How can I check that my ligation reaction worked?
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?
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.
Answer Id: 4010
How can I determine if my ligase is still active?
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.
Answer Id: 4011
How can I test for inhibitors in my ligation reactions?
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.
Answer Id: 4012
What concentration of insert and vector do you recommend for most ligation reactions?
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.
Answer Id: 4013
Do both my insert and vector need to be phosphorylated for ligation?
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
Why do I sometimes get low cDNA yield when using SuperScript® Reverse Transcriptase?
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
What are some of the problems associated with sticky-end cloning?
The amplified DNA needs to be purified from the PCR mixture components prior to cloning. The dNTPs carried over from the PCR are competitive inhibitors for ATP in the ligation reaction.
If during synthesis of the PCR primers their chemical integrity has been compromised by either a base substitution or modification, the enzyme recognition site may in actuality not exist. If this is the case, PCR products will be resistant to digestion with restriction enzymes. It may be necessary to use a higher concentration of the restriction enzyme and to incubate at the appropriate temperature overnight to ensure cutting.
Answer Id: 1102
How many degenerate sites are allowed in a 25-mer present at a total concentration of 1 micromolar (100 pmoles/100 mL)?
Primer mixtures with 256-fold and 32-fold degeneracies have been used [see Mack DH, Sninsky JJ (1988) Proc Natl Acad Sci USA 85:6977-6981 and Lee et al. (1988) Science 239:1288-1291.] We recommend that users synthesize pools of no more than 32- to 64-fold degenerate primers, making additional pools separately to account for all possible degeneracy. A matrix should be set up so that degeneracy is no more than 2-fold at each site, with all sites in the matrix run at the same time. Inosine may also be used for the degenerate positions in the primer. Performing touchdown PCR may help increase the specificity of degenerate primer PCR amplification.
Answer Id: 1103
In the TaqMan® Rodent GAPDH Control Reagents kit (P/N 4308313), what rodent is the Control RNA isolated from?
I want to make peptide amides, but your amide resin has no amino acid attached. Why not? Do I need to do anything special?
There is no amino acid attached because one is not needed. The amide linker has a free amine which is protected by an Fmoc group. Upon removal of the Fmoc group, an amide bond may be formed with the incoming activated amino acid. Nothing special needs to be done, although you must tell your synthesizer you are using an amide support and/or the first amino acid is not on the support. Standard cleavage protocols may be used.
Answer Id: 1236
What can cause broad dips or cyclical peaks in the baseline when using the Procise™ System?
How can I improve peak resolution between ASP and DTT or ASN when using the Procise™ System?
You can move ASP (and GLU) away from DTT and to later retention times by reducing slightly the pH of Solvent A3. This is best accomplished by adding a small amount of TFA (R3)--about 50 to 100 ul/liter of Solvent A3. In the Procise™ System cLC, when ASP and ASN are too close together, the problem can usually be resolved by replacing the guard column (part no. 401883). Decreasing initial %B in the gradient (e.g., from 10% to 8%) will move both DTT and ASP to later retention times.
Answer Id: 1253
I am using the Procise™ System and have increased the final %B in the gradient to separate LYS from LEU, but now all the late PTH amino acid peaks are crushed together. What can I do?
Why doesn't the 433A manual or the "quick start card" mention the need for an extra AA cartridge in the guideway at the start of a sequence?