Restriction and ligation: GFP amplicon insert into pUC19

Amplify GFP with a recognition site on each primer's 5' tail. Cut pUC19 and GFP amplicon with HindIII and SphI, gel-purify the 2684 bp backbone and the 719 bp insert, ligate them, and confirm the clone by colony PCR and sequencing.

Vector: pUC19, 2686 bp
Insert: GFP, 719 bp, amplified
Enzymes: HindIII-HF (R3104) and SphI-HF (R3182)
Backbone: 2684 bp
Junctions: GCATGC and AAGCTT
Product: pUC19-GFP, 3403 bp
Selection: ampicillin or carbenicillin

Keep in mind

One ligation joins two fragments: pUC19 backbone (2684 bp) and GFP amplicon insert (719 bp).

No enzyme was named, so the pair was chosen: HindIII and SphI. 89 other pairs were refused, 75 on the vector site rule, 12 on the insert site rule and 2 on the backbone rule. The pair that got furthest was KpnI and XmaI, and the backbone's two ends anneal to each other (two 3' GTAC ends), so it closes on itself and the insert could go in either way round; the chooser passes over such a pair, and naming it plans it with a dephosphorylation instead.

The insert is amplified rather than cut out, so its two ends come from its primers -- forward primer, 6 spacer bases and the SphI site; reverse primer, 6 spacer bases and the HindIII site. The spacer is what lets an enzyme cut a site that close to the end of a fragment, and the 741 bp amplicon is digested in place of a plasmid.

This method's junction is not scarless: the two ends came from a recognition site and ligating them puts it back, so the product reads GCATGC at 442 (SphI) and AAGCTT at 1165 (HindIII).

GFP reads on the opposite strand from lac promoter, 65 bp away, so that promoter does not transcribe it.

There is no ribosome binding site annotated ahead of GFP, so the clone is not expected to make its protein.

The insertion interrupts lacZα, so correct clones are white and empty vector is blue on X-gal and IPTG.

bufferpass
digest temperaturepass
digest clean-uppass
methylationpass
self-ligationpass
pUC19 backbonepass
GFP amplicon insertpass
junctionspass
reading framepass
ligation ratiopass
diagnostic digestpass
primerswarn

primers warn: 6 designed, 2 with a warning, 0 failing: GC on 2; not judged: full-primer Tm, 3' end stability

Materials

Name	Supplier	Catalogue	Storage	Per run	Note	Source
pUC19 plasmid			-20 °C		cut to make the backbone
GFP fragment			-20 °C		template for the PCR
Q5 DNA Polymerase and its reaction buffer	New England Biolabs		-20 °C
dNTP mix			-20 °C		10 mM of each base
HindIII-HF	New England Biolabs	R3104	-20 °C		cuts the vector and the insert once each, leaving a 5' overhang
SphI-HF	New England Biolabs	R3182	-20 °C		cuts the vector and the insert once each, leaving a 3' overhang
rCutSmart Buffer, 10X	New England Biolabs		-20 °C		both enzymes are supplied in it, so they digest together in it
T4 DNA Ligase and its 10X reaction buffer	New England Biolabs	M0202	-20 °C		thaw a fresh buffer aliquot: its ATP goes over freeze-thaws
Agarose, 1X TAE or TBE, and a DNA stain
NEB 1 kb Plus DNA Ladder	New England Biolabs	N3200				^[1]
NEB 100 bp DNA Ladder	New England Biolabs	N3231				^[2]
Monarch Spin DNA Gel Extraction Kit	New England Biolabs	T1120			recovers each band off the gel
Monarch Spin PCR & DNA Cleanup Kit	New England Biolabs	T1130
NEB 5-alpha Competent E. coli (C2987)	New England Biolabs		-80 °C	50 µL per transformation
SOC or NEB 10-beta/Stable Outgrowth Medium				950 µL per transformation
LB agar plates with ampicillin or carbenicillin, 80 µg/mL X-gal and 200 µM IPTG				one plate per transformation		^[3]
Plasmid miniprep kit					for the clones that go to sequencing
OneTaq Quick-Load 2X Master Mix with Standard Buffer	New England Biolabs	M0486	-20 °C	12.5 µL per reaction

