Immunologicalscreening of