How can I check that my ligation reaction worked?

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Ligation reactions are best analyzed by actual transformation of bacteria, since not all of the high molecular weight forms created in a reaction (seen in gel analysis) will transform cells efficiently.

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If ligation/transformation reactions did not yield any colonies, or if a high number of background colonies are observed, what control reactions should be used to determine the problem?

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Several ligation controls may be necessary to identify the source of ligation problems.

Recommendations for problems with no colonies after transformation:
1. Test the transformation efficiency of the competent cells using a supercoiled vector, or the control DNA provided with Invitrogen™ competent cells. Perform a transformation reaction and plate the number of cells that is expected to generate 50-100 colonies per plate, based on the anticipated transformation efficiency of the competent cells. The expected number of colonies should be seen, indicating that the competent cells are transforming with high efficiency. The control DNA provided with Invitrogen™ competent cells is supercoiled monomer; vector DNA preparations that contain other forms will not transform as efficiently. Transformation efficiencies will be up to 10-fold lower for ligated vectors than for intact control DNA.
2. Restriction endonuclease-digested, re-ligated vector. Set up a ligation reaction using the same amount of vector DNA that is used in the experimental ligations and use it to transform competent cells. Re-ligation of vectors with cohesive ends should result in less than or equal to 50% of the number of colonies obtained with supercoiled vector DNA, indicating that the components of the ligation reaction are working; re-ligation of vectors with blunt ends should yield 10% to 20% of the number of colonies obtained with supercoiled vector DNA. This is an appropriate control only with vectors that have been digested with a single restriction endonuclease; double-digested vectors may not re-ligate because the ends are incompatible and the small DNA fragment that is released from between the two sites is sometimes lost during ethanol precipitation of the DNA.

For observation of a high number of background colonies:
1. Restriction endonuclease-digested vector. Perform a transformation with an amount of restriction enzyme-digested vector DNA equivalent to that contained in the fraction of the ligation reaction being used for the experimental transformations. Few or no colonies should be seen, indicating complete restriction endonuclease digestion of the vector. The presence of colonies indicates incomplete digestion of the vector that will cause a background of colonies containing non-recombinant vector in the experimental transformations.
2. Restriction endonuclease-digested, dephosphorylated, re-ligated vector. Set up a ligation reaction using the same amount of vector DNA that is used in the experimental ligations and use it to transform competent cells. Few or no colonies should be observed, indicating complete dephosphorylation of the vector - a dephosphorylated vector should not be re-ligated by T4 DNA ligase.
3.  No DNA transformation control. Perform a mock transformation of competent cells, to which no DNA is added. No colonies should be seen, indicating that the selection antibiotic on the agar plates is potent and that the competent cells are pure.

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How can I determine if my ligase is still active?

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Ligation reagents may be tested by performing a ligation reaction with a molecular size marker such as the 1 Kb DNA Ladder or lambda DNA/Hind III Fragments. Compare the ligation reaction products to unlighted DNA on an agarose gel. The ligation reaction should contain a high molecular weight smear and few low-molecular weight bands. If the marker ligation does not work, use fresh ligase.

Another reason for low activity could be degradation of the ATP in the reaction buffer; use 5X ligase buffer that is less than 24 months old. The buffer should be stored at -20°C.

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How can I test for inhibitors in  my ligation reactions?

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To test for the presence of ligation inhibitors, perform a ligation reaction in which some of the vector or insert DNA is included along with some marker DNA such as lambda DNA/Hind III Fragments. If ligation of the DNA marker fragments occurs alone but is not observed when other DNA is added, then a diffusible inhibitor is present in the vector or insert DNA.

To purify and remove inhibitors, extract the DNA with buffer-saturated phenol, then extract with chloroform:isoamyl alcohol, and precipitate with ammonium acetate and ethanol. Be sure that the DNA is free of phenol and that the phosphate concentration is less than 25 mM and the NaCl concentration is less than 50 mM. Also, be sure that the DNA is free of contaminating DNA that might compete for ligation to the insert or vector (e.g., linker fragments, DNA fragments from which the insert was completely purified).

