E. coli chromosomal DNA is prepared following the method of Heath et al. ( J. Bacteriol., 174, 1992). Cells are embedded in agarose, then treated with deter gents and enzymes which remove the cell wall, proteins and other cellular material. The purified chromosomal DNA remains embedded in the agarose plug.
Preparation of cell plugs
1-Grow an overnight culture of E. coli MG1655 in L broth. The OD600 of an overnight culture of this strain should be approximately 1.7.
2-Make a 1.6% solution of InCert (FMC) agarose in water. Dissolve the agarose by boiling, then cool in a 50 °C water bath.
3-Pellet cells by spinning 15 min at 5,000 rpm in the Sorvall centrifuge (SA600 rotor).
4-Resuspend cells in an equal volume of PIV buffer.
5-Spin cells as in step 3.
6-Resuspend cells in 1/2 volume PIV buffer.
7-Warm cells to 37 °C.
8-Mix the cells with an equal volume of molten (50 °C) agarose and pipet into the wells of a 24-well microtiter plate. Add 1/2 ml of the cells/agarose mixture to each well.
9-Place plate at 4 °C to allow the agarose to harden.
1-Add 1 ml per well EC lysis solution without lysozyme and RNase.
EC lysis solution:
2-Shake gently at room temperature approximately 15 min.
3-Aspirate off the EC lysis solution (be careful not to suck up the agarose plug!).
4-Add 1 ml per well EC lysis solution with lysozyme and RNase.
5-Seal plate inside a heat-sealable bag.
6-Place bag (submerged) inside a shaking water bath. You will have to hold the bag down with lead donuts or other weights. Shake gently overnight at 37 °C .
Proteinase K digestion
1-Aspirate off the EC lysis solution.
2-Add 1 ml per well ESP buffer without proteinase K.
3-Shake gently at room temperature approximately 15 min.
4-Aspirate off ESP buffer.
5-Add 1.5 ml per well ESP buffer with proteinase K.
6-Seal plate inside a heat-sealable bag.
7-Place inside a shaking water bath as above, and shake gently overnight at 50 °C.
1-Aspirate off the ESP buffer.
2-Rinse each well with 1 ml sterile TE buffer (pH 7.5). Aspirate off.
TE (pH 7.5):
3-Add 1.5 ml per well sterile TE.
4-Shake at room temperature 30 min.
5-Aspirate off the TE.
6-Repeat steps 3-5 for a total of 6 washes.
7-After removing the last wash, add 1 ml sterile TE to each well and store plugs at 4 °C.
The DNA is digested while still embedded in agarose.
1-With a sterile spatula or pair of tweezers, carefully remove a DNA plug from its well and place it on a sheet of parafilm.
2-With a sterile razor blade, slice each plug in half, then slice each half-plug in half again, lengthwise.
3-Place two slices (which together would make up one half-plug) inside a sterile Eppendorf tube containing 900 µl sterile 10 mM Tris-HCl (pH 7.5). The pieces should fit into the bottom of the tube.
4-Place the tube in a rack and shake gently at room temperature approximately 45 min.
5-Remove the TE (be careful not to damage the gel slices) and add 900 µl of the appropriate restriction enzyme buffer. Shake gently at room temperature for approximately 1 hr.
6-Repeat step 5.
7-Remove buffer. Add 200 µl fresh restriction enzyme buffer and enzyme. Use approximately 30 to 60 units of enzyme per reaction.
8-Place tubes in 37 °C incubator overnight.
9-The next day, stop the reaction by adding 10 µl of 0.5 M EDTA to each tube. Store reactions at 4 °C.
Pulsed field gel electrophoresis is run using the BioRad CHEF MAPPER TM system. See the instruction manual for complete information on set-up and use of the system.
1-Prepare a solution of 1% SeaKem GTG agarose (FMC) in
0.5x TBE. Do not add ethidium bromide to the gel solution.
2-Pour the gel according the instructions in the BioRad manual. Use 100 ml agar ose for the small gel tray.
3-Chill gel at 4 °C before loading and running.
Molten agarose samples are loaded as follows:
1-Pre-warm Pasteur pipets by placing several inside a test tube, capping the tube, and placing it in a 70 °C water bath.
2-Re-melt (if necessary) the 1% agarose solution that was used to pour the gel and equilibrate it to 70 °C in the water bath.
3-Remove a gel slice from the restriction digest buffer and rinse with water. Place gel slice inside a clean Eppendorf tube.
4-To melt the gel slice, place tube in the 70 °C water bath, leaving it in just long enough to melt the gel (2 or 3 minutes should be enough time to melt it). Flick the tube often to mix the contents and to ensure the entire sample is melted.
5-As soon as the sample is melted, load it on the gel. Use a
pre-warmed Pasteur pipet to load the sample and work quickly, being careful not to let any air bubbles get entrapped in the wells. Load samples to just flush with the top of the gel.
