10. Combining Biobricks
The outline procedure is covered in the Registry Help Pages under 'Standard Assembly'. The following is a more detailed procedure relating specifically to my lab.
Digesting the DNA
Prepare digests of the two biobricks with the appropriate enzymes as listed above.
- Water: 10 μl
- Plasmid DNA: 6 μl
- Buffer: 2 μl
- Enzyme 1: 1 μl
- Enzyme 2: 1 μl
- total volume: 20 μl
Incubate at 37˚C for 2 hours.
- If Biobrick A is being inserted upstream of Biobrick B, then Biobrick A should be digested with EcoRI and SpeI using buffer E. Biobrick B should be digested with EcoRI and XbaI in buffer H.
- If Biobrick B is being inserted downstream of Biobrick A, then Biobrick A should be digested with SpeI and PstI using buffer B, and Biobrick B should be digested with XbaI and PstI using buffer H.
Purifying the fragments from a gel
Add 5 microlitres of loading buffer to each digest, and run it in two adjacent lanes on an agarose gel. Stain with SYBR-Safe: NOT ethidium bromide. Visualise under blue light. Do not expose the gel to ultraviolet light. Using a scalpel, excise the vector band from one digest, and the insert band from the other, depending which is being inserted into which. Take the minimum amount of agarose with the DNA.
Transfer the two pieces of agarose into pre-tared 1.5 ml microcentrifuge tubes and note the weight of the agarose in mg in each tube.
The remainder of this procedure is a variant of the DNA purification procedure given in section 4, and uses the same materials (including a waterbath pre-heated to 55˚C).
1. To each tube, add three times that many microlitres of 6 M sodium iodide (one mg of gel equals one microlitre, so for 100 mg gel, add 300 microlitres sodium iodide, for example).
2. Place the tubes in a waterbath at 55˚C for 10 minutes. Check that the agarose has completely melted.
3. Add 5 microlitres of glass bead suspension. Mix and incubate on ice for 10 minutes.
4. Incubate on ice for 15 minutes (or longer; the exact time is not important). If all goes well, the DNA will stick to the glass beads.
5. Spin the tube briefly in a microcentrifuge to get the beads to the bottom of the tube. The speed is not critical, and 30 seonds or so should be plenty, since glass beads settle very well.
6. Remove the supernatant to a waste beaker. Ultimately this can be discarded to the drains.
7. Add 250 microlitres of ice-cold wash buffer. (This is stored in the freezer and should be kept on ice when out of the freezer). Mix by inversion. Do not attempt to resuspend the glass beads. Spin the tube briefly and remove the supernatant to your waste beaker. Remove as much of the supernatant as possible. The purpose of this step is to wash away sodium iodide.
8. Repeat step seven twice more.
9. If there is still some liquid stuck to the sides of the tube, spin again briefly and remove it. Make sure that you have removed as much of the liquid as you possibly can.
10. Add 10 microlitres of EB (elution buffer). Resuspend the glass beads by pipetting up and down. Make sure that they are well resuspended.
11. Incubate in a 55˚C waterbath for 10 minutes or so, mixing again by flicking the tube at the 5 minute mark (this is probably not essential, but the beads do tend to settle very quickly).
12. Spin at high speed for 1 minute.
13. Transfer the supernatant to a clean labelled tube. Try to avoid getting any glass beads in it. Discard the tube with the glass beads.
14. The DNA should be stored at -20˚C. You can check that the purification procedure has worked properly by running 5 microlitres of the DNA on a gel, as before. I don’t usually bother to do this unless the subsequent procedure fails and I have to figure out what went wrong.
Mix in a 1.5 ml microcentrifuge tube:
- vector DNA: 4 microlitres
- insert DNA: 4 microlitres
- 10 x ligase buffer: 1 microlitre
- T4 DNA ligase: 1 microlitre
- total volume 10 microlitres
Mix, spin briefly and incubate in the 16˚C waterbath (in the working cold room) overnight. Then use 5 microlitres to transform E. coli. If you are lucky, most of the colonies will have the desired combination. You can screen by PCR or plasmid DNA miniprep as previously described.
If you don’t get any colonies, or if you screen a dozen or so colonies and none has the right combination, this is the time for troubleshooting. I suggest running 5 microlitres of the ligation, insert DNA and vector DNA on a gel, to check that the DNA purification worked and that the ligase was active. If all these look OK, the competent cells are probably the problem. Try a control transformation with 1 microlitre of supercoiled plasmid DNA. Plating 100 microlitres should give several hundred colonies if the cells are highly competent.
Tagging procedure for increased efficiency
This is a plan to obtain increased cloning efficiency when preparing constructs with multiple biobrick parts. It involves the use of a pseudo-biobrick 'tag' encoding lacZ' with its own promoter, with an extra SpeI site at the 5' end after the prefix. Thus the tag has SpeI sites at both ends. Imagine that you want to add sequentially biobricks A to D to generate ABCD. The procedure would be:
1. Insert the tag downstream of biobrick D. Note that D will be digested SpeI-PstI, the tag XbaI-PstI. Select the blue colonies. (This assumes that biobrick D itself does not contain lacZ').
2. Insert D-tag downstream of C. Again, D-tag is not digested with SpeI in this procedure. Again, select blue colonies.
3. Insert CD-tag downstream of B. Again, select blue colonies.
4. Insert BCD-tag downstream of A. Again, select blue colonies.
5. You now have ABCD-tag. Digest this with SpeI, purify and self-ligate (self-ligation is much more efficient that ligating two separate pieces of DNA). The tag will be cut out. Transform and select white colonies. The result will be biobrick ABCD, with an extra scar just before the terminal SpeI site, equivalent to a nominal extra very short biobrick (just a few bases long).
Whether or not this procedure is worth the extra effort depends on how many problems you are having getting the right product at each biobrick addition step. If you are using the triple ligation procedure, it may be that you are getting the right clone with such high frequency that this would be a waste of time.