UNIT 3: cDNA LIBRARY SCREENING

UNIT 3: cDNA LIBRARY SCREENING

INTRODUCTION:

In this course you will only practice screening a commercially prepared cDNA library. However, it is important that you understand how such a library could be prepared. In brief, the first step in preparing the library is to isolate mRNA from the cells or tissues of choice. It is important to insure that the mRNA is free of genomic DNA contamination since otherwise clones will be isolated corresponding to genomic sequences, i.e. the clones will contain introns and/or other sequences which are not found in mature mRNA. Although it is possible to construct libraries with mRNA contaminated with ribosomal or tRNA, it is desirable to reduce the contamination by these species to as low a level as possible. Otherwise many of the "cDNA" clones will be derived from rRNA or tRNA sequences. Several methods for isolating mRNA are discussed in the Northern blotting section of the manual (Chapter 10).

The next step in library construction is to convert the mRNA to cDNA using the enzyme reverse transcriptase (RT), which is purified from the capsid of retrovirus. The most commonly used reverse transcriptases are Moloney Murine Leukemia Virus RT (MuLV-RT, M-MuLV or MmLV-RT) and Avian Myeloblastosis Virus RT (AMV). Cloned versions of these RT, such as Superscript RT (Gibco-BRL) are also available. These cloned versions sometimes have properties that make them even more desirable for library construction. The RT reaction requires a primer and dNTPs. Since most mRNAs conveniently have a polyA tail, the most commonly used primer is oligo-dT. However, other primers can be useful in specific circumstances, such as random hexamers, or oligo-dT containing one or more non-T bases at the 3' end (e.g. oligo-dT CA). These later oligo constructs are known as anchor primers. Oligo-dT primers are biased toward the 3' end, whereas random primers prime at any site along the mRNA and thus increase the probability of being able to reverse transcribe the 5' end of the mRNA. A potential problem with random primers is that they readily prime off of rRNA, tRNA or genomic contaminants. Anchor primers are advantageous when one wants to transcribe only a subset of the poly-A mRNA thereby reducing the complexity of a given library.

At this point the cDNA is present as a DNA-RNA hybrid. In order to obtain larger quantities of the DNA it will be necessary to clone it. To make this possible, the RNA strand needs to be removed and replaced by DNA to produce a double stranded DNA. This is most efficiently accomplished by treating the RNA-DNA hybrid with the enzyme ribonuclease H (ribonuclease hybridase) which specify nicks the RNA in RNA-DNA hybrids. These nicks represent sites for second strand synthesis so that following RNase H treatment, if one adds DNA polymerase I (Pol I), any remaining RNA will be digested by the 5'-3' exonuclease activity of Pol I. The hydrolyzed RNA will be replaced by the 5'-3' DNA polymerase activity of Pol I resulting in a nicked double stranded DNA. These nicks can be sealed by DNA ligase to generate an intact double stranded DNA.

Finally a short linker (a sequence that needs to be digested with an appropriate restriction enzyme to generate sticky ends) or an adapter sequence which is synthesized with a sticky end are blunt end ligated onto the ends of the double stranded DNA. The reaction is carried out at a very high ratio of adapter/linker to double stranded DNA, that the reaction can be made very efficient. The resulting DNA is then ready to be ligated into the appropriately cut cDNA vector and screened.

PROTOCOL 3.1: ANTIBODY SCREENING OF LAMBDA PHAGE cDNA LIBRARY USING ECL^TM CHEMILUMINESCENT DETECTION

INTRODUCTION:

Traditional methods of library screening involve the use of radiolabeled DNA probes which detect the gene of interest by DNA:DNA hybridization. In order to synthesize the DNA probe, usually one first has to know the sequence of amino acids of the protein in order to guess at the order of nucleotides in the gene. Unfortunately, in order to determine the amino acid sequence of the protein, long and extensive protein purification procedures are involved that usually yield very small amounts of purified protein. Peptides are generated from the pure protein, fractionated to purity and sequenced. Often times, it is impractical to purify enough protein for amino acid sequencing and more often protein sequencing is beyond the scope of the laboratory's ability or expertise. An alternative method to identify genes of interest was developed which allows one to screen a library with an antibody probe which directly detects a portion of the protein encoded by the gene. Practically, it has been much easier to raise an antibody to a protein than it has been to determine its amino acid sequence, but in order to use these antibody probes, several criteria must be met. First, the cDNA must be inserted into position in the genome of the vector that is transcribed and translated into protein. Second, the antibody probe must be directed against one protein. Therefore "mono" specificity of the antisera is essential for the successful identification of a clone from the cDNA library. Even if the antisera is monospecific and a clone is identified, some type of secondary confirmation of the clone is necessary to establish its identity. The ability of the clone to react with the antisera is NOT ENOUGH! Criteria used to confirm the identity of clones will be discussed in class.

In practice, usually it is necessary to clean up the antibody preparation before it is used for screening of the library. Often a polyclonal antisera will contain additional endogenous antibodies directed against E. coli determinants. This can contribute to a high background when screening the library. Lysates of E. coli can be added directly to the diluted antibody preparation in an effort to "subtract out" background contributed by antibodies against E. coli determinants. Alternatively, the antibody of interest can be affinity purified on an antigen affinity column. The antibody preparations can be used several times in succession for multiple screenings and often times, the preparation becomes better with multiple uses.