6-Load an appropriate molecular weight standard.
7-Fill all empty wells and, if necessary, top off sample wells with molten 1% agar ose.
1-Fill gel chamber with 2 l 0.5x TBE. Turn on cooler and set it to 14 °C. Turn on pump, set the pump speed to 7, and recirculate the buffer until it equilibrates to 14 °C.
2-Place gel inside the gel chamber and secure it with a clamp on each corner.
3-Program run according to the instructions under two-state mode, and start run. Run parameters will vary depending on the size of the fragments you are trying to resolve. As an example, the following run parameters were used to resolve an 800 kb fragment.
4-Before you walk away from the instrument, check that:
·the run parameters are correct and electrophoresis is taking place (the bubble test)
Staining and visualizing the gel
1-Stain the gel in 0.5x TBE + 0.5 µg/ml ethidium bromide for approximately 1/2 hr.
2-Destain in 0.5x TBE (approximately 1/2 hour).
3-Visualize gel under UV light. For a preparative gel, visualize only with long-wave length UV light, and expose it to UV light for the shortest amount of time possible.
Excision of bands and electroelution
DNA bands are cut out of the gel and the DNA is then electroeluted from the gel into a dialysis bag.
1-Electroelution is carried out at 4 °C, so set up a standard electrophoresis cell and power supply in the walk-in refrigerator.
2-With a sterile razor blade, cut the band out of the gel. Place the gel slice in 0.5x TBE.
3-Prepare appropriate lengths of dialysis tubing by rinsing in distilled water, then in 0.5x TBE. Always wear gloves when handling dialysis tubing and never let it dry out. Dialysis tubing can be bought ready to use from GIBCO-BRL (catalog #5961-014 for 1/4" diameter and #15961-022 for 3/4" diameter).
4-Clamp off one end of the dialysis tubing with a dialysis clip.
Insert the gel slice, fill with 0.5x TBE, remove any air bubbles, and clamp off the other end with a second dialysis clip. Check to see that there is no leakage.
5-Place the dialysis bag inside an electrophoresis cell filled with 0.5x TBE. Orient the dialysis bag so that the length of the bag is parallel to the electrodes and the DNA will move out of the gel and onto the sides of the bag.
6-Check to see that the dialysis bag is completely submerged, place the lid on the gel box, and start the electroelution. Electroelute at 60 volts, 4 °C, for at least 18 hr.
7-Turn off the power and reverse the polarity of the current by switching the placement of the electrodes. Electrophorese in this orientation for 30 sec at 60 volts.
8-Take the dialysis bag out of the gel box, carefully remove one dialysis clip, and pipet the liquid out of the bag into a sterile Eppendorf tube. A Pasteur pipet works well for this pipetting.
9-Rinse the inside of the bag and gel slice with TE and combine this rinse with the first liquid.
Concentration of the electroeluted DNA
The dilute DNA solution is concentrated by ultrafiltration using a Centricon-30 concentrator from Amicon. For detailed instructions on using Centricons, read the instruction booklet.
1-Transfer the electroeluted DNA to a Centricon-30 (the Centricon-30 holds a maximum of 2 ml).
2-Centrifuge at 10 °C, 5600 rpm in the Sorvall SA600 rotor. Use thick-walled rubber adaptors in the rotor wells to hold the Centricons.
3-Spin until the volume has been reduced to the deadstop volume (approximately 40 µl). It takes appproximately 30 min to reduce 2 ml to 40 µl.
4-If the original volume of electroeluted DNA was larger than
2 ml, add the remainder to the Centricon and spin again. Continue doing this until all of the electroeluted DNA is reduced to a volume of approximately 40 µl.
5-Add 1.9 ml sterile TE to the Centricon and spin until the volume is again reduced to the deadstop volume. This step exchanges the buffer from TBE to TE.
6-Recover the concentrated DNA by inverting the Centricon over the collection cup and spinning at 2200 rpm for 2 min (SA600 rotor).
7-The DNA is now ready for shearing.
Quantifying the electroeluted DNA
The concentration of the electroeluted DNA will probably be too low to determine spectrophotometrically without sacrificing most of the sample. The following method uses very little sample and will give a rough approximation of the DNA concentration.
1-Prepare standards containing a range of DNA concentrations. The source of the DNA for these standards is unimportant (for example, plasmid DNA will work just as well as chromosomal), but the DNA must be fairly free of contaminants like RNA, proteins, etc. A good range of concentrations for these standards is something like the following: 0, 0.03, 0.06, 0.12 and 0.24 µg per µl
2-Mix 1.5 µl standard with 1.5 µl 2 µg/ml ethidium bromide solution. Spot this mixture on a UV light box.
3-Repeat this procedure with the electroeluted DNA.
4-Visualize the spots under UV light. Take a picture and compare the fluorescence of the electroeluted DNA with that of the standards to approximate the concentra tion of the electroeluted DNA.