Tuesday, 12 November 2013

How to subclone

What is subcloning and what is it used for?
A common method to study the function of a gene is either to knock it out or to overexpress it (knock in). This can be done in mice or in cultured cells. Exercise physiologists use this method for example to test whether a potential regulator of adaptive responses does indeed trigger the hypothesized adaptations when its concentration is increased.

In many cases the DNA constructs can be obtained from colleagues. Alternatively many DNA constructs can be purchased from Addgene (www.addgene.org). However, not all vectors are suitable for all cells and sometimes you wish to add your own gene into a vector in order to study it. For this DNA need to be cut out a longer piece of DNA or cDNA and inserted into a suitable vector. This process is termed subcloning.

In this blog entry Abdalla Diaai (Figure 1), who is a PhD student in our group, explains how to subclone and answers some key questions about subcloning. Abdalla has recently subcloned several Hippo gene inserts into a vector that is suitable for the use in satellite cells. Abdalla and Dr. Roby Urcia, a post doc in our group, are the cloning experts in our team.

Figure 1. Abdalla in the DNA lab of the Musculoskeletal Group in Aberdeen.

Over to Abdalla: OK, In figure 2 I have drawn a schematical overview which illustrates how a DNA insert (shown in red) is taken out of a donor plasmid and inserted into a recipient plasmid. Plasmids are pieces of circular DNA which can be inserted into cells as vectors for DNA inserts that may encode a gene knock in (overexpression) or knock out construct.
Figure 2.In the first step the DNA of interest (red) in the donor plasmid needs to be cut out. The molecular scissors are termed restriction enzymes and they cut DNA at specific, short DNA sequences. In this examples no suitable restriction sites are found near the gene of interest so we are introducing EcoRI and NotI (these sites are cut by enzymes termed EcoRI and NotI) sites upstream and downstream of the insert so that we can cut it out. These restriction sites are added by PCR using unique primers which contrain the EcoRI and NotI restriction sites respectively. After that the DNA of interest is cut out by adding the EcoRI and NotI restriction enzymes. We also use the same enzymes to open up the recipient plasmid which already has these sites and then ligate our gene of interest into the recipient plasmid to yield the subcloned plasmid. Easy! 

As with many molecular biology method quite a bit of know how is required to successfully sub clone. Here I will answer a few key questions.

Question: How do I cut DNA?
Answer: For this you can use restriction enzymes which cut DNA at specific DNA sites. In Figure 2 EcoRI and NotI are restriction enzymes which cut DNA at EcoRI and NotI restriction sites. Restriction enzymes are the scissors of molecular biologists and restrictions sites are the specific, short DNA motifs, which are cut by these scissors.

Question: How do you find restriction enzymes in a DNA sequence?
Answer: Several websites are available to identify the short sites in a DNA sequence where specific restriction enzymes cut. I am using Serial Cloner which is a free download (http://serial-cloner.en.softonic.com/) and after trying it a bit you should be able to identify restriction sites.

Question: The Serial Cloner programme comes up with many restriction sites. Which ones should I use?
Answer: When selecting restriction sites to cut out your gene of interest the key criteria are:
-       Restriction sites must lie outside the insert (gene of interest);
-       The restriction sites need to be in the desired location in your recipient plasmid (plasmids often have multiple cloning sites (abbreviated as ‘MCS’) with several restriction sites). However, they should not be elsewhere in your recipient plasmid or you will cut it where you do not want to cut it.
-       Ideally choose restriction enzymes that work in the same restriction enzyme buffer as this makes things easier.
-       Try to identify two different restriction sites and enzymes for your subcloning because otherwise the recipient plasmid might self-ligate (i.e. form a ring of DNA without the gene of interest or insert in it!). You can get around that problem by using a phosphatase like Shrimp Alkaline Phosphatase which catalyzes the dephosphorylation of 5´ and 3´ ends of DNA. Such dephosphorylation prevents religation of linearized plasmid DNA.

If you cannot find enzymes that meet these criteria then there are alternatives such as:
-      You can modify the MCS of your recipient plasmid and add new restriction sites by using ‘annealed-oligos’ which enable you to add short stretches of DNA into your recipient plasmid. More information can be found on (http://www.addgene.org/plasmid_protocols/annealed_oligo_cloning/).
-      Use the annelead oligos as primers and PCR to add restriction sites to the MCS of your recipient plasmid.

How to run the PCR to amplify your construct and how to purify the PCR product?
In order to have sufficient insert DNA (gene of interest) you will need to run a PCR to amplify your insert DNA. For this it is important to use a high fidelity taq polymerase (Platinum® Pfx DNA Polymerase-Invitrogen) to minimize the number of errors. The longer the expected PCR product is, the more important the fidelity of the polymerase becomes. You should select an annealing temperature based on the melting temperature (Tm) of the portion of the primer that binds to the sequence to be amplified (the hypridization sequence), not the Tm of the entire primer. After the PCR reaction is complete, isolate your PCR product from the rest of the PCR reaction using a kit, such as the QIAquick PCR Purification Kit (Qiagen). The PCR product is now ready for restriction digestion. If you amplify your PCR product from a plasmid, it’s advisable to digest your PCR product in DpnI as this digests methylated DNA. It is important to run the product on a gel. This allows you to visualize that your PCR product is the anticipated size and that your band is strong (indicating that the PCR reaction worked and that you have a sufficient amount of DNA).

