(for cloning and subcloning of DNA fragments)
1. Digest your DNA that you wish to clone and the vector (e.g. a plasmid) you are cloning into with the appropriate restriction enzymes. Use a large quantity of vector - I usually use 5-7ml of DNA from a Promega Mini Wizard kit preparation in a 40-50ml reaction. Other kits (such as Quiagen plasmid mini) may have MUCH higher DNA yields so you should probably quantify your DNA the first time you use a particular preparation kit or technique before proceeding.
DO NOT use dirty DNA for your vector(as in the speedy mini-preps protocol) if you are not gel extracting your vector. Your ligation will NOT work if you use dirty vector DNA and it is not cleaned up by gel extraction.
2. Run your restriction digest on an agarose gel of appropriate percentage.
You should gel extract your vector ONLY if:
Your insert fragment is from a PCR reaction AND/OR
The restriction enzymes you used cannot be heat inactivated
There are no restriction sites for you to use to cut re-ligated vector after the ligation
Otherwise you should run only a small amount to check for complete digestion–like 5ml in wells from a small comb.
If you are gel extracting your vector you should run and extract ALL of the digest (tape the wells together if necessary)
If you are NOT gel extracting the vector, heat inactivate the restriction enzymes. Usually this is 65C for 20 minutes but be sure to CHECK the enzyme manufacturer’s catalog for specific inactivation instructions.
3. Gel extract the insert and vector fragment (if necessary).
4. Quantify the insert and vector for their DNA concentrations. Some people claim they can use a spectrophotometer for this but in our lab we have obtained the best results by running the fragments on the gel and doing gel densitometry analysis.
To do gel densitometry:
a. Run 5ml of your DNA preparation and a DNA standard –preferably one in which the amounts of each fragment in the ladder are known (e.g. NEB 1KB ladder) on an agarose gel with a small comb.
b. After running the agarose gel –Stain with ethidium bromide.
c. Visualize the gel under UV using the Bio-Rad gel doc apparatus attached to the Macintosh computer (Quantity One software)
d. Take a picture using relatively low exposure-make the bands light
e. SAVE the image as a file on the computer
f. Open up the densitometry menu and select density in box
g. Draw a box around your band and a band in the standard that you know the DNA quantity of –preferably one of similar molecular weight) and record the number listed as average quantity (this is not a quantity however it is a measure of band intensity) –make sure you check that areas of the gel with no DNA in them have a lower intensity reading If not you may need to invert the image.
h. Calculate your DNA quantity by making a proportion of the known DNA quantity to the band intensity. Make a similar proportion using your sample of unknown concentration-cross multiply and divide.
i. This will tell you the number of nanograms (if that is the measurement of your known DNA band) of your DNA band. You need to turn this into a concentration by dividing with the volume that you loaded on the gel.
STEP 3 Calculate DNA ratios and volumes
5. Now that you know the concentration of your DNA you can continue with your ligation calculations.
Most of my ligations are done at a MOLECULAR ratio of 3:1 insert to vector respectively. However if you have trouble with ligations you can modify this ratio to 9:1 or 1:9. But in most cases 3:1 works okay.
6. Notice that I said molecular ratio. You must now calculate what I have called the “factor” in determining how much of insert and vector to use in your ligation. Simply divide your DNA concentration by the length of the fragment in kilobases (you can also use base pairs instead but you’ll typically end up with very tiny decimal factors – it doesn’t really matter either way) Some protocols tell you to use the actual molecular weight of DNA and you can get a true number of molecules of DNA in your preparation but in this case it is unnecessary because you are really only interested in the relative number of molecules. The ratio is what is important.
7. Calculate how much vector DNA you want in the reaction. Usually I like to have around 100 ng of vector in a particular reaction. If the vector is not gel extracted 40-80 ng usually works fine and it’s best not to use over 100ng. If the vector is gel extracted more vector is almost always better –the closer you can get to 100ng the better.
8. Make another ratio of the volumes of the insert and vector multiplied by their respective factors. Set this equal to whatever ratio you have chosen and solve. The answer you get is the volume of insert you should use. If you don’t have enough insert you may need to adjust your quantities. You may need to change your total reaction volume based on the volumes of the insert and vector.
STEP 4 Set up the ligation reaction
9. Set up the ligation reaction using the volumes you calculated. Add 10X ligase buffer to a 1X final concentration (If your ligase buffer contains ATP you should divide the buffer into single use aliquots).
I typically add 0.3 ml of Ligase to a 20-25 ml reaction – as with all enzymes in molecular biology add it last and keep on ice. If you are using non-gel extracted vector be SURE to include a positive control ligation (vector only – no insert) and a negative control reaction –same reaction with vector DNA – no ligase. These two will tell you if your ligase is functioning properly and how much uncut vector background you have. You should also set up these reactions for gel extracted vectors as well but I often don’t use the actual gel extracted vector but just another cut plasmid instead.
10. Once your reaction is assembled you should place it at 13-16C –simply turn on the small thermo-cool incubator near the 42C waterbath the incubator should be properly set but check to make sure.
There is one exception to this – if your reaction includes a blunt end it should be left at room temperature. This is because the ligase enzyme has higher activity at higher temperatures and there is no need for the colder temps to allow the overhangs of stick ends to anneal.
11. Leave your reaction overnight. The next day, take your reaction out and heat inactivate the ligase –this is not really necessary if your vector was gel extracted. If your vector was gel extracted you should cut the reaction with a restriction enzyme that will cut re-ligated vector but not molecules that have your insert. I often leave the digest for several hours
12. After heat inactivation of the ligase and/or restriction digestion cool the reaction on ice and get out your competent cells - Proceed to transformation.