Monoclonal antibodies can also be used to screen expression libraries. They have the unique advantage that (in theory) they recognize only one protein determinant. Unfortunately, this can also be a detriment since the antibody molecule will recognize only one site on the protein (in theory). If that antigenic determinant is coded for by sequences at the very 5' end of your mRNA and you have failed to synthesize cDNA that extended to the 5' end, it is possible that no clone will be able to express that antigenic determinant. Furthermore, some monoclonal antibodies only recognize conformational epitopes, that may not be present when a heterologous protein is expressed in bacterial cells. For these reasons it is better to use a polyclonal antisera or battery of monoclonal antibodies to screen the library since it increases the odds that at least one cDNA clone will express the necessary antigenic determinant (Ausubel et al. 6.7-6.8).

In addition to concerns about the antibody, the host cell may also have an effect on the success of antibody screening. Certain proteins may be strongly degraded during expression by particular host proteases and hence more difficult to detect using antibody screening. The choice of expression vector, and the means used to induce expression, can also effect the success of antibody screening. For example, certain expression vectors may produce fusion proteins which are relatively insoluble, particularly at high levels of expression. In this case, the fusion proteins may be so insoluble as to make transfer or attachment of protein to filters problematic. Finally, even if the protein is successfully expressed and transferred to the filter, it may be in a confirmation not recognized by the antibodies one is using. In this case, one may have to attempt to renature the protein by incubating the blotted filters in a series of gradually decreasing concentrations of a protein denaturant such as urea or guanadinium HCl in order to promote folding of the protein in a more native-like confirmation.

PROCEDURE:

Initial screen:

1. Determine concentration of phage/ml by titering phage on host bacteria that you will be using to screen. For Lambda ZAPII the bacterial strain is XL1-Blue MRF’. (see Titering of phage library in Chapter 2.) You should not rely on the titer of the library if it has not been recently determined since often the titer of the library will decrease when the library is stored for long periods of time.

2. Usually 15,000-50,000 phage are screened per 150 mm plate. Depending on the expected frequency of the clone, it is usually necessary to screen at least 200,000 to 300,000 recombinant phage in order to be reasonably certain that you will find the clone of interest.

3. Prepare host bacteria for plating:

a. The NIGHT BEFORE you want to screen your library, start a 50 ml culture of host bacteria by inoculating culture with colony from plate. DON'T FORGET TO ADD MALTOSE AND MgSO4. Grow overnight with shaking at 37^oC (Final concentrations = 0.2% maltose, and 10 mM MgSO4)

b. The next morning, spin down bacteria at 2000 x g for 10 minutes. Resuspend pellet in 0.5 volume of 10 mM MgCl₂ or SM.

4. Plate the library for screening:

a. Place 300 ml of plating cells in a sterile 16 x 150 capped tube.

b. Add phage (diluted in SM) to the cells and allow bacteria to adsorb phage for 10 minutes at 37^oC.

c. While phage are adsorbing, melt soft agarose in microwave and cool to 55^oC in water bath.

d. Mix cells and 8 ml soft agarose and IMMEDIATELY pour onto 150 mm L-plate (prewarmed to between 37^oC to 42^oC) and rock until surface of plate is evenly covered with phage/bacteria/agarose mixture. Use 4 ml of soft agarose for 100 mm plates.

e. Let plate stand at room temperature to harden.

f. Place in 42^oC incubator (inverted).

g. Soak filters in 10 mM IPTG-

-place 150 mls of sterile filtered 10 mM IPTG in Petri dish.

-wet nitrocellulose filter by laying it on the surface of the liquid.

-once wet, submerge filter.

-add next filter, and so on.

-allow to soak for at least 30 seconds to several minutes.

-blot on Whatman paper to remove excess liquid

-leave at room temperature to dry.

-label filters with pencil or ball point pen.

h. Monitor the growth of phage. At ~3 to 3.5 hours plaques will begin to form. Overlay the plates with IPTG-soaked nitrocellulose filters only after the plaques have begun to form.

i. Overlay plates by gently holding filters with forceps or gloved hands at opposite edges and centering filter over plate.

j. Let filter contact plate in the center and gradually pull itself to the surface of the plate. Place in 37^oC incubator inverted for 3.5 h. Leaving the filters on overnight may also work well if the protein is not subject to rapid degradation.

DO NOT PICK THE FILTER OFF THE PLATE AND LAY IT BACK DOWN AGAIN ON PLATE IF YOU FIND IT WAS NOT CENTERED. This will distribute phage over surface of the plate and confuse the screening procedure.

Try not to trap air bubbles between the filters and the surface of the plate. If bubbles do form, leave them, do not try to remove. Incubate the plates/filters (inverted) overnight at 37^oC unless the particular strain of bacteria you are using require a slightly different temperature.

5. After 3.5 h or in the morning, MARK PLATES WITH 18 GAUGE NEEDLE

a. Asymmetrically mark filters by puncturing with 18 gauge needle. Twirl the needle between your fingers to puncture the filter and the agar. It is ESSENTIAL that this be done because you will use these marks to align filters to plates for picking of phage plaques.

b. Carefully remove filters from plate. Mark protein side as “up” and keep protein side up during all subsequent washes and antibody incubations. At this point a second filter can be placed on the plate and incubated for an additional 3 h before removal.

c. Place filter in PBS/0.1% Tween-20.

d. Place plates in cold room or refrigerator until ready to core plaques.

e. Rinse excess bacteria off filters, by washing 2 to 4 times for 5 minutes in either cold TBS or PBS containing 0.1 % Tween-20, and 5 to 10% non-fat dry milk. Note: Tris buffered saline (TBS) can be substituted for phosphate buffered saline (PBS) in any of these steps including the mixtures with Tween-20. Usually 5% non-fat milk works well as blocking agent, but many people use 10%.

