PREPARATIVE HIGH TITER BACTERIOPHAGE LYSATES

Production of 2.5 ml High Titer Lysate
  1. Place an agar plug containing a plaque-purified l bacteriophage (see Screening Bacteriophage Libraries with a DNA Probe or with an Antibody Probe Protocol, and the Benton Davis Blot Protocol) into a 0.5 ml microcentrifuge tube with 100-200 µl of TMG. Vortex for two minutes, let sit for 15 minutes, and re-vo rtex. Centrifuge briefly in a microfuge to pellet the agar plug and bacteria.
  2. Transfer supernatant to a sterile glass tube (13 X 100mm) containing 100 µl of a saturated E. coli (LE392, Y1090, etc.) culture (fresh overnight or stored in 10 mM MgSO4 not for more than one week).
  3. Incubate for 15 min at 37°C without shaking.
  4. Add 2.5 ml L Broth (containing 2mM MgSO4) to each tube and shake at 37°C overnight.
  5. The next morning, the solution should be fairly clear with some particulate debris due to bacterial lysis. Add a few drops of chloroform to each tube and shake for 5 more minutes at 37°C. This will lyse any remaining bacteria.
  6. Pellet the cellular debris by centrifugation at 1200xg for 10 minutes at 4°C (2500 rpm in an IEC PR-J or Sorvall RC-5b centrifuge).
  7. Decant the supernatants into s terile, snap-cap, Falcon (#2063) culture tubes (the opaque ones).
  8. Add 20µl of 1M MgSO4 to bring the concentration to 10mM.
  9. Determine the titer by spot testing serial dilutions of the solution on a lawn of E. coli (100µl saturated bacterial culture plus 3 ml top agar for 100 mm Petri dishes). Try 10-2 to 10-10 dilution in steps of 1/100 (diluted in TMG). Spot 2ul of each dilution. Incubate the plate overnight at 37°C.
  10. The next morning, determine the highest dilution that lyses the E. coli (either totally OR partially) and base calculation on this dilution. (e.g. if the 10-6 dilution only partially cleared a spot, add up the individual # of plaques present and multiply this number x106 and the by 5 00 to get the number of bacteriophage/ml in the original stock). In this case, the 10-2 and 10-4 dilutions will be totally cleared and the 10-8 and 10-10 dilutions not lysed at all. It takes about 100 PFU (plaque forming units) to totally clear a spot of this size, so a partially cleared spot will be some percentage of this.
Preparative High Titer Lysates
  1. You will need 40 µl of a bacteriophage stock that is 1 x 1010per ml (i.e. 4 x 108 bacteriophage based on the 2.5 ml titer) per liter of L broth. If the titer of your lysate is less than 109/ml, do another overnight amplification first to increase the titer before proceeding.
  2. Add the appropriate volume of the phage stock to 20 mls of saturated E. coli culture (preferably from a fresh, overnight culture, centrifuged and resuspended in 10 mM MgSO4) and incubate at 37°C for 15-20 minutes without shaking.
    NOTE: It is best to start this portion of the protocol the first thing in the morning since it usually takes 6 to 8 hours (sometimes longer) for lysis of the bacteria. It is desirable to stop the cultures soon after the bacteria have lysed in order to maximize the yield of phage DNA, as bacteriophage will still attach to bacterial receptors on lysed cells and inject their DNA into the media. If it is not possible to complete this portion of the protocol in a day's work, then wait until the end of a day to inoculate the cultures with the bacteria and the bacteriophage. Allow them to grow and lyse overnight.
  3. Transfer the bacteriophage and bacteria to one liter of L Broth (containing 2mM MgSO4; (2 ml of 1M MgSO4 per liter of broth) that has been pre-warmed to 37°C . Incubate with shaking 6-8 hours until lysis occ urs. Monitor bacterial growth and lysis in a spectrophotometer at A600 about every 45 minutes. The OD should drop from the peak, and then level off to about 1/3 of the maximum OD.
  4. Once the O.D. begins to level off, add chloroform to 2% (20 mls per liter) and NaCl to 0.5 M (29.22 grams per liter). Incubate with shaking for 5 more minutes at 37°C to lyse remaining bacteria.
  5. Chill this lysate in an ice bath and add MgSO 4 to 10mM (10.4 ml of 1M per liter of culture). The magnesium stabilizes the coat proteins on the bacteriophage. This is a convenient place to stop work for the day, storing the flasks overnight in the cold room.
