What is the minimum length required for a circular DNA ligation?

What is the minimum length required for a circular DNA ligation?

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What is the minimum length of a DNA molecule for the ends to come in close enough contact that they can ligate. Assume there are free divalent cations in solution. Does anyone have any idea?

I stumbled across this paper demonstrating it is between 150-200 base pairs.

DNA flexibility studied by covalent closure of short fragments into circles D Shore, J Langowski, and R L Baldwin PNAS 1981 78 (8) 4833-4837

Full-length human cytomegalovirus terminase pUL89 adopts a two-domain structure specific for DNA packaging

A key step in replication of human cytomegalovirus (HCMV) in the host cell is the generation and packaging of unit-length genomes into preformed capsids. The enzymes involved in this process are the terminases. The HCMV terminase complex consists of two terminase subunits, the ATPase pUL56 and the nuclease pUL89. A potential third component pUL51 has been proposed. Even though the terminase subunit pUL89 has been shown to be essential for DNA packaging and interaction with pUL56, it is not known how pUL89 mechanistically achieves sequence-specific DNA binding and nicking. To identify essential domains and invariant amino acids vis-a-vis nuclease activity and DNA binding, alanine substitutions of predicted motifs were analyzed. The analyses indicated that aspartate 463 is an invariant amino acid for the nuclease activity, while argine 544 is an invariant aa for DNA binding. Structural analysis of recombinant protein using electron microscopy in conjunction with single particle analysis revealed a curvilinear monomer with two distinct domains connected by a thinner hinge-like region that agrees well with the predicted structure. These results allow us to model how the terminase subunit pUL89’s structure may mediate its function.

Featured Video

Avoiding RNase Contamination

SplintR Ligase, also known as PBCV-1 DNA Ligase or Chlorella virus DNA Ligase, efficiently catalyzes the ligation of adjacent, single-stranded DNA splinted by a complementary RNA strand. This previously unreported activity may enable novel approaches for characterization of miRNAs and mRNAs, including SNPs. SplintR is ideally suited for many target enrichment workflows with applications in next-generation sequencing and molecular diagnostics. The robust activity of the enzyme and its affinity for RNA-splinted DNA substrates (apparent Km = 1 nM) enable sub-nanomolar detection of unique RNA species within a complex mixture, making SplintR ligase a superior choice for demanding RNA detection technologies.

The enzyme is active over a broad range of ATP concentrations (10 µM &ndash 1 mM) and pH (6.5&ndash9). Optimal activity is observed using Mg 2+ > 5 mM, and a pH between 7.5 and 8.0. The activity is enhanced at higher temperatures (up to 37°C), and by supplementation with 5 mM Mn 2+ . The reaction is inhibited by salt concentrations in excess of 100 mM.

The enzyme tolerates all base pair combinations at the ligation junction, but is partially inhibited by dC/G and dG/C base pairs at the donor (phosphorylated) side ligation junction, particularly when the + 2 base was also a C/G base pair.

Ligation of DNA splinted by RNA
(A) Outline of the ligation assay: a 5´-phosphorylated, 3´-FAM labeled DNA &ldquodonor&rdquo oligonucleotide and an unmodified DNA &ldquoacceptor&rdquo oligonucleotide are annealed to a complementary RNA splint. This substrate is reacted with a ligase to form a mixture of unreacted starting material (I), adenylylated DNA (II), and ligated product (III). These products are denatured, separated by capillary electrophoresis and detected by fluorescence. (B) Ligation of the RNA-splinted substrate in SplintR Ligase Reaction Buffer for 15 minutes at 25°C with (a) no enzyme, (b) 1 &muM T4 DNA Ligase and (c) 100 nM SplintR Ligase. Indicated peaks correspond to starting pDNA (I), AppDNA (II) and ligated product (III) as determined by co-elution with synthetically prepared standards. (C) The fraction of ligated product catalyzed by either SplintR Ligase or T4 DNA Ligase was analyzed by performing sets of ligations with both ligases at concentrations between 10 pM and 10 &muM for 15 minutes at 25°C. SplintR Ligase is clearly much more efficient at ligation of RNA splinted DNA than T4 DNA Ligase.

Product Source

Reagents Supplied

The following reagents are supplied with this product:

SplintR® Ligase M0375SVIAL -20 1 x 0.05 ml 25,000 units/ml
SplintR Ligase Reaction Buffer B0375SVIAL -20 1 x 1 ml 10 X
SplintR® Ligase M0375LVIAL -20 1 x 0.25 ml 25,000 units/ml
SplintR Ligase Reaction Buffer B0375SVIAL -20 1 x 1 ml 10 X


Application Features

  • Ligation of ssDNA splinted by complementary RNA sequences
  • Detection of RNA using ligation of DNA probes
  • SNP or splice variant detection
  • RASL-seq

Unit Definition

One unit is defined as the amount of enzyme needed to ligate (to 50% completion) 2 picomoles of a tripartite FAM-labeled DNA:RNA hybrid substrate in a 20 &mul reaction at 25°C in 15 minutes in 1X SplintR Ligase Reaction Buffer.

1X SplintR Ligase Reaction Buffer
50 mM Tris-HCl
10 mM MgCl2
1 mM ATP
10 mM DTT
pH 7.5 @ 25°C