Immunoscreening Using ECL^TM

The construction of cDNA libraries in vectors designed to express protein molecules enables the isolation of individual clones based on the properties of these proteins. The majority of recombinants isolated in this way have been selected by screening libraries for clones expressing a protein that reacts with a particular antibody. By using a horseradish peroxidase (HRP) conjugated secondary antibody, positives can be selected following detection by luminol based enhanced chemiluminescence. The speed and sensitivity of the ECL Western blotting system enables rapid screening and economical use of valuable primary antibodies. When coupled with HRP conjugated secondary antibodies, a rapid and powerful system for expression screening is realized. One problem with this procedure is the extreme sensitivity. If your clone is expressing a lot of protein, the primary and/or secondary antibody needs to be cut way back. The reason is that you can get so much secondary bound that the light emitting substrate is exhausted almost immediately. This means that the filter will have given off all its light before you can get it on film. This would lead you to believe that there are no positives. Care must therefore be taken to optimize the antibody concentrations before screening. See the section on antibody optimization in the Western Protocol for some guidance.

Methods:

Libraries were plated onto appropriate host strains and allowed to grow for 3 hours. IPTG impregnated supported nitrocellulose filters (OptiTrans, S&S) were overlain to induce expression for a further 3 hours, after which the membranes were removed, washed and left in PBS overnight. IPTG impregnated filters were prepared by wetting in 10mM IPTG by laying them on the surface of the solution. Once wetted the solution was swirled quickly to completely submerge the filter. The filters were then laid out to dry on paper towels. Filters must be dry before overlaying onto a plate and can be stored dry at room temperature for 2 months. Unused IPTG solution can be stored at -20^oC. Rapid ECL Western protocol was used for immunodetection which involved the following steps:

1. Dilute the primary antibody (typically 5000 to 50,000 fold) in cold PBS/0.1% Tween-20, 5 to 10% Non-fat dry milk. This dilution has to be determined empirically for each antibody.

2. Incubate filters from 2 to 24 hr at 4^oC on a horizontal shaker platform. Multiple filters can be placed in single container as long as there is sufficient liquid so that mixing between the filters occurs,

3. Wash in PBST 3 x 10 minutes in PBS/0.1% Tween-20. The primary antibody must be diluted until it is insignificant before proceeding the secondary antibody.

4. Dilute secondary antibody, typically 1:5,000 to 1:10,000 in PBS/0.1% Tween-20, 5 to 10% Non-fat dry milk and incubate filters for 1 to 6 hours at 4^oC. (Again, the antibody dilution has to be determined empirically for each primary and secondary antibody.)

5. Wash in PBST for 2 x 10 minutes.

Signal Generation/Detection

Read through this whole section before proceeding. It is necessary to work quickly once the blots have been exposed to the detection solutions. All steps can be carried out in a dark room; it is only necessary to switch off the light after step 5. Equipment that is needed includes an X-ray film cassette, a roll of Saran Wrap (other 'cling-films' may not be suitable), a timer and blue-light sensitive autoradiography film, for example Hyperfilm-ECL (RPN 2103). If possible, wear powder-free gloves because powder spots cause blank areas on films.

1. Mix an equal volume of detection solution 1 with detection solution 2 to sufficiently cover the blot, in this case 5 ml of both solution 1 and 2.

2. Place a clean sheet of Saran Wrap on the bench and smooth it out with a clean Kimwipe.

3. Use forceps to remove the filter(s). Be careful to only clamp the edges of the filter to prevent crushing. Crushing sometimes causes high background.

4. Remove excess wash buffer by holding the filter vertically and touching one of the lower corners to a small pile of Kimwipes. Wait until all the excess moisture is wicked from the filter.

5. Place the filter in the middle of the Saran Wrap with the protein side up.

6. Using pipette draw up mixed developing solution and apply solution to corners and middle of the filter. Add enough so that the entire filter is covered and you can see a positive meniscus. Surface tension will hold the fluid on top. Stop adding the solution if it begins to flow out on the surrounding Saran Wrap.

7. Incubate for 1 minute at room temperature.

8. While the sample is incubating open a sheet protector and place it on the bench.

9. Using fresh Kimwipes, drain off excess detection buffer as described in Step 3..

10. Place filter protein side down onto the sheet protector and rapidly lower the upper sheet. Gently smooth out air pockets. Wipe off any excess fluid around the edges so the entire sheet is dry. This prevents the film from sticking to the sheet protectors.

11. Place the wrapped filter(s), protein side up, in the film cassette. Work as quickly as possible; minimize the delay between incubating the blots in substrate and exposing them to the film.

12. Switch off the lights in the dark room and place a sheet of autoradiography film (for example Hyperfilm-ECL) on top of the blots, close the cassette and expose for 1 to 3 hours.

NOTE: If there is likely to be a strong signal, some people expose one piece of film for only 1 min., then remove film and immediately replace it with a fresh piece of unexposed film, reclose the film cassette and start a timer. The first film is developed to estimate the intensity of the signal. This allows an estimate of how long the second film should be incubated for optimal exposure.

MATERIALS:

1. Expression library in Lambda ZAP, Lambda ZAP II, or Lambda UNI-ZAP XR

2. Host bacteria- XL1-Blue for Lambda ZAP

XL1-Blue MRF' for Lambda ZAP II

XL1-Blue MRF' for Lambda UNI-ZAP XR

3. 1 M MgCl₂

Compound Amount/ 100 ml

Deionized H₂O 90 ml

MgCl₂^.6H₂O 20.3 g

· Dissolve and adjust to volume with deionized H₂O

· Sterilize by autoclaving for 20 minutes at 15 lb/sq. in. on liquid cycle.

4. 20% Maltose (FW=360.3)

Maltose, an inducer of the gene (lamb) that codes for the bacteriophage l receptor, is often added to the medium during growth of bacteria that are to be used for plating bacteriophage l. Add 1 ml of a sterile maltose solution for every 100 ml of medium.

