T4 DNA ligase requires a 5’ phosphate on one of the DNA molecules to be ligated in order to join DNA, for this reason it is often necessary to phosphorylate DNA molecule prior to adding it to ligation, for example when blunt cloning a PCR product.
Nuclease free water: 4.5 ul
PCR product or other DNA (cleaned-up): 4 ul (*if your PCR product has 5' recessed or blunt ends, heat it to 70 degrees C for 5 minutes and cool on ice before adding it to the reaction)
10 x T4 DNA Ligase buffer: 1 ul (* ligase buffer is used because it contains ATP)
T4 Polynucleotide kinase: 0.5 ul
Author: Dr Ryan Cawood
Sometimes it is necessary to make the ends of a DNA molecule blunt, for instance:
DNA should be dissolved in 1x restriction digest buffer (for example, NEBuffers 1-4) or T4 DNA Ligase Reaction Buffer supplemented with 33 µM each dNTP [final concentration].
Add 1 unit of Klenow per microgram of DNA and incubate for 15 minutes at 25°C.
Stop the reaction by adding EDTA to a final concentration of 10mM and then heating at 75°C for 20 minutes.
DNA should be dissolved in 1x restriction digest buffer (for example, NEBuffers 1-4) & supplemented with 100 µM dNTPs [final].
Add 1 unit T4 DNA Polymerase per microgram DNA and incubate for 15 minutes at 12°C.
Stop the reaction by adding EDTA to a final concentration of 10 mM and heating to 75°C for 20 minutes.
Suspend DNA (0.1 µg/µl) in 1X Mung Bean Nuclease Buffer or a restriction enzyme buffer (for example, 1X NEBuffer 1, 2 or 4).
Add 1.0 unit of Mung Bean Nuclease per µg DNA.
Incubate at 30°C for 30 minutes.
DO NOT attempt to heat inactivate Mung bean nuclease because single stranded regions of DNA may appear before the enzyme is inactivated resulting in unintended degradation. Inactivate the enzyme by spin column purification or by phenol/chloroform extraction and ethanol precipitation.
Author: Dr Ryan Cawood
Blue / White colony screening is a strategy to quickly and easily distinguish between recombinant and non-recombinant colonies. It requires a special vector and a special strain of E. coli. It is particularly helpful in tricky cloning strategies such as blunt ended cloning or DNA library preparation.
The first gene in the E. coli lac operon is lacZ, which encodes ß-galactosidase (ß-gal). The active form of ß-gal is a tetramer and hydrolyses lactose into glucose and galactose. Deleting amino acids 11-41 of ß-gal (called the lacZ?M15 mutation) means the enzyme is unable to form a tetramer and is non-functional (Langley et al. 1975). It was discovered that supplying amino acids 1-59 (the a-peptide) of ß-gal in trans (separately) allowed the truncated ß-gal to form tetramers and function again (Ullmann et al. 1967; Langley et al. 1975). Rescuing ß-gal by supplying the a-peptide in this way was termed a-complementation. Later, Vieira and colleagues (Vieira & Messing 1982) realised that a-complementation could be used to screen E. coli colonies for the presence of inserts. They cloned the a-peptide coding region into a pUC plasmid and then introduced a multiple cloning site (MCS) into the middle of that region. When a piece of DNA is ligated into the MCS, it disrupts the a-peptide, rendering the ß-gal non-functional.
5-bromo-4-chloro-indolyl-ß-D-galactopyranoside (x-gal) is a colourless analogue of lactose. When ß-galactosidase hydrolyses x-gal, it creates a blue product (5,5'-dibromo-4,4'-dichloro-indigo). In blue white screening, an E. coli strain is transformed with a ligation reaction and spread onto agar plates containing x-gal. A blue coloured colony indicates that the a-peptide in the plasmid is intact (no insert) whereas a white colony indicates that the a-peptide is disrupted (insert present).
