Another relevant factor is the experience of the operator. We have performed the same ligation mix with several operators, and obtained different results, probably depending on the individual hand pressure over the Drigalsky spatula. Glass beads have been used and the differences were reduced in part. We can’t discern yet other factors involved.
In any experiment, don´t forget the controls! In a cloning experiment they are essential indicators of your cloning procedure and allows you to identify problems and troubeshouting. We recommend:
Successful cloning depends on good competent cells like CVX5α™ Chemically Competent Cells. Cell competence is the DNA uptake efficiency calculated as the number of colonies formed per microgram of DNA (cfu/ µg). In some labs this factor is underestimated and even unknown.
A control cell competence should be carried out in parallel, as competent cells are very fragile. Do not thaw competent cells with your hands. Keeping competent cells out of an ice bath even for extremely short times strongly affects their transformation efficiency. Also, avoid pipetting up and down while mixing the competent cells, which should be made by gently flicking.
Cell competence higher than 1 × 106 cfu/μg DNA is enough for standard transformations.
Ligation efficiency depends on the T4 DNA Ligase selected once both insert and vector are in optimal conditions. All pSpark® DNA Cloning Kits are provided with a robust T4 DNA Ligase. Due to the extremely high cloning efficiency, ligations can be performed in as little as 10 minutes with the supplied protocol, while other ligases employ at least 1 hour or overnight ligation.
When only 10 minutes of ligation are used, the number of colonies obtained is about the same as a T/A cloning reaction made in parallel under optimal conditions (60 minutes to overnight ligation at 16 ºC). Generally, incubation at 22 ºC for 60 minutes produces about 5 times more transformants than a ligation for 10 minutes. Overnight incubation does not improve nor is detrimental for transformation.
Do not heat-inactivate a ligation with PEG as heat inactivation of Ligase dramatically reduces transformation efficiency of DNA heated in the presence of PEG.
Vector to insert molar concentration ratios are critical to ensure efficient cloning. If initial experiments with your PCR product are suboptimal, ratio optimisation may be necessary. Ratios of insert to vector from 5:1 to 1:1 provide good initial parameters. A highly relevant property of pSpark® DNA cloning Kits is that they use about 5-10 times less insert DNA than any other Cloning kit available on the market.
pSpark® DNA Cloning Kits have been optimised using an insert to vector ratio of 5:1. Optimal ratio of insert to vector is from 3:1 to 5:1. However, ratios of 0,2:1 to 50:1 have been used successfully.
For large inserts (more than 7 kb) insert to vector ratios of 3:1 to 1:1 produce more colonies than insert to vector ratios from 5:1 to 3:1 that are optimal for inserts of 0,1 kb to 4 kb. For large inserts (> 7 kb) an insert to vector ratio of 3:1 represents a good starting point for cloning into pSpark® DNA cloning systems.
Although, rarely, sometimes our cloning process failed because of the ends left by the polymerase used and the linear vector selected. It is indispensable to choose the appropriate vector to clone a PCR-amplified fragment.
pSpark® Cloning technology has been designed to clone PCR-blunted fragments produced with proofreading polymerases. It is part of a new generation of cloning vectors that ensure the highest blunt cloning efficiency.
Previous TA Cloning® technology had been designed to clone PCR products generated with Taq DNA polymerases (like Horse-Power™ Taq DNA Polymerase). Taq or Tth DNA polymerase add a single base overhangs at each 3´-end and thus, this DNA can be cloned only into pSpark® TA DNA Cloning Kits.
The PCR product should be analysed on an Agarose gel before its use in the ligation reaction to verify both the quality and quantity of your PCR product. A PCR that contains one single homogeneous band with only primers-dimers can be purified by any high-quality PCR clean-up kit for cloning into pSpark® DNA Cloning Kits.
Generally, high-yield PCR amplifications will produce also a good yield after purification from agarose gels. As a consequence, several hundreds, or even thousands, of white positive colonies is the most common result obtained using pSpark® DNA Cloning Kits.
In several cases, crude unpurified PCR products can be used directly for ligation into pSpark® DNA cloning Kits but, even in those cases we recommend to run an aliquot of PCR amplified DNA by gel Electrophoresis to check both quality and quantity.
Our Expert team have found that 1 µL of a very low yield PCR could be transformed directly without purification, and most of the times it generates colonies with the desired insert. Cloning of crude PCR products should be checked by colony PCR because white colonies could be formed by cloning contaminating bands and even primer dimers.
Often, some cloned fragments do not interrupt the beta galactosidase open reading frame generating blue colonies that have the insert cloned. If the cloned fragment encode a toxic protein for E. coli, the colonies could be small and showing a slower growth.
Some inserts have a complex secondary structure that decrease the cloning efficiency. pSpark® DNA Cloning Kits are compatible with any E. coli strain. Several E. coli strains have been developed to help stabilization of unstable DNA.
