cDNA library screening

cDNA LIBRARY SCREENING

Last Update: December 2006

PREPARE SOLUTIONS

1. 10mM MgSO₄, 0.2% Maltose LB (100 mL):

Mix 1.0 g of Bacto-Tryptone, 1.0 g of NaCl, 0.5 g of Yeast Extract, and 1.0 mL of 1M MgSO₄. Adjust pH to 7.5 with NaOH (Autoclave). Add 2 mL of 10% Maltose

2. Cells for phage infection:

Y1090-ZL

3. NZY Media (1.0 L):

Mix 10 g of NZ amine (Casein hydrolysate) , 5 g of NaCl, 5 g of Bacto-yeast Extract, and 2 g of MgSO₄-7H₂O. Adjust pH to 7.5 with NaOH

4. Denaturation solution (500 mL is good for 45 membranes):

Mix 50 mL of 5M NaOH (0.5M), 150 mL of 5M NaCl (1.5M), and 300 mL of dH₂O

5. Wash solution (500 mL is good for 45 membranes):

Mix 50 mL of 20X SSPE (2X) with 450 mL of dH₂O

6. Neutralization solution (500 mL is good for 45 membranes):

Mix 250 mL of 1M Tris.HCl, pH 8.0 (0.5M) , 150 mL of 5M NaCl (1.5M), and 100 mL of dH₂O

7. 10X STE buffer (30 mL):

Mix 6 mL of 1M Tris-HCl, pH 7.5, 6 mL of 5M NaCl, 6 mL of 0.5M EDTA, and 12 mL of dH₂O

8. 50X Denhardts (500 mL):

Mix 5 g of Ficoll , 5 g of Polyvinylpyrrolidone , 5 g of Bovine serum albumin in 500 mL of dH₂O

9. Prehybridization/Hybridization solution (120 mL):

Mix 60 mL of Formamide (4^oC), 30 mL of 20X SSPE, 4.8 mL of 50X Denhardts, 0.6 mL of 20% SDS, and dH₂O to 120 mL. Add 2.4 mL of treated salmon sperm DNA* (10mg/ml) to every 120 mL of solution

10. * Treated salmon sperm DNA:

Boil DNA in a screw cap eppendorf tube for 5 mins. Leave on ice for >5 mins

11. Wash solution II (2 L, use 500 mL per wash for each container):

Mix 100 mL of 20X SSC (1X), 10 mL of 20% SDS (0.1%), and dH₂O to 2 L

12. 0.7% NZY Top Agarose:

Mix 3.5 g of Agarose with 500 mL of NZY Media (Autoclave)

13. NZY agar:

Mix 15 g of Bacto-agar with 1 L of NZY Media (Autoclave)

14. 20% Maltose:

Mix 20 g of maltose with dH₂O to 100 mL. Filter through a 0.45m filter

15. SM buffer (1 L):

Mix 5.8 g of NaCl, 2 g of MgSO₄-7H₂0, 50 mL of 1M Tris-HCl (pH 7.5), and 100 mg of Gelatin type A from Porcine skin (Autoclave). Store in 50 mL aliquots. Put all reagents in the bottle to be autoclaved since the reagents don't go into solution easily

16. 5M NaCl (1 L):

Mix 292 g of NaCl and dH₂O to 1 L (Autoclave)

17. 5M NaOH (1 L):

Mix 200 g NaOH with dH₂O to 1 L

18. 20X SSPE (1 L):

Mix 175.3 g of NaCl, 27.6 g of NaH₂PO₄.H₂O, 7.4 g of EDTA and dH₂O to 1 L. pH to 7.4 with NaOH (Autoclave)

19. 20% SDS (1 L):

Mix 100 g of SDS with dH₂O to 1 L. pH to 7.2 with HCl

20. 20X SSC (1 L):

Mix 175.3 g of NaCl, 88.2 g of Sodium Citrate, and dH₂O to 1 L. pH to 7.0 with NaOH (Autoclave)

21. Wash solution I (2 L, use 500 mL per wash for each container):

Mix 200 mL of 20X SSC (2X), 20 mL of 20% SDS (0.2%), and dH₂O to 2 L

22. Wash solution III (1 L):

Mix 5 mL of 20 X SSPE (0.1X), 5 mL of 20% SDS (0.1%), and dH₂O to 1 L

PROCEDURE - Step 1 - Prepare Y1090-ZL cells

1. Pick a single colony of Y1090-ZL and grow an ON at 30^oC (~230 rpms) in 50 mLs of 10mM MgSO₄, 0.2% Maltose LB

2. Check the OD₆₀₀. Grow at 37^oC until the OD₆₀₀ = 05-0.7. If the OD₆₀₀ goes over, dilute the culture

3. Centrifuge the culture for 10 minutes at 2,000 rpms

4. Resuspend the pellet in 10mM MgSO₄:

4.1 Resuspend an initial stock solution with 20-40 mLs of 10mM MgSO₄. The OD₆₀₀ = about 0.7-1.0. The stock can be stored at 4^oC for 2-3 weeks

