Fragment Restriction Digestion

This is a protocol for a preparative digest, which is the cutting of DNA to prepare it for ligation with another piece of DNA, not simply to confirm the identity of the DNA. Usually these pieces of DNA are circular plasmids but they could also be PCR fragments.

You should aim to set up the digest of both your fragment and vector at the same time. This will save you time later.

This protocols assume a plasmid solution in Tris-EDTA (TE)/Elution buffer (EB) or nuclease free H2O that is 200-300 ng/µl and about 3-5 kb. Larger plasmids may require you to use more volume because at the same concentration there will be less plasmid copies.


DNA Fragment Digestion Reagents

In this particular example, the following could be done:

  • 10-20 µl of plasmid (200-300 ng/µl, total amount 3-5 µg)
  • 7.5 µl of 10X Restriction buffer
  • 7.5 µl of 10X Bovine Serum Albumin (BSA, final concentration is usually 100 µg/ml)
  • 1.5-2 µl of Restriction enzyme 1(10-20 u/µl)
  • 1.5-2 µl Restriction Enzyme 2 (Optional)
  • Make up to 70 µl total with TE or water


DNA Fragment Restriction Digestion Protocol

  1. Start mixing the reagents in a 1.5ml eppendorf, first the TE or water, then the plasmid/DNA, then the restriction buffer and BSA, and then finally the enzyme.

  2. Remember that the enzyme is stored in glycerol and will sink to the bottom of the tube, so mix the reaction thoroughly with a pipette.

  3. Incubate the reaction at the right temperature (usually 37°C but always make sure that you are not dealing with an exception) and start your alarm clock (count up).

  4. Now make an agarose gel, make a large well in the gel for the fragment. This can be done by placing tape over multiple small tabs of a comb, or by using a comb with pre-designed large wells.

  5. After 40 minutes/1 hour add about 5-10µl of loading dye to the total reaction and load the entire sample carefully into a large well of the agarose gel. Also load a molecular weight marker in an adjacent standard lane so you can see the size of your DNA.

  6. Run the gel for 40-60 minutes at approximately 10 volts per cm of gel length. 12cm long gel = 120 volts.

  7. Proceed to the gel extraction.


Note 1: This reaction is scaled down to allow easy loading into a larger gel comb slot. See agarose gel for more information on making the gel. You can scale it down further but try to avoid having >5% glycerol content as this can cause enzyme STAR activity.

Note 2: Many companies supply 100X BSA. Dilute this down to 10X into 100 µl aliquots with water and store at -20°C.


Tip 1: It is advised you read both the vector and fragment sections before starting. You can do both at the same time.

Tip 2: Restriction enzymes are very sensitive to temperature change so don’t put them on ice. Use a -20°C cool box. Avoid holding them at the bottom of the tube and try to be quick. They will last a long time if you look after them.


Author: Dr Ryan Cawood


Plasmid Restriction Digestion

Plasmid Vector Restriction Digest Protocol Background Information


A preparative digest is the cutting of DNA to prepare it for ligation with another piece of DNA, not simply to confirm the identity of the DNA.

You should aim to set up the digest of both your fragment and vector at the same time. This will save you time.

This protocol assumes a plasmid solution in Tris-EDTA (TE)/Elution buffer (EB) or nuclease free H2O that is 200-300 ng/µl and about 3-5 kb. Larger plasmids may require you to use more volume because at the same concentration there will be less plasmid copies.


Plasmid Restriction Digest Reagents

In this particular example, the following could be done:

10-20 µl of plasmid (200-300 ng/µl, total amount 3-5 µg)
10 µl of 10X Restriction buffer
10 µl of 10X Bovine Serum Albumin (BSA, final concentration is usually 100 µg/ml)
1.5-2 µl Restriction enzyme 1(10-20 units/µl)
1.5-2 µl Restriction Enzyme 2 (Optional)
Make up to 70 or 100µl total volume with TE or water


Other reagents:

5 X Loading dye


Plasmid Restriction Digest Protocol

  1. Prepare 2 Eppendorf tubes with 7 µl of water and 2 µl loading dye and write 0 and 40 (or 60, minutes) on them.

  2. Then start mixing the reagents for the digest in a new seperate sterile 1.5ml Eppendorf, first add the TE or water, then the plasmid/DNA, then the restriction buffer and BSA, and mix thoroughly. No enzyme is added at this point.

  3. Take 1-2 µl of the mixture and add it to the loading dye in the tube labelled 0 from step 1.

  4. Then add the enzyme. The enzyme is stored in glycerol and will sink to the bottom of the tube, so mix it thoroughly with the pipette tip.

  5. Incubate the reaction at the right temperature (usually 37 °C, but always make sure that you are not dealing with an exception, like SwaI at 25 °C) and start your alarm clock (count up).

  6. Now make an agarose gel. A gel will take about 20 minutes to set.

  7. After the reaction has run for 40-60 minutes, take another 1µl from the digest and mix it with the tube labelled 40 from step 1.

  8. Now you can de-phosphorylate the vector using alkaline phosphatase (CIP). You may wish to deactivate the restriction enzymes at this point, this is standard practice. We have performed the dephosphorylation many times with the restriction enzymes still active for the length of the dephosphorylation reaction, it does not appear to affect the cloning efficiency, but it may affect certain enzymes we have not used.

  9. Whilst the de-phosphorylation is incubating, load the tubes labelled 0 and 40 (or 60) on the agarose gel. Also load a molecular weight marker so you can see the size of your DNA.

  10. Load your DNA fragment (to be ligated into the vector, from the protocol for restriction digesting the fragment) into a large well of a gel at this point as well.

  11. Run the gel for 40-50 minutes at approximately 10 volts per cm of gel length. 12cm gel = 120 volts.

  12. If your fragment (from Step 9) is in the same gel as your vector samples (0 and 40) then don’t put the gel in the UV transilluminator yet as this will damage the DNA fragment. If it isn’t, then check the vector samples in the gel on the transilluminator and decide what to do next. Roughly 3 possibilities exist if the reaction:

    • has not worked at all (i.e. the DNA in the 0 tube looks the same as that in the 40 tube), there is no point in waiting longer because the reaction has failed. If the enzyme has worked on another plasmid before, perhaps re-precipitate the plasmid your are trying to cut, or do a new phenol extraction prior to precipitation and try again. Otherwise, check if the enzyme is still OK by using another plasmid.