Competent cells of an E. coli strain with the lacZ?M15 mutation. Common Blue white compatible strains include: XL1-Blue, XL2-Blue, DH5a F', DH10B, JM101, JM109 and STBL4. Please see the 'Competent cells' protocol for details about preparing E. coli cells for transformation.
A vector with the a-peptide coding region and MCS. Common blue white compatible vectors include: pGEM-T, pBluescript, pUC18 and pUC19 (see button below).
Your ligation reaction (i.e. your insert of choice into a blue white compatible vector, above).
Control plasmid (e.g. pBluescript).
Antibiotic for selecting for the vector.
X-gal 20 mg/ml. X-gal can be purchased ready dissolved or as a powder. It may be dissolved in DMSO or DMF at a concentration of 20 mg/ml. X-gal must be stored at -20°C and protected from light (by wrapping foil around the stock container).
Isopropyl ß-D-1-thiogalactopyranoside (IPTG) 10 mM (needed to induce ß-gal expression in the E.coli)
Prepare some LB agar plates containing the appropriate antibiotic to select for your chosen plasmid. Please see the 'Making agar plates' and 'Antibiotic concentrations' protocols.
Onto each plate to be used for blue white screening, spread:
and allow the plates to dry with the lid slightly open before use. This can be performed either next to a bunsen burner on the bench or in a laminar flow hood. Using a hood may dry out the plates if they are left for too long.
Transform your ligation reaction(s) into competent E. coli cells as usual. Spread the transformation reaction onto an x-gal IPTG plate (prepared as above). Incubate the plate overnight at 37°C. Once the colonies have grown, the plate may be incubated at 4°C for 1 hour. This helps the blue colour to develop making it easier to discern the negative colonies.
It is a good idea to include a control. Transform an aliquot of E. coli with an intact a-peptide-containing-plasmid (pUC19 for instance, see button below). The colonies on this control plate should all be blue. If they're not, then the x-gal may not have been spread evenly or the antibiotic may not be working properly.
This screen does not give any information about the direction of an insert, just its presence or absence. If the insert is quite short and maintains the frame of the a-peptide, it is possible (but unlikely) that it will produce a functional a-peptide fusion giving blue colonies even when the insert is there (false negatives). However, these colonies will likely be a lighter blue than the true negatives. It is also possible (but again unlikely) to get a white colony with no insert (false positive). This could result from nuclease degradation of the linearised vector disrupting the a-peptide before re-ligation. Therefore, it is always a good idea check your insert by sequencing too.
Oxford Genetics Other
Expression Plasmids
Langley, K.E. et al., 1975. Molecular basis of beta-galactosidase alpha-complementation. Proceedings of the National Academy of Sciences of the United States of America, 72(4), pp.1254–1257.
Ullmann, A., Jacob, F. & Monod, J., 1967. Characterization by in vitro complementation of a peptide corresponding to an operator-proximal segment of the beta-galactosidase structural gene of Escherichia coli. Journal of molecular biology, 24(2), pp.339–343.
Vieira, J. & Messing, J., 1982. The pUC plasmids, an M13mp7-derived system for insertion mutagenesis and sequencing with synthetic universal primers. Gene, 19(3), pp.259–268.
Author: Dr Richard Parker-Manuel and Dr Ryan Cawood
Agarose gels are an easy, cheap and effective method of separating, and viewing, DNA. The agarose concentration of the gel you make is determined by the size of the DNA fragments you intend to resolve, and view. For the majority of DNA work, a gel of 1% agarose is fine and is good for DNA sizes that are between 400bp and 10kb (roughly) and where the two DNA fragments you want to resolve are not of a similar size. If they are similar sizes then you may have to change the concentration. For example, for two fragments of 500 and 700 bp a 2-3% gel would help to separate them. For two fragments that are 10-11kb you would probably need a 0.5% agarose gel, and run it for a long time at a slightly lower voltage (to prevent melting).