Equipment: Thermocycler with a heated lid, Agarose gel rig, power supply and a gel imager, Scalpel or gel cutting tips, and a clean cutting surface, Microcentrifuge, Spectrophotometer (NanoDrop) or fluorometer (Qubit), Heat block or water bath at 42 °C, Shaking incubator and a plate incubator at 37 °C

Oligos

Order sheet: primers.tsv

Name	Sequence (5'→3')	Length	Tm (°C)	For	Working stock	Checks
GFP forward	`AAACACGCATGCATGAGTAAAGGAGAAGAACTTTTCAC`	38	62.1	Amplify GFP	10 µM	warn
GFP reverse	`AAACACAAGCTTCTATTTGTATAGTTCATCCATGCCAT`	38	62.0	Amplify GFP	10 µM	warn
Colony PCR forward	`GCCAGTGAATTCGAGCTCGGTAC`	23	61.9	Screen colonies by PCR	10 µM	pass
Colony PCR reverse	`AGCGGATAACAATTTCACACAGGAAACAGC`	30	62.1	Screen colonies by PCR	10 µM	pass
Sequencing forward	`GTGCTGCAAGGCGATTA`	17	62.8	Confirm the clone by sequencing	10 µM	pass
Sequencing reverse	`CGCAACGCAATTAATGTGAG`	20	62.2	Confirm the clone by sequencing	10 µM	pass

Checks on 2 of 6 oligos

GFP forward warn: GC 35% (band 40-60)
GFP reverse warn: GC 35% (band 40-60)

Make the fragments

1Amplify GFP

Caution: Keep the polymerase on ice.

Thaw the buffer, dNTPs and primers on ice, then vortex and spin them down.
Mix the master mix, dispense it into each tube, then add the GFP template.
Run the program below: 63 °C annealing and 20 s extension for a 741 bp product.

GFP PCR

Reactions mix includes 10% extra

Component	Stock	Final	1 rxn (µL)	Mix for 1 (µL)
Q5 Reaction Buffer	5X	1X	10	11
dNTP mix	10 mM each	200 µM each	1	1.1
Forward primer	10 µM	500 nM	2.5	2.75
Reverse primer	10 µM	500 nM	2.5	2.75
Template DNA			1	each tube
Q5 DNA Polymerase	2 U/µL	1 unit	0.5	0.55
Nuclease-free water		to 50 µL	32.5	35.8
Total			50	53.9 mix only

Put 49 µL of mix in each tube, then add 1 µL Template DNA.

GFP PCR · 27 min 30 s plus ramps

Step	Temperature	Time	Cycles
Initial denaturation	98 °C	30 s	1
Denature	98 °C	10 s	×30^[4]
Anneal	63 °C	20 s
Extend	72 °C	20 s
Final extension	72 °C	2 min	1
Hold	4 °C	∞	1

GFP PCR

Expected result

One band at 741 bp.

Troubleshooting

No band: Drop the annealing temperature by 3 °C and check the GFP template is there.^[4]
Several bands: Raise the annealing temperature, or gel-purify the band of the right size.^[4]

Notes

Each primer's 5' tail is a spacer and a recognition site, SphI on the forward end and HindIII on the reverse end. The tail is not on the template, so it does not anneal in the first cycles and the annealing temperature above is read from the annealing regions alone.
The spacer is what lets the enzyme cut a site this close to the end of a fragment: cleavage is measured at one to five bases, and six is the answer for an enzyme the table does not list.^[5]

2Check the PCRs on a gel

Use a 2% agarose gel.
Load 5 µL of each reaction beside the ladder.
Run until the dye front is two thirds down the gel.

