tabs don't switch, no results), the browser is likely using a cached JavaScript file.

• Confirm the ID is correct (e.g., NCBI nucleotide accession like A02710; rsIDs like rs4988235).
• Network/firewall policies may block external API calls; use manual FASTA upload instead.
• Try again later if the upstream service is rate-limiting or temporarily unavailable.

• Enable Non-specific priming control and increase the minimum complexity threshold.
• Run TotalRepeats on your target to identify repeat coordinates, then add /.../ exclusion markup around them.
• For multiplex panels: tighten the primer Tm window.

• Ensure sequences are in the correct assembly order in the FASTA file.
• Verify that adjacent fragments share ≥20 bp overlapping sequence with a Tm ≥50°C for exonuclease processing.
• If using vector sequences at the ends, confirm they contain the appropriate homology regions matching the adjacent insert ends.
• Widen the Tm window (e.g., 58–64°C) or increase the maximum primer length to allow more candidate primers.

• Try increasing the Max F2-B2 amplicon size (default 200 bp; try up to 350 bp).
• Widen the Tm range slightly (e.g., 58–63°C) to allow more inner-primer candidates.
• Disable Non-specific priming control temporarily to confirm a set can be generated on the current sequence, then re-enable it.
• Use [ ... ] markup to direct the tool to the most unique subregion of a long target sequence.