Benton Davis Blots

Day 1

Prepare in advance:

Chill plates containing bacteriophage-lysed E. coli for at least 1 hour at 4℃ (2 hours works well).

Use 2 nitrocellulose filters for primary screening, and 1 filter after that per Petri dish (S & S BA85 or Amersham Hybond-C 0.45 µm nitrocellulose; 132 mm diameter for 150 mm Petri dishes, or 82 mm diameter for 100 mm diameter petri dishes). Handle these as little as possible and only by the edges using gloved hands and forceps. Number each filter with India ink or pencil to correspond with the number of each respective plate.

25 G needle and small drop of India ink on piece of parafilm.

Tray with base solution .

Whatman 3MM filter paper or gel blot paper to blot filters dry after removal from base and neutralizing solution.

Tray with neutralizing solution.

Whatman #1 filter paper to layer between filters.

80℃ vacuum drying oven (preheated 1 hour).

Hybridization buffer (sterile 5x SSPE; 10x filter sterilized Denhardt's solution; sterile 0.1% SDS; use sterile H2 O - do not autoclave hybridization buffer; warm to 65℃ before use).

Container, or bag with heat sealer, for hybridization (we have used one Kapak/Seal-a-Meal bag for as many as 40 filters on a given day).

2x SSPE (30-50 ml) diluted in water from a 20x SSPE stock.
Protocol:

Slowly lay nitrocellulose filters on the agar surface from the center to the side. Avoid trapping bubbles between filter and agar . Handle filters only by their edges with forceps. Mark position of filter on the plate by piercing the filter with a 25G needle dipped briefly in India ink in three asymmetrically positioned spots per filter.

For primary screenings, blot one filter on each petri dish for one minute and the second filter for five minutes on the same plate. After primary screening, blot each dish with only one filter for two minutes.

Remove filters from the petri dishes with flat-end tweezers, lifting from one edge and rolling the filter off the agar. Avoid tearing filter and lifting agar from the surface. Chilled plates help to avoid the latter problem.

Place filter in base solution for 1 minute, agar side up. The base solution (and the neutralizing solution below, may be used to saturate a Whatman 3MM filter paper, and the nitrocellulose filters laid on this, rather than floating freely in a container of each solution.

Blot filters on dry Whatman 3MM or gel blot paper to remove excess base.

Transfer filters to neutralizing solution for two minutes.

Blot filters on dry Whatman 3MM paper and stack between Whatman #1 filter paper.

Dry in 80°C vacuum oven 1-2 hours. No moisture should be on the glass oven window. It takes about 2 hours for the large filters and an hour for the small filters. After baking, nitrocellulose membranes are very fragile.

Before hybridization, wet filters briefly in 2x SSPE and stack filters on top of one another. (You can omit this step if you have only a few filters to hybridize. This pre-wetting step helps reduce bubbles which may become trapped between filters when large numbers of filters are added to a bag.)

Allow excess solution to drip from the filters and transfer to Kapak bags.

Add hybridization buffer (preheated to 65℃) to wet filters and remove air bubbles within bag. Ten ml of buffer per bag is usually sufficient for a few (about 6) filters.

Heat seal bag and allow to warm in 65℃ oven for 15 minutes.

Denature the radiolabelled DNA probe (1-2 x 106 cpm/filter) at 95℃, 5-10 minutes.

Add probe at corner of sealed bag using a 25G needle and syringe. Heat seal below the injection site. Briefly rinse corner of bag around injection site to remove any probe on outside of bag.

Hybridize the filters at 65℃ in a slowly rotating H 2 O bath overnight.

Day 2

Prepare in Advance:

5X SSPE, 65℃ (0.5 - 1L)

1-2 L 2x SSPE

1-2 L 1x SSPE

Whatman 3MM chromatography paper to blot filters

14 C-India ink (10 µCi/ml) or other means such as a light sensitive marker to align developed film with original filters

Hybridization solution (5x SSPE; 10x Denhardt's solution; 0.1% SDS)

Cassette with intensifying screen

Whatman #1 filter paper cut to size of X-ray film with corners numbered with 14 C-ink
Protocol:

Open Kapak and transfer solution to 32 P liquid waste.