Expected result

GFP: one band at 741 bp.

PCR products

L NEB 100 bp DNA Ladder (N3231)
1 GFP: 741 bp

Troubleshooting

A smear or an extra band: Gel-purify the band of the right size; a wrong template in the assembly gives wrong clones.^[6]

Notes

2% agarose resolves bands of these sizes.^[7]

3Purify every amplicon

Caution: Keep the eluate under 25% of the digest below; it carries salt.^[8]

Run each reaction over a Monarch Spin PCR & DNA Cleanup column and elute in the smallest volume the kit allows.

Expected result

Clean DNA, free of polymerase, primers and dNTPs.

Troubleshooting

Low recovery: Elute in a larger volume and leave the buffer on the column longer.^[9]

Notes

A column recovers 70% to 90% of the reaction and takes the polymerase, the primers and the dNTPs away, so the digest cuts the amplicon and nothing else.^[9]
Nothing further has to take GFP away: it reads neither enzyme's site, so its ends cannot join the backbone. A DpnI digest earns a step only against a Dam-methylated plasmid template.

Cut and recover

4Digest pUC19 with HindIII and SphI

Caution: Keep the DNA solution under 25% of the reaction; a column eluate carries salt, and salt leaves the digest incomplete.^[8]

Caution: Keep the enzymes under 10% of the reaction, which keeps glycerol under 5%.^[8]

Caution: Use the buffer the supplier supplies each enzyme in wherever you can.^[8]

Caution: Use the fewest units and the shortest incubation that digest completely.^[8]

Caution: Keep the DNA free of organic solvents, and use Mg²⁺ rather than another metal.^[8]

Mix the reaction below, 1000 ng of pUC19 first.
Incubate at 37 °C for 60 minutes.

pUC19 digest

Reactions mix includes 10% extra

Component	Stock	Final	1 rxn (µL)	Mix for 1 (µL)
pUC19^[10]		0.604 pmol (1000 ng)	1	each tube
10X rCutSmart Buffer^[11]	10X	1X	5	5.5
HindIII-HF (R3104)^[11]		10 units	1	1.1
SphI-HF (R3182)^[11]		10 units	1	1.1
Nuclease-free water		to 50 µL	42	46.2
Total			50	53.9 mix only

Put 49 µL of mix in each tube, then add 1 µL pUC19.

pUC19 digest

Expected result

pUC19 backbone: 2684 bp, which is what goes on (HindIII and SphI ends).
pUC19 offcut: 2 bp.

Troubleshooting

An uncut band remains: Add more units or incubate longer, and check the enzymes' methylation sensitivity against the strain the plasmid was grown in.^[6]
Extra bands: Star activity: use fewer units, a shorter incubation and the supplied buffer.^[6]

Notes

Both enzymes are supplied in rCutSmart Buffer, which is the rule for digesting two of them together.^[12]

5Digest GFP amplicon with HindIII and SphI

Caution: Keep the DNA solution under 25% of the reaction; a column eluate carries salt, and salt leaves the digest incomplete.^[8]

Caution: Keep the enzymes under 10% of the reaction, which keeps glycerol under 5%.^[8]

Caution: Use the buffer the supplier supplies each enzyme in wherever you can.^[8]

Caution: Use the fewest units and the shortest incubation that digest completely.^[8]

Caution: Keep the DNA free of organic solvents, and use Mg²⁺ rather than another metal.^[8]

Mix the reaction below, 1000 ng of GFP amplicon first.
Incubate at 37 °C for 60 minutes.

GFP amplicon digest

Reactions mix includes 10% extra

Component	Stock	Final	1 rxn (µL)	Mix for 1 (µL)
GFP amplicon^[10]		2.19 pmol (1000 ng)	1	each tube
10X rCutSmart Buffer^[11]	10X	1X	5	5.5
HindIII-HF (R3104)^[11]		10 units	1	1.1
SphI-HF (R3182)^[11]		10 units	1	1.1
Nuclease-free water		to 50 µL	42	46.2
Total			50	53.9 mix only

Put 49 µL of mix in each tube, then add 1 µL GFP amplicon.