    • has worked but the vector is not completely cut after 40 minutes (visible as more than one band present at the approximate molcular weight you are expecting from the vector), estimate from the progress how much longer it will take. Remember that if you haven’t inactivated the enzyme yet it will still have been cutting whilst the gel was running. You could increase the amount of enzyme but be careful of increasing the glycerol content >5% as this can cause STAR activity. A new 1µl sample can then be run on a gel (in loading dye) to make sure.

    • was complete after 40 minutes, then you can stop the reaction if the alkaline phosphatase reaction has finished. Leaving the restriction enzymes active during the gel running usually does not have any detrimental effect but if you have any problems later you could consider that it the DNA may be over-digested, although this is rare.



Note 1: Many companies supply 100X BSA. Dilute this down to 10X into 100 µl aliquots with water and store at -20°C.

Note 2: The total reaction volume is decided by whether you will need to load the vector onto a gel to extract it from another piece of DNA. This is only necessary if the other fragment can contaminate the final product. In this instance a 100 µl reaction will probably be too big to load into a large well.


Tip 1: It is advised you read both the vector and fragment sections before starting. You can do both at the same time.

Tip 2: Restriction enzymes are very sensitive to temperature change so avoid putting them on ice. Ideally use a -20°C cool box. Also, avoid holding them at the bottom of the tube with your fingers and try to work quickly whilst handling the enzyme stocks. They will last a long time if you look after them.


Author: Dr Ryan Cawood


Dephosphorylating DNA

DNA Dephosphorylation Background Information


The purpose of de-phosphorylating the vector is to prevent it from ligating back on itself during the ligation step by removing the 5' phosphate groups that are required by DNA ligase to join the phosphodiester DNA backbone together. Various alkaline phosphatases exist, including Calf Intestinal Phosphatase (CIP), Shrimp and Antarctic phosphatases. The most commonly used is calf intestinal phosphatase but it is difficult to heat inactivate. For this reason, Antarctic phosphatase was developed which can be heat inactivated, although we recommend using CIP. The temperature and buffers of the different enzymes can be different, refer to the manufacturers instructions.


DNA Dephosphorylation - Protocol 1 (for dephosphorylating DNA in a restriction digestion reaction/solution):

50-100 µl of DNA (5µg) in a restriction digest reaction/solution
1-2 µl of CIP enzyme (1unit/µl)

1) Add 1-2 µl of CIP to your restriction digest. CIP is stable and active in most restriction digestion buffers. For example, NEB CIP works in all 4 NEB buffers. Be careful of STAR activity caused by excessive glycerol if you haven’t inactivated the restriction enzymes in the reaction. Incubate the sample for 30-60 minutes at 37°C.


DNA Dephosphorylation Protocol 2 (for dephosphorylating DNA in TE or H2O):

20-40 µl of DNA (5µg) in TE or H2O
5 µl of 10xCIP buffer
1-2 µl of CIP enzyme (1unit/µl)
Make up to 50 µl

1) In a sterile 1.5ml eppendorf, add the DNA, then the CIP buffer and then the 1-2 µl of CIP. Mix thoroughly with a pipette tip and incubate for 30-60 minutes at 37°C.


Tip 1: Make sure your fragment you are going to ligate into the dephosphorylated vector possesses 5’phosphate groups. Standard oligos/primers and PCR products are generally not phosphorylated and must be treated with T4 polynucleotide kinase (see phosphorylating 5’ ends). It is usually easier to add restriction sites to the ends of a PCR product (plus a few extra base pairs at the ends), rather than phosphorylating a fragment.

Tip 2: CIP is stored in a glycerol buffer for stability, but this means it sinks to the bottom of aqueous solutions. When adding the CIP, watch it drop into the DNA mixture by doing it with the tube held up in front of you, and then ensure it is properly re-suspended before incubation. 



Author: Dr Ryan Cawood


Gel Extraction of DNA

Gel Extraction of DNA Fragments Protocol

  1. To make an agarose gel please see the making a gel protocol. For loading DNA fragments for extraction you will normally require a large well in the gel. This can either be achieved by using specifically designed gel combs or by placing some tape over multiple tabs on the comb to make a larger well. Carefully trim off any excess tape with a scalpal. Autoclave tape and electrical tape work well, masking tape is terrible and leaves fibres in the well after you take the comb out. 

  2. To excise the band you will need a clean scalpel or razor blade. An important consideration is the amount of agarose you take with DNA. The higher the ratio is between the quantity of DNA to agarose the less contaminants you will get carrying through to the ligation.

  3. The standard method used to extract DNA from agarose gels is by using UV light. This is not ideal because the UV will damage the DNA and can cause mutations and can reduce the ligation efficiency. An alternative is to use a Clare Chemical Dark Reader Trans-illuminator but it can sometimes be difficult to see low quantity bands. If you have a signficant amount of DNA these systems are ideal. Alternatively, you can do a blind excision by running a small quantity of your cut DNA in the lane adjacent to your large well. Then you cut the gel vertically/lengthways between the two lanes, then visualise the small quantity of DNA in the small lane (whilst wearing the appropriate UV protective clothing). Then you can either measure the distance from the well to the DNA fragment you want using a ruler and then cut the DNA out of the large lane without ever putting it in the UV by measuring it. Be very careful with this technique as some bands can be quite small lengthways/top-to-bottom in the gel and can be missed if you are not accurate. Alternatively, the DNA in the small lane can be excised and the remaining gel can be lined up against the DNA in the large well to allow you to identify the position of the DNA in the gel. After excision you can look at the remaining gel to see if you got the band, do it quickly because if you didn't get the DNA you dont want to damage it with the UV.   

  4. Be careful of DNA fragments that are close together. The key of this technique is to seperate the vector backbone from the fragment to prevent contamination. Just because you cant see something when you are excising doesn’t mean it isn’t there, you will only see the peak of a Gaussian distribution of DNA and whilst the bands may look separate they may not be and you may get some DNA from bands that are close by by accident.