How to digest (cut) your DNA?
Set up restriction digests for your PCR product and recipient plasmid. Because you lose some DNA during the gel purification step, it is important to digest plenty of starting material. It’s recommended to use your entire PCR reaction and 4 μg of recipient plasmid. You can add up to 10 units of your restriction enzymes per 1 μg of DNA. Be careful as some enzymes have star activity where restriction enzyme cuts DNA at random sites other than restriction site.  Most restriction enzymes digest efficiently at 37°C for 2 h. After you have digested your gene of interest and in a second reaction your recipient plasmid using the same restriction enzymes you can then go on to ligate your insert into the recipient plasmid. For this you need to isolate your DNA first.

How to isolate your vector by gel purification?
Run your digest DNA on an agarose gel and conduct a gel purification to isolate your DNA. When running a gel for purification purposes it is important to have nice crisp bands and to have space to cut out the bands. Because of this it is recommended that you use a wide gel comb, to run the gel slowly and to leave lanes empty in-between samples. In addition to a DNA ladder standard, it is also important to run an uncut sample of each plasmid to help with troubleshooting if your digests do not look as you would expect.

Once you have cut out and purified your digested plasmid bands away from the gel, it is important to determine the concentration of recovered DNA. Do not use a Nanodrop as sometimes it gives false results. Instead quantify your DNA by running it on a gel against standard ladder of known mass (Figure.3).

Figure 3. In this gel 0.5, 1, 2 and 4 µl of a 1 kb DNA size ladder were loaded with 8 µl of distilled water and 3 µl loading dye plus the DNA of the experimenter. The intesnity of the DNA of interest (rightmost two bands) was compared to the DNAs at different concentrations. Each band on the ladder has a different mass as longer DNAs are heavier. In this case the DNA of interest has been estimated to be around 36 ng/µl. 

Tips for better gel extraction results
To obtain the DNA from a gel the following tips are useful:
1)     Trim the gel slice as much as possible.
2)     Minimize exposure on the UV light as it can damage the DNA. 
3)     Always perform the additional wash to remove residual gel debris and then perform a second ethanol (PE) wash to make sure all the salt is removed (if using gel extraction kit).

How to ligate your insert into your recipient vector?
Conduct a DNA Ligation to fuse your insert to your recipient plasmid. It’s recommended to use around 100ng of total DNA in a standard ligation reaction. You ideally want a recipient plasmid to insert concentration ratio of approximately 1:3. It is also important to set up negative controls in parallel. For instance, a ligation of the recipient plasmid DNA without any insert will tell you how much background you have of uncut or self-ligating recipient plasmid backbone.

How to transform?
Proceed with the transformation according to the manufacturer’s instructions for your so-called competent cells. For most standard cloning, you can transform 1-2 μl of your ligation reaction into competent cells. Additionally, if your final product is going to be very large (>10 kb) you might want to use electro-competent cells instead of the more common chemically-competent cells. The number of bacterial colonies resulting from your transformation will give you the first indication as to whether your ligation worked. Your recipient plasmid plus insert plate should have significantly more colonies than the recipient plasmid alone plate. Always run positive control plasmid to make sure that your transformation was successful. Also this will help you to troubleshoot if the gel results look different than expected.

How to isolate the finished plasmid?
Finally, you will need to pick individual bacterial colonies and check them for successful ligations. Pick 3 -10 colonies depending on the number of background colonies on your control plate (the more background, the more colonies you will need to pick) and grow overnight cultures for DNA purification. After purifying the DNA, conduct a diagnostic restriction digest of 100-300 ng of your purified DNA with the same restriction enzymes you used for the cloning. Run your digest on an agarose gel. You should see two bands, one the size of your vector and one the size of your new insert. You can also run a PCR reaction using the same primers to confirm the insert orientation (Figure 3).
Figure.3. Confirmation of the ligation by PCR (left) and diagnostic double digestion (right).

How to verify your plasmid by DNA sequencing?
PCR based cloning carries a much higher risk for mutation than restriction enzyme based cloning. DNA replication by PCR has error rates that range from roughly 1 per 500 bp to roughly 1 per 10 million bp depending on the polymerase used. Because of this, no matter which taq polymerase you use, it is important that you sequence the final product. When designing sequencing primers avoid GC rich parts of your new plasmid. GC content of primers should be around 40-50%. Sequencing machines can only pick up to 500 bp. So, it’s advisable to design more than one primer to cover the whole sequence.

Useful websites

Finally, there is a lot of information in this blog entry and I may have forgotten to explain everything in detail. If you have any questions then please e-mail me under r01aadm@abdn.ac.uk.