4.2 Working solution: aliquot a portion of the stock to a separate tube and dilute to OD₆₀₀ = 0.5-0.7 using 10mM MgSO₄

PROCEDURE - Step 2 - Determine Library titer and Library plating

1. Grow ON culture of Y1090-ZL cells in 50 mL of 10mM MgSO₄, 0.2% Maltose LB at 28-30^oC shacking

NOTE: the point of the lower temperature is not to exceed the OD₆₀₀ of ~0.5; alternatively, growing cells at 37^oC for ~3hrs from 5.0 mL of ON cells should give the same results. The advantage of the ON at 30^oC is that the cells can be used the same day they were made since they will be ready early in the morning

2. OD₆₀₀ and grow at 37^oC until the OD₆₀₀ reaches 0.5-0.7 (The ON culture should have reached this level by now). Dilute if the OD₆₀₀ is too high (>1.0)

NOTE: An OD₆₀₀ of 0.5-0.7 should contain cells at their exponential growth phase and thus most of them should be alive. Changes in the amount of cells used can be done to compensate for different OD₆₀₀ readings

3. Dilutions should be done to determine the titer and thus the amount of pfus (plaque forming units) in the original stock prior to plating the plates for the screen

A sample set of dilutions is as follows:

3.1 Dilute 10 mL of library in 990 mL of SM buffer. Plate 100 mL (stock equivalent: 1 mL)

3.2 Dilute 100 mL of library in 900 mL of SM buffer. Plate 100 mL (stock equivalent: 0.1 mL) (dilution: 1:10)

3.3 Dilute 100 mL of library in 900 mL of SM buffer. Plate 100 mL (stock equivalent: 0.01 mL) (dilution: 1:100)

3.4 Dilute 100 mL of library in 900 mL of SM buffer. Plate 100 mL (stock equivalent: 0.001 mL) (dilution: 1:1,000)

3.5 Dilute 100 mL of library in 900 mL of SM buffer. Plate 100 mL (stock equivalent: 0.0001 mL) (dilution: 1:10,000)

Assuming a library with a titer of 100,000 pfus/mL (the original titer of the cDNA library used for this screen), the 1:1000 dilution should yield 10 plaques per plate. An average of the last three dilutions should give a good estimate of the titer

4. Add 100 mL of each dilution to 500 mL of Y1090-ZL prepared cells

5. Incubate at 37^oC for 20 minutes

6. Add all mixture to 9 mL/4 mL (big/small plates) of pre-warmed (at 50^oC for >1 hr) NZY 0.7% Top Agarose. Immediately mix by inverting the tube twice and pour in pre-warmed (>1 hr at 37^oC, ideally dry 1-2 day old plates left at room temperature) NZY agar plates. Swirl plates to ensure proper spread of the solution

7. Leave at room temperature until the Top Agarose solidifies (~10 minutes)

8. Incubate at 37^oC for 5-51/2 hours until the plaques appear. Leave at 4^oC ON

Leaving an ON at 37^oC of the reactions gave positive results. 100,000 pfu/plate were used for the original screen; this is not necessary and may cause problems due to coalescing of the plaques

Use big plates for the first screen (20 plates at 100,000 pfus each) and then small ones for the subsequent screens

PROCEDURE - Step 3 - Plaque lifts

1. Mark with a pencil each membrane to distinguish them

NOTE: It is a good practice to write this mark at least in two positions in the membrane to ensure proper identification/localization. After all the process has been carried out, some membranes may be damaged and miss portions. A good way of doing this is to write arabic numbers (1, 2, 3...) at three locations in the membrane. This can also be used as a guide for the SIDING of the membrane: by placing the numbers on the opposite side of where the DNA is going to be fixed, these numbers can be used to know at all times through the experiment where the DNA is; specially helpful when transferring between solutions or x-linking

2. Mark plates using India ink or some other non-diffusing dye as a positional reference

3. Carefully place the nitrocellulose membrane on top of the plate, not letting any bubbles to form between the nitrocellulose and the plate (DO NOT lift membrane once this has been in contact with the phage)

4. Leave filters for >10 minutes on the plates

NOTE: The phage are being transferred at this point. Longer times are fine. A good idea is to start the timer after placing the first filter and estimating the amount of time it takes to put one filter. Then at the end, coordinate the removal of each filter with the amount of time it has been on the plate for uniformity in the transfer