  5. Proceed to the cleaning up a DNA fragment from agarose gels protocol.


Note: This is a relatively dangerous proceedure as far as molecular biology goes. UV light, carcinogenic ethidium bromide, and a scalpel are a recipe for disaster. Please abide by all local safety rules and regulations and consult with the local safety officer if you are unsure about the necessary precautions to take.



Author: Dr Ryan Cawood


Clean Up of Gel Fragments

Specialised Kits for Gel Extraction of DNA


For gel extracting a DNA fragment from an agarose gel, it usually isn’t worth using anything other than a bought kit. They are relatively cheap and highly effective. Other methods exist that don’t involve a kit (like dialysis tubing and squashing the gel fragment between parafilm to squeeze out the DNA in solution) but often using these methods result in high levels of contaminants and low DNA yields.



DNA gel extraction kits usually consist of either beads or spin columns. The sizes of DNA fragments that each kit can accomodate can vary, so check that the kit will extract the size range of DNA you are working with. Most kits cover from 100bp – 5 Kb but some have lower or higher capacities.



The principle of both spin column and bead kits is to bind the DNA to something (usually a silica resin) and then wash away the contaminants, either by pelleting the beads, or washing the column. The final elution step releases the DNA from the resin/beads/column to provide you with a clean DNA fragment. However, even the best extraction kit can still carry though some contaminants and, for this reason, using large volumes of a solution that has been produced by gel extraction in a ligation should be avoided. Sometimes ‘adding less is more’ when it comes to adding your fragment to your ligation. Try to avoid making a ligation consist of more than 30% volume of liquid isolated by gel extraction.



Gel Extraction Kit Variability



We have tested six dfferent DNA gel extraction kits in our laboratories, head-to-head, because the process is so essential to our work. We have found that whilst some kits constantly provide high yields and pure DNA, others sometimes provide good yields but sometimes fail, and others consistenly under-perform.



Low DNA spec readings after DNA extraction?



If you gel extract a fragment from a plasmid, you should probably expect the DNA concentration numbers (if using a spectrophotometer) to be low, because you will have started with a large plasmid (most likely around 5kb), then cust out a much smaller sub-fraction (perhaps 1kb), and the kits will never purify 100% back of the DNA from the gel. So a yield of 20ng/ul in 35ul total volume for 1Kb fragment isolated from 5ug of plamsid DNA (5Kb) would not be too bad (70% of the DNA has been recovered).



We frequently find that often the level of contaminants, or DNA prep quality, is a more important determinant of cloning success, rather than the DNA yield itself. We have also consistenly observed that sometimes adding more fragment to a ligation reduces the cloning success compared to the same reaction with slightly less. Although, this could simply be because the extra DNA ends are titrating the ligase away from the vector ends, adding too much gel extracted material may be contributing because of low levels of contaminants in the prep from the gel.



Our recommendation is that once you find a kit that works for your lab, dont change it!



Author: Dr Ryan Cawood





DNA Ligation Protocol

DNA Ligation Protocol Background Information


During the cloning process, all roads lead to the ligation, and so all of the steps that precede it can affect its efficiency significantly. It is important to read the tips and notes at the bottom of this protocol before beginning.
Ligation reactions are set up along with controls. Usually you have two controls which are:

  1. The vector alone without ligase (controls for uncut vector)

  2. The vector with ligase (controls for insufficient dephosphorylation when used in conjuction with control 1)
    And then:
  3. The real ligation which is vector + fragment + ligase.


If you got colonies on control 1 then your restriction digest of your vector did not work because you got colonies even without ligase. If you got colonies on control 2 but not 1 then your alkaline phosphatase treatment didn’t work. This is because your digest worked well (no colonies on control 1) but the ligase was able to re-circularise the plasmid because of the 5’ phosphates were not removed during dephosphorylation. If you got colonies on 3 and none (or just less) on 1 and 2 then you should congratulate yourself, pick some colonies and go home for a cup of tea to celebrate.

Reaction conditions can vary between labs. Best results are achieved at 16°C overnight but this means adding an extra day to the whole process (making it a 4 day cloning cycle). Performing a ligation at room temperature for 1-2 hours usually gives good results and shortens the cloning to 3 days for a complete cycle. This is provided you don’t mind a long first day.

The two most difficult types of ligations are ligating PCR products and blunt ligations. You should expect these to be less efficient than standard cloning of a fragment from one vector to another.


DNA Ligation Protocol

A typical reaction could be set up as follows:


Sample Vector Fragment 10x Ligase Buffer Ligase (1U/µl) Water
 Ligation 1µl 3µl 2µl 1µl 13µl
 Control 1 1µl - 2µl 1µl 16µl
 Control 2 1µl - 2µl - 17µl



  1. Start setting up the reaction by adding the components that are shared for all reactions i.e. add the water then the buffer and then the vector to each of the tubes.

  2. Then add the fragment to the ligation tube.

  3. Finally, add the ligase to the ligation tube and the correct control tube. Do not vortex the reaction as DNA ligase is shear sensitive. Instead mix with the tip you added the ligase with by stirring well and/or flicking the tube gently repeatedly with your finger.

  4. Incubate for either 1-2 hours at room temperature, or 16°C overnight

  5. Some people heat inactivate the ligation at 65°C for 20 minutes but this is not strictly necessary.

  6. Proceed to the transformation of E. Coli.

Tip 1: The BSA in the ligase buffer can precipitate out on freeze thawing, visible as a white precipitate at the bottom of the buffer. Re-suspend by vortexing and temporarily warming it between your fingers or at 37°C.

Tip 2: The ligase buffer contains ATP, which degrades after multiple freeze thaw cycles. Keep the buffers stocks in small aliquots and throw away after >3 cycles. Buffer can be spiked with ATP if necessary. New ligase buffer has a distinctive smell, if the buffer has lost this smell it might be time to replace the vial you are using. (Disclaimer: This is not recomendation to put ligase buffer up your nose....)