Compound Amount/ 100 ml

Maltose 20 g

Deionized H₂O 100 ml

· Dissolve and sterilize the solution by filtration through a 0.22-micron filter.

· Store the sterile solution at room temperature.

5. LB Medium (Luria-Bertani Medium)

In a clean dry 2L Erlenmeyer flask add the constituents listed below. If you turn the Bacto-tryptone and Yeast extract bottles on their side and gently tap them to loosen up the contents before you open the bottle, the contents will shift so they can be more easily scooped out of the bottle without generating a large amount of dust.

Compound Amount/1000 ml

Bacto-tryptone 10 g

Yeast extract 5 g

NaCl 10 g

· Add 900 ml H₂O. Stir until the solutes have dissolved.

· Adjust to pH 7.5 with 5 N NaOH.

· Adjust the volume of the solution to 1 liter with deionized H₂O.

· Sterilize by autoclaving for 20 minutes at 15 lb/sq. in. on liquid cycle.

6. Blocking Solutions:

There are a number of possible blocking solutions. Which one works best for each antibody has to be worked out empirically. The most common is TBS or PBS containing 0.1% Triton X-100 and 5 to 10% Carnation non-fat dry milk. 0.05% Sodium Azide is sometimes added to prevent bacterial growth.

Compound Amount/ 100 ml

TBS or PBS 90 ml

Carnation Non-fat Dry Milk 10 g

Triton X-100 0.1 ml

7. SM

This buffer is used for storage and dilution of bacteriophage l stocks.

Compound Amount/ 1000 ml

NaCl 5.8 g

MgSO₄ • 7H₂O 2.0 g

1 M Tris • Cl (pH 7.5) 50 ml

2% gelatin solution 5 ml

Deionized H₂O to volume

· Sterilize the buffer by autoclaving for 20 minutes at 15 lb/sq. in. on liquid cycle.

· After the solution has cooled, dispense 50-ml aliquots into sterile containers.

· SM may be stored indefinitely at room temperature.

8. LB Medium Plates (Luria-Bertani Medium)

Compound Amount/1000 ml

Bacto-tryptone 10 g

Yeast extract 5 g

NaCl 10 g

Bacto agar 15 g

· Add 950 ml H₂O. Stir until the solutes are evenly dispersed.

· Adjust to pH 7.5 with 5 N NaOH.

· Adjust the volume of the solution to 1 liter with deionized H₂O.

· Sterilize by autoclaving for 20 minutes at 15 lb/sq. in. on liquid cycle.

· Let the agar cool to around 55 or 56^oC. Add any necessary antibiotic and 10 ml of sterilized 1 M MgCl₂, mix and pour being careful not create excess bubbles during the mixing process. Too many bubbles can ruin the surface of a plate as it cools.

9. 500 mM IPTG (FW = 238.3)

Compound Amount/1 ml Amount/10 ml

Isopropylthio-b-D-galactosidase 119 mg 1.19 g

(IPTG)

· Deionized H₂O to volume.

· Sterilize by filtration through a 0.22 mm disposable filter.

· Dispense the solution into 1-ml aliquots and store them at -20^oC.

10. 82 mm and 132 mm diameter (S & S nitrocellulose filters BA 85)

11. TBS/Tween-20

TBS with 0.01% Tween 20 (Polyoxyethylenesorbitan monolaurate Sigma P-1379)

12. Soft Agarose (0.7%) in LB w/ 10 mM Mg⁺⁺

Compound Amount/1000 ml

Bacto-tryptone 10 g

Yeast extract 5 g

NaCl 10 g

Bacto agar 7.0 g

Deionized H₂O 950 ml

· Stir until the solutes are evenly dispersed.

· Adjust to pH 7.5 with 5 N NaOH.

· Adjust the volume of the solution to 1 liter with deionized H₂O.

· Sterilize by autoclaving for 20 minutes at 15 lb/sq. in. on liquid cycle.

13. Tris-Buffered Saline, pH 7.4 (TBS)

Compound Amount/ 1000 ml

Deionized H₂O 800 ml

NaCl 8 g

KCl 0.2 g

Tris base 3 g

· Adjust pH to 7.4 with HCl

· Add deionized H₂O to 1 L

· Dispense the solution into aliquots and sterilize them by autoclaving for 20 minutes at 15 lb/sq. in. on liquid cycle.

· Store at room temperature.

14. 1X Phosphate Buffered Saline, pH 7.4 (PBS)

Compound Amount/ 1000 ml

Deionized H₂O 800 ml

NaCl 8 g

KCl 0.2 g

Na₂HPO₄ 1.44 g

KH₂PO₄ 0.24 g

· Adjust pH to 7.4 with HCl.

· Add deionized H₂O to 1 L.

· Dispense the solution into aliquots and sterilize them by autoclaving for 20 minutes at 15 lb/sq. in. on liquid cycle.

· Store at room temperature.

15. India ink

PROTOCOL 3.2: SCREENING OF EXPRESSION LIBRARY WITH ANTIBODY PROBES AND BCIP

INTRODUCTION:

This is an alternative protocol to the ECL antibody screening procedure. The BCIP (5-bromo-4-chloro-3-indolyl phosphate) procedure is based on an alkaline phosphatase second antibody rather than horseradish peroxidase which is used in the ECL procedure. BCIP is a substrate for alkaline phosphatase which reacts with NBT (Nitro Blue Tetrazolium) to form a dark colored precipitate on the filters. This procedure is relatively inexpensive and works well for many primary antibodies. However it is not as sensitive as the ECL procedure and often has high background problems due to E. coli alkaline phosphatase. Others have reported that the backgrounds can be reduced by adding an inhibitor of bacterial alkaline phosphatase levamisole (Sigma) to the filter during development. Levamisole does not inhibit calf intestinal phosphatase which is usually used to make the secondary antibody/conjugate.