Storage Buffer

10 mM Tris-HCl
300 mM NaCl
1 mM DTT
0.1 mM EDTA
50% Glycerol
pH 7.4 @ 25°C

Heat Inactivation

  1. SplintR Ligase is inhibited by monovalent cations. We strongly recommend ensuring these common reactants (NaCl, KCl) are kept to below 50 mM in the reaction. The enzyme is supplied in a storage buffer containing 300 mM NaCl, for storage stability. A minimum 6-fold dilution of the enzyme by addition to the reaction is recommended, with the optimal dilution being &ge 10-fold.
  2. If dilution of enzyme for storage is needed, we recommend using Diluent A (NEB #B8001).
  3. Reactions with SplintR should be performed between 16&ndash37°C. We recommend initial testing be performed at 25°C.
  4. The enzyme is supplied as a 10.5 &muM solution. We suggest maintaining the enzyme below 1 &muM in the reaction with a suggested range of 100 nM to 1 &muM. For many applications, starting with a 2-fold excess of enzyme over ligatable ends is ideal. For example, in the experiment described in the accompanying figure a substrate concentration of 100 nM was found to be ideal. In that workflow, 250 nM enzyme gave complete ligation in 15 minutes on all sequences tested.
  5. If the reaction is not proceeding as efficiently as desired, we strongly recommend extending the incubation time rather than increasing the concentration of enzyme in the reaction beyond 1 &muM.
  6. An alternative option for recalcitrant substrates is to use a low concentration of ATP. A low ATP buffer can give higher yields of ligation product for substrates that have low ligation efficiencies in the standard SplintR Ligase Reaction Buffer, such as substrates with runs of G:C base pairs at the ligation junction. We suggest 1X T4 RNA Ligase Reaction Buffer (NEB #B0216) supplemented with ATP (NEB #P0756) to a final concentration of 10 &muM.
  7. We recommend an RNA splint of at least 20 complementary bases. We have found 10 bp of dsDNA/RNA on either side of the junction to be sufficient for all substrates tested. The splint does not have to be centered on the ligation junction, however, with as few as four bases on one side of the junction giving complete ligation for a splint with 20 bases of total complementarity, depending on substrate sequence. If regions of overlap <10 bp are desired, some testing will be required to determine the minimum length of the ds region for your specific sequence.
  8. In order to maximize fidelity in ligations by SplintR DNA Ligase, please consider the following:


Ligatable nick formation requires POLγA exonuclease activity

The exonuclease activity of POLγ can be inactivated by a single amino acid substitution in the second exonuclease motif in the POLγA subunit (D274A, hereafter referred to as ‘EXO-’) 29 . The EXO- mutation causes a strand-displacement activity, that is, polymerase continues to displace downstream DNA encountered during DNA synthesis 15,22 . We speculated that failure to stall at the 5′-end of downstream DNA would lead to the formation of a 5′-flap, which is incompatible with ligation. To address this possibility, we used a linear substrate with a 20-nt long single-stranded gap and investigated how actively synthesizing WT and EXO- POLγ behave when encountering a free 5′-DNA end. The template used was constructed from an 80-nt-long oligonucleotide annealed to a 30-nt radioactively labelled upstream primer and a 30-nt downstream blocking oligonucleotide (Fig. 1a). Precise gap filling and creation of ligatable ends will extend the primer to 50 nts. At early time points (1–3 min), WT POLγ did not halt at the 5′-end of the downstream blocking primer, but extended up to 6 nts into the duplex region (Fig. 1b). After 5 min, bands at positions 50–52 became the dominant species, suggesting that POLγ had backed up towards the nick. EXO- POLγ behaved similarly to WT at the earliest time point, but later the enzyme continued to polymerize into the double-stranded region, eventually completely displacing the downstream strand to produce a main product of 81 nts in length by 5 min (Fig. 1b, lanes 13–15). Please note that the extra nucleotide (the template is only 80 nts long) results from the terminal transferase activity of EXO- POLγ (shown in Supplementary Fig. 1).

(a) Diagram of the linear gapped substrate used in strand-displacement assays, with products shown below. (b) Strand displacement by WT and EXO- POLγ over time. WT polymerase performs limited strand displacement, up to 6 nts within the downstream duplex region, before backing up towards the nick position (arrowhead at 50 nts). EXO- polymerase rapidly displaces the downstream oligonucleotide completely (arrowhead at 81 nts). Reactions were started by the addition of POLγ. Time points are indicated above in minutes (− represents reactions where no POLγ was added). MM, molecular marker.

To further analyse strand displacement, we used the same substrate, but now labelled the downstream oligonucleotide (Supplementary Fig. 2). Confirming the above results, we observed increased strand displacement with EXO- POLγ compared with WT POLγ at all time points. By 30 min of DNA synthesis, almost 50% of the downstream oligonucleotide was completely displaced by EXO- POLγ compared with <5% for WT POLγ. We therefore concluded that the exonuclease activity of POLγ is required to limit POLγ strand-displacement activity and to promote idling near the nicked position.

Efficient DNA ligation requires POLγ exonuclease activity

We hypothesized that the aberrant formation of 5′-flaps by EXO- POLγ would impair ligation. To investigate this possibility, we performed coupled DNA synthesis and ligation assays using purified recombinant WT and EXO- POLγ proteins on DNA substrates. To this end, we primed an ∼ 3-kb circular ssDNA substrate with a 32-nt DNA oligonucleotide that had been radioactively labelled at the 5′-terminus (Fig. 2a). WT POLγ was able to polymerize until it reached the 5′-end of the primer, creating a nicked, circular dsDNA molecule (Fig. 2a, middle, and Fig. 2b, lane 2). When ligase was added, the nick was sealed (Fig. 2a, right, and Fig. 2b, lane 3). The EXO- mutant could also produce a full-circle, double-stranded product (Fig. 2b, lane 4), but did not generate ligatable ends, as no closed circular DNA molecules were observed (Fig. 2b, lane 5).

(a) Diagram of the circular substrate and resulting products in replication-coupled ligation reactions. (b) POLγ needs functional 3′–5′exonuclease activity to produce a closed circular DNA molecule. In the coupled second strand, synthesis and ligation assay WT (lane 3) but not EXO- (lane 5) is able to form a ligated product. The presence of EtBr in the agarose gel facilitates differentiation between nicked and closed circular dsDNA molecules. (c) Diagram of the linear gapped substrate and resulting products in replication-coupled ligation reactions. (d) Ligation (80 nt product, indicated with arrow head) is severely reduced in reactions using EXO- compared with WT POLγ. Molecular marker in lane 1. (e) Quantification of ligation efficiency was measured as the amount of ligation product formed (bands indicated with arrowhead) against the amount of starting substrate in the absence of ligase (lane 3). WT was set to 100%. Mean values±s.e.m., n=3, P≤2.5 × 10 −3 (Student’s unpaired t-test).

