Make plates: 21g NZY+ 15g Agar + 1L H2O, make 2 Litters, Autoclave
for 30 min.
Plate about 75 ml into each 150mm bacterial plate. Leave on bench for 2 days
and warm up at 37 C before using.
Top agrose: 10.5g NZY + 3.5g agrose + 500 ml H2O, Autoclave for 30 min, Store in two 500 ml bottles.
Host cells: Strike XL-1 Blue cells on a LB plate (NO Amp), incubate
for overnight.
Pick a single colony and incubate in 50 ml LB medium containing 10 mM MgCl2
and 0.1% Maltose at 37 C for overnight. Then, Sipn at 4000 rpm for 15 min, and
resuspend the bacterial pellet in 25 ml 10 mM MgCl2. You may store at 4 C for
up to a week.
Phage titration: Warm up your plates in 37C. Melt the top agrose in
the microwave and cool down to 50 C.
Most of the phage we have are about 109/ml. Take 1 ul of origional stock
and put it into 100 ul of phage dilution buffer (10 mM MgCl2, 10 mM Tris- HCl,
PH 7.4). Aliquot 5 ul of the diluted phage into 1 ml of host cells (in a 15
ml tube) and mix. In order to count the phage titer, you can take 10 ul the
phage/host cell mixture and put them into 1 ml of host cells. Incubate at 37
C for 20 min. Mix 8 ml of top agrose with the phage/bactoria and pour them on
to wermed plate. After cooling down (10 min), incubate the plate in 37 C for
overnight (upside down).
Plate phages: Mix 50,000 phages with 1 ml host cells in 15 ml tube. Make 10 to 20 tubes for each screening. Incubate at 37 C for 20 min. Mix 8 ml of top agrose with the phage/bactoria and pour them on to wermed plates. After cooling down (10 min), incubate the plates in 37 C for about 8 hours (depending on the size of plaques). Cool the plates in cold room for 2 hours to 2 days.
Filter lift: Label each filter with plate number and date.
Put the filetr on the plate carefully.
Punch 6-7 holes (for orientation) through the filter and the agar on the plate
using a needle and indian ink.
Take the filter out from a connor using a forcept.
Put the filter onto denaturing Buffer (1.5 M NaCl + 0.5 M NaOH) for 2 min
Neutralize for 5 min by submerging the filter in 1.5 M NaCl + 0.5M Tris- HCl
(PH 8.0).
Rinse the tilter in 2 X SSC.
Blot briefly on Whatmann 3 -MM paper.
UV-crosslink the DNA to the filters.
Store the plates at 4 C.
PreWashing: Prewet the filter for a few minutes in 1 x SSC, 0.1% SDS.
prewash the filter at 42 C for 0.5 to 2 hours in the prewash solution (5 X SSC,
0.5 % SDS).
Prehybridization: After getting rid of dripping liquid, put the filters
into containers. Add hybridization buffer (0.5 M Na3PO4, PH7.2; 1% BSA, 7 %
SDS, 1 mM EDTA) untill it just covers the filters. Incubate at 50 to 65 C for
2 to 12 hours.
Probe: 50 ng DNA fragment + H2O = 11 ul.
Boil for 5 min. and put immediately onto icewater.
Add 2 ul random priming buffer + 1 ul 0.5 mM dNTP (-dCTP) + 5 ul 32P dCTP +
1 ul Klenow.
Incubate for 30 min at 37 C
Pass the probe onto G50 column and check if you get over 50% of incooperation.
Denature the probe (DsDNA) by boiling in water for 5 minutes and put immediately
onto icewater.
Hybridization: Add 1 to 10 x 106 Cerenkov cpm/ml of denatureed probe
to the hybridization solution. Then add filters one by one into the container
(it is important to mix each filter with solution). Heat the solution in 65
C if necessary. For 1 to 2 filters, use 10 ml hybridization buffer; for 3-10
filter, use 20 ml buffer; for 11 to 20, use 30 ml. Incubate at 50 to 65 C for
12 to 36 hours.
Wash: all SSC wash solutions contain 0.1% SDS.
[SSC] Time Temperature
2 x SSC 5-10 min 20 C
2 x SSC 20 min 50 to 65 C
0.5 x SSC 20 min 50 to 65 C
0.2 x SSC 30 min 50 to 65 C
0.1 x SSC 30 min 50 to 65 C
You can repeat any washing step if necessary, you can also stop washing at any point if your background is low enough (100 cpm by survey meter).
Exposure: Take a 3 MM paper (cut to the film size) or an old film and
wrap with wrapping film.
Brifely dry the filters and add the filters onto the wrapping film.
Cover the filters with another layer of wrapping film.
Stick the orientation markers between filters.
Add the X-ray film and expose at -70 C for 1 to 3 days.
Develop the film.
Pick plaques: To orient the filters, line up the film and mark
the plate numbers and the "dots" where the needle poked through onto
the film based on the orientation markers.
Determine the "putative" clone with the strongest signal (round with
a little tail, fussy, not very sharp).
Line up the film with the plate based on the "dots" on light box.
Use the large end of a pasteur pipet to cut through the plate where the putative
clone lines up with the film spot.
Put the agar piece into 1 ml pf SM buffer and 20 ul of chloroform and Votex
Elute the phages at 4C for overnight or RT for 3 hours.
Secondary: Dilute the eluted phages 1 to 1000 into 1 ml of phage dilution
buffer.
Infect 3 ul of the diluted phages with 1 ml of host cells and plate the palte
as before.
Repeat filter lifting, prehybridization, hybridization, washing, etc as before
Pick Plaques: Line up film, tilter and plate as before
Pick one or two well isolated positive plaques using the small end of a pasteur
pipet and put them into 0.5 ul of SM buffer and 20 ul of chloroform.
Excision: Grow an overnight culture of SOLR cells in LB medium at 37
C.
Make a 1/300 dilution of SOLR cells in LB and drow at 37C till CD600 = 0.5 -1.0.
Add 200 ul XL-B host cells into a 15 ml tube.
Add 200 eluted phages from secondary screening.
Add 1 ul ExAssist helper phage (> 1 X107 pfu/ml).
Mix and incubate at 37C for 15 min in the shaker.
Add 3 ml of LB medium in the the tube.
Incubate in the 37 C shaker for 2.5 hours.
Transfer 1.5 ml of cells into a microcentrifuge tube.
Spin at 12 Krpm for 2 min.
Transfer the supernatant into a new tube and heat the tuhe at 70C for 15 min
Spin again at 12 Krpm for 2 min.
Transfer the supernatant into a new tube, which can be store at 4C.
Mix 100 ul of the supernatant (single strand phage) with 200 ul of freshly grown
SOLR cells (OD = 0.5 -1.0) in 1.5 ml tube.
Incubate the tube at 37C for 15 min.
Add 50 ul of the culture on the cornor of a LBamp plate and strik.
Incubate overnight at 37 C.
Pick colonies
Plasmid miniprep.
Digestion
Sequencing