PROCEDURE:

Initial screen:

1. Determine concentration of phage/ml by titering phage on host bacteria that you will be using to screen. For Lambda UNI-ZAP XR, the bacterial strain is XL1-Blue MRF' and for lambda gt11 it is Y1090. (See Titering of phage library.) You should not rely on the titer of the library if it has not been recently determined since often the titer of the library will decrease when the library is stored for long periods of time.

2. Once the concentration of phage/ml has been accurately determined, calculate what types of dilutions are necessary in order to plate 10,00 - 15,000 phage per 100 mm plate or 30,000 - 50,000 on 150 mm plate. Make all dilutions in SM buffer.

3. Prepare host bacteria for plating:

a. The NIGHT BEFORE you want to screen your library, start a 50 ml culture of host bacteria, e.g. XL1-Blue MRF' or Y1090 by inoculating the media with a colony from a plate. DON'T FORGET TO ADD MALTOSE AND MgSO4. Grow overnight with shaking at 37^oC (Final concentrations = 0.2% maltose, and 10 mM MgSO4)

b. The next morning, spin down bacteria at 2000 x g for 10 minutes. Resuspend pellet in 0.5 volume of SM.

4. Plate the library for screening - Usually 15,000-50,000 phage are screened per 150 mm plate. Depending on the expected frequency of the clone, it is usually necessary to screen between 200,000 to 300,000 recombinant phage in order to be reasonably certain that you will see the clone of interest.

a. Place 200 ml of plating cells in a sterile 16 x 150 capped tube if using 100 mm plate and 600 ml if using 150 mm plates.

b. Add 10,000 - 15,000 phage for 100 mm plate or 30,000 - 50,000 for 150 mm plate (diluted in SM) to the cells and allow bacteria to adsorb phage for 10 minutes at 37^oC.

c. While phage are adsorbing, melt soft agarose in microwave and cool to 55^oC in water bath.

d. Mix cells and 2.5 ml of soft agarose for 100 mm plate or 7.5 ml soft agarose (pH 7.5) for 150 mm plate and IMMEDIATELY pour onto L-plate (prewarmed to 37^oC) and rock until surface of plate is evenly covered with phage/bacteria/agarose mixture.

e. Let plate stand at room temperature to harden.

f. Place in 42^oC incubator (inverted) for 3.5 hours.

g. Soak filters in 10 mM IPTG-

-label filters with pencil or ball point pen

-place 150 mls of sterile filtered 10 mM IPTG in Petri dish

-wet nitrocellulose filter by laying it on the surface of the liquid

-once wet, submerge filter

-add next filter, and so on

-allow to soak for at least 5 minutes

-blot on Whatman paper to remove excess liquid. Allow to dry at room temperature

h. Monitor the growth of phage. At ~3 to 3.5 hours plaques will begin to form. Overlay the plates with IPTG-soaked nitrocellulose filters

i. Overlay plates by gently holding filters with forceps or gloved hands at opposite edges and centering filter over plate.

j. Let filter contact plate and gradually pull itself to the surface of the plate.

DO NOT PICK THE FILTER OFF THE PLATE AND LAY IT BACK DOWN AGAIN ON PLATE IF YOU FIND IT WAS NOT CENTERED. This will distribute phage over surface of the plate and confuse the screening procedure.

-Try not to trap air bubbles between the filters and the surface of the
plate. If bubbles do form, leave them, do not try to remove.

-Incubate the plates/filters (inverted) for 3.5 h longer. Some investigators let the plate go overnight at 37^oC. This may, however, increase the chance of degradation.

5. After either the 3.5 h incubation or the overnight incubation, Mark Plates With India Ink

a. Asymmetrically mark filters by stabbing with needle in three or more locations around the edge of the filter. If you twirl the needle back and forth between your fingers as you puncture the membrane a cleaner hole results. Make sure the needle goes down into the agar enough to leave a definite hole that can be used later to align the filter. It is ESSENTIAL that this be done because you will use these marks to align filters to plates for picking of phage plaques.

b. Carefully remove filters from plate and place filter in TBS/Tween 10% non-fat dry milk.

c. Place plates in cold room.

d. Rinse excess bacteria off of filters 2 times for 5 minutes in TBS/Tween, 10% non-fat dry milk.

e. Block filters by washing at room temperature for 1 hour in Blocking solution. The blocking solution is generally 5 to 10% non-fat dry milk. Other possible blocking agents include 1% BSA, 1% gelatin, 20% calf serum, etc. in TBS.

Use 7.5 ml for 82 mm filter and 15 ml for 132 mm filter.

f. Incubate filters for at least 2 hours to overnight with PRE ADSORBED antigen-specific antibody probe diluted in Blocking solution. High titer, high affinity antibodies produce better signals than low titer, low affinity antibodies. In general antibodies that produce good signals on Western blots will produce good signals in the screening procedure at similar dilutions. Most primary antibodies can be used in the 1:1000 to 1:10,000 dilution range. Some antibodies will give high backgrounds at this dilution and will have to be diluted further.

g. Rinse filters once with TBS/Tween and immediately pour off the solution to remove most of the primary antibody solution. Wash filter 2 times with 15-20 ml TBS/Tween for 10 minutes per wash.

h. Incubate filters for at least 1 hour at room temperature with secondary antibody (i.e. Goat anti-rabbit or Goat anti-mouse Ig coupled with alkaline phosphatase or HRP) in TBS. A 1:5000 to 1:7500 dilution is recommended. Allow 7.5 ml per 82 mm filter and 15 ml per 132 mm filter.