GFP amplicon digest

Expected result

GFP amplicon insert: 719 bp, which is what goes on (SphI and HindIII ends).

Troubleshooting

An uncut band remains: Add more units or incubate longer, and check the enzymes' methylation sensitivity against the strain the plasmid was grown in.^[6]
Extra bands: Star activity: use fewer units, a shorter incubation and the supplied buffer.^[6]

Notes

Both enzymes are supplied in rCutSmart Buffer, which is the rule for digesting two of them together.^[12]
The two tails come off as 11 and 11 bp ends. They run off the bottom of the gel and neither can close the circle, so the insert is the only band to cut out.

6Separate the digests on a gel and recover the two fragments

Caution: Keep the gel off the ultraviolet box for as long as you can; use blue light where there is one.

Use a 1% agarose gel and load each whole digest beside the ladder.
Run until the bands below are apart.
Cut out the 2684 bp pUC19 backbone band and the 719 bp GFP amplicon insert band.
Recover each slice on a Monarch Spin DNA Gel Extraction column and elute in the smallest volume the kit allows.

Expected result

pUC19: 2684 bp and 2 bp.
GFP amplicon: 719 bp.
Each slice recovers 70% to 90% of the DNA that was in it.
2 bp runs past the bottom of this ladder.

Digests

L NEB 1 kb Plus DNA Ladder (N3200)
1 pUC19: 2684, 2 bp
2 GFP amplicon: 719 bp

Troubleshooting

Two bands did not separate: Run the gel further, or use a percentage that resolves that size range.^[13]
Little DNA comes off the column: Elute in a larger volume, leave it on the column longer, and keep the agarose percentage low.^[13]

Notes

A gel is what separates the two pieces of each digest from one another, so a piece that is not wanted cannot religate into the backbone.
It also takes the enzymes away, so nothing has to be heat inactivated before the ligation.^[14]
1% agarose resolves bands of these sizes.^[7]

7Measure every concentration

Measure each purified amplicon by A260 (NanoDrop) or with a fluorometer (Qubit).
Enter each concentration in the next table's Stock column.

Expected result

pUC19 backbone: 0.02 pmol is 33.06 ng, so 33 ng/µL or more fits in 1 µL.
GFP amplicon insert: 0.06 pmol is 26.57 ng, so 27 ng/µL or more fits in 1 µL.

Troubleshooting

Too dilute to fit in the reaction: Concentrate the amplicon, or scale the reaction up.

Assemble

8Ligate the insert into the backbone

Caution: Keep the two fragments together at 1 to 10 ng/µL. Below that a fragment closes on itself instead of joining its partner.^[15]

Mix the reaction below, the DNA first and the ligase last.
Hold at 25 °C for 10 minutes, or overnight at 16 °C.
Then run the heat step below, and chill on ice.

Ligation, T4 DNA Ligase (NEB #M0202)

Reactions mix includes 10% extra

Component	Stock	Final	1 rxn (µL)	Mix for 1 (µL)
pUC19 backbone^[10]	ng/µL	0.02 pmol (33.06 ng)	1	each tube
GFP amplicon insert^[10]	ng/µL	0.06 pmol (26.57 ng)	1	each tube
T4 DNA Ligase Buffer^[15]	10X	1X	2	2.2
T4 DNA Ligase^[15]	400 U/µL	400 units	1	1.1
Nuclease-free water		to 20 µL	15	16.5
Total			20	19.8 mix only

Put 18 µL of mix in each tube, then add 1 µL pUC19 backbone, 1 µL GFP amplicon insert.