5. Transfer membranes to a 3MM paper presoaked with Denaturing solution and leave for 5 minutes

IMPORTANT: timing is important to get as many membranes processed in the least amount of time. Once the first membrane has been transferred from the plate and into the denaturing solution, start your timer. Estimate the amount of time between each transfer of a membrane. Depending how many membranes are being processed at a time (up to 24 per plastic flat used in this experiment), make sure that the first membrane DOES NOT stay longer than 6-7 minutes in the denaturing solution (5 minutes is ideal, yellowing around the nitrocellulose is indicative of longer times); this could mean that by the time the last membrane as placed in the solution, the first one must be transferred to the next step

6. Transfer membranes to a 3MM paper presoaked with Neutralization solution and leave for 5 minutes. >10 minutes is fine, the point is to neutralize, so there is no time limit

7. Transfer membranes to a 3MM paper presoaked with Wash solution and leave for 10 minutes

8. UV X-link. Place the membranes in a 3MM paper damp with wash solution. Using a Stratalinker, press Autocrosslink and the DNA will be x-linked in about 1 minute

9. Air dry the membranes for >1 hour at room temperature; store at room temperature in a plastic bag

PROCEDURE - Step 4 - Prepare probe

Prepare probe as described in Random Primed labeling of DNA

DNA cut from an agarose gel and phenol extracted twice (DNA Fragment Isolation from LM agarose) showed incorporation rates of up to 80% (>60%)

PROCEDURE - Step 5 - Prehybridization and Hybridization reactions

1. Prehybridization: prehybridize in 120 mL of prehybridization solution per plastic container for 3 hours at 42^oC

NOTE: Prehybridization can be done ON. The fast way (that also worked fine) is to label the probe while prehybridizing, so the hybridization reaction can be left ON on the same day the prehybridization reaction was started

2. Dump prehybridization solution

3. Add 120.0 mL of pre-warmed (42^oC) hybridization solution to each container

4. Boil the labeled and purified probe (part C.) for 5 mins and then leave on ice for >5 minutes

5. Add 1 million CPMs of the labeling reaction (Part C.) per mL of the hybridization solution

6. Let hybridization go ON at 42^oC

NOTE: up to 17 hours of hybridization showed positive results without any background

PROCEDURE - Step 6 - Washes (Pre-warm the solutions prior to starting the washes)

1. Wash twice at room temperature using wash solution I for 10 mins each

2. Wash twice at 65^oC using wash solution I for 30 mins each

3. Wash twice at 65^oC using wash solution II for 30 mins eac

4. Check the membranes for radioactivity. If it is too low, stop here; otherwise, if the signal is still too high, do the last, stringent, wash

5* (optional, but recommended). Wash ONCE at 65^oC using wash solution III for 10-30 mins

6. Remove membranes and let dry for >10 minutes

PROCEDURE - Step 7 - Exposures

1. Tape the membranes to a new 3MM paper, recording the location of each one

2. Mark the orientation of the film (use some fluorescent marker)

3. Cover membranes with Saranwrap

4. In the dark, place film on top of the membranes and tape the film to prevent it from moving out of position

5. Leave at -80^oC

An exposure of 3 days is typically adequate. Don't use cassettes without intensifying screens (it takes too long to get an adequate exposure)

6. Develop film

PROCEDURE - Step 8 - Plaque isolation

1. Using the developed films (part F.), align the plates with the developed film and remove a positive plaque or region (plug). This can be done using the broad region of a Pasteur pippet or preferentially a blue (1 mL) pippet tip that has had it's end point cut (and autocalved)

2. Place the plug in 1.0 mL of SM buffer and store at 4^oC

3. Elute the phage from the plug by rocking ON at 4^oC

4. This solution is used as the source of phage for the next round of screening

PROCEDURE - Step 9 - Clone purification

H. Repeat A. through G until a single plaque can be isolated that when spread on a plate and probed will show that all the plaques are positives (usually 3 screens are enough to achieve this)

PROCEDURE - Step 10 - Excision of plasmids from phage

1. Isolate single plugs as described in G

2. Vortex vigorously for 10 secs and leave at room temperature for 5 mins

3. Use 25 mL of supernatant and mix with 100 mL of DH10B cells

DH10B cells: grow from a single colony in 1.5 mL of LB at 37^oC (7-8 hours)

4. Mix and incubate at room temperature for 5 mins

5. Spread on LB-10mM MgCl₂-Amp (100 mg/mL) plates and let dry for >10 mins

Optional: Add 40 mL of X-gal (20 mg/mL, 100% DMF) to each plate

6. Add 4.0 mL of 1M (200 mg/mL) IPTG in each plate

Both reagents can be used together or X-gal alone (for blue/white selection of inserts within the B-galactosidase gene). Leave the solution to dry (>10 minutes) before plating

7. Leave plates ON at 37^oC

8. Screen the colonies obtained to identify the desired clone

9. Good luck :)