Next, we investigated coupled gap filling and DNA ligation using a substrate in which a 30-nt unlabelled upstream primer and a 30-nt radioactively labelled downstream blocking oligonucleotide were annealed to an 80-nt long template strand (Fig. 2c). Addition of WT POLγ or EXO- in the presence of mtSSB and T4 DNA ligase led to the formation of an 80-nt long ligated product (Fig. 2d, lanes 6 and 8). The ligation efficiency was however about ten times higher for the WT POLγ than for EXO- (Fig. 2e). The assay was repeated with mitochondrial Lig3 with similar results (Supplementary Fig. 3). We could therefore conclude that the exonuclease activity of POLγ is required for efficient DNA ligation in vitro.

Identification of exonuclease-deficient POLγ from patients

A number of disease-causing amino acid substitutions have been identified in the exonuclease domain of POLγ 31 and we wanted to investigate whether these mutations also affected exonuclease activity and the formation of ligatable ends. To this end, we expressed and purified seven mutant POLγA variants in recombinant form (R232H, G268A, R275Q, H277L, G303R, L304R and S305R (Fig. 3a)). We first investigated the DNA-binding activity of the mutants in an electrophoretic mobility shift assay (EMSA), in which POLγA was incubated together with a short, radioactively labelled primed DNA template. In the absence of POLγB, the R275Q, G303R, L304R and S305R mutants all displayed reduced DNA-binding affinity compared with WT POLγ (Fig. 3b). In combination with POLγB however, all the mutants were able to bind DNA (Fig. 3b) and to synthesize a short stretch of DNA with efficiency similar to that observed for WT POLγ (Supplementary Fig. 4).

(a) Schematic diagram of the exonuclease domain of POLγA and the position of the patient-associated mutations. The EXO- mutation D274A is indicated in grey. Black boxes represent the exonuclease motifs I, II and III. (b) The DNA-binding efficiencies of the POLγA mutants were tested on a primed template either as monomers or heterotrimers with POLγB (as illustrated on the left side). The R275Q, G303R, L304R and S305R mutants had markedly reduced binding activity as monomers. (c) Similar to EXO- POLγ, the G303R, L304R and S305R mutants have no apparent exonuclease activity. R232H has increased exonuclease activity compared with WT. The substrate is shown in the lower right quadrant. (d) DNA polymerization by the different POLγ proteins on a circular, 3,000-nt ssDNA template (as shown in bottom right quadrant). Synthesis rates are moderately slower for the R232H and R275Q mutants, and markedly compromised for the G303R, L304R and S305R proteins.

We next monitored exonuclease activity using a radioactively labelled 32-nt-long oligonucleotide annealed to pBluescript SK+ ssDNA, creating a 31-bp double-stranded region with a one-nucleotide mismatch at the 3′-end (lower right panel in Fig. 3c). The template was incubated with the WT and mutant POLγ variants for the indicated times. Of the seven proteins, G303R, L304R and S305R exhibited severe exonuclease deficiency, similar to the EXO- mutant protein, whereas the G268A and R275Q mutants had mildly impaired exonuclease activity. Interestingly, we also found a mutation, R232H, which had markedly increased exonuclease activity compared with WT. Similar results for exonuclease activity were also obtained for all mutants using a 5′-labelled primer-template substrate in the absence of dNTPs (Supplementary Fig. 4, lanes with 0 μM dNTPs).

DNA synthesis efficiency on a longer substrate

To investigate whether our POLγ mutants could synthesize longer stretches of DNA, we used the ∼ 3,000-nt circular ssDNA template described above (Fig. 2a). Similar to WT POLγ, the EXO-, G268A and H277L mutants produced full-length products within 5 min (Fig. 3d, compare lanes 2, 7, 17 and 22). We observed reduced polymerization for the R232H and R275Q mutants, and even weaker activity with G303R, L304R and S305R. The L304R and S305R mutants were only able to produce full-length product after 30 min incubation (Fig. 3d, lanes 30 and 45), whereas no full-length products were observed with the G303R mutant (Fig. 3d, lane 40).

Strand-displacement activities of POLγ mutants

Whereas the strand-displacement activity of EXO- polymerase was clearly visible in the long-stretch DNA synthesis assay (Fig. 3d, smear above main product in lanes 8–10), we could not see a similar effect for the disease-causing mutants. We decided to further analyse the strand-displacement activities of the different mutant polymerases over time, using the linear gapped substrate (shown in Fig. 1a). Precise gap filling will result in a band of 50 nt, whereas strand-displacement will generate products up to 80 nt. Using this template, none of the patient-related mutant proteins had strand-displacement activities as severe as the EXO- (Fig. 4a). We did however notice that three mutants, L304R, G303R and S305R, generated bands around 50 nts that appeared broader than the corresponding band generated by the WT (Fig. 4a, compare lanes 4–6 with 34–36, 46–48 and 52–54), which could be the result of entry into the duplex region and the creation of a short 5′-flap. To address this possibility, we used sequencing gels for increased resolution (Fig. 4b). In this analysis, we observed stalling at position 50–56 (that is, at the nick or 1–6 nt within the downstream dsDNA region). In keeping with its increased exonuclease activity, the R232H mutant displayed reduced strand-displacement activity, pausing at the nick (Fig. 4b, compare lanes 2–3 with 6–7). The H277L, G268A and R275Q mutants behaved similar to WT POLγ by pausing at the nick position or 1–2 nt downstream. The remaining three mutants, L304R, G303R and S305R (lanes 12-13, 16–19), stalled in a broader range within the downstream dsDNA region, creating longer 5′-flaps.