Tip 3: Heating your ligation (before adding enzyme) to 37°C for a few minutes to open up the sticky ends, or help to linearise transiently re-cirularised vectors when using single restriction site ligations. Cool the ligation back to room temperature before adding the enzyme to avoid damaging the ligase when you add it. We dont do this, but we have heard it can help from others who do.

Tip 4: Treating PCR products with Proteinase K before ligation is reported to help with the ligation. Gel extracting the PCR product would make this unnecessary, as would using a PCR clean up kit to purify it.

Tip 5: Avoid exposing your DNA to UV when extracting it from a gel. This can reduce ligation efficiency dramatically in some cases.

Note 1: Avoid using more than 20-30% of the total ligation volume (normally the total volume is 20 µl) of gel-purified material. If you do, you may have too much salt and other contaminants in your reaction and the ligation efficiency may be reduced.

Note 2: The amount of vector should be barely visible if you were to run it on an agarose gel, so about 20-30 nanograms. Fragments should be more abundant, but not more than 5-10-fold concentrated compared to the vector. We normally use a 1:2 or 1:3 vector to fragment ratio.


Calculating the insert to vector ratio 

The insert to vector molar ratio can have a significant effect on the outcome of a ligation and subsequent transformation steps. Molar ratios can vary from a 1:1 insert to vector to a ratio of 10:1. It may be necessary to try several ratios in parallel for best results. The calculation below will tell you the amount of fragment to us relation to the vector at a ratio of 6:1 (fragment:vector). Change the 3 to anything to work out alternative concentrations.

Author: Dr Ryan Cawood


Bacterial Transformation

Bacterial Transformation

Bacterial Transformation General Information

The easiest ways to get DNA into bacteria are heat shock and electroporation. The principle of heat shock is exactly what it states, you have to shock the bacteria by heating. If it isn’t a shock then it doesn’t work, so keeping the bacteria cold until then is essential. This cannot be over emphasised. When the bacteria go from cold to hot it creates holes in the cell membranes and allows the DNA to enter the cell. Electroporation is a similar principle except the electricity makes the holes. These processes mimic, or are at least based on, the natural process of bacterial competence. Typically electroporation gives more colonies, but not always. For most DNA cloning applications heat shock works fine.

This protocol page was updated in February 2014 to provide the transformation protocol that is more comonly used by most laboratories. Both methods shown here work with equal efficiencies for standard cloning. 


Bacterial Transformation Heat Shock Protocol


  1. Thaw one tube of your pre-made competent cells per DNA/ligation reaction or control reaction on ice and push the tube deep into the ice. Thawing takes about 5-10 minutes.
  2. Keep the cells as cold as possible and avoid touching the part of the tube containing the cells; a small amount of heat can significantly decrease the transformation process.
  3. Pre-chill 15ml Falcon Tubes (Sigma-Aldrich: SIAL0791) on ice and transfer 3-4 μl of the ligation reaction (or control reaction) into each tube.
  4. Add 95 μl of competent cells into each ligation reaction and incubate on ice for 20 minutes (minimum). Longer is OK but we have only tested up to 45 minutes.
  5. Heat shock at 42°C for 90 seconds in either a heat block or water bath.
  6. Then add 1 ml of LB or SOC medium without antibiotic and incubate the cells in an incubation shaker at 37°C, 227RPM for 1 hour.
  7. Pour all the LB containing the transformed competent cells onto an agar plate containing the correct antibiotic.
  8. Leave the plate upright to dry with the lid slightly off in a class 1 hood or in a 37°C incubator for about 5-10 minutes. Do not do this in a hood that is used for mammalian tissue culture. Your colleagues will not be happy. Do not leave the plate to dry for too long as some of the bacteria may die.
  9. Incubate the plate overnight at 37°C. The colonies that will appear originate from single transformed cells and are resistant to the antibiotic due to the presence of the plasmid. Each colony will contain millions of identical copies of the same cell, hence the term clone.
  10. Now move on to Picking your colony.

Bacterial Transformation Heat Shock Protocol - Our old Protocol

  1. Thaw one tube of your pre-made competent cells per DNA/ligation reaction or control reaction on ice and push the tube deep into the ice. Don’t just place it on the ice or just in it; keep the tube as cold as you can. Thawing takes about 5-10 minutes.
  2. Avoid touching the part of the tube containing the cells, a small amount of heat can signficantly decrease the transformation process.
  3. Add 2-5 μl of each ligation reaction (or control reaction) to the 100μl of competent cells. As you pipette out the mixture stir the tip very quickly. Do not pipette up and down. The tip is probably at room temperature so remove it as quickly as possible.
  4. Incubate on ice for 15 minutes (minimum). Longer is OK but we have only tested up to 45 minutes.
  5. Heat shock at 37°C for 3 minutes in either a heat block or water bath.
  6. Then add 1ml of LB medium without antibiotic and incubate the cells for a further 15 minutes at 37°C.
  7. Pour all the LB containing the transformed competent cells onto an agar plate containing the correct antibiotic.
  8. Leave the plate upright to dry with the lid slightly off in a class 1 hood or in a 37°C incubator for about 5-10 minutes. Do not do this in a hood that is used for mammalian tissue culture. Your colleagues will not be happy. Replace the lid and turn the plate over when it is close to being dry. Do not leave the plate to dry for too long as some of the bacteria may die as the agar dries out, you will also end up with a very thin plate of agar! 
  9. Incubate the plate overnight at 37°C. The colonies that will appear originate from single transformed cells and are resistant to the antibiotic due to the presence of the plasmid. Each colony will contain millions of identical copies of the same cell, hence the term clone.
  10. Now move on to Picking your colony.


Authors: Dr Ryan Cawood / Weiheng Su


Picking Bacterial Colonies

Method of Picking Bacterial Colonies


This step is easy and doesn't really warrant a separate protocol but given that it is an important part of the process we have included a brief description. 

After you have performed a transformation with the appropriate controls you should be left with three bacterial plates. If everything has gone correctly you should have lots of colonies on the ligation plate, less colonies (or in a fantasy world no colonies) on the plate with no fragment but with ligase, and even less colonies (or again, no colonies) on the plates with no fragment and ligase.