Ligation · 20 min plus ramps

Step	Temperature	Time	Cycles
Ligate	25 °C	10 min	1^[15]
Heat inactivation	65 °C	10 min	1^[15]

Ligation

pUC19-GFP as the ligation closes it: the GCATGC and AAGCTT junctions it spells, and where HindIII and SphI cut it.

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Expected result

The product carries GCATGC at 442, joining pUC19 backbone to GFP amplicon insert.
The product carries AAGCTT at 1165, joining GFP amplicon insert to pUC19 backbone.
1 to 5 µL of this goes into the cells; the rest keeps at -20 °C.

Troubleshooting

No colonies later: At least one fragment has to carry a 5' phosphate; vary the ratio, and use a fresh buffer aliquot, because its ATP goes over freeze-thaws.^[6]
Empty vector on the plate: The backbone band carried some uncut plasmid; run the gel further and cut the band clean.^[16]

Notes

Picomoles, not nanograms: the table asks for a molar ratio, and the shorter fragment weighs less at the same ratio.
Junction positions are 1-based, on the product.

Transform and screen

9Transform and plate

Caution: Competent cells die if they warm up; keep them on ice until the shock.

Thaw 50 µL of NEB 5-alpha Competent E. coli (C2987) on ice for 10 minutes.
Add 2 µL of the assembly and flick the tube four or five times.
Hold on ice for 30 minutes.
Heat shock at 42 °C for 30 seconds.
Return to ice for 5 minutes.
Add 950 µL of outgrowth medium and shake at 37 °C for 60 minutes at 250 rpm.
Spread 50 µL of a 1:5 dilution on a warmed plate and grow overnight at 37 °C.

ThawOn iceHeat shockBack on iceOutgrowth

Expected result

No supplier states a colony count for this method, so run the controls below and read the plate against them rather than against a number.
Correct clones are white and empty vector is blue: the insertion interrupts lacZα, which is then not there to complete the host's own.
Uncut vector, 100 pg to 1 ng: the cells are viable and the antibiotic is right.^[6]
Cut vector, no ligase: under 1% of the colonies the uncut vector gave.^[6]
Vector-only ligation: the same as the cut-vector control, its ends being unable to rejoin.^[6]
A transformation efficiency under 10⁴ cfu/µg means the cells, not the ligation.^[6]

Troubleshooting

No colonies: Check the antibiotic and the cells' efficiency, and plate the rest of the outgrowth.^[6]
A lawn: Plate a smaller volume or a greater dilution next time.

Notes

The plate reads colour only with an alpha-complementing host, such as NEB 5-alpha Competent E. coli (C2987). A host that cannot complement gives white colonies whatever the clone carries.
The clone is not expected to make GFP: it reads on the opposite strand from lac promoter and no ribosome binding site is annotated ahead of it.

10Screen colonies by PCR

Touch a well-separated colony with a sterile toothpick and stir it into the tube until the liquid clouds.
Streak the same toothpick onto a numbered plate, so the clone survives the PCR.
Run the program below and load 5 µL on a 2% gel.

Colony PCR

Reactions mix includes 10% extra

Component	Stock	Final	1 rxn (µL)	Mix for 1 (µL)
OneTaq Quick-Load 2X Master Mix with Standard Buffer (M0486)	2X	1X	12.5	13.8
Forward primer	10 µM	200 nM	0.5	0.55
Reverse primer	10 µM	200 nM	0.5	0.55
Nuclease-free water		to 25 µL	11.5	12.7
Total			25	27.5

Put 25 µL of mix in each tube.

Colony PCR · 1 h 10 min plus ramps

Step	Temperature	Time	Cycles
Lysis	94 °C	5 min	1
Denature	94 °C	30 s	×30^[17]
Anneal	56.9 °C	30 s
Extend	68 °C	1 min
Final extension	68 °C	5 min	1
Hold	10 °C	∞	1

Colony PCR

Expected result

Each of the 2 junctions is read: the flanking pair crosses them all.
Correct clone: 828 bp.
Empty vector: 111 bp.