(a) Time course of strand displacement by purified recombinant WT and mutant POLγ proteins. The reactions were performed as depicted in Fig. 1a. Arrows: solid lines indicate gap filling broken lines indicate strand displacement. Lanes are numbered 1–54. (b) Strand-displacement reactions as above resolved on a sequencing gel. The failure by G303R, L304R and S305R to reverse to the nick position at the 10 min time point is evident (arrowhead at 50 nts). The sizes of an oligonucleotide molecular marker are indicated to the left. (c) Quantification of ligation efficiencies (ligation product formed as percentage of substrate) of the different POLγ proteins relative to WT. The ligation assay and quantification was performed as in Fig. 2c–e. Mean values±s.e.m., WT was set to 100%, asterisks represent significant differences compared with WT (n=3 *P≤0.05, **P≤0.01, ***P≤0.001 one way analysis of variance).

POLγ mutations affect replication-coupled ligation

Three of the investigated mutants thus displayed increased strand displacement (L304R, G303R and S305R), whereas one displayed decreased strand displacement (R232H) compared with WT. We investigated whether these mutations also impaired ligation, as demonstrated for EXO- POLγ (Fig. 2). To this end, we performed coupled replication and ligation assays on a gapped linear substrate as illustrated in Fig. 2c,d. The experiments were performed in triplicate and the amount of ligated products was quantified. The graph in Fig. 4c compares the ligation efficiency for the different mutants relative to WT POLγ. Ligation was severely reduced with 5′-flap producing POLγ mutants (L304R, G303R and S305R). In contrast, the R232H mutant with decreased strand-displacement activity was even better than WT POLγ in producing ligatable ends.

Increased levels of mtDNA nicks in mutator mouse cell lines

Our studies demonstrated that mutations that change the exonuclease activity of POLγ impair ligation in vitro. To investigate whether these problems could also be detected in vivo, we used mouse embryonic fibroblasts (MEFs) derived from the mutator mouse. These homozygous knock-in mice express EXO- (D257A) in place of WT POLγ. If EXO- POLγ produces 5′-flaps and impairs ligation in vivo, we would expect to observe higher levels of nicked mtDNA in these animals. To address this possibility, we analysed genomic DNA isolated from WT and mutator MEFs by agarose gel electrophoresis and Southern blotting. The gel analysis was performed in the presence or the absence of ethidium bromide (EtBr) to differentiate between nicked and closed circular dsDNA molecules. In the absence of EtBr in the agarose gel, full-length mtDNA from WT and mutator cells migrated as a single band of equal size. As expected, DNA isolated from mutator cells also contained a shorter, linear DNA fragment, which was only detected with the probe against the major arc of mtDNA (Fig. 5a). In the presence of EtBr, full-length mtDNA could be separated into closed or nicked circles, which are unable to form supercoils in the presence of EtBr. Whereas a majority of mtDNA molecules isolated from WT cells were in the closed conformation, the opposite was true for mtDNA isolated from mutator cells, which mostly contained nicked circles. As controls, to verify the migration pattern of closed and nicked forms of mtDNA, the DNA was cut with different restriction enzymes (Supplementary Fig. 5). We could thus conclude that exonuclease-deficient POLγ causes the formation of nicks in vivo.

(a) MtDNA was analysed on a 0.4% agarose gel in the absence (left panel) or the presence (right panel) of EtBr. The presence of EtBr in the agarose gel facilitates differentiation between nicked and closed circular dsDNA molecules. The mtDNA was detected by Southern blotting using probes against the major or minor arc. Results were confirmed three times with two independent batches of genomic DNA preparations. (b) A schematic representation of the PCR template including the D-loop region. Arrows and black bars show the PCR amplification regions. (c) The relative ratio of DNA copies of H-/L-strands in WT and mutator (EXO-) mice. Asterisks represent significant differences compared with WT (n=3 *P≤0.05, Student’s t-test). (d) Model of mtDNA linear deletion formation. Failure to ligate at OriH leads to a nick in the H-strand (left panel). The nick does not inhibit initiation of a new round of replication from OriH, as the intact L-strand is used as template for H-strand synthesis. DNA synthesis initiated at OriH may therefore continue full circle and create a new DNA molecule with a nick at OriH (right upper panel). In contrast, DNA synthesis initiated at OriL will result in linear, deleted fragments. In this later case, L-strand DNA synthesis will initiate from OriL and use the H-strand as template. DNA synthesis will reach the nick near OriH and a double-strand break will be formed. The molecule formed will also contain an unstable ssDNA region that will be degraded, leaving a linear double-stranded product spanning the major arc (right lower panel). Failure to ligate at OriH will thus result in the same round of replication generating two different replication products a circular, nicked mtDNA and a linear, deleted fragment spanning OriH and OriL. Solid back line, H-strand dashed black line, nascent H-strand solid red line, L-strand dashed red line, nascent L-strand.

As initiation of mtDNA replication takes place at OriH and then proceeds unidirectionally, termination of one of the two new daughter molecules should also occur at OriH. Thus, failure to ligate after completion of DNA replication should result in mtDNA molecules that contain a nicked H-strand, but an intact L-strand near OriH. To address this possibility, we employed strand-specific PCR and monitored the levels of intact H- and L-strands in the OriH region of WT and mutator mice. Our upstream primer was located in a region not present in the linear deletion, ensuring that we were only monitoring circular mtDNA (Fig. 5b). For precise quantification, we used droplet digital PCR (ddPCR). As controls, we first quantified the levels of intact H- and L-strands in three different regions: ND5 and COXII in the major arc and 16S rRNA in the minor arc. We found equal levels of the two strands at all these locations (Fig. 5c). We next analysed a region surrounding OriH and found that the ratio between the H- and L-strand was reduced in the mutator mice as would be expected if there is a nick in the H-strand (Fig. 5c, OH-1). Owing to the displacement activity of EXO- POLγ, the nicks will be located in a broad zone rather than at a single point. In agreement with the notion, the strand bias was even stronger when we analysed a longer region, spanning the entire D-loop region (Fig. 5c, OH-2). We also analysed the OriL region and found equal levels of the two strands. This is also what is expected according to the strand-displacement replication model, as explained in the discussion (see below). Together, these results provide in vivo support for the notion that the exonuclease activity is required for efficient ligation and also demonstrate that the H-strand is frequently nicked in the OriH region of mutator mice.