Often the background will be quite high on the control plates and this does not necessarily mean that the ligation hasn't worked as long as you have more colonies on the ligation plate. The number of colonies you pick for further screening is determined by the ratio of colonies from the control plate (that had ligase in the reaction) to the number of colonies on the ligation plate.

For example, if you have 10 times more colonies on the ligation plate compared to the control plate then in theory (if you are performing standard sticky ended directional cloning) 9 out of 10 of colonies you pick will have the correct fragment ligated into it (this doesn't normally actually work out to be true because of things like multimers or concatemers etc). In this case picking 5-10 colonies should be more than enough.

If you only have twice as many colonies on the ligation plate compared to the control, then picking 10 or more would be wise. The plate can be put in the fridge and more colonies can be picked if necessary. Don't go nuts when picking colonies, picking 50 is a waste of time unless you are doing a very difficult ligation. 

It is important to acknowledge a failure if there is no difference between the plates. In this case, don't bother picking the colonies. Repeat the previous steps and try to decrease the background by either digesting for longer, dephosphorylating for longer, exposing to less UV,  or try someone else's enzymes. Quite often an enzyme has just stopped working.     


Picking Colonies Protocol:

  1. After taking the plates out of the 37ºC incubator place them upside down (i.e. the way they were in the incubator) on the bench top.

  2. Using a pipette boy or similar instrument, pipette 3-5 ml of LB media containing the correct concentration of antibiotic into sterile 25 ml or 50 ml tubes (the number of tubes depends on how many you want to grow) or similar tube with a screw top and label them. The volume to volume ratio of the bacterial culture to the amount of air in the tube is important to allow the bacteria to grow to sufficient density, as an approximate guide never have less than a 1:3 ratio of liquid to air but ideally more.

  3. In one hand take a sterile pipette tip on the end of a pipette, with the other hand pick up the upside down plate containing the bacteria from the ligation. Turn the plate over in your hand so that the bacteria are now facing upwards towards you and touch the tip of the pipette tip gently to a bacterial colony that is completely isolated from any other colony.

  4. Now place the same tip with bacteria on it into one of the tubes containing LB media (from step 2) and move the tip around a bit to release some of the bacteria into the liquid. Some people simply eject the pipette tip into the media but if you do this you will need to recover it the next day.       

  5. Culture the tubes overnight in an incubated orbital shaker at 37ºC at 190-225 rpm. The next day you can either continue to the miniprep protocol on this website, or follow the manufacturer's instructions for DNA isolation using a miniprep kit.        


Note: The above procedure should be performed with sterile technique. Ideally in close proximity to a Bunsen burner, if this is not available a class 1 hood will suffice. In reality, it is probably more important not to cross contaminate each sample with each other because you are growing them all in the same antibiotic, than worrying about external bacteria contaminating your samples (because they shouldn't have the resistance cassette, unless they come from a previous experiment off your bench).   


Author: Dr Ryan Cawood


Miniprep Guide and Protocol

What is a miniprep?


The ability to work with DNA is dependent on the ability to isolate it from bacteria. The amount of DNA that is required for analysis determines the preparation technique that is used. For small quantities of DNA (<5µg) a miniprep is performed and normally provides plenty of DNA for analysis. If more DNA is required, a maxiprep can be performed, and if even more DNA is required a Mega or Giga prep can be used.

A miniprep is most often used for determining if a bacterial clone contains the correct piece of recombinant DNA. After picking colonies (typically 5-15), and growing each one in 3-5ml of LB media overnight, the bacteria are pelleted and then lysed. The genome of bacteria is tethered to the inner surface of the plasma membrane and as such stays with the membranous portion of the cell during the extraction. Small plasmids are not tethered in this manner and can be easily isolated from the cell debris.

Many laboratories use only spin columns to perform minipreps. Each miniprep usually works out at about £1($1.5) but the cost varies with the supplier. Many laboratories cannot afford this and will regularly perform the miniprep protocol that is listed on this website. The protocol doesn’t take long, or cost much, but typically the yield is lower and the DNA is of a poor quality than that isolated using spin column kits. However, for screening purposes the technique works well.

For most in vitro studies in mammalian culture more DNA is required than is isolated in a miniprep. Usually after a correct clone has been identified, the DNA can be re-introduced into bacteria and isolated using a maxiprep protocol. It is also possible to keep a small portion of each of the cultures that were used in the minipreps and then grow the culture containing the correct clone overnight. This saves time by not having to re-transform the bacteria again and pick a colony.


Why perform a miniprep?

To allow the diagnostic restriction digestion of recombinant DNA clones to determine if a ligation has been successful. Alternatively, a miniprep can be used to provide a small amount of DNA for any other purpose.


How does a miniprep work?

This depends on the method used. For spin columns, the process is as follows:

  1. Bacterial cell lysis in alkaline conditions followed by clearing (normally by centrifugation). In this step the bacterial chromosome is removed because in nature bacteria attach their chromosomes to the inside of the membrane. Plasmids are generally free in the cytoplasm. 

  2. Absorption or binding of the DNA to silica in presence of high salt concentrations in a column.

  3. Washing of the DNA that is bound to the silica in the column.

  4. Elution of the purified DNA.  


What are the Important Factors?

  1. The concentration of the bacteria in the culture

  2. The copy number of the plasmid (defined by the origin of replication in the plasmid)

  3. The kit you are using (some are better than others, or at least take longer than others)


DNA Miniprep Protocol Reagents

Many laboratories use miniprep spin colunm kits for screening but this protocol can be used if for what ever reason these kits are not available. It is a fast and cheap routine procedure that is based on the use of fairly standard chemicals, a waterbath, pipettes, eppendorf tubes and a microcentrifuge. However, the DNA it produces can be somewhat lower in quality and quantity than the equivalent DNA isolated using a spin column kit. For diagnostic cloning however this should not be a problem.