The size of PCR fragment to be cloned is an important element to keep in mind in a cloning reaction. Smaller fragments are easier to clone than larger fragments. Larger fragments require more DNA than smaller fragments to obtain a similar cloning efficiency.
pSpark® DNA Cloning Kits use much less DNA for cloning than any other kit available on the market. As a rule of thumb, use 5 ng of insert per kilobase, that is 2.5, 5, 10, 15, 20 or 25 ng of insert for an optimal ligation of an insert of 500 bp, 1,000 bp, 2,000 bp, 3,000 bp, 4,000 bp or 5,000 bp, respectively.
In order to clone 7 kb PCR products good competent cells are necessary (more than 5×107 cfu/µg) and, in some cases, a DNA cloning vector with low copy number to improve insert stabilization.
Some cloning vectors available on the market require a special primer design so they can be compatible to each other. Any primer could be used for cloning into pSpark® DNA Cloning Kits. This includes, but it is not limited to, unphosphorylated and phosphorylated primers, primers purified by any technique such as desalted only, reverse cartridge purified, HPLC or PAGE purified, primers with modified bases and primers with any sequence at their 5´-ends. Neither special sequences at the 5´-ends of the primers are needed thus, avoiding cloning artifacts due to the presence of long sequences in the primers specially when using complex template DNA such as, genomic DNA.
Cloning of DNA fragments into a linear vector is a common process in Molecular Biology labs. Three days is the time it takes a specialized laboratory to obtain a desired recombinant clone but the efficiency of this process varies from lab to lab. Although the procedure is simple, be careful adding the small volumes that are usually employed. It is important to use controls that confirm us that everything is OK at each step:
Usually a 0.5- 1 minute DNA denaturation at 94-95 ºC is enough for most templates. GC-rich templates require longer denaturation times.
Although most DNA Purification methods generate a good DNA quality for the PCR reaction, it is important to know that certain compounds could be inhibitory agents (e.g. EDTA, Phenol, Proteinase K, Heparin, Hemo group, SDS, Bromophenol blue, Xylen cyanol).
In general terms, the optimal amounts of DNA are 10-250 ng for low complexity templates (e.g. plasmids) while for high complexity templates (e.g. genomic DNA), 250-500 ng must be used.
Use sterile fresh water to eliminate a probable contamination source.
Generally included in all PCR programme as 5-15 minutes at the same extension temperature step. Any Polymerase formulation containing Taq DNA polymerase will produce PCR fragments with 3´adenine overhangs. If a blend formulation is used, a mixture of both PCR fragments with blunt ends as 3′ A overhangs ends are obtained.
Usually the number of cycles used in a standard PCR are 25-30 cycles. But this parameter could be increased (up to 40 cycles) when you have scarce DNA or low copy number template.
The temperature and extension time depends on enzymatic optimal activity temperature and enzyme processivity, respectively. Also, time extension depends on amplicon length. Unlike Horse-Power™ Taq DNA Polymerase and HotBegan™ Hot Start Taq-DNA Polymerase required 1 minute for kb, FastPANGEA™ High Fidelity DNA Polymerase required 30 seconds for every kb to be amplified.
The annealing temperature depends on primer Tm. Although there are different methods to calculate primers Tm, annealing temperature is an empiric value. Most of times, annealing temperature is around 3-5 ºC lower than the calculated primer–template Tm. But sometimes, it is necessary to use a temperature gradient to find the optimal annealing temperature. When PCR additives must be used, annealing temperature has to be adjusted. For example, 10% DMSO reduces the annealing temperature 5-6 ºC.
Annealing time is generally 0.5-2 minutes.
This step ensures a complete denaturation of the template. This first denaturation step with Horse-Power™ Taq DNA Polymerase occurs at 94-95 ºC, 3-5 minutes but it has to be longer (up 10 minutes) if GC content requires it. HotBegan™ Hot Start Taq DNA Polymerase doesn´t require a denaturation step longer than 5 minutes.
PCR cycling conditions depend on the enzyme processivity, template and primers.
Although a concentration of 1,75 -2 mM is suitable for most applications, Mg2+ is an important element to modify in a PCR optimisation. Commonly, the optimisation range used is from 1 to 4 mM. If the Mg2+ concentration is too high, non-specific spurious bands may appear, while if the Mg2+ concentration is too low, the PCR yielding could be reduced. If you are working with RAPDs, 3,5-5 mM Mg2+ could be necessary. Also quantitative PCRs using SYBR-Green require 0,5 mM Mg2+ more than a standard PCR (3 mM final concentration).
PCR buffers are often an invaluable component in the PCR reaction.
The PCR reaction is a versatile and highly robust process that fails in some circumstances. Frequently, this failure is attributed to the “current” or “new” polymerase without considering that the novel PCR is not optimised. Thus, is very important to know how to adjust the different components of the reaction mixture or the PCR programme.