  6. Since this is a HIGH titer lysate, discard anything that is disposable in a contaminated waste box, or add bleach to any solutions before discarding them. Try to avoid generating aerosols whenever possible.
  7. Centrifuge cellular debris in opaque, POLYPROPYLENE, 500 ml centrifuge bottles. Do NOT use clear polycarbonate bottles because even trace amounts of chloroform will destroy them. Avoid transferring chloroform when decanting the lysate into the bottles since CHCl3 extracts PEG which will be added in the next step. The chloroform will settle out on the bottom of the flask, so just avoid pouring out the last bit of lysate. Do not overfill the bottles or the liquid will flow out of a fixed angle rotor. Centrifuge at 4200 xg ( 5000 rpm in a Beckman JA-10 or Sorvall GS-3 rotor) for 20 minutes at 4°C. Use 0-ring inserts with the screw cap lids on the bottles to avoid leaks and cross-contamination of clones.
  8. Pour off the supernatants into a glass beaker taking care not to disturb the pellet. Estimate the volume and add PEG (approximate mw. 8000) to 10% (105 grams/liter) while stirring the solution on stir plate in the cold room. Use CLEAN stir bars that have been soaked in 95% ethanol. Let stir for a minimum of 2 hours (overnight or longer is fine).
  9. Centrifuge the PEG precipitate in CLEAR, POLYCARBONATE, 500 ml bottles at 8300 xg (7000 rpm in a Beckman JA-10 or Sorvall GS-3 rotor) at 4°C for 20 minutes. We prefer to use clear centrifuge bottles at this point since it is easier to see the precipitate on the walls of the bottle, thereby increasing the yield.
  10. Pour off the supernatant completely and drain the bottles upside down on paper towels. Wipe the inside lip of the bottle with a Kimwipe to remove the last bit of liquid. It is important to remove as much PEG as possible in preparation for CsCl banding of the bacteriophage.
  11. Resuspend pellet in a minimal volume of TM buffer using a sterile, disposable, 10 ml pipet. Because the PEG precipitate was centrifuged in more than one bottle, add about 6 ml of TM buffer to the first bottle. Dissolve the precipitate and then transfer this solution to the second bottle. Dissolve the contents of the second bottl e, then transfer and dissolve the precipitate in the third bottle, etc. Finally, add an additional 4 mls of buffer to the first bottle to dissolve any remaining precipitate and repeat the transfer to the other bottles. You should end up with no more than 11 ml of solution in the final bottle. Break clumps of PEG precipitate by drawing the solution up and down in the pipet.
  12. Split the solution equally into three 30 ml silanized Corex tubes. WHILE VORTEXING, add an equal volume of chloroform t o each tube (approximately 4 ml) to extract the PEG. Use a vortexer for test tubes (not the Thermolyne Maxi-Mixer) so that the phases will mix well during the chloroform addition. You must have the tube vortexing while adding the CHCl3 or the phases will not properly mix for efficient extraction. Vortex the tube 3 times for 15 seconds each.
  13. Cover the tubes with Saran wrap and tape them around the top (CHCl3 eats parafilm). Centrifuge at 12,000 xg (10,0 00 rpm in a Sorvall rotor) for 10 minutes at 4°C. Use rubber sleeves to protect the Corex tubes. Swinging bucket rotors are preferred for better interfaces, but fixed angle rotors may be used.
  14. The bacteriophage will be in the upper aqueous phase. The PEG will form a thick, white interface and the CHCl3 will be on the bottom of the tube. Pipet off the top phase carefully with a sterile, glass, Pasteur pipet. Avoid tilting the tube to keep the chlor oform under the PEG. Transfer the aqueous phase in a sterile, 50 ml, disposable Falcon centrifuge tube (blue capped). Remove any CHCl3 that may have been transferred. Determine the volume and add 0.75 grams of CsCl per ml of solution. Mix well by hand, but avoid foaming.
  15. After the CsCl is dissolved, check the refractive index of the solution. It should be close to 1.380 for proper banding of the bacteriophage in the middle of the tube. Adjust acco rdingly by adding either more buffer to dilute or more cesium to concentrate your solution. (Use the cesium chloride calculator on the computer to determine volume or grams to add).