If restriction endonucleases are present, causing redigestion of ligated products, your ligation will also be inhibited. After digestion of the vector and insert DNA, remove restriction endonucleases by extraction with buffer-saturated phenol, extraction with chloroform:isoamyl alcohol, and ethanol precipitation.

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What concentration of insert and vector do you recommend for most ligation reactions?

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Recommendations would vary depending on the size of the vector and insert or the nature of the insert, but for most plasmid cloning or subcloning reactions, a vector concentration of 1-10 ng/µl is recommended. Inserts should generally be 2- to 3-fold excess in molar concentration relative to the vector.

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Do both my insert and vector need to be phosphorylated for ligation?

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At least one molecule in a ligase reaction (i.e., insert or vector) must be phosphorylated. Ligation reactions are dependent on the presence of a 5' phosphate on the DNA molecules. The ligation of a dephosphorylated vector with an insert generated from a restriction enzyme digest (phosphorylated) is most routinely performed. Although only one strand of the DNA ligates at a junction point, the molecule can form a stable circle, providing that the insert is large enough for hybridization to maintain the molecule in a circular form.

Answer Id: 4014

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Why do I sometimes get low cDNA yield when using SuperScript® Reverse Transcriptase?

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Low cDNA yield can result due to several different reasons. Please see a few listed below:

(1) Poor quality mRNA: visualize total RNA on a denaturing gel to verify that the 28S and 18S bands are sharp. OD 260:280 ratio should be 1.7.

(2) Template degraded by RNase contamination: maintain aseptic conditions.

(3) Inhibitors of SuperScript® II RT may be present: remove inhibitors by ethanol precipitation of the RNA preparation before the first-strand reaction. Include a 70% (v/v) ethanol wash of the RNA pellet. Test for the presence of inhibitors by mixing 1 μg control RNA and comparing yields of first-strand cDNA.

(4) RNA preparation may have coprecipitated with polysaccharides: precipitate RNA with lithium chloride to purify RNA.

(5) mRNA concentrations were overestimated: quantitate the mRNA concentrations by measuring the A260 if possible.

(6) If using 32P-isotope, it may be too old: use isotope less than 2 weeks old.

(7) Not enough enzyme was used: use 200 U SuperScript® II RT/μg RNA.

(8) SuperScript® II RT activity was decreased by incorrect reaction temperature: perform the first-strand reaction at a temperature between 37 degrees C and 50 degrees C.

(9) DTT was not added to first-strand reaction.

(10) TCA precipitations were performed incorrectly: adequately dry GF/C filters before immersion into scintillant.

(11) SuperScript® II RT was improperly stored: store at -20 degrees C. Do not store the enzyme at -70 degrees C.

(12) The reaction volume was too large: the reaction should be done in volumes less than or equal to 50 μL.

Answer Id: 4015

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What are the best conditions for double digests with restriction endonucleases?

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First, please note that we changed our restriction enzymes and buffer formulations in 2010. The REact® buffers 1-10 were discontinued in favor of a smaller group of universal buffers: H, K, L, M, T. The new buffers are not compatible with older restriction enzymes, and it is not recommended to do a double digest with an old enzyme (with REact® buffer) and a new enzyme.

When performing any double digest, there may be buffer incompatibilities and enzyme steric hindrance problems. These can be avoided by performing sequential digests, separated by buffer exchange or chloroform extraction and ethanol precipitation. However, if these issues are understood and a double digest will be performed, you can evaluate enzyme combinations using our buffer compatibility chart. You can find the chart by searching “Recommended universal buffers for double digestion” on the Life Technologies website.

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Why is ATP present in the reaction buffer for T4 DNA Ligase?

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ATP is necessary for enzymatic function. It is involved in phosphorylating the ligase prior to the ligation reaction. Ligation efficiency is markedly reduced by removing ATP from the reaction. It is important, therefore, to handle the buffer appropriately in order to minimize degradation of ATP.

