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Choice of bacterial strain

The bacterial E. coli strains we use when plating out cDNA and genomic libraries are normally LE392 and NM538, but some times we also use XL1-blue cells.

• Streak out bacteria on an LB plate. Pick a single colony and transfer to a flask with 50 mL LB. NB! remember to add maltose to the LB, 0.2% final concentration. Grow over night.
• Spin down cells in a 50 mL Falcon tube and resuspend in 20-25 mL 10mM MgSO₄. The bacteria can be stored at 4° C for up to one month, but if you are using the bacteria to check the titer of the library or to plate out the lambda library (primary plates), ALWAYS use fresh bacteria.

Checking the titer of the library

• Make a dilution series of the library. Depending on the titer of your library, normally around 10⁸ - 10⁹ pfu/mL, take out 10 µL of the library stock solution and transfer to 990 µL SM solution, this will be the 10⁷ stock solution which you normally plate the library from. Make a 1/100 dilution of the 10⁷ stock, (10⁵ mix), a 1/100 dilution of the 10⁵ mix (10³ mix), and a 1/100 dilution of the 10³ mix (10¹ mix).
• Transfer bacterial cells (e.g. LE392) to three eppendorf tubes, 100 µL in each. Add 2 µL of the 10⁵ mix, 2 µL of the 10³ mix, and 2 µL of the 10¹ mix. Incubate for 15 min at 37° C to let the phage particles adhere to the E. coli cells.
• Melt the top agarose in microwave oven, transfer 3 mL to 15 mL tubes.
• Transfer the phages and cells to the top agarose, 3 mL in a 15 mL tube. (The tube with top agarose is kept in a water bath at 50° C). Invert the tube a couple of times and pour the top agar on a lambda plate. Wait until the top agarose has solidified and incubate the plates at 37° C until you see the plaques.
• Count the plaques and calculate the titer of the library.

Top agarose, 500 mL

• 5 g bacto tryptone
• 2.5 g NaCl
• 3.5 g agarose
• Add MgSO₄ to a final concentration of 10 mM.

Lambda plates, 1 L

• 10 g bacto tryptone
• 5 g NaCl
• 14 g agar

Plating out the library

It is important to use fresh bacterial cells. We normally plate out from 10-15 plates (140 mm) depending on the library. That is, from 300.000 to 500.000 pfu.

Example for plating out 12 plates, 300.000 pfu:

• Take out 3.6 mL of E. coli cells (grown with 0.2% maltose), and transfer to a 15 mL tube.
• Add 300,000 pfu from the library, x µL.
• Incubate 30 min at 37° C.
• Transfer 300 µL of phages/cells to 7 mL top agarose (tubes with topagarose is kept at 50° C). Invert the tube a couple of times and pour the top agarose over the lambda plate, 140 mm.
  
  It is important that the lambda plate is a bit "dried out" before you use them. Pouring the topagarose onto a "wet" newly made plate can give problems when you take filter lifts later.
  
  
• Plate out all the phages (the 11 remaining plates). Let the top agarose solidify and incubate the plates at 37° C.
  
  Watch the plates carefully when the bacteria grow. cDNA libraries are normally much faster to produce plaques, and you can normally see them after 4 hours. Wait until the plates are almost confluent. That takes 5-7 hours with cDNA libraries and 6-10 hours with genomic libraries.
  
  When the plates are ready you can store them at 4° C over night. If the plates are almost confluent you might expect some diffusion of phages but that is normally not any problems.

Taking lifts of the plates (transferring the phages to nylon membranes)

We normally use Amersham Hybond N filters/membranes. They are robust and can be re-screened many times. When you take lifts of the primary plates, the complete library, we recommend that you take duplicate lifts. For high stringency screening, (hybridization and washing at 65° C), this may not be necessary but for low stringency screening it is a must.

• Label the filters thoroughly. Place the filter on top of the top agarose for about 1-2 min. Then, using a needle, mark the filter with needle holes. (These holes are necessary when you later align the filters and exposed films.) We also mark the needle holes at the back side of the lambda plate with a marker. This makes it easier to align the exposed films against the plates.
• The duplicate filter is placed on the top agarose for 2-5 min, and marked similarly as the first filter.
• Let the filter dry. The lambda phage is now transferred to the filter.

Denaturation solution: 1.5 M NaCl, 0.5 M NaOH.

Neutralization solution: 1.5 M NaCl, 0.5 M Tris-Cl pH 8.0.

• Soak the filter in denaturation solution for 2-5 min. Transfer the filer to 3M paper where it partially dries up.
• Soak the filter in neutralization solution for 5 min. Transfer the filer to 3M paper where it partially dries up.
• Wash the filter two times in 2 X SSC.
• Let the filters dry.
• UV fixate for 1-2 min (the time depends on the strength of your UV transilluminator).
• The filters are now ready for use. Before use they should be stored in a dry place at RT.

Keep the library plates well wrapped in a plastic bag at 4° C to avoid drying out. The library can be kept at 4° C for months, but the titer will gradually decrease.