TES buffer (10 mM Tris-HCl pH 8.0, 5 mM EDTA, 250 mM sucrose, filter sterilized)
Lysozyme stored at 10mg/ml in water.
5M NaClO4
TE Buffer (10mM Tris-HCl pH 8.0, 0.1mM EDTA)


DNA Miniprep Protocol

  1. Start with fresh overnight liquid cultures (A single colony picked and inoculated into 3ml LB medium the evening before).

  2. Add approximately 1.5 ml of each culture into an eppendorf tube. Number the eppendorf tubes according to the numbers written on the culture tubes.

  3. Centrifuge the tubes for 1 minute at maximum speed and then discard the supernatant by throwing it out. Make sure only minor quantities of LB medium are left with the bacterial pellets.

  4. Re-suspend the pellets in 150 µl TES by pipetting the solution up and down with a 200µl pipette close to the pellet (systematically going from one side of the pellet to the other until nothing is left sticking to the wall)

  5. Quickly add 20µl of Lysozyme solution (10mg/ml in distilled water, usually kept as small stocks frozen at -20).

  6. Incubate for 5 minutes at room temperature.

  7. Pipette 300 µl of distilled water quickly to the suspensions, it should mix while the water is sprayed into the tube. Don't shake the tubes afterwards. This should be done as quickly as possible and the tubes are immediately placed in a heat block at 73°C.

  8. Incubate for 15 minutes.

  9. Centrifuge at maximum speed (13000 rpm) for 15 minutes. Sometimes, the pellet is too big (more pellet than supernatant), if so then centrifuge for another 15 minutes, be patient, it will pellet down eventually. When the pellet is big, usually the culture was too young (less than 12 hours old).

  10. Pour the supernatant into another eppendorf tube (don't forget the numbering). Add 5M NaClO4 (approximately 10% of the supernatants volume, usually 10% of 300µl. If some of your tubes have less supernatant, add some TE to those so that they look like the majority). Close the lids and mix by shaking the tubes. This can be done either in the rack using the lid to stop the tubes falling out or by shaking the tubes individually.

  11. Add 400 µl isopropanol, shake as before and centrifuge for 15 minutes at maximum speed.

  12. \"Throw\" out the liquid and centrifuge once more for 2 minutes. Remove the last bit of liquid with a 200 µl pipette.

  13. Dry for 15 minutes in a 37°C room or incubator with the lids open.

  14. Add 50 µl of TE and put on a shaker for 5 minutes then store the DNA or proceed to the screening colonies protocol.


Author: Dr Ryan Cawood


This is a teaching page and contains only basic information and advice. Oxford Genetics Ltd take no responsibility or liability for any information described within this page. All protocols and procedures should be discussed with the relevant safety staff within your facility.


Screening Colonies by PCR

Protocol for Screening Bacterial Colonies by PCR


To detect which bacteria have the correct recombinant DNA, it is possible to screen the colonies from the agar plate using a PCR method without growing them overnight. One method of doing this is simply to touch a colony with a sterile tip, then dip the tip briefly into a PCR mix in a PCR tube, then dip the same tip into some media to grow them overnight. Typically, this will only work efficiently if you have a relatively small region to amplify (<500bp) to detect your recombinant plasmid, and it relies on the 98ºC stages of the amplification to lyse the bacteria to release the DNA. To ensure that the E.coli are lysed efficiently prior to performing the PCR amplification, the protocol below can be used as a more reliable alternative.  


  1. Prepare 5 mg/ml Proteinase K stock solution

    Dissolve 25 mg PCR grade Proteinase K in 5ml of T0.1E (10 mM Tris-HCl, 0.1 mM EDTA, pH 8.0)]
    Store in aliquots at –20 ºC.

  2. For each sample prepare 25 µl lysis solution:

    Mix 1 µl of 5 mg/ml Proteinase K stock and 24 µl T0.1E in an eppendorf

  3. Pick colonies; for each colony:

    Pick the colony using a sterile pipette tip.
    Spot some cells onto a labelled LB plate (with appropriate antibiotic)
    Deposit the remaining cells into a 25 µl aliquot of the lysis solution
    Incubate the labelled plate overnight at 37 ºC to recover colonies.

  4. Lyse the cells:

    Incubate the samples at 55 ºC for 15 min.
    Incubate the samples at 80 ºC for 15 min.
    Cool the samples on ice.
    Mix and spin down briefly.

  5. Use 1 µl of lysate in a PCR reaction.


Authors: Dr Richard Parker-Manuel and Dr Ryan Cawood


Screening by Restriction Digestion

Screening of recombinants using restriction digestion


PCR screening can be used as an alternative method to screening by miniprep and restriction digestion. However, any correct colony identified by PCR will still require restriction digestions to determine if the DNA is correct. If you would like to screen by PCR then click here for the protocol.

This digest is meant as a quality control, or to test different clone recombinants, and requires only a small amount of plasmid, to be digested for a standard time (1 hour) with an amount of enzyme that is in excess. The total reaction is intended to be loaded onto an agarose gel. There is no need to do a time course as for the preparative digest.

200ng of plasmid is usually sufficient and 1 unit of enzyme in a total reaction volume of 10 µl. Remember that larger plasmids may require you to use a bit more DNA if you are cutting out small fragments. 200ng of a 3 kb plasmid is 5 times more copies than of a 15 kb plasmid and so there is 5 times less copies of any fragments you cut out.

If the plasmid is not from a relatively clean maxiprep or from a silica resin based miniprep kit then you may want to use more enzyme. If the DNA is prepared using the miniprep protocol on this website you might want to double or triple the amount of enzyme.


Protocol for a clean plasmid with 1µg/µl:

0.2-0.5 µl plasmid
1 µl 10x Restriction buffer
0.1-0.3 µl Restriction enzyme 1
0.1-0.3 µl Restriction enzyme 2 (Optional)
Make up to 10 µl with TE or water


Protocol for phenol chloroform extracted miniprep plasmid (DNA concentration much lower and usually unknown):

3 µl plasmid
1 µl 10x Restriction buffer
0.1-0.3 µl Restriction enzyme 1
0.1-0.3 µl Restriction enzyme 2 (Optional)
Make up to 10 µl with TE or water

  1. Calculate the total number of samples you have to screen and then add one (n+1).

  2. Make a master mix using the guidlines above with everything except the DNA to screened. So if you have ten samples use 11 µl 10x Restriction buffer, 1.1 µl enzyme and 95.7 µl water.

