Extracting Genomic DNA from E. coli
(1) Grow 15 mL overnight cultures of the desired strains with 0.2 % glucose and any appropriate antibiotic.
(2) Harvest 10 mLs of the culture in a 15 mL tube in a swinging bucket centrifuge (~3-5,000 x g).
(3) Aspirate the growth medium with a 200 uL tip on a vacuum hose (connected to a liquid trap).
(4) Resuspend the cells in 1 mL (1/10th harvest volume) of Wash Buffer.
(5) Transfer cells to a 1.7 mL microfuge tube and re-harvest (~5,000 x g).
(6) Aspirate wash buffer with a 200 uL tip connected to a vacuum line.
(7) Thoroughly resuspend the cells in 100 uL of Lysis Buffer (now 100X).
(8) Add 467 uL of TE Buffer.
(9) Incubate at room temp. for 10 minutes.
(10) Add 30 uL of 10 % SDS - mix gently to avoid foaming.
(11) Add 3 uL of Proteinase K (from stock in freezer at ~20 mg/mL) and mix gently.
(12) Incubate at 37 C for 30-60 min (can just set in rack in incubator). Periodically mix tubes by inversion.
(13) Add 185 uL of 3 M NaCl (final NaCl = 700 mM) and mix.
(14) Add 90 uL of 10 % CTAB (this detergent precipitates when stored, so re-melt before starting).
(15) Add 700 uL of chloroform (this runs out of the pipette fast, so have the bottle near tube).
(16) Mix by vortexing to form an emulsion, but not to much (will break DNA more and more).
When vortexing tubes with chloroform, alcohols, or phenol, hold the tube with a Kimwipe at the tube/cap interface. These solvents will escape the tube and you want to catch that liquid. Discard the Kimwipe afterward.
(17) Centrifuge 2-3 minutes at 10,000 x g (CHCl3 will weaken tubes, so don’t spin full speed).
(18) Transfer the aqueous phase to a clean microfuge tube (pre-labeled!) using a 200 uL tip (multiple draws).
- Pull the sample slowly from the top meniscus with each withdrawal. Resist the temptation to push sample back out, it will squirt and stir up the material. If that happens, re-spin for a minute or so. Approach the interface carefully. Tip the tube as the aqueous volume decreases to encourage a drop to form to one side. Slowly draw sample from the top of this drop. Leave some behind (~20-30 uL). These transfers can be difficult because some of the DNA gets trapped in the interface where the junk is and when you draw it into the pipette tip you will pull some junk with it. You can sacrifice yield (leave some DNA behind), you don’t need all of it.
(19) Add 700 uL of phenol/chloroform mixture, vortex briefly to form an emulsion.
(20) Centrifuge 3 mins at 10,000 x g.
(21) Transfer aqueous phase to a clean tube (pre-labeled).
(22) Add 700 uL chloroform, vortex briefly and re-spin.
(23) Transfer aqueous phase to clean tube (pre-labeled).
(24) Add 700 uL (greater than equal volume) of 100 % isopropanol.
(25) Vortex briefly to mix (look for loss of Schlieren).
(26) Incubate on ice for at least 10 minutes (can go longer or days in the -20 freezer).
(27) Pellet DNA at full speed in centrifuge of 10 minutes. Make note of tube orientations, hinges out.
(28) Aspirate the supernatant with a vacuum & clean tip.
- When removing solvents from precipitations and washes, orient the tubes such that the outer wall during the spin is on the bottom as you tip the tube. Because you spun with the tube hinges on the outside, the caps will be on the bottom as you aspirate the liquid. This helps keep the cap out of the way, because you will focus on accessing the top of the tube. Move the vacuum pipette gently along the top wall (where you material is not), just contacting the solvent enough to draw it in. As you approach the pellet (or, if it’s invisible, where the pellet should be), pause the pipette tip at the top edge of the meniscus and let the solvent get dragged into the tip without pushing the tip against the pellet. This will avoid sucking the pellet onto the tip. If that happens, don’t panic. Just gently remove the tip from the vacuum line and wiggle the pellet off into the remaining solvent.
You may have to re-seat the tip on a pipetter t get it to release. Re-spin to get the precipitated material collected in the bottom again. If needed, you can add more wash solvent to help collect the material.
(29) Wash pellet with 700 uL of 75 % ethanol. Vortex well to release pellet.