Colony PCR

L NEB 100 bp DNA Ladder (N3231)
1 Correct clone: 828 bp
2 Empty vector: 111 bp

Troubleshooting

No band in any lane: The colony was too much material: touch a smaller one, or dilute it.^[18]

Notes

The long first step at 94 °C lyses the cells; there is no purified template.
The insert's two ends do not anneal to each other, so it cannot go in the other way round and there is no reversed lane to read.

11Check a miniprep by digesting it with HindIII and SphI

Miniprep two or three of the colonies the PCR called correct.
Mix the reaction below, 1000 ng of miniprep first.
Incubate at 37 °C for 60 minutes.
Run the whole digest on a 1% agarose gel beside the ladder.
Sequence only a miniprep that gives the clone's bands.

Diagnostic digest

Reactions mix includes 10% extra

Component	Stock	Final	1 rxn (µL)	Mix for 1 (µL)
pUC19-GFP^[10]		0.477 pmol (1000 ng)	1	each tube
10X rCutSmart Buffer^[11]	10X	1X	5	5.5
HindIII-HF (R3104)^[11]		10 units	1	1.1
SphI-HF (R3182)^[11]		10 units	1	1.1
Nuclease-free water		to 50 µL	42	46.2
Total			50	53.9 mix only

Put 49 µL of mix in each tube, then add 1 µL pUC19-GFP.

Diagnostic digest

Expected result

A correct clone: 2684 bp and 719 bp.
A colony carrying pUC19 instead: 2684 bp and 2 bp.
2 bp runs past the bottom of this ladder.

Diagnostic digest

L NEB 1 kb Plus DNA Ladder (N3200)
1 pUC19-GFP: 2684, 719 bp
2 pUC19: 2684, 2 bp

Troubleshooting

Every miniprep gives the vector's bands: The background is uncut or religated vector; run the controls under the transformation to find which.^[6]
One band, at the plasmid's full length: Only one site cut. Check the enzymes' methylation sensitivity against the strain the miniprep was grown in.^[6]

Notes

The junctions put both recognition sites back, so the pair that made the clone is what cuts the insert out of it again.
1% agarose resolves bands of these sizes.^[7]

12Confirm the clone by sequencing

Miniprep two or three colonies that read as correct.
Send each miniprep with both sequencing primers.
Check each read against product.dna; keep a clone only when every junction and insert reads true.

Expected result

Sequencing forward anneals 100 bp from its own junction and has to read 823 bp to cover the far one.
Sequencing reverse anneals 123 bp from its own junction and has to read 846 bp to cover the far one.
The junctions read as GCATGC and AAGCTT, and the parts match GFP amplicon insert.

Troubleshooting

The read starts too close to the junction: Move the primer further out; the first bases after a primer are unreadable.^[19]

Notes

Only a read across the junctions confirms the assembly, whatever the screen said.
A provider whose read is shorter than the lengths above needs a further primer inside the inserts.