i. Wash 2 to 3 times in TBS/Tween for 5 minutes per wash.

j. Wash once in TBS for several minutes.

k. Blot the filters damp dry on filter paper and transfer to the color development or chemiluminescent substrate solution.

l. For colorimetric assay, place filters in developing solution until positive plaques become visible then continue with the steps below. For chemiluminescent detection place filters in the appropriate substrate solution for alkaline phosphatase or horseradish peroxidase depending on the secondary antibody complex being used. Place between sheet protectors, affix luminescent stickers, and expose to film. Develop film and place over original blot aligning the luminescent stickers and the film. Mark the outline of the filters, the ID for the filter, and the location of the needle holes.

m. Stop development by placing filters in water for 5 minutes if doing colorimetric detection.

n. Immediately circle spots with soft lead pencil if doing colorimetric detection. Sometimes the spots will fade as the filter dries out. So immediate marking will be helpful in locating the spot when you want to pick the plaques.

o. Place filter underneath a transparency.

p. Mark orientation marks with pen and then mark location of spot.

q. Use this transparency with light box and original plate to locate area from which you want to remove phage.

MATERIALS:

The majority of reagents are the same as those listed for section 1.3.

1. Blocking Solutions:

Compound Amount/ 100 ml

TBS or PBS 90 ml

Carnation Non-fat Dry Milk 10 g

Triton X-100 0.1 ml

Another blocking solution that is used is given below:

Compound Amount/ 100 ml

TBS 90 ml

1% gelatin (Sigma G-6269) 1 g

1% BSA (Sigma RIA grade Fraction V A-7888) 1 g

2. Nitro blue tetrazolium (NBT, 0.05 g/ml, FW=817.6) Sigma N-6876 Grade III

Compound Amount/ 10 ml

Nitro blue tetrazolium chloride 0.5 g

70% dimethylformamide 10 ml

· Dissolve and store at 4^oC.

3. BCIP (5-bromo-4-chloro-3-indolyl phosphate disodium salt) Sigma B-8503

Compound Amount/ 10 ml

100% Dimethylformamide 10 ml

BCIP 0.5 g

· Dissolve. The solution may be stored for several weeks at 4^oC.

4. Developing solution:

Final Concentration Stock Solution Amount/ 200 ml

Deionized H₂O 174 mls

100 mM 1 M Tris, pH 9.5 20 mls

100 mM 5M NaCl 4 mls

5 mM 1M MgCl₂ 1 ml

· The add, 0.8 mls NBT (nitroblue tetrazolium) (75mg/ml) dissolved in 70% N,N dimethylformamide

· mls BCIP (5-Bromo-4-Chloro-3-indoyl phosphate: p-toluidine salt) dissolved in 100% N,N dimethylformamide

5. Anti mouse or anti-rabbit Alkaline Phosphatase or Horseradish Peroxidase secondary antibody

6. India ink

7. Photocopier transparencies or plastic folders

8. Chemilumnescent substrates

9. Film

10. Sheet protectors

11. Glow in the dark stickers

PROTOCOL 3.3: SCREENING PLAQUES WITH NON-RADIOACTIVE ECL DNA PROBES

INTRODUCTION:

Sometimes one needs to screen a cDNA library with DNA probes rather than antibodies. For example if you picked a partial clone with an antibody screen you might want to rescreen the library with a DNA probe to identify clones carrying additional parts of the sequence. In other cases you might have some partial amino acid sequence information about a protein, but might not have an antibody to it. In this case you might wish to screen a cDNA library with an oligonucleotide probe. While the details of screening with an oligonucleotide probe (Protocol 3.6) may be slightly different from those used with a larger DNA probe, the basic design of the experiment is similar to what we present here.

Although traditionally screening protocols use ³²P labeled DNA probes, we present here a modified protocol in which the ECL system is used instead of radioactivity. Because we will be trying to detect DNA rather than protein, the plating and filter overlay procedure is slightly different from that used in an antibody screening. The detection step is of course completely different since we will be using hybridization rather than antibody binding to detect target plaques.

PROCEDURE:

1. The first steps of the procedure are identical to what we have already presented in Protocol 3.3 and will not be repeated here. We assume that you have already plated out your library at an appropriate dilution. Unlike the antibody screening protocols in which the lac operon must be induced with IPTG while the phage are still multiplying to maximize the yield of fusion protein, there is no need here to carry out such a step. Therefore the plaques can be allowed to develop overnight.

2. In the AM remove the plates from the incubator and overlay them with a membrane filter. BE SURE NOT TO MOVE THE FILTER AFTER YOU LET IT TOUCH THE AGAROSE SURFACE OR THE PLAQUES WILL BE SMEARED. Nitrocellulose filters can be used, but the sturdier nylon filters are preferred. PVDF filters work well with the ECL Direct labeled DNA probes.

3. Mark the filter and agarose asymmetrically using a sterile 18 gauge needle dipped in India ink or other means so that later you will be able to align the autoradiograph of the plaques with the actual plaques on the agarose. You should let the filters stay in contact with the agarose for about five minutes and then carefully remove it. If you see the agarose begin to tear away from the plate, place the plate at 4^oC for 30 minutes to stiffen the agarose. Some researchers like to prepare a duplicate filter so that they can be more certain that any dark spots noted on the autoradiogram are reproducible. If you do this, you should let the second filter stay in contact with the agarose for about 10 minutes.

4. Place the disc, plaque side up, for 3 to 5 minutes onto two sheets of Whatman 3MM paper which has been saturated in 0.5 M NaOH. This step serves to denature the immobilized DNA.

5. Place the filter into a dish containing 400 ml 5X SSC and agitate on an orbital mixer for 1 min. If any agarose remains, grasp disc on one edge using blunt forceps and agitate vigorously by hand in the SSC.