(30) Spin to collect the DNA (3 mins, full speed).
(31) Wash the pellet with 700 uL of 95-100 % ethanol.
(32) Re-spin, remove solvent. Try to get most out, just pause over the pellet with the tip and let the ethanol mostly evaporate into the vacuum.
(33) Set the tubes on their sides on a Kim-wipe. Avoid stirring up dust and paper fragments into the tube. You can cover the tubes with a Kim-wipe to prevent stuff from settling in the tubes. Dry for at least an hour - the DNA should be bright white and have no moisture left in it. You can leave overnight - dry DNA is stable.
(34) Resuspend DNA in 50 uL of DNA buffer.
(35) Quantify DNA using a 100-fold dilution in a spectrophotometer.
(36) CLEARLY write concentration on tube, along with date, initials, and details of the genotype.
(37) This can be frozen at -20 C (in a non-frost-free freezer) or at -80 C.
Recipes for Required Reagents
Wear gloves, eye protection, and lab coats when making and working with these.
50 mM bis-Tris (this is not Tris, it’s under “B”)
100 mM NaCl
pH 6.5 (use HCl, forming bis-Tris-Cl)
- Pre-rinse glassware and stir bar (~100-150 mL beaker and the largest stir bar that fits). Stir it up in ~40 mLs, adjust pH, pour into 50 mL Falcon tube (conical bottom). ASK FOR ASSISTANCE WHEN CALIBRATING AND USING THE pH METER. Add clean water from a diH20 squirt bottle to 50 mLs (use the molded line at 50 mL, not the paint). Label tube and cap, initial, and date.
Lysis Buffer - Make this each time it's needed, start with this and melting CTAB solution before harvesting cells
1 mL of BPER-2 (a commercial non-ionic detergent that doesn’t unfold proteins)
5 mM EDTA (add from 250 mM stock)
0.1 mg/mL lysozyme
1 ug/mL RNase A
- Lysozyme is stored lyophilized in the -20. It doesn’t survive freeze-thaw transitions well, so it is prepared each time for high activity. Warm the lysozyme bottle to room temp before opening (can hold in fist to warm quickly). This step prevents water from condensing in the bottle on the reagent. There should be a few beads of desiccant in the bottle, they should not be dark blue. If they are, add a few more pink ones from the desiccant bottle before you put it back (I might have that backward). Lysozyme will not refold well when placed in BPER directly, so a soluble folded stock is made first. You can weigh out the lysozyme, but that’s a hassle and the accuracy isn’t necessary. Put a 200 uL tip on a pipetter, dig it into the lysozyme horizontally, and transfer a tip’s worth of lysozyme to a tube with 400 uL of Wash Buffer in it. The lysozyme will re-hydrate and fold into it’s active form. This solution will be ~10 mg/mL (exact amount changes as the amount on the tip varies, but it’s close). Gently mix the lysozyme into solution, spin it briefly at full speed in the centrifuge to move insoluble material to the bottom, and draw from the top liquid to make your 0.1 mg/mL final, in this case, 10 uL. Just add the lysozyme and the RNAse A into the 1 mL of BPER, the final amounts won’t be exact, but it doesn’t matter.
10 mM Tris base
1 mM EDTA (from 250 mM stock)
adjust pH to 8.0 using diluted HCl (this forms Tris-Cl)
Stir up in ~40 mL, adjust pH, make 50 mL in a Falcon tube with water.
10 % SDS
Measure 4 g of SDS and dissolve in 40 mL water. The extra room in the tube is to help vortexing it into solution. CAUTION: SDS powder can form a fine dust when being weighed out. Inhaling it is very unpleasant.
Stored as a stock in glycerol in the -20. Remove the tube from the freezer (don’t stand there with the door open), bring it to your bench, and pipette the amount you need. It’s OK if it gets a little warm. Place back in freezer.
3 M NaCl
A good chemical to have on hand for lots of applications in you rack. Make 50 mLs. Just weigh and add water and mix.
10 % CTAB (cetyl trimethylammonium bromide)
Dissolved in water - 1 gram in 10 mL is plenty. May take heating to go into solution. When stored, it will precipitate, so place the tube in a warm sand bath when you begin for the day. Vortex it back into solution when warm. It will stay soluble at your bench while you work with it. CTAB is used in some protocols to selectively precipitate DNA. In high salt, DNA stays soluble (this protocol). Here, the CTAB is used to form a co- precipitate with oligosaccharides, which are invisible in the spectrophotometer, but would contaminate the DNA and interfere with downstream manipulations.