Gels can be made with either TAE or TBE buffers. There are differences between them that may, or may not, be important to you. See the notes and tips on the buffers page.
Making Agarose Gels - Protocol:
Tip 1: Many lab bottles have a blue or clear plastic rim which can go missing or get burnt by flaming. These blue rims prevent the liquids running down the bottle when pouring and keep an air tight seal after autoclaving. Make sure the rim is intact.
Tip 2: Some companies sell low melt agarose which can help with making the gel. Remember that some gel runs, particularly in TAE, can cause excessive heat. This may damage your gel on extended high voltage runs.
Tip 3: Running gels at above 10 volts per gel cm length can cause the gel to melt.
Tip 4: Try to make your gel from the buffer you intend to run it in, if not, you may get a conductivity imbalance. Also, old buffers in tanks that have been repeatedly topped up can have a higher salt concentration than a gel made from the same buffer because of evaporation. This can also cause a conductivity imbalance and lead to poor gels.
Note 1: Using SYBR Safe DNA Gel Stain is a safer, non-carcinogenic alternative to ethidium bromide. We have not verified its activity.
Author: Dr Ryan Cawood
It is often useful to amplify a region of DNA to contain new restriction sites on the ends to allow easier downstream cloning. When designing primers for this purpose it is important to add extra nucleotides to the 5' end of each primer to allow the restriction enzymes to cut the DNA efficiently. Most enzymes like a minimum of three nucleotides but some will require four.
It is also important to consider that the melting point (TM) of the primers will vary depending on the stage of the reaction. During the first two rounds of the PCR reaction the primers will only bind the target with a sub-section of the primer, however, once the new DNA (with restriction sites on the ends) has been produced after the first few rounds of the reaction, the primers will bind along their whole length (with a higher TM). Normally, this is not a problem and a single melting point can be used, but if the amplification is inefficient, perhaps start with a lower melting point (that of the section that binds the target DNA) for the first few rounds of the reaction.
Author: Dr Ryan Cawood
The basic concept of annealing oligos is to heat two oligonucleotides up such that they denature, then follow this by a period of cooling to allow the two oligos to base pair together.
This process is often to used to prepare short DNA sections for:
Oligo and primer stocks are often resuspended at 100µM (100 picomoles/ul) concentrations.
Author: Dr Ryan Cawood
Precipitating DNA can be hit and miss. Sometimes you end up the same quantity and its more concentrated and cleaner, sometimes you end up with nothing. If you do it right then it should work fine but working with low concentrations of DNA (<20ng/µl) in large volumes can be tricky.
Tip 1: Performing the precipitation at 4°C is not strictly necessary for it to work but it can increase the efficiency, especially of low concentration samples (<20ng/ µl).
Note 1: DNA precipitation can be used to clean a DNA sample, not just concentrate it. It will remove any contaminating salts from the solution.
Author: Dr Ryan Cawood
This recipe is designed to make approximately 50-60 plates of Luria-Bertani (LB) agar plates for growing E. Coli with standard plasmids. To watch the video of this protocol click here.
10g Tryptone
5g Yeast extract
10g NaCl
15g Agar powder
To 1 litre with water
Potential Hazard: Powders can be sensitising and you can develop allergies to them. Avoid inhalation of aerosolised powder.
Measure out the amounts above and add to a 1 litre bottle with a good working lid (See below). Fill with clean deionised water (ideally at least 18 megaohms). Tighten lid and shake to mix the liquid and powder, don’t expect to dissolve it all but simply to free the powder from the bottom and remove the major clumps. If not mixed properly the powder can bake on the bottom of the bottle. Undo the lid about half a turn; add some autoclave tape and then autoclave.