  3. Add an aliquot (minus the DNA quantity) of the mastermix to the number of tubes you have samples for. In this case 10.

  4. Add the DNA and mix with the tip to make sure it digests.

  5. Incubate for 1 hour at 37°C

  6. Add 1 µl of loading dye and load onto an agarose gel.


Note: Some enzymes react at temperatures above or below 37°C. Check before you set up the reaction.   


Author: Dr Ryan Cawood


How Does DNA Sequencing Work?

Sanger Dideoxy Sequencing Background Explanation:


DNA sequencing was originally developed in the 1970s. A variety of other methods were also developed around the same time but the chain termination method developed by Frederick Sanger rapidly became the method of choice.

DNA sequencing uses both normal nucleotides (A, T, G and C) but also chain terminating dideoxynucleotides (ddA, ddT, ddG, and ddC). When DNA polymerase incorporates a dideoxynucleotide of any base into a chain that is being extended, it causes the polymerase to stall and the chain can no longer be extended. This is pivotal for DNA sequencing reactions to work.  


In order to sequence a piece of DNA the things that are required are:

  1. A primer that is specific to the region of DNA to be sequenced.

  2. A DNA template to be sequenced.

  3. Normal deoxynucleotides (A, T, G, and C)

  4. Dideoxynucleotides (ddA, ddT, ddG and ddC)

  5. DNA Polymerase


Just like when a normal cellular DNA polymerase extends DNA, during a sequencing reaction the growing DNA strand is extended in the 5’ to 3’ direction.

DNA sequencing reactions always contain more normal nucleotides than dideoxynucleotides. In the original method developed by Frederick Sanger, the sequencing reaction would be performed in four separate tubes, where each tube contained lots of normal nucleotides and a small amount of either dideoxy G, T, A or C. The reason for this will become clear below.  

The sequencing method works in the following way. The primer binds to the DNA template in the region to be sequenced. The polymerase then starts to extend the primer in the 5’ to 3’ orientation. As the primer extends, it more frequently incorporates normal nucleotides because there are more of those in the reaction mixture. So if the primer sequence was AAAATTTTGGGGCCCC and then the next nucleotides in the DNA to be sequenced were ATCGAAA then the primer (plus the new nucleotides) would now have the sequence AAAATTTTGGGGCCCCATCTAAA. Then if we imagine that in this reaction we have small amount of dideoxy guanosine in it, and that the next base is a G, we can assume that most of the time the extending chain will incorporate a normal G nucleotide (because there are more of them in the reaction). But occasionally the chain will incorporate a dideoxynucleotide and the chain will terminate with the sequence AAAATTTTGGGGCCCCATCTAAAG. However, those extending chains that did not terminate at the first G will continue to the next one, where the same possibilities apply, the chain will either terminate or carry one.  For example, AAAATTTTGGGGCCCCATCTAAAGTAAAACCCG may be the next terminated DNA fragment. So where ever you have a G in the DNA sequence, you will have some chains that have terminated and are of a specific length. Now also imagine you have the same reaction for the other nucleotides A, C and T. Where each tube contains a population of DNA fragments that terminate at each of the nucleotides for which there is dideoxynucleotide in the reaction.

Now we run those four reactions of an agarose gel that separates the DNA by size. In the case above, we would see that the G reaction would produce DNA fragments that are 24 base pairs (bp) long and 33bp long. Whereas if we did the same reaction with a C dideoxynucleotide in the reaction, we would have had chains that terminated at 19bp, 30bp, 31bp and 32 bp long. The reaction with dideoxy T would produce fragments of 18bp, 20bp and 25 bp and the A reaction would produce fragments of 17bp, 21bp, 22bp, 23bp, 26bp, 27bp, 28bp and 29bp. If these are all run on a gel in adjacent lanes, it then becomes possible to see how we can read the DNA sequence. See below:




If you read the letters up from the bottom, you will see that it tells you the DNA sequence of the reactions described above. However, this method was quite time-consuming and required four different reactions to get a single sequence. For this reason, sequencing reactions are now performed with dideoxynucleotides where each different nucleotide is labelled with a different fluorphore. So each terminated chain has a colour that is associated with the last base in its chain. Then if the reaction is run on a gel you can see how the sequence can be read in the same way as before. It is now common to run the sample through a capillary tube and a computer detects the colour of each DNA fragment as it runs out of the end of the capillary. Using this technique it is common to obtain DNA sequence reads in excess of 1000 base pairs.  




In this diagram the dideoxynucleotides are labelled as: ddA = Green, ddT = Red, ddC = Blue, and ddG = Black.


Author: Dr Ryan Cawood


Clean Up of DNA

The reaction conditions of many recombinant DNA procedures are incompatible, for example, between a restriction digest and a ligation. This requires that you remove the enzyme and buffer from the previous step so that you can start the next step. This can be done through various methods including using extraction and purification kits, phenol-chloroform extraction, or using miniprep kits with a modified method. Probably the most important consideration for most labs is cost. A phenol-chloroform extraction is the cheapest and using a miniprep spin column being the most expensive. However, in terms of safety this is the exact opposite!


Using specialist kits to clean up DNA

Many specialist kits exist for cleaning up reactions. Many people use PCR clean up kits for the cleaning up of enzymatic reactions but do check the size exclusion limits on the column and the binding capacities. Some PCR kits don’t bind DNA <200bp or >4-5kb.


Phenol-Chloroform DNA Clean Up Protocol

Below is method for using Phenol and Chrolform to extract DNA from an enzymatic reaction but please be aware of following safety issues.

Potential Hazard: Phenol is very dangerous and causes skin burns immediately on contact. It is toxic on contact, inhalation and 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.


Reagents Required

TE (10mM Tris-HCl pH 8.0, 0.1mM EDTA)
Equilibrated Phenol
5M Sodium perchlorate (5M NaClO4)


Phenol Chloroform DNA Clean Up Protocol

  1. 50µl of “dirty” DNA is your starting material in a 1.5ml eppendorf tube.