A YAC is built using an initial circular DNA plasmid, which is typically cut into a linear DNA molecule using restriction enzymes DNA ligase is then used to ligate a DNA sequence or gene of interest into the linearized DNA, forming a single large, circular piece of DNA. [2] The basic generation of linear yeast artificial chromosomes can be broken down into 6 main steps:

  1. Ligation of selectable marker into plasmid vector: this allows for the differential selection of colonies with, or without the marker gene. An antibiotic resistance gene allows the YAC vector to be amplified and selected for in E. coli by rescuing the ability of mutant E. coli to synthesize leucine in the presence of the necessary components within the growth medium. TRP1 and URA3 genes are other selectable markers. The YAC vector cloning site for foreign DNA is located within the SUP4 gene. This gene compensates for a mutation in the yeast host cell that causes the accumulation of red pigment. The host cells are normally red, and those transformed with YAC only, will form colorless colonies. Cloning of a foreign DNA fragment into the YAC causes insertional inactivation of the gene, restoring the red color. Therefore, the colonies that contain the foreign DNA fragment are red. [4]
  2. Ligation of necessary centromeric sequences for mitotic stability [5]
  3. Ligation of Autonomously Replicating Sequences (ARS) providing an origin of replication to undergo mitotic replication. This allows the plasmid to replicate extrachromosomally, but renders the plasmid highly mitotically unstable, and easily lost without the centromeric sequences. [3][6]
  4. Ligation of artificial telomeric sequences to convert circular plasmid into a linear piece of DNA [7]
  5. Insertion of DNA sequence to be amplified (up to 1000kb)
  6. Transformation yeast colony [8]

In March 2014, Jef Boeke of the Langone Medical Centre at New York University, published that his team has synthesized one of the S. cerevisiae 16 yeast chromosomes, the chromosome III, that he named synIII. [9] [10] The procedure involved replacing the genes in the original chromosome with synthetic versions and the finished synthesized chromosome was then integrated into a yeast cell. It required designing and creating 273,871 base pairs of DNA - fewer than the 316,667 pairs in the original chromosome.

Yeast expression vectors, such as YACs, YIps (yeast integrating plasmids), and YEps (yeast episomal plasmids), have an advantage over bacterial artificial chromosomes (BACs) in that they can be used to express eukaryotic proteins that require posttranslational modification. By being able to insert large fragments of DNA, YACs can be utilized to clone and assemble the entire genomes of an organism. [11] With the insertion of a YAC into yeast cells, they can be propagated as linear artificial chromosomes, cloning the inserted regions of DNA in the process. With this completed, two processes can be used to obtain a sequenced genome, or region of interest:

This is significant in that it allows for the detailed mapping of specific regions of the genome. Whole human chromosomes have been examined, such as the X chromosome, [13] generating the location of genetic markers for numerous genetic disorders and traits. [14]

YACs are significantly less stable than BACs, producing "chimeric effects": artifacts where the sequence of the cloned DNA actually corresponds not to a single genomic region but to multiple regions. Chimerism may be due to either co-ligation of multiple genomic segments into a single YAC, or recombination of two or more YACs transformed in the same host Yeast cell. [15] The incidence of chimerism may be as high as 50%. [16] Other artifacts are deletion of segments from a cloned region, and rearrangement of genomic segments (such as inversion). In all these cases, the sequence as determined from the YAC clone is different from the original, natural sequence, leading to inconsistent results and errors in interpretation if the clone's information is relied upon. Due to these issues, the Human Genome Project ultimately abandoned the use of YACs and switched to bacterial artificial chromosomes, where the incidence of these artifacts is very low. In addition to stability issues, specifically the relatively frequent occurrence of chimeric events, YACs proved to be inefficient when generating the minimum tiling path covering the entire human genome. Generating the clone libraries is time consuming. Also, due to the nature of the reliance on sequence tagged sites (STS) as a reference point when selecting appropriate clones, there are large gaps that need further generation of libraries to span. It is this additional hindrance that drove the project to utilize BACs instead. [17] This is due to two factors: [18]

  1. BACs are much quicker to generate, and when generating redundant libraries of clones, this is essential
  2. BACs allow more dense coverage with STSs, resulting in more complete and efficient minimum tiling paths generated in silico.

However, it is possible to utilize both approaches, as was demonstrated when the genome of the nematode, C. elegans. There majority of the genome was tiled with BACs, and the gaps filled in with YACs. [17]

Synthesis of circular DNA templates with T4 RNA ligase for rolling circle amplification

Currently, isothermal methods of nucleic acid amplification have been well established in particular, rolling circle amplification is of great interest. In this approach, circular ssDNA molecules have been used as a target that can be obtained by the intramolecular template-dependent ligation of an oligonucleotide C-probe. Here, a new method of synthesizing small circular DNA molecules via the cyclization of ssDNA based on T4 RNA ligase has been proposed. Circular ssDNA is further used as the template for the rolling circle amplification. The maximum yield of the cyclization products was observed in the presence of 5−10% polyethylene glycol 4000, and the optimum DNA length for the cyclization constituted 50 nucleotides. This highly sensitive method was shown to detect less than 10 2 circular DNA molecules. The method reliability was proved based on artificially destroyed dsDNA, which suggests its implementation for analyzing any significantly fragmented dsDNA.


The establishment and persistence of hepadnavirus infection is dependent upon the viral cccDNA, which is a non-replicating episomal viral genome deposited in the nucleus of infected cell after conformational conversion from viral rcDNA [9]. Due to the limited gene-coding capacity of hepadnavirus genome, the virus needs to borrow host functions to complete its lifecycle [8]. The cellular DNA repair is a well-conserved surveillance and restoration system to detect and heal the damage in chromosomal DNA, by which maintains the stability and integrity of the host genome for replication and transcription [33, 34]. It is plausible that hepadnaviruses hijack the cellular DNA repair apparatus for cccDNA formation by disguising the rcDNA as a “damaged” DNA [9, 20]. The two gaps on rcDNA would be recognized as lesions for DNA repair by the host, and the DNA termini and their associated modifications are expected to undergo trimming, elongation, and ligation, during cccDNA formation (Fig 1). However, the host DNA repair pathway responsible for cccDNA formation remain largely unknown, and thus far, only a few host DNA repair enzymes have been reported to be involved in cccDNA formation, including the tyrosol-DNA phosphodiesterase 2 (TDP2) [17], polymerase κ (POLK) and λ (POLL) [21]. In this study, we screened 107 host DNA repair factors to assess their individual effect on HBV cccDNA formation by lentiviral shRNA knock-down, and identified and validated the host DNA LIG1 and LIG3 as key factors for hepadnavirus cccDNA formation by using a battery of in vitro and cell-based assays. Such work hence provides new insights into the mechanisms underlying hepadnavirus cccDNA formation in hepatocyte nucleus.