After autoclaving ensure that the lid is done up tight and do not allow to cool below 45-50 degrees. Agar generally sets at about 40 degrees. As a guide if you can hold the bottle comfortably in your hand then it is ready to pour and may actually be too cool already. Don’t add the antibiotic when the agar is too hot, this can affect the antibiotic stability and half life, particularly Ampicillin. Add the correct amount of antibiotic (see below) and mix by gentle swirling. Avoid getting bubbles as this will result in bubbly or uneven plates. Pour about 12-15ml per plate. A simple method is to stack the plates up with the large sections (lids) upright. Lift the entire stack up using the lid of the bottom plate and using your other hand pour in the agar. Pour enough to fill the bottom and then a little bit more. Then put the stack back down and lift again with the next lid in the stack. Sounds inconsistent but with time this is very reproducible.
Tip 1: Many lab bottles have a blue or clear plastic rim which can go missing or get burnt by flaming. These blue rims prevent the agar running down the bottle when pouring the plates and keep an air tight seal after autoclaving. Make sure the rim is intact.
Tip 2: 1 litre is quite a lot of agar and antibiotic, scale down the recipe to suit your needs. I generally make 0.5 litres at a time ( about 30 Plates).
Note 1: Antibiotic concentrations:
Stored in water: add 1µl per 1ml of media
Kanamycin – Stock 50mg/ml, final concentration 50µg/ml
Ampicillin - Stock 100mg/ml, final concentration 100µg/ml
Streptomycin – Stock 50mg/ml, final concentration 50µg/ml
Spectinomycin - Stock 100mg/ml, final concentration 100µg/ml
Stored in Ethanol: Add 5µl per ml media
Chloramphenicol - Stock 34mg/ml, final concentration 170µg/ml
Tetracycline HCL - Stock 10mg/ml, final concentration 50µg/ml
Note 2: Pre-mixed LB powders are available from many suppliers and don’t actually work out much more, if any, expensive.
Author: Dr Ryan Cawood
TE 50/1 buffer (50mM Tris-HCL pH 8.0, 1mM EDTA pH 8.0)
Lysozyme at 10mg/ml in water.
0.5M EDTA pH 8.0
Ribonuclease A solution at 20 mg/ml in distilled water and stored in 50µl aliquots in -20°C.
10% Triton X 100 in water
Equilibrated Phenol (pH 8.0 with 0.1% 8-hydroxyquinoline)
5M NaClO4
Isopropanol
TE (10mM Tris-HCl pH 8.0; 0.1 mM EDTA pH 8.0)
SS34 Sorvall 50ml tubes and rotor or similar
30ml Corex tubes and HB4 rotor or similar
Potential Hazard: Phenol is very dangerous and causes skin burns immediately on contact. It is toxic on contact, by inhalation and by consumption. Burns should be treated immediately with 20% Poly Ethylene Glycol (PEG) 300 or 600 water solution.
Read the protocol before starting and make sure you have all the stock solutions ready. The lysozyme solution is usually made during the centrifugation period. The rest has to be ready before you start.
Author: Dr Ryan Cawood
TE
RNase A at 20mg/ml in water
Phenol
Chloroform
5M NaClO4
Isopropanol
Potential Hazard: Phenol is very dangerous and causes skin burns immediately on contact. It is toxic on contact, by inhalation and by consumption. Burns should be treated immediately with 20% Poly Ethylene Glycol (PEG) 300 or 600 water solution.
Potential Hazard: Chloroform is toxic by inhalation, oral consumption and skin contact. Wear gloves and use a fume hood. If skin is contacted then wash immediately with plenty of water.
Author: Dr Ryan Cawood
Inoue and colleagues developed this method in 1990. It works well for many strains commonly used in cloning. The original method calls for growing the overnight E. coli cultures at 18°C. We find that it still works well if they are grown overnight at 37°C.
You will need:
0.5 M PIPES buffer:
Inoue Transformation Buffer:
Component Final concentration Amount per litre
MnCl2.4H2O 55 mM 10.88 g
CaCl2.2H2O 15 mM 2.2 g
KCl 250 mM 18.65 g
0.5 M PIPES pH 6.7 10 mM 20 ml
H2O Up to 1 litre
SOB:
Component Final concentration
Tryptone 20 g/l
Yeast extract 5 g/l
NaCl 0.5 g/l
KCl 2.5 mM
MgCl2 10 mM*
*The MgCl2 must be added after autoclaving from a 1 M stock.