  2. Add 50 µl of TE

  3. Add 50 µl of equilibrated phenol.

  4. Vortex (the sample should become “milky”). The idea is to get an emulsion between the phenol and the water phase. Be careful to ensure the lid is properly shut.

  5. Proteins will migrate to the phenol phase, whereas the DNA will stay in the water phase.

  6. Add 100 µl of chloroform and vortex. Be careful to ensure the lid is properly shut. Now that the DNA and proteins are partitioned, the idea is to separate the two phases.

  7. Centrifuge the tube for 5 minutes at maximum speed (12-13000 rpm) in a microcentrifuge.

  8. Recover the upper (aqueous) phase and transfer to a new 1.5 ml tube containing 100 µl of chloroform.

  9. Vortex and centrifuge again for 2 minutes at maximum speed in a microcentrifuge.

  10. Recover the upper (aqueous) phase in a new 1.5 ml tube and then add 10 µl of 5M NaClO4 and mix.

  11. These last two steps to recover the aqueous upper-phase are a bit tricky, perhaps the most difficult of all recombinant DNA techniques, and it is worthwhile training it on fake samples (just TE) before you start. Shaky hands are not helpful.

  12. Add 110 µl of Isopropanol, vortex and centrifuge 15 minutes at maximum speed in a microcentrifuge.

  13. Carefully discard the supernatant and centrifuge again for 1 minute at maximum speed.

  14. Insert either a fine pipette tip or fine glass pasteur pipette to the opposite side of the pellet (which should be barely visible) and suck out the liquid. Keep sucking even when the liquid has gone, to remove all liquid from the walls of the tube. You should also try to collect all the droplets that you see on the walls of the tube by moving the pipette and using the capillary forces. It is important to have only a dry pellet left.

  15. Leave the tube open for 1 minute at room temperature and then resuspend the pellet it in an appropriate volume of TE (approx 30-50µl).

  16. Keep the tube on ice and vortex now and then, it does take a few minutes before all of the plasmid in the pellet is dissolved again.  

  17. Now you can move to the next manipulation, for example a ligation.


Tip: Disposing of phenol waste can be difficult in many labs. Ideally you would place all Phenol waste and contaminated tips in a sealed shatterproof container, or the container it arrived in and the place this container in plastic container with wooden fragments in the base to catch any spillage in the unlikely event it occured. Pouring it down the sink is not an option.  Phenol/chloroform mixtures can be treated as halogenated waste solvent and disposed of accordingly. Solid and liquid phenol must be disposed of as hazardous waste substances. Small quantities of solid waste (e.g. gels, contaminated paper towels etc) could be placed into a suitable, leak-tight container and then into a yellow bag and treated as clinical waste for incineration. Basically, contact the person in charge of safety and disposals in your building and ask them what you need to do so that you comply with local regulations.



Author: Dr Ryan Cawood



Making Agarose Gels

Agarose gels and Electrophoresis

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.

1)    Add either autoclave tape or electrical tape to the ends of your gel setting tank to hold the gel in whilst setting. Don’t use masking tape as this leaks when wet. Then add your combs so it is ready for pouring. Remember to tape some of the wells of the comb together to make a bigger well if you want to gel extract a DNA fragment (which might mean loading up to 100 µl). Some tanks have rubber stoppers that fit on the end, if this is available it may be less messy than using tape.

2)    Weigh the correct amount of agarose into a conical flask or 500ml bottle with a functioning lid and rim (see Tip 1). Most tanks have a recommended gel volume; usually between 40-120 ml. therefore use between 0.4-1.2 grams for the 1% gel.

3)    Add the correct amount of buffer to the bottle or flask and mix by swirling.

4)    If using a bottle, undo the lid slightly to allow steam to escape. Do not heat with the lid closed. 

5)    Heat the agarose solution in a microwave until it begins to bubble. Then remove and swirl to mix the solution. It will be very hot so use protective gloves.

6)    Microwave until the agarose particles are dissolved. If you swirl the mixture and look at in the light you will see any un-dissolved agarose particles moving around.

7)    If using a bottle you can immediately tighten the lid up when you take it from the microwave. Then if you run the bottle under cold water the gel liquid will begin to boil and bubble as it cools. This happens because you cool the air in the bottle which makes a vacuum and allows more efficient heat exchange between the liquid and the gas. Be careful not to cool it too much, you should just be able to hold it comfortably.

8)    If using a conical flask then leave it to cool on the bench side until it is just comfortable to hold. Swirl occasionally to ensure the bottom doesn’t set first.

9)    Add 1µl of Ethidium Bromide (Potential Hazard: ETBR is a potent carcinogen, wear cloves and dispose of tips and liquid properly) stock per ml of gel. Stock is at a concentration of 500 µg/ml ETBR in water. Final concentration of 0.5 µg/ml i.e. use one 1µl of ETBR stock per ml of gel. For a safer alternative, see note 1.

10)    Swirl gently to mix but avoid getting bubbles.

11)    Pour the gel from one corner slowly; don’t make it too thick as you might get a lot of auto fluorescence when you image it.

12)    You should wait a little while before you use it (approx 40 minutes - 1 hour). It will become opaque when set.

13)    The gel starts to set after 15 minutes, and if you are in a hurry, you may then place the gel in a fridge and wait only a further 15 minutes.

14)    To run a sample, the gel is placed in an electrophoresis tank containing buffer so that the gel is just immersed (a few millimetres). The DNA will run from negative to positive, so ensure you place it in the correct orientiation.

15)    Load your sample by placing your tip just inside the top of a well and slowly ejecting the sample. The glycerol (if homemade) or Ficoll (if pre-bought) in the loading dye will make the sample sink to the bottom.
16)    Load a DNA ladder in an adjacent lane. Which one depends on the size ranges of your samples.

17)    Run the gel at 10V/cm of gel.

18)    Fragments are observed using UV light and a permanent photographic record is made if required.

19)    Loading too much DNA can cause problems to separate the bands so try to avoid this is if possible.

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.


Page written and produced by Dr Ryan Cawood