Sources

New England Biolabs #N3200 1 kb Plus DNA Ladder product page · capture 2026-03-07 · Wayback Machine · docs/research/primer-design-and-pcr.md cited at band sizes
New England Biolabs #N3231 100 bp DNA Ladder product page · capture 2025-10-31 · Wayback Machine · docs/research/primer-design-and-pcr.md cited at band sizes
Potapov, V. et al. (2018) Comprehensive profiling of four base overhang ligation fidelity by T4 DNA Ligase and application to DNA assembly. ACS Synth. Biol. 7, 2665-2674 · docs/research/golden-gate-assembly.md https://doi.org/10.1021/acssynbio.8b00333 cited at methods
New England Biolabs #M0491 Q5 High-Fidelity DNA Polymerase protocol page · capture 2024-08-04 · Wayback Machine · 2026-09-12 cited at thermocycling conditions · annealing temperature
New England Biolabs usage guideline, Cleavage Close to the End of DNA Fragments · 2026-09-18 · docs/research/restriction-ligation.md https://www.neb.com/en-us/tools-and-resources/usage-guidelines/cleavage-close-to-the-end-of-dna-fragments cited at cleavage close to the end
New England Biolabs, Troubleshooting Guide for Cloning · 2026-09-18 · docs/research/restriction-ligation.md https://www.neb.com/en-us/tools-and-resources/troubleshooting-guides/troubleshooting-guide-for-cloning cited at colonies contain the wrong construct · incomplete restriction enzyme digestion · extra bands · few or no transformants · transformation controls · too much background
New England Biolabs, Agarose Gel Resolution · capture 2024-09-18 · Wayback Machine · docs/research/primer-design-and-pcr.md cited at optimum resolution for linear DNA
New England Biolabs, Restriction Endonuclease Technical Guide · version 5.0 - 7/17 · neb-online.de mirror · docs/research/restriction-ligation.md https://www.neb-online.de/literatur/pdf/Restriction_Endonuclease_Technical_Guide.pdf cited at salt inhibition, p. 10 · avoiding star activity, p. 5
New England Biolabs #T1130 Monarch Spin PCR & DNA Cleanup Kit instruction manual · version 1.0 06.24 · 2026-09-18 · docs/research/restriction-ligation.md cited at troubleshooting, low DNA yield · typical recovery
New England Biolabs NEBioCalculator · script main-6f3020c533.js, capture 2026-08-28 · Wayback Machine · docs/research/primer-design-and-pcr.md cited at dsDNA molecular weight
New England Biolabs usage guideline, Optimizing Restriction Endonuclease Reactions · 2026-09-18 · docs/research/restriction-ligation.md https://www.neb.com/en-us/tools-and-resources/usage-guidelines/optimizing-restriction-endonuclease-reactions cited at typical reaction conditions
New England Biolabs usage guideline, Double Digests · 2026-09-18 · docs/research/restriction-ligation.md https://www.neb.com/en-us/tools-and-resources/usage-guidelines/double-digests cited at choosing a buffer
New England Biolabs #T1120 Monarch Spin DNA Gel Extraction Kit instruction manual · version 2.0 10.25 · 2026-09-18 · docs/research/restriction-ligation.md cited at agarose concentration, p. 6 · troubleshooting, low DNA yield
New England Biolabs usage guideline, Heat Inactivation · capture 2021-04-20 · Wayback Machine · 2026-09-18 · docs/research/restriction-ligation.md https://www.neb.com/en-us/tools-and-resources/usage-guidelines/heat-inactivation cited at enzymes heat does not inactivate
New England Biolabs #M0202 protocol, Ligation Protocol with T4 DNA Ligase · 2026-09-18 · docs/research/restriction-ligation.md https://www.neb.com/en-us/protocols/dna-ligation-with-t4-dna-ligase-m0202 cited at General Guidelines, DNA · protocol
New England Biolabs FAQ, How can I reduce the number of vector-only background colonies? · capture 2019-11-23 · Wayback Machine · 2026-09-18 · docs/research/restriction-ligation.md cited at reducing vector-only background colonies
New England Biolabs #M0480 OneTaq DNA Polymerase protocol · protocols.io version 2, 2022-02-21 · 2026-09-12 https://dx.doi.org/10.17504/protocols.io.bd24i8gw cited at thermocycling conditions
New England Biolabs application note, Robust Colony PCR from Multiple E. coli Strains using OneTaq Quick-Load Master Mixes · Y. Xu, 11/13 · 2026-09-12 · docs/research/primer-design-and-pcr.md cited at template
Azenta/Genewiz, Sanger sequencing FAQ and technical notes · 2026-09-12 · docs/research/primer-design-and-pcr.md cited at sequencing primer design