For most researches, it is sufficient if the transfected genetic material is only transiently expressed. It is usually not integrated into the nuclear genome, so the foreign DNA will be diluted or degraded through Mitosis. To accomplish a stable transfection, a marker gene is necessary, which gives the cell some selectable advantage, for example resistance towards a certain Antibiotic. When the antibiotic is added to the transfected cells, only those cells with the DNA integrated into their genomes will be able to proliferate.
24-48 h after Transfection, the efficiency should be analyzed. The optimal time interval depends on the cell type and the specific transfected genes. It is advisable to observe the state of the cells and see if there are dead cells and changes in morphology. With a viability study it is possible to know its toxicity grade. Transfected gene products may be toxic. You can use reporter genes to easily monitor transfection efficiency. Reporter genes are luciferase (a green fluorescent protein) and b-galactosidase. Check the transfection efficiency using an appropriate assay for the reporter gene.
The optimal Transfection time depends on the Transfection Reagent, Nucleic Acid, and cell line used. For optimisation, test Transfection times from 30 minutes to 4 hours or overnight. There are cells that lose viability with extended time, especially if the transfection is happening in Serum free medium.
Before stable Transfection, you will need to determine the amount of Antibiotic necessary to eliminate untransfected cells. This may vary from one cell type to another. You must do a survivor curve with different drug concentrations, and after a few days to see what is the minimum concentration at which cells die. It is often used as selection drug Geneticin (G-418), Hygromycin, Puromycin and Blasticidin.
Charge ratios of DNA: Transfection Reagent depends upon the type of DNA, Transfection Reagent, and target cell line. It is advisable to test different ratios of DNA /Transfection reagent, as recommended by the manufacturer. Each ratio may be optimal for certain cell types or applications but not others. Once the ratio has been optimised, the optimal amount of DNA and Transfection Reagent will vary depending on the culture dish and number of cells.
When the cells are adherent is recommended seeding them the day before Transfection at a density such that the next day they are at least 60-70 % confluent. Some cell types have higher toxicity when transfected at low density. It is recommended to use a 24-well plate format to optimise Transfection conditions for a particular cell type before initiating a stable transfection. To transfect cells in different tissue culture formats, vary the amounts of Transfection Reagent, DNA cells, medium used in proportion to the relative surface area. Follow the recommendations of specific Transfection Reagent.
Nucleic acids need to be of high quality, free of proteins, other contaminating nucleic acid and chemical contamination, i.e, a A260/A280 ratio of 1.8-2.0. DNA suspension will be in sterile water or TE Buffer to a final concentration of 0,2-1 mg/mL. Endotoxin-free DNA is not necessary for a standard Transfection protocol, although it is indispensable endonuclease-free DNA purification for Transfection of viral and Packaging Systems to produce viral supernatants. The optimal amount of DNA to transfect cells depends upon the type of DNA and target cell line, and number of cells used. Increasing the amount of DNA does not necessary result in a better transfection. It is advisable to initially test different amounts of DNA, as recommended by the specific transfection reagent.
Cells must be grown in an appropriate medium and supplemented with the corresponding serum and growth factors. This medium must be fresh. On the day of Transfection, it is crucial to know whether to change the culture medium, which will depend on the Transfection Reagent selected, or if the presence of serum and other additives may interfere with the process of DNA release inside the cell. The Transfection mixture is prepared in free-serum culture medium and without additives so they don´t interfere with the complex DNA-transfection reagent formation. The addition of antibiotics to media during transfection may result in cell death because cells are more permeable to them.
There are a lot of Transfection methods in the market such as chemical reagents, cationic lipids and physical methods. So far, the most popular gene transfection reagents are cationic lipids and cationic polymers, but they have several limitations. One given reagent may work better with certain cells than others.
On the other hand, any Transfection Reagent or Transfection method cause cell death. CANFAST™ Transfection Reagent is a new generation cationic polymer, a high-quality gene transfection reagent and non-toxic for cells. It is effective with many different types of cultured cells and makes easier for the DNA / transfection reagent complex to cross the membrane. CANFAST™ should be used for transient expression and long-term studies.
Transfection is the process of introducing Nucleic Acids into cells deliberately. For maximum efficiency, Transfection must be optimised on the culture conditions and in the transfection protocol, including optimisation of the relation mg DNA vs mL Transfection Reagent, medium used, length of incubation, etc.
To transfect cells, it is very important to consider those culture conditions that can influence transfection efficiency, such as cell health, exponential growth, cell passage number and degree of confluency. The cells to be transfected must show optimal morphological conditions and it is necessary to verify that they are not contaminated with mycoplasma or with other microbial contaminants.
They must be in an exponential growth phase and cells must not be allowed to remain confluent for more than 24 hours. Besides, excessive passages of cells must be avoided because it decreases transfection performance. For stable transfection studies, during selection, the selective medium must be changed several times a week to eliminate dead cells and debris until colonies can be visualized.