This chemical is highly volatile, very runny, and dangerous to inhale. Working with small amounts at the bench is OK, but larger volumes should be handled in the chemical hood. In the hood, pour ~50-100 mLs into a clean glass bottle for use at the bench. When working with it, wear safety glasses, ALWAYS LEAVE A HAND GRIPPING THE BOTTLE: when removing the cap, accessing the bottle, and re-capping. Don’t let go during the whole operation, so have your tubes with the caps open in a rack next to the bottle and transfer aliquots out. This mechanism prevents the bottle from tipping over when being used. Don’t get distracted, focus on your operation. Deliver above the tubes, not in the tubes (an exception to the general pipetting method). The exact amount delivered will vary. When done pipetting, leave your hand on the bottle, eject the tip, set the pipetter down, and re-cap. Then move the bottle to a safe place before proceeding. You don’t want this spilled. The chloroform is used here to phase separate the CTAB/sugars in one step, and also to remove leftover phenol in another. It is a useful chemical for partitioning small hydrophobic molecules and for storing viruses that lack membranes because nothing can grow in solutions saturated with it (you will see it used in phage protocols). Remember to hold the tube with a Kimwipe when vortexing.
A commercial mixture of the two chemicals. Phenol denatures proteins and separates as a phase from aqueous solutions. Most proteins get trapped at the interface and in the phenol layer. You have digested your proteins with an enzyme, so some peptides will be the aqueous phase (if they don’t have sufficient hydrophobic character). We keep two kinds in the refrigerator: one around pH 8.0 for DNA work, and one acidic mix for RNA work. You want the higher pH one. This is a dangerous chemical. Wear a lab coat and wear eye protection. Operate with it as you do for chloroform, except you will draw from the stock bottle, which will be cold and slippery. The phenol solutions are stored under an aqueous buffer solution, so the phenol is saturated with water and the pH is controlled. You don’t want the buffer on top, your tip has to carefully and slowly go through the buffer layer into the phenol layer. Don’t go so deep that the tip submerges in the buffer layer, draw from about a quarter to a half centimeter below the buffer. When you draw up the phenol, some will run down the outside of the tip, let that fall back into the bottle before moving away from the bottle. Don’t let go of the bottle. Focus. Move the tip over your tube and deliver the phenol. Some will remain in the tip and on the outside, just leave that, don’t try to wiggle it out or rub it on the side of the tube, deliver above the tubes, not in the tubes. CHANGE TIPS between each delivery - this prevents contamination of the stock bottle with your sample. Don’t worry about closing the tubes after each addition: get them all done, recap the bottle, move it to the safe place in the refrigerator (in a secondary container). THEN come back to the bench and re-cap the tubes. Don’t pick up the tubes to re-cap: hold the rack stably and close each one. The goal with this and the chloroform and other nasty solutions is to minimize the reach-for-and-grab steps, which are the causes of spills. Remember to hold the tube with a Kimwipe when vortexing.
100 % Isopropanol
Carefully pour ~40 mLs into a 50 mL Falcon tube. With high salt, DNA (and proteins and RNA and .. nearly everything big) will precipitate when the isopropanol concentration is at or over 50 %. More doesn’t hurt, so we usually add excess. Remember to hold the tube with a Kimwipe when vortexing.
75 % Ethanol
Carefully pour some ethanol into a 50 mL tube, take note of the volume, and add water to make the final 75 %. Our ethanol is ~95%, just assume it’s 100 % for ease of calculation. This is used to wash the remaining salt from the DNA (the water allows for salt solvation, but DNA remains insoluble in 75 %). Remember to hold the tube with a Kimwipe when vortexing.
95 % Ethanol
Pour an aliquot and label the tube. This is another handy reagent to always have on hand. It is used here to wash away the remaining 75 % ethanol and salt. It also allows for faster drying. Some labs use acetone for the last wash, which speeds up drying, but adds another hazard. Remember to hold the tube with a Kimwipe when vortexing.
- The Kimwipe holding is more important for the chloroform, but it prevents spraying solvent away form the tube and onto your hands. If you ever work with radioactive samples, you will be double-gloved as well. You then assume the Kimwipe is contaminated after vortexing and you ditch your outer gloves to be sure.