6. Place the filter, DNA side up on Whatman 3MM paper until the excess moisture is absorbed.

7. Place the filter DNA side up in Stratalinker (Stratagene) and push auto cross link to fix the DNA to the filter. Alternatively the filter can be placed while still damp DNA side down on a piece of clean Saran wrap over a transilluminator. It will usually take about 5 minutes to UV fix the DNA to the filter on the transilluminator. The exact time that will give the best hybridization signal needs to be determined empirically. If you prefer you may bake the filter at 80^oC for 1 hour. If you are using a nitrocelluose filter this step must be carried out in a vacuum oven to prevent explosions.

The preparation of probe, pre-hybridization, hybridization, washing, and film development procedures are given in Protocol 5.8

PROTOCOL 3.4: PLAQUE PURIFYING CLONE(S) OF INTEREST

SOURCE: (Sambrook et al. 2.63-2.64)

INTRODUCTION:

Once a plaque is identified by antibody or DNA screening it is essential to purify it away from contaminating plaques. The following is the basic procedure which is repeated as many times as necessary (usually twice) to obtain a pure plaque.

Procedure

1. Remove phage plug with cut off end of 1000 ml Pipetteman tip

2. Place plug into 500 ml of SM buffer with 20 ml of CHCl₃.

3. Allow phage to diffuse into SM buffer for at least 2 hours or overnight

4. Transfer supernatant to new tube

5. Determine titer of phage or make a guess. Usually, there are ~2 x 10⁶ phage per plaque, and depending on the size of the plug that was picked, calculate between 10 and 30 plaques per plug.

6. Repeat above procedure for second screen EXCEPT use smaller 100 mm plates and plate between 500 and 1000 phage for the second screen. For a third screen (when single plaque should have been picked), 20 to 50 phage particles per plate are sufficient.

MATERIALS:

1. Phage

2. SM

3. Chloroform

PROTOCOL 3.5: EXCISION OF pBLUESCRIPT CONTAINING INSERTS FROM LAMBDA UNI-ZAP XR OR ZAP II.

SOURCE: ZAP cDNA SYNTHESIS KIT INSTRUCTIONS, STRATEGENE, 1994

General Remarks:

After a clone is identified (with antibody or DNA probes), the cDNA insert will probably be moved into a variety of different vectors for different purposes (M13 for sequencing, plasmids for expression, etc.). Although lambda phage is a good vector for the construction of cDNA and genomic libraries, it is not the best vector for the isolation of large amounts of insert cDNA. Since the insert cDNA usually comprises only 5% of the DNA isolated per phage recombinant molecule (2000 bp insert/41,000 bp vector), one usually must grow up large amounts of the phage DNA in order to isolate only small amounts of insert. Large amounts of lambda phage DNA can be isolated from plate lysates (similar to amplification procedure - Protocol 3.1), followed by purifying the bacteriophage by DEAE adsorption or CsCl gradient centrifugation, and purifying the DNA from the bacteriophage. However, these steps can be long and tedious. Once the recombinant bacteriophage DNA is isolated, the insert can be removed by restriction digestion. Alternatively PCR techniques using primers flanking the vector insertion site can be used, but one must be somewhat concerned about the possibility of introducing mutations during the amplification process.

To circumvent these problems, hybrid vectors have been created that allow for the "in vivo" biological excision of the insert and subcloning into a plasmid vector in one simple step. Lambda UNI-ZAP XR is a hybrid of 3 recombinant vector molecules. The basic structure is a typical lambda phage vector with all of the genes necessary for replication and controlled expression of fusion proteins. Inserted into this phage vector is a linearized plasmid vector which carries an antibiotic selectable marker gene, an origin of replication (ori) and a unique restriction site in the middle of a regulatable gene fragment (beta-galactosidase) for the insertion of cDNA inserts. The plasmid also contains an f1 origin of replication and an f1 termination of replication. The f1 origins of replication/termination allow for the single stranded replication of any DNA that is between these two sites. (See the diagram below for details).

This elegant system is used as follows. After a cDNA of interest is identified (with antibody or DNA probes) in the intact Lambda ZAP vector, the lambda phage is plaque purified. The purified lambda phage is then infected into a bacterial host in a normal fashion. A single-stranded helper phage is coinfected into the lambda phage infected cells. The single stranded phage (ExAssist) replicates in the bacteria BUT in addition to recognizing its own f1 origin of replication, it also uses the f1 origin of replication that is part of the Bluescript plasmid cloned in the l ZAP vector. Single stranded copies are produced from the lambda phage template. These copies contain all of the plasmid components in addition to the insert cDNA. The single stranded DNA is packaged into

f1 phage particles and secreted from the bacterial cell as "phagemids". These packaged single stranded phagemids (containing the cDNA) are isolated and infected into fresh host bacterial cells (SOLR). The single stranded plasmid DNA circularizes inside the SOLR bacterial host cells and effectively becomes a plasmid molecule. The infected bacteria are plated on antibiotic plates, and only cells carrying the plasmid with the antibiotic resistance gene can grow on the selective media. Cells harboring the plasmid containing the cDNA insert of interest can be grown, the plasmid/insert DNA isolated, and insert cDNA characterized. Therefore, the cDNA insert has been biologically excised and transferred to a host bacterium in one simple step with a minimum amount of effort.

