Experiment 1: Testing the binding of specifically designed DNA strands to glass slides
Glass slides were prepared (put link to Glass Slide Preparation Protocol) by being cleaned and functionalised (with HCl and GOPTS respectively).
Specifically designed oligonucleotides containing zinc finger binding domains were introduced to the slides.
These oligonucleotides comprise of a general adaptor strand, a specific short strand and a specific long strand.
Terminal amine groups within the oligonucleotides bind (by a nucleophilic addition reaction) to the epoxy group of GOPTS, sticking the DNA to the glass slides.
The presence of an EcoR1 cut site in the oligonucleotide allows us to have an extra level of control in our experiments. Although the zinc finger proteins will stay attached to their binding domains, cutting the oligonucleotide at this site allows cells to become ‘unstuck’ from the slides.
Chemical transformation
Puts your plasmid into cells
1. Take chemically competent cells (-80C freezer) and thaw on ice/water mix.
2. Add plasmid DNA to 50uL of competent cells: for minipreps 0.5-1 uL of DNA is enough, for ligations, use 5-10 uL. Mix thoroughly.
3. Leave on ice for 30-45 minutes. Turn on the waterbath set to 42C, so it reaches the target temperature over this time.
4. Heat shock the cells at 42C for 30 seconds. This will create pores in the competent cells through which the plasmid can enter into the cell.
5. Put back on ice for 2 minutes. This allows the cell to recover and begin repairing the pores, preventing cell death.
6. Add 950uL of growth media (e.g. SOC, LB, etc.) bringing the volume up to ~1000uL
7. Incubate with shaking at 37C for 45-60 minutes.
8. Plate 200uL on appropriate antibiotic: If using a ligation (or anything likely to have low efficiency) centrifuge the cells first at 8000rpm for 2 minutes and resuspend in 200uL of media then plate everything. If there are still some cells left after plating, the rest can be kept up to 3 weeks in a 4 degree fridge.
9. Incubate overnight at 37C.
Ethanol Precipitation Protocol
Simple method for gel extraction
Ethanol Precipitation of DNA Reagents Needed:
• 3 M sodium acetate pH 5.2 or 5 M ammonium acetate
• DNA
• 100% ethanol
Extracting gel
After running a gel and identifying the band that contains the DNA that you wish to extract, simply use a scalpel to cut around the band leaving as little excess agarose gel as possible.
Protocol
1. Measure the volume of the DNA sample.
2. Add 1/10 volume of sodium acetate, pH 5.2, (final concentration of 0.3 M) - These amounts assume that the DNA is in TE only; if DNA is in a solution containing salt, adjust salt accordingly to achieve the correct final concentration.
3. Mix well.
4. Add 2 to 2.5 volumes of cold 100% ethanol (calculated after salt addition).
5. Mix well.
6. Place on ice or at -20 degrees C for >20 minutes.
7. Spin a maximum speed in a microfuge 10-15 min.
8. Carefully decant supernatant.
9. Add 1 ml 70% ethanol. Mix. Spin briefly. Carefully decant supernatant.
10. Air dry or briefly vacuum dry pellet.
11. Resuspend pellet in the appropriate volume of TE or water.
Double restriction digestion for NEB restriction enzymes
Cuts a select piece of DNA from a plasmid
Protocol
Set up the reaction as follows:
- 1ug DNA
- 5uL 10x digest buffer (use NEB cloner to find which buffer works best with which enzyme)
- 1uL or 10 units of first enzyme
- 1uL or 10 units of second enzyme
- Up to 50uL nuclease-free water
Incubate at 37C for 1 hour. If the enzymes being used are both time save qualified, this can be reduced to 5-15 minutes, but incubating for longer is still recommended.
Add the reagents into the mix from largest volume to smallest, always finishing with adding the enzymes in last.
If multiple restriction digests are being set up, a master mix containing everything but the sample DNA can be made with the condition that the concentrations of the different sample DNA are similar or equal.
Bacterial immunofluorescence protocol
For preparing slides to be visualised under fluorescent microscopy
1. For every cell type that needs testing, grow a culture of bacterial cells in 5mL LB
(+antibiotics) overnight at 37 ˚C.
2. Next morning, take OD600 of the cultures (OD of 1 for E. coli corresponds to ~10^8 cells/mL),
and dilute into 2 fresh 5mL LB tubes (+antibiotics) to OD600 of ~0.01. To one of these tubes,
add IPTG to end concentration of 1mM. Incubate both tubes in a 37 ˚C shaker.
3. After ~3 hr of incubation, start monitoring OD of the cultures every half hour. We want to fix
these cells at an OD600 of ~0.5.
4. As soon as a culture reaches OD600 of ~0.4-0.5, spin down 1mL of the culture in an Eppendorf
tube at 8000xg (=rcf) for 1 min, and carefully discard the supernatant (be careful so as to
only remove the supernatant, without disturbing the cells in the pellet).
5. Re-suspend the pellet in 1mL 1xPBS by pipetting up and down 5 times. Spin down the cells at
8000xg (=rcf) for 1 min, and carefully discard the supernatant.
6. Repeat the PBS wash in Step-5 two more times, but this time only use 0.5mL PBS.
7. Now, re-suspend the cells in 0.5mL 1xPBS by pipetting.
8. Mix 500 uL Blocking buffer with the annealed oligo (5.13uL) for each cell type in a separate
Eppendorf tube, then add this to your cells.
9. Spin down the cells at 8000xg (=rcf) for 1 min, and carefully discard the supernatant.
10. Do 1x PBS wash (0.5mL PBS).
11. Now, fix the cells (in the tube itself) by resuspending in 1xPBS+4%(para)formaldehyde (we used glutaraldehyde but it fulfills the same purpose) (500uL). Incubate at room temperature for 20 min.
12. Do 1x PBS washes (0.5mL PBS)
13. Drop 50uL of the resuspension on a coverslip (round coverslips preferred), and incubate at
37 ˚C until it is completely dry. Once dry, save the coverslip at RT until all the cultures have
been processed similarly.
14. Add a drop of the mounting medium (ProLong Diamond Antifade Reagent, Fisher
#15372192) on a glass slide and place the coverslip on top of it (bacterial-side-down).
15. Seal the edges of the cover-slip with nail-polish, and save in the fridge (4˚C), for later
visualization.