As part of the cellular DNA replication and repair machineries, DNA ligases complete joining of DNA strands by catalyzing the phosphodiester bond formation. Specifically, LIG1 ligates the Okazaki fragments during chromosomal DNA synthesis, and it is involved in the ligation steps of homologous recombination repair (HRR), long-patch base-excision repair (BER) and nucleotide excision repair (NER) LIG3 is responsible for sealing single strand DNA breaks during the process of short-patch BER and NER [26]. The involvement of DNA ligases in cccDNA formation indicated that the process of rcDNA termini generates ends that can be ligated and DNA ligases are the end-effectors for sealing the breaks of rcDNA. It is worth noting that previous studies have shown that LIG3, but not LIG4, is essential for nuclear DNA replication in the absence of LIG1 [35, 36] and LIG1 is a backup enzyme for LIG3 in BER and NER DNA repair pathways [37, 38]. Such functional redundancy between LIG1 and LIG3 may explain the unaffected cell viability and the incomplete inhibition of cccDNA formation by knocking down/out LIG1 or LIG3 only (Figs 4A, 5–7 and S8A) or knocking down both (Figs 4B and S8B). In previous studies, the redundant functions in cccDNA formation have also been observed between POLK and POLL [21], and perhaps between TDP2 and an unknown TDP2-like protein(s) [17]. It is of note that the potential role of TDP2 in cccDNA formation remains controversial. While one study demonstrated that knock-down of TDP2 inhibited, or at least delayed, DHBV cccDNA formation [17] another study suggested that TDP2 might even serve as a negative regulator of HBV cccDNA formation rather than a facilitator [18], and a recent study showed that TDP2 chemical inhibitors did not inhibit HBV infection in cell cultures [39]. In our shRNA screen, POLK or TDP2 lentiviral shRNA did not significantly reduce cccDNA formation in HepDES19 cells (S1 Fig), indicating that cellular functional redundancy for each enzyme might also exist in our experimental system. Further validation and mechanistic studies are required to reconcile these results. On the other hand, it is also possible that the first round cccDNA formation from virion DNA during infection and the intracellular cccDNA amplification pathway may have preference for different DNA repair enzymes, or there is hepatic cell line- or clone-specific requirement of host DNA repair factors/pathways for cccDNA formation. We had attempted to CRIPSR out both ligases in HepDG10 cells but only achieved partial double knock-down (Fig 4B), suggesting that at least one of LIG1 and LIG3 is required by the cells, and perhaps by hepadnaviruses as well. However, our data does not completely rule out a possibility that a LIG1/3-independent ligation mechanism might be involved in cccDNA formation, such as DNA topoisomerase I (TOP1) which has been suggested to play a role in rcDNA circularization through its DNA endonuclease and strand transferase activities [40]. Based on the previous and current data, it can be also inferred that hepadnaviruses have evolved to take advantage of the functional redundancy of host DNA repair machinery for a successful cccDNA formation.

The mechanism underlying the different efficiency of cccDNA formation between HBV and DHBV remains largely unknown, but likely in a virus-specific but not host-specific manner [16, 25, 30]. Based on that, we created the cell-free and the human hepatoma cell-based DHBV system to facilitate the identification and validation of host and viral regulators of cccDNA formation (Figs 2 and 3). In addition to the possible determining factors for cccDNA formation in the steps of rcDNA maturation, deproteinization, nuclear importation and uncoating, whether the cellular DNA repair system differentially recognizes and repairs nuclear HBV and DHBV rcDNA into cccDNA remains obscure. In this study, we found that both viruses employ LIG1 and LIG3 for cccDNA formation (Figs 4–7, S8 and S11), suggesting that the different repair process of HBV and DHBV rcDNA, if any, should be at the steps upstream of rcDNA end joining.

Though LIG1 and LIG3 have overlapping functions, knocking out/down of LIG3 resulted in relatively lower level of cccDNA than LIG1 knock-out/down (Figs 4–7 and S8), which suggests that LIG3 may play a more important role in cccDNA formation. In line with this notion, the two separated nicks/gaps in rcDNA are reminiscent of single strand breaks, which are preferable substrates for LIG3 in BER- and NER-mediated single strand break repair (Fig 1). Moreover, during the primary screen, two other BER components, APEX1 and POLB [41], emerged as candidates for positive regulator of cccDNA formation (S1 Fig), further suggesting the potential involvement of short-patch BER in cccDNA formation, which awaits further systematic investigations.

With the protein and RNA attachments at the 5’ end of minus- and plus-strand, respectively, hepadnavirus rcDNA is not a typical DNA break substrate for the major known repair pathways, and it is unknown whether the two gaps in rcDNA are repaired simultaneously or separately, including the final ligation step. A recent study revealed a nuclear rcDNA species with a covalently closed minus strand but an open plus strand, indicating that the nick on minus strand may be sealed first during cccDNA formation [28]. However, we did not observe an increased accumulation of protein-free rcDNA after blocking cccDNA synthesis in LIG1 or LIG3 knock-out cells (Figs 4, 6, 7B and S8). This phenomena may be due to a fact that the processed rcDNA ready for ligation is unstable. Previous studies have shown that the Hirt DNA samples from DHBV replicating cells contain high levels of nicked cccDNA which might be generated intracellularly or during the Hirt extraction [16, 30]. We also found that the protein-free rcDNA in Hirt extraction from HepDG10 cells were largely nicked cccDNA (S9 Fig), indicating that the observed reduction of protein-free rcDNA in LIG1/3 knock-out cells might be a consequence of cccDNA reduction (Figs 4 and 7). Nonetheless, further characterization of the nuclear rcDNA in LIG1/3 knock-out cells will provide further information for understanding the biological processes of rcDNA termini prior to the final ligation step during cccDNA formation.