This helper phage is itself a cleverly constructed recombinant virus. It contains an amber mutation in two genes essential for phage replication. As a result, this phage can only grow in strains of E. coli containing an amber suppressing t-RNA. So when the helper phage is added to such a suppressing E. coli strain infected with a lZAP vector (which contains a phagemid with an f1 origin of replication), the helper phage is able to grow and supply the proteins needed to excise the phagemid from the l ZAP vector. However, when the resulting mixture of phagemids, i.e., phagemids with or without inserts and the helper phage are subsequently added to a non-suppressing E. coli host such as SOLR cells, the helper phage are unable to grow. This reduces possible recombination between the helper phage and the Bluescript phagemids that otherwise might occur. In addition, because the SOLR cells are l resistant, there is less chance of contaminating the final Bluescript colonies with l phage.

PROCEDURE:

Note: This procedure is for obtaining plasmid clones from isolated phagemid plaques. See the Stratagene manual for instructions on the modifications needed for mass rescue which can be used to convert a lambda library into a plasmid library.

Day 1

1. Core plaque of interest from agar plate and transfer it to a sterile microcentrifuge tube containing 500 ml of SM buffer and 20 ml of chloroform. Vortex the tube to release the phage particles into the SM buffer. Incubate 1-2 hours at room temperature or overnight at 4^oC. (This phage stock is stable for up to 1 year at 4^oC.)

2. In 15 ml tubes, grow 5 ml overnight cultures of both XL1-Blue MRF’ (tet^r, 15 mg/ml) and SOLR (kan^r, 50 mg/ml) cells in LB broth at 30^oC.

Day 2

3. Make a 1/100 dilution of both XL1-Blue MRF’ and SOLR cells into separate tubes using 0.25 ml of the overnight and 25 ml of LB broth. Grow at 37^oC with shaking for 2-3 hours to mid-log phase (OD₆₀₀=0.2-0.5).

4. Gently spin-down the XL1-Blue MRF’ cells (1500 X g). Resuspend at OD₆₀₀=1.0 in 10 mM MgCl₂. (Note: resuspending to 1/2 the original volume with 10 mM MgCl₂ works almost as well).

5. Allow the SOLR cells to grow to OD₆₀₀=0.5-1.0, while continuing with steps 6-10. Before the SOLR cells reach OD₆₀₀³1.0, remove the cells from the incubator and let them incubate at room temperature.

Note: Use cells the same day.

6. In a 50 ml conical tube or large test tube combine

200 ml of OD₆₀₀ =1.0 XL1-Blue MRF’ cells

100 ml of phage stock containing >1 X 10⁵ phage particles from step 1)

1 ml of ExAssist helper phage (1 X 10⁶ pfu/ml)

7. Incubate mixture at 37^oC for 15 minutes.

8. Add 5 ml of 2X YT broth and incubate to 2-2.5 hours at 37^oC with shaking. In the case of single plaque excisions, the reactions can be safely performed overnight, since clonal representation is not relevant.

Note: Cloudy growth may not always be seen.

9. Place 2 ml of the culture from step 8 in a microfuge tube and cap tightly. Spin 15 minutes at 2000 X g. Transfer the supernatant to a fresh tube. Heat the tube at 70^oC for 15 minutes and then spin again for 15 minutes at 4000 x g. Heating will kill the bacteria, but does not affect infectivity of the phagemid.

10. Decant the supernatant into a sterile tube. This stock contains the excised phagemid Bluescript packaged as filamentous phage particles, the original lambda zap phage, and some limited number of helper phage. The phage particles can be stored at 4^oC for 1-2 months.

11. To plate the excised phagemids, add 200 ml of freshly grown SOLR from step 5 (OD₆₀₀ = 1.0) to two 1.5 ml tubes. Add 10 ml of the phage stock from step 10 above to one tube and 100 ml of the phage stock to the other tube. (Note: In workshop we are only doing the 10 ml infection. Thus only one tube of SOLR cells is needed).

12. Incubate tubes at 37^oC for 15 minutes.

13. Plate 1 and 100 ml from each tube on LB-ampicillin plates (50 mg/ml) and incubate overnight at 37^oC. Usually 1 ml gives an large number of colonies. Note: You cannot spread 1 ml of bacterial cells efficiently. Instead you need to pipette 100 ml of LB media onto the center of the plate, then add your 1 ml of cells to it. At that point you can spread the bacteria.

Another method to assure single colonies is as follows. Dip a sterile loop into the bacterial culture at the end of step 12 then streak a plate as you would normally do to isolate a single colony from a bacterial stock. This should give you a single colony even if the 1 ml rescue is too dense.

Due to the high-efficiency of the excision process, it may be necessary to titrate the supernatant to achieve single-colony isolation.

Colonies appearing on the plate contain the Bluescript double-stranded phagemid with the cloned DNA insert. Helper phage will not grow, since they are unable to replicate in Su- (non-suppressing) SOLR strains and do not contain ampicillin resistance genes. SOLR cells are also resistant to lambda phage infection, thus preventing lambda phage contamination after excision.

To maintain your Bluescript phagemid, streak the colony on a new LB-ampicillin plate. For long-term storage, prepare a bacterial glycerol stock and store at -20^oC.

MATERIALS:

1. Plaque pure lambda ZAP clone in SM

2. ExAssist helper phage

3. Host Bacteria: XL1-Blue MRF’ and SOLR

4. 15 mg/ml tetracycline in ethanol (1000x)

5. 2X YT Medium

Compound Amount/ 1000 ml Amount/ 2000ml

Bacto-tryptone 16 g 32 g

Bacto-yeast extract 10 g 20 g

NaCl 5 g 10 g

Deionized H₂O 900 ml 1800 ml

· Shake until the solutes have dissolved.

· Adjust the pH to 7.0 with 5 N NaOH.

· Adjust the volume with deionized H₂O.

· Sterilize by autoclaving for 20 minutes at 15 lb/sq. in. on liquid cycle.

6. LB ampicillin plates (100 mg/ml)