Answer Id: 2940

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Which is better to use, T4 or E. coli DNA ligase?

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It depends on your application. For ligation of dsDNA fragments with cohesive ends, either enzyme can be used. E. coli DNA ligase requires the presence of beta-NAD, while T4 DNA ligase requires ATP. However, only T4 DNA ligase can join blunt-ended DNA fragments - E. coli ligase is unable to join such fragments.

E. coli DNA ligase is generally used to eliminate nicks during second-strand cDNA synthesis. T4 DNA ligase should not be substituted for E. coli DNA ligase in second-strand synthesis because of its capability for blunt end ligation of the ds cDNA fragments, which could result in formation of chimeric inserts.

Answer Id: 2949

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What are the recommended conditions for cohesive-end ligations?

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Generally, ligations are done in a 20 μl volume. Use a total of 10 to 100 ng of DNA per reaction with an insert to vector ratio of 3:1. Add 0.1 units (Weiss) ligase to the reaction. Incubate at room temperature for 30-60 minutes.

Optimal ligation may occur at other ratios (e.g. 1:5, 1:10). If possible, assemble several ligation reactions of varying insert to vector ratios in order to reveal the optimal ligation conditions.

Life Technologies™ offers T4 DNA ligase at two concentrations: 1 U/μl (15224-017) and 5 U/μl (15224-041). When performing blunt or TA cloning ligations, the higher concentration of ligase is generally preferred since ligating a blunt or single base overhang requires more enzyme.

Answer Id: 2950

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What are the recommended conditions for blunt-ended ligations?

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Generally, ligations are done in a 20 μl volume. Use a total of 100 to 1000 ng of DNA with an insert to vector ratio of 3:1. Add 1.0 units (Weiss) to the reaction. Incubate at room temperature for 4 h or overnight at 14-16°C.

Ideally, assemble several reactions with varying ratios of vector:insert (i.e. 3:1, 5:1, 10:1, 20:1, etc.) to determine the optimal ratio for ligation.

Life Technologies™ offers T4 DNA ligase at two concentrations: 1 U/μl (15224-017) and 5 U/ul (15224-041). When performing blunt or TA cloning ligations, the higher concentration of ligase is generally preferred since ligating a blunt or single base overhang requires more enzyme.

Answer Id: 2951

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02180771a9b609a26dcea07f272e141f_FAQ

Why are alkaline phosphatases used in cloning protocols? What is a typical protocol for dephosphorylation of nucleic acids?

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Alkaline phosphatases are used to dephosphorylate the 5' ends of DNA. In cloning, it is used to prevent self-ligation of vector DNA. Standard ligation of DNA with ligase requires a 5' phosphate to be present on at least one of the ends being joined. When a DNA insert containing phosphates on both 5' termini is added to a dephosphorylated vector, the insert will be efficiently ligated into the vector, but the vector will not be able to self-ligate. Thus, dephosphorylation of vector lowers the number of background colonies containing vector without insert.

Answer Id: 2955

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Why is it necessary to dilute ligated DNA products before adding them to competent bacterial cells?

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Components of the ligation reaction (enzymes, salts) can interfere with transformation, and may reduce the number of recombinant colonies or plaques. We recommend a five-fold dilution of the ligation mix, and adding not more than 1/10 of the diluted volume to the cells. For best results, the volume added should also not exceed 10% of the volume of the competent cells that you are using.

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Which T4 DNA Ligase protocol do you recommend when ligating an insert containing one cohesive (sticky) end and one blunt end?

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For cloning an insert with one cohesive end and one blunt end, use the conditions for blunt ends.  The sticky end may ligate quickly, but the blunt end ligation will still be inefficient. You should use the more stringent protocol to optimize the blunt end ligation. This usually means using more enzyme (5 U), a lower reaction temperature (14C) and a longer incubation time (16-24 hours). 

Answer Id: 4268

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