In parallel with the bona fide rcDNA-to-cccDNA formation during hepadnavirus infection, the viral dslDNA byproduct is also repaired into cccDNA with indel mutations at the joint region [16, 23]. Although the dslDNA-derived cccDNA is generally defective of initiating a new round of viral DNA replication, it remains functional to express HBsAg and thus may play a role in viral pathogenesis. Based on the linear format of dslDNA and the indel mutations of its cccDNA derivative, it is hypothesized that dslDNA is a substrate for host error-prone NHEJ DNA repair system, and we have previously reported that another NHEJ component Ku80 is required for DHBV cccDNA formation from the dslDNA but not rcDNA [25]. LIG4 is the DNA ligase responsible for performing the last step of double strand DNA end joining in the NHEJ pathway [24]. In this study, we have demonstrated that LIG4 plays an essential role in cccDNA formation from DHBV dslDNA, and no functional redundancy was observed between LIG4 and other ligases (Fig 8). In addition, it has been reported that the chromosome DNA double strand breaks are targets for DHBV DNA integration [42], which indicates that the NHEJ machinery, including LIG4, is also responsible for the integration of hepadnavirus dslDNA into host genome.

Altogether, our study revealed a critical role of cellular DNA ligases in hepadnavirus cccDNA biosynthesis. Another possible function of DNA ligases in hepadnavirus life cycle can be to maintain the integrity of cccDNA, provided the preexisting cccDNA undergoes DNA damage and the host cell is able to repair it. Based on our observations, the DNA ligase inhibitors, which are currently under development for anti-cancer therapy [43], may be developed into host-targeting antiviral means to treat chronic hepatitis B by blocking cccDNA formation and/or repair.


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What is the minimum length required for a circular DNA ligation? - Biology

Directionality, in molecular biology and biochemistry, is the end-to-end chemical orientation of a single strand of nucleic acid. The chemical convention of naming carbon atoms in the nucleotide sugar-ring numerically gives rise to a 5′-end and a 3′-end (usually pronounced "five prime end" and "three prime end"). The relative positions of structures along a strand of nucleic acid, including genes and various protein binding sites, are usually noted as being either upstream (towards the 5′-end) or downstream (towards the 3′-end). (See alsoupstream and downstream.)

This naming convention is important because nucleic acids can only be synthesized in vivoin the 5′-to-3′ direction, as the polymerase that assembles new strands only attaches new nucleotides to the 3′-hydroxyl (-OH) group, via a phosphodiester bond. Directionality is related to, but independent from sense. In coding DNA, codons read 5′–ATG–⋯–3′ on the sense strand, and 3′–TAC–⋯–5′ on the complementary antisense strand. Thus only theantisense strand will be transcribed to sense (5′–AUG–⋯–3′) mRNA. By convention, single strands of DNA and RNA sequences are written in 5′-to-3′ direction.

The 5′-end (pronounced "five prime end") designates the end of the DNA or RNA strand that has the fifth carbon in the sugar-ring of the deoxyribose or ribose at its terminus. Aphosphate group attached to the 5′-end permits ligation of two nucleotides, i.e., the covalent binding of a 5′-phosphate to the 3′-hydroxyl group of another nucleotide, to form aphosphodiester bond. Removal of the 5′-phosphate prevents ligation. To prevent unwanted nucleic acid ligation (e.g. self-ligation of a plasmid vector in DNA cloning), molecular biologists commonly remove the 5′-phosphate with a phosphatase.

The 5′-end of nascent messenger RNA is the site at which post-transcriptional cappingoccurs, a process which is vital to producing mature messenger RNA. Capping increases the stability of the messenger RNA while it undergoes translation, providing resistance to the degradative effects of exonucleases. [ citation needed ] It consists of a methylated nucleotide (methylguanosine) attached to the messenger RNA in a rare 5′- to 5′-triphosphate linkage.

The 5′-flanking region of a gene often denotes a region of DNA which is not transcribed into RNA. The 5′-flanking region contains the gene promoter, and may also contain enhancers or other protein binding sites.

The 5′-untranslated region (5′-UTR) is a region of a gene which is transcribed into mRNA, and is located at the 5′-end of the mRNA, but which does not contain protein-coding sequence. The 5′-untranslated region is the portion of the DNA starting from the cap site and extending to the base just before the AUG translation initiation codon. While not itself translated, this region may have sequences, such as the ribosome binding site and Kozak sequence, which determine the translation efficiency of the mRNA, or which may affect the stability of the mRNA.

The 3′-end of a strand is so named due to it terminating at the hydroxyl group of the third carbon in the sugar-ring, and is known as the tail end. The 3′-hydroxyl is necessary in the synthesis of new nucleic acid molecules as it is ligated (joined) to the 5′-phosphate of a separate nucleotide, allowing the formation of strands of linked nucleotides.

Molecular biologists can use nucleotides that lack a 3′-hydroxyl (dideoxyribonucleotides) to interrupt the replication of DNA. This technique is known as the dideoxy chain-termination termination method or the Sanger method, and is used to determine the order of nucleotides in DNA.

The 3′-end of nascent messenger RNA is the site of post-transcriptional polyadenylation, which attaches a chain of 50 to 250 adenosine residues to produce mature messenger RNA. This chain helps in determining how long the messenger RNA lasts in the cell, influencing how much protein is produced from it.

The 3′-flanking region is a region of DNA that is not copied into the mature mRNA, but which is present adjacent to 3′-end of the gene. It was originally thought that the 3′-flanking DNA was not transcribed at all, but it was discovered to be transcribed into RNA and quickly removed during processing of the primary transcript to form the mature mRNA. The 3′-flanking region often contains sequences that affect the formation of the 3′-end of the message. It may also contain enhancers or other sites to which proteins may bind.