Probe labeling

It is normally enough to label 20-50 ng probe, and use approx. 50 µCi ³²P alpha dCTP. Any kind of random labeling kit will work, just follow the manufacturer recommendations. After the hybridization we normally clean up the probe on a nick column (Sephadex G-50 size exclusion column from Pharmacia). This normally gives a clean probe eluted in a 500 µL TE. Before you use the probe remember to boil it! (You need a single stranded probe in the hybridization.) Boil the probe for a couple of minutes and place it on ice until you add it to the hybridization solution.

Filter Hybridization

We always do a 1-2 hour pre-hybridization before we start the real hybridization with the probe.

Pre-hybridization buffer: 6 X SSC, 5X Denhards, 0.5% SDS, 10 µg/mL salmon sperm DNA.

Pre-hybridization is normally done at 60 - 65° C in a water bath with shaker.

Hybridization

We normally do the hybridizations in a glass dish or a glass beaker, using a water bath to control the temperature. The hybridization volume can be from 20-100 mL. The larger the volume the longer you need to run the hybridization. One to two days is normal if the volume is above 50 mL. Over-night hybridization is enough if the volume is less than 50 mL.

Hybridization solution: 6X SSC, 0.5% SDS, plus the ³²P labeled probe.

Procedure

• Boil the probe before you add it to the pre-warmed hybridization solution (approx. 60° C).
• Pour off the pre-hybridization solution and add the probe/hybridization solution.
• Seal the hybridization container (a sealed plastic bag, or hybridization beaker or a Hybaid glass, or whatever you might find to do the hybridization in).
• Hybridization temperature depends on your probe and what you are screening after.
  
  
  • High stringency screening: 65° C.
  • Medium stringency screening: 55-62° C.
  • Low stringency screening: 50-55° C.

After hybridization, pour the probe into a 50 mL tube and put it in a -20° C freezer. Shield the probe. The probe can be reused a couple of times.

Washing the filters

We normally wash the filters in SSC/SDS solutions after the hybridization is done. The washing solution and temperature depends on your probe and stringency conditions.

• High stringency wash: 0.1X SSC, 0.1% SDS, at 65° C.
• Medium stringency wash: 0.5 - 1.0X SSC, 0.1% SDS, at 55-62° C.
• Low stringency wash: 2X SSC, 0.1% SDS, at 50-55° C.

Normally you wash the filters 2 - 3 times in a volume of 200-300 mL. Check the activity of the filters with a Geiger counter. Too high activity normally means that the background signals are high. The activity should normally be "low", that's unless you have 10-100 of positive clones per filter.

Wrap the filters in Saran wrap, and put them in a film cassette. You get the best results if you have screens in the cassette. Leave the cassette in a -20° C freezer for 1-2 days. Develop the film, and if you have done everything right and you library is good, you should have some positive clones to play with.

After you have developed the film you normally see a faint contour of the membranes together with the dark spots of the positive clones. If you do not see the contour of the membranes you will have to make some radioactive ink (S³⁵ or P³³), or use a Stratagene marker and label the Saran wrap you have wrapped around the membranes. If you can see the contours of the membranes, put the developed film on top of the membranes and align the film so they match. (This is best done on a light table, the needle holes in the membranes is also easier to spot that way). Mark all the needle holes onto the film with a marker. Find the lambda plate which corresponds with this membrane and align these marks with the marks on the plate. If you have done a good job they should overlap 100%. You are now ready to pick a positive clone... and probably 10-100 other neighbor clones. To be sure you do not miss, use the back of a pasteur pipette to pick the clone. If it is difficult to get the plug with the positive lambda loose try to suck it loose. (Try not to act like a vacuum cleaner or you might end up swallowing it, no joking...). Place the plug in 1 mL of SM solution. Let the plug soak overnight. If it is a fresh library you have to make a dilution, 1/100 is normally OK. If it is a old library (more than one year), the titer has normally fallen so low that you can plate out the phage without diluting it. (cDNA libraries often have higher titers than genomic libraries.)

Transfer 100 µL of LE392 (or the cells you used when you plated the library) and add 2 µL of the lambda phage. Incubate for at least 15 min at 37° C, add the mixture to a 12 mL tube with 3 mL top agarose, invert the tube a couple of times and plate it out on a 82 mm. lambda plate. Incubate the plates at 37° C until you have a good plaque size. The secondary plates are now ready for lifts.

These next steps (taking lifts of the secondary plates, hybridization and all that) are similar to the ones described above. If you are able to pick clean plaque pure phage from secondary plates, you are lucky. If not, start over again, pick the positive clones but this time make sure you plate out less phage on the plate. You should try to plate out less than 100 pfu on the tertiary plates.

When you have picked a plaque pure clone from a lambda ZAP library you are ready to roll (just use the manufacturer descriptions on how to do the excision of the bluescript plasmid). If you are screening a genomic library, however (a lambda FIX vector or similar), you should be 100% sure you have a plaque pure clone. If you start up running lambda preps with a clone which is not 100% clean you may end up with some nasty surprises. To be sure that does not happen you should do a last check, a "plaque pure screen", where you plate out the clone you think is plaque pure and re-screen it. From the plate where all the clones are positive you pick your master clone! Make a plate lysate from this clone. That is, plate out a confluent plate from this clone:

• Add 2 mL of SM to the confluent plate and place the plate on a rotary shaker for a couple of hours.
• Transfer the phage solution to a eppendorf tube. Add a couple of drops of chloroform to the tube and store it at 4° C.