Pipet 10-20 ml of 65℃ 5x SSPE into the bag, gently mix, add this solution to liquid waste.

Transfer filters to tray and cover with typically a few hundred ml of 5x SSPE. Incubate in 65℃ oven 30-45 min, gently shaking every 10 min.

Discard this 5x SSPE in 32 P liquid waste container.

Transfer filters to 2x SSPE for 20 min at room temperature, shaking.

Repeat step 5 once, discarding solution between treatments.

Transfer filters to 1x SSPE for 20 min, room temperature while shaking.

Repeat step 7 once.

Blot filters to remove excess liquid and air dry for 30 min, room temperature.

Transfer filters to paper labeled with 14 C ink or light sensitive ink. Tape a small area of each filter to hold it in place and wrap with Saran wrap. For 14 C-India ink, cut holes in Saran wrap covering the ink to allow 14 C emissions to expose film.

In the dark room, open film cassette, put 1 piece of X-ray film (Kodak X- Omat R film, XR-5) on intensifying screen, insert filters oriented towards the film, seal cassette and wrap in foil (safety lights can be on).

Expose film at -70℃.

Remove film (safety lights on OK) in dark room and develop. For hand processing, place in film holder; develop 5 min, rinse briefly in H ₂ O, and fix for 2 min. Rinse with running water 10 min and hang to dry.

positive hybridizing plaques are found, align the filters with the autoradiograph and pull the entire agar plug surrounding the hybridizing plaque using the wide end of a Pasteur pipet. Transfer plug to TMG . Vortex vigorously to dislodge top agar in the solution and disperse the bacteriophage.

Secondary and Tertiary Screening:
After vortexing, spin the sample in a microcentrifuge to pellet the agar and dilute an aliquot of the supernatant in TMG to 10-2 , 10 -3 , and 10 -4 dilutions. Spot 2µl volume of these dilute solutions on a lawn of E. coli (100µl E. coli with 3 ml top agar for small petri dishes) to determine the titer. These are approximate titers and are frequently off by 2-5-fold from that estimated from plating out an entire plate.

With a fresh dilution of the supernatant, plate out 800 pfu per small plate. After the E. coli have lysed, blot the petri dishes and hybridize to detect positive plaques (1 filter/plate). Remove agar plug using the thin end of a Pasteur pipet, and transfer to TMG as described above. Determine the titer using the 2 µl spot test. Plate out 100 - 200 pfu per small plate for the final screening. All of the plaques on this plate should be positive. Remove agar plugs and transfer to 100 µl of TMG. These are "plaque purified" bacteriophage.

RECIPES

Base Treatment Solution (1.5M NaCl, 0.5M NaOH) for Southern and Benton Davis blots.

For 2 liters:

175.35 g NaCl

40 g NaOH

QS to 2 liters with water and autoclave.

Neutralizing Solution (0.5 M Tris-HCl, 3M NaCl, pH 7.4) for Southern and Benton Davis blots.

For 2 Liters:

121.1 g Tris

350.7 g NaCl

Adjust pH to 7.4 with concentrated HCl (c. 70 ml?) and QS to 2 liters with water.

100x Denhart's Solution (2% BSA, 2% Ficol, 2% PVP in 3x SSPE)

For 100 ml:

2% (w/v) BSA (Sigma Fraction V)

2% (w/v) Ficol (400,000 mw)

2% (w/v) Polyvinylpyrrolidone (40,000 mw)

Dissolve in 3x SSPE (diluted with autoclaved water) and filter sterilize. This solution may be stored at room temperature, 4℃ or -30℃.

10% SDS Stock(w/v)

10 g SDS

QS to 100 ml with water. (Autoclaving is optional. We do it).

20X SSPE (3 M NaCl, 0.2 M NaH₂ PO₄ , 20 mM EDTA, pH 7.0)

350.7 g NaCl

55.2 g NaH₂ PO ₄(H 20) (--or-- 48 g anhydrous NaH ₂PO ₄, mw = 120.0)

14.89 g EDTA

Dissolve in boiling water, cool to room temperature, & adjust pH to 7.0 with NaOH. QS to 2 liters with water and autoclave.

14 C India Ink

Prepare at 10 µCi/ml of ink. We usually use a labelled amino acid or glucose.

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.