  16. Transfer the solution to a cellulose nitrate tube [Beckman "Ultraclear" tubes also work well] (9/16" x 3 1/2"; holds about 18 ml). Top off tube with CsCl in the same buffer and density. Fill the tube almost to the top or it will collapse in the ultracentrifuge. Balance tubes to within 50 mg.< BR>
  17. Centrifuge in a SW 40Ti rotor at 10°C, 37,000 rpm for 24 to 36 hours. Leave the brake on the ultracentrifuge OFF, or the gradient may be disturbed at the end of the run. (It will take about 30 minutes to stop without the brake). Remove the rotor and clean the inside of the ultracentrifuge. Run the dry cycle to evaporate any condensate that may form in the chamber. The dry cycle takes about 30 minutes. Return later to turn off the power.
  18. Using a ring stand, clamp the centri fuge tube directly above a dissecting microscope light to illuminate the band from below. Turn OFF the room lights for best viewing. Remove any PEG layer that may be on top with a metal spatula or cotton-tipped swab. Then pipet off most of the top of the gradient (above the band) with a Pasteur pipet and discard into a waste beaker. Place a sterile 9" Pasteur pipet just on top of the band and begin drawing it into the pipet in one smooth motion (usually if the rubber bulb is totally depressed at th e start, this can be accomplished in one shot). Try to do this while making slow, horizontal, circular motions with the pipet in the band. Transfer the bacteriophage to a sterile, snap-cap, Falcon tube. There should be only a very thin band left left in the centrifuge tube at this point.
  19. Collect the remaining (<10%) bacteriophage band in a separate tube. Keep this as starting material for future high titer lysates. If the bacteriophage did not band well (very diffuse instead of a sharp, fat band), it is better to recentrifuge the bacteriophage rather than removing a lot of the gradient to recover them. You risk contaminating your DNA preparation with other proteins and bacterial chromosomal DNA if you do this, as the bacterial DNA that is present will be in the bottom third of the gradient.
Extraction of Bacteriophage DNA
  1. Pre-heat a few 100 ml of bacteriophage dialysis buffer to 37°C.
  2. Measur e the volume of bacteriophage using a sterile serological pipet. Add to this 1/20 volume of a freshly made 20mg/ml solution of Pronase in Pronase buffer , and 1/20 volume of 0.5M EDTA.
  3. Dialyze this solution in 100 volumes of the pre-heated bacteriophage dialysis buffer for 60-90 minutes at 37°C. Use freshly boiled dialysis tubing (wear gloves when handling). Leave extra space at the top of the tubing for expansion due to the high concentra tion of CsCl in the sample.
  4. After dialysis, decant the bacteriophage into a 15 ml silanized Corex tube. Twist the dialysis tubing from the top down (wear gloves) to remove all of the liquid.
  5. Add an equal volume of phenol (TE or STE saturated) to the tube. Cover the tube with Saran wrap, place thumb over the top, and mix by inversion 20-25 times. Remove Saran wrap, wipe the top of the tube clean with a Kimwipe, cover with fresh Saran wrap, a nd tape to the tube.
  6. Transfer the Corex tube to a rubber tube adapter and centrifuge in an RC-5b Sorvall (preferably in a swinging bucket rotor) for 10 minutes at 7000 rpm at room temperature. (Some baby powder on the outside of the Corex tubes may help remove the tube from a tight rubber sleeves after centrifugation).
  7. Using a sterile Pasteur pipet, remove the phenol (bottom) phase. When inserting the pipet through the upper, aqueous phase, keep the rubber bulb depressed. When th e tip is immersed in the phenol phase, blow some air out of the pipet to remove any of the aqueous phase that may be present in the tip. This also tends to remove DNA that may have stuck to the sides of the pipet. (The upper, cloudy, aqueous phase will contain the DNA, unless there is so much DNA that the phases invert. This is easy to check by adding a few drops of TE buffer to a given phase and seeing whether or not they are miscible.)
  8. Add another volume of phenol to the DNA phase and repeat steps 4 through 6.
  9. Add 2 volumes of ether to the aqueous phase and extract the residual phenol by inversion.
  10. Remove the upper ether phase and discard it. Repeat the ether extraction twice.
  11. Allow ether to evaporate by leaving tubes in a fume hood for an hour, or by chasing with some nitrogen gas.
  12. You may now dialyze the DNA in TE buffer (24 hours with 1-2 changes), or ethanol precipitate the DNA directly (my preference since some DNA is lost during d ialysis. For ethanol precipitation, centrifuge the pellet at 10K rpm, 30 minutes, 4°C in the RC-5b and resuspend the DNA in TE buffer.