Day 1
Authors: Dr Richard Parker-Manuel and Dr Ryan Cawood
Site directed mutagenesis (SDM) is a useful technique for introducing a specific mutation into a plasmid at a specific site. The mutation can be a substitution, insertion or deletion. There are many applications for SDM, for instance to assess the function of certain amino acids in an enzyme, to investigate the effect of altering bases in a promoter or removing restriction sites from a plasmid.
Here we describe a PCR based site directed mutagenesis method. The basic principle is to design a pair of PCR primers back to back, so that the entire plasmid is amplified by PCR. One of these primers incorporates the desired mutation. The PCR creates a linear product whose ends can then be joined together (after phosphorylation) with DNA ligase. The circularised vector is transformed into E. coli.
Design a pair of primers incorporating your desired mutation into the 5' end of one of them. Design them so that the complementary region has a Tm of around 60 degrees C.
Example:
Note that only the forward primer contains the mutation, so you could easily make a series of different mutations by keeping the rev primer the same but using different forward primers.
The above example is a 3 bp substitution but an insertion can be made in the same way. If you require a very large substitution or insertion, then mutant bases may be introduced at the 5' end of both primers.
Deletions of any size can be made by spacing the forward and reverse primers apart on the template.
PCR primers normally come without a phosphate group on their 5' termini due to the way they are synthesised. This means that the ends of a PCR product cannot simply be ligated together, they must be phosphorylated first. There are two main options for this: 1) you could order the primers with a phosphate already added to the 5' end, or 2) you could phosphorylate the PCR product using polynucleotide kinase (PNK). Phosphorylated primers are a good idea if you are only doing a few SDMs and don't have any PNK in the freezer (plus it cuts out a step). Using PNK is more cost effective if you're doing a lot of SDMs (probably 10 or more)
It is important to use a proofreading polymerase to avoid introducing any other mutations. That said, if you were still worried about introducing mutations you could sub-clone the mutated fragment back into the same backbone after the mutagenesis.
Because the method is based on PCR, it tends to work better on smaller plasmids. It is best to follow the instructions that come with the polymerase, but set up a PCR similar to this example:
35.5µl water
5µl 10x polymerase buffer
1.5µl Forward primer (0.3µM final)
1.5µl Reverse primer (0.3µM final)
5ul dNTPs (200 µM final)
1ul Template DNA (a 1 in 100 dilution of a mini-prep)
0.5µl polymerase
Set the reaction up on ice, and transfer the tube to a pre-heated PCR block straight after mixing the reaction.
PCR program:
1. 98°C 60 seconds
2. 98°C 8 seconds
3. 55-65°C 20 seconds
4. 72°C (the elongation time depends on the plasmid size and the type of polymerase used)
repeat/cycle steps 2-4 a further 27-30 more times
5. 72°C 5 minutes
Hold at room temperature.
Run the entire reaction on an agarose gel. Excise the band and clean up the DNA using a gel extraction kit, eluting it in 30µl.
*This step can be omitted if you used phosphorylated oligos in the PCR.
4.5µl Nuclease free water
4µl PCR product
1µl 10x T4 DNA Ligase buffer*
0.5µl T4 Polynucleotide kinase
Incubate at 37°C for 40 mins.
*Ligase buffer is used because it already contains ATP and PNK is active in it.
Set the ligation reaction up on ice.
6.7µl Nuclease free water
2µl PNK Reaction (from step 5)
0.8µl 10x T4 DNA Ligase buffer
0.5µl T4 DNA Ligase
Incubate at 16°C overnight or at room temperature for 2 hours.
Authors: Dr Richard Parker-Manuel and Dr Ryan Cawood.