The 3′-untranslated region (3′-UTR) is a region of the DNA which is transcribed into mRNA and becomes the 3′-end of the message, but which does not contain protein coding sequence. Everything between the stop codon and the polyA tail is considered to be 3′-untranslated. The 3′-untranslated region may affect the translation efficiency of the mRNA or the stability of the mRNA. It also has sequences which are required for the addition of the poly(A) tail to the message, including the hexanucleotide AAUAAA.

T4 DNA Ligase

For details on NEB's quality controls for DNA ligases, visit our Ligase Quality page.

T4 DNA Ligase Competitor Study - Nuclease Contamination
T4 DNA Ligase from multiple suppliers was tested in reactions containing a fluorescent labeled single stranded, double stranded blunt, 3&rsquooverhang or 5&rsquo overhang containing oligonucleotides. The percent degradation by contaminating nucleases is determined by capillary electrophoresis and peak analysis. The resolution is at the single nucleotide level.


Product Source

Reagents Supplied

The following reagents are supplied with this product:

T4 DNA Ligase M0202SVIAL -20 1 x 0.05 ml 400,000 units/ml
T4 DNA Ligase Reaction Buffer B0202AVIAL -20 2 x 0.5 ml 10 X
T4 DNA Ligase M0202TVIAL -20 1 x 0.01 ml 2,000,000 units/ml
T4 DNA Ligase Reaction Buffer B0202AVIAL -20 2 x 0.5 ml 10 X
T4 DNA Ligase M0202LVIAL -20 1 x 0.25 ml 400,000 units/ml
T4 DNA Ligase Reaction Buffer B0202AVIAL -20 2 x 0.5 ml 10 X
T4 DNA Ligase M0202MVIAL -20 1 x 0.05 ml 2,000,000 units/ml
T4 DNA Ligase Reaction Buffer B0202AVIAL -20 2 x 0.5 ml 10 X

Application Features

Unit Definition

One unit is defined as the amount of enzyme required to give 50% ligation of HindIII fragments of &lambda DNA (5´ DNA termini concentration of 0.12 µM, 300- µg/ml) in a total reaction volume of 20 &mul in 30 minutes at 16°C in 1X T4 DNA Ligase Reaction Buffer.

400,000 cohesive end units/ml and 2,000,000 cohesive end units/ml

Reaction Conditions

1X T4 DNA Ligase Reaction Buffer
Incubate at 16°C

1X T4 DNA Ligase Reaction Buffer
50 mM Tris-HCl
10 mM MgCl2
1 mM ATP
10 mM DTT
(pH 7.5 @ 25°C)

Storage Buffer

10 mM Tris-HCl
50 mM KCl
1 mM DTT
0.1 mM EDTA
50% Glycerol
pH 7.4 @ 25°C

Heat Inactivation

Companion Products

Materials Sold Separately

  1. ATP is an essential cofactor for the reaction. This contrasts with E. coli DNA ligase which requires NAD.
  2. To dilute T4 DNA Ligase that will subsequently be stored at -20°C, 50% glycerol storage buffer (Diluent Buffer A,NEB #B8001S) should be used to dilute for immediate use, 1X T4 DNA Ligase Reaction Buffer can be used.
  3. Ligation can also be performed in any of the four restriction endonuclease NEBuffers or in T4 Polynucleotide Kinase Buffer if they are supplemented with 1 mM ATP.
  4. Room Temperature Ligation
    For convenience, ligations may be done at room temperature (20-25°C). For cohesive (sticky) ends, use 1 µl of T4 DNA Ligase in a 20 µl reaction for 10 minutes. For blunt ends, use 1 µl of T4 DNA Ligase in a 20 µl reaction for 2 hours or 1 µl high concentration T4 DNA Ligase for 10 minutes. Alternatively, NEB's Quick Ligation Kit (#M2200S) [30 reactions] or (#M2200L) [150 reactions]) is uniquely formulated to ligate both blunt and cohesive (sticky) ends in 5 minutes at room temperature.
  1. Engler, M.J. and Richardson, C.C. (1982). P.D. Boyer(Ed.), 5, 3. San Diego: Academic Press.
  2. Remaut, E., Tsao, H. and Fiers, W. (1983). Gene. 22, 103-113.
  3. Sambrook, J., Fritsch, E.F. and Maniatis, T. (1989). Molecular Cloning: A Laboratory Manual, (2nd Ed.). 1.53-1.73.
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  1. What are some potential problems with the ligation reaction using T4 DNA Ligase that can lead to transformation failure?
  2. What are some other problems that should be considered when troubleshooting a transformation problem?
  3. What problems can be encountered in the restriction digest that can cause ligation using T4 DNA Ligase or subsequent transformation to fail?
  4. What controls should be run to test the cells and DNA when using T4 DNA Ligase?
  5. When should T4 DNA Ligase be the enzyme of choice?
  6. Can the T4 DNA Ligase be used with the Quick Ligase buffer?
  7. What is the definition of a Weiss Unit and a Cohesive End Unit?
  8. What is the difference between the two definitions and why does NEB use the Cohesive End Unit?
  9. How much DNA should be used in a ligation using T4 DNA Ligase?
  10. Can T4 DNA Ligase be used in other NEBuffers, including rCutSmart?
  11. Can T4 DNA Ligase be heat inactivated?

Product Citation Tool

Additional Citations

  • Sexton T, Kurukuti S, Mitchell JA, Umlauf D, Nagano T, Fraser P (2012) Sensitive detection of chromatin coassociations using enhanced chromosome conformation capture on chip Nat Protoc 7(7), 1335-50. PubMedID: 22722369, DOI: 10.1038/nprot.2012.071
  • Thuronyi, B.W., Koblan, L.W., Levy, J.M. et al (2019) . Continuous evolution of base editors with expanded target compatibility and improved activity Nat Biotechnol. 37, 1070-1079.. DOI: 10.1038/s41587-019-0193-0


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