  13. Transfer the DNA to a 1.5 ml microfuge tube for storage.
  14. Determine the DNA concentration on the spectrophotometer (see "Nucleic Acid Conversion Factors") for procedure. A relatively pure DNA preparation will have a 260:280 ratio of at least 1.7, (typically 1.8 or 1.9 for this procedure). If the ratio is 1.6 or less, extract again with phenol and then ether. If this is not done, the DNA may not be efficiently digested with restriction endonucleases, and you run the risk of loosing DNA due to potential nuclease digestion.

RECIPES

Dialysis Membrane Treatment: (5% sodium bicarbonate, 1 mM EDTA) for nucleic acid electroelution.
700 ml     500 ml     400 ml
< /DL>
NaHCO3          35 g        25 g         20 g
0.5 M EDTA   1.4 ml     1.0 ml      0.8 ml
Cut lengths of tubing a little longer than the width of an agarose gel. Boil membrane for 20 minutes in a beaker covered with a watch glass. Rinse extensively with autoclaved water before use.
TMG (10 mM Tris-HCl, pH 7.5; 10 mM MgSO4; 0.1% gelatin)
For 100 ml:
1 ml 1 M Tris-HCl pH 7.5
1 ml 1 M MgS04
0.1 g gelatin
QS to 100 ml with water and autoclave.
1 M Magnesium Sulfate
For 100 ml:
24.65 g of MgS04(H2O)7< /SUB> (i.e. heptahydrate; mw = 246.5)
QS to 100 ml with water and autoclave.
L Broth (LB; Luria-Bertani)
10 g tryptone
5 g yeast extract
5 g NaCl
1 L water
Autoclave
Bacteriophage Resuspension (TM) Buffer
50 mM Tris-HCl, pH 7.8 (25 mls of a 1M stock)
10 mM MgSO4 (5 mls of a 1M stock)
Q S to 500 ml with water and autoclave for 25 minutes, slow exhaust.
Pronase Buffer
25 mM Tris-HCl, pH 7.5 (1.25 mls of a 1M stock)
0.1 mM EDTA (10 µl of a 0.5M stock)
QS to 50 ml with water and autoclave for 25 minutes, slow exhaust.
0.5 M EDTA, pH 8.0
For 250 ml:
Heat water, add 46.53 g of EDTA, disodium salt (372.24 mw). Cool to room temperature (may use an ice bath) and adjust pH with NaOH. QS to 250 ml with water and autoclave.
Bacteriophage Dialysis Buffer
0.1 M Tris-HCl, pH 7.5 (50 mls of 1M stock)
0.5 M NaCl (50 ml of 5M stock)
2.5 mM EDTA (2.5 ml of a 0.5M stock)
QS to 500 ml with water and autoclave.
STE (10 mM Tris-HCl,pH 7.8; 10 mM NaCl; 1 mM EDTA)
Heat water and add 0.727 g of Tris base
1.2 m l 5 M NaCl
1.2 ml 0.5 M EDTA
Adjust pH to 7.8. QS to 600 ml with water and autoclave.
TE (10 mM Tris-HCl pH 7.8; 1 mM EDTA)
For 600 ml solution:
0.727 g Tris
1.2 ml of 0.5M EDTA
pH to 7.8 with approximately 7 drops of concentrated HCl with a pasteur pipet (ACID - wear rubber gloves), or about 4 ml of 1 M HCl. QS to 600 ml with water and autoclave.
L Broth Top Agar < DD>Make L Broth and add 7.5 g of agar per liter.
PEG Use approximate molecular weight = 8000.

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Copyright 1993, 1996, 1997, and 1998 by Mark Barton Frank, Ph.D.
Proper citation for data acquired from this document is: "Besta, R. M. and Frank, M. B. Preparative High Titer Bacteriophage Lysates. In: Frank, M. B. ed. Molecular Biology Protocols. (http://omrf.ouhsc.edu/~frank/htl.html). 1997. Oklahoma City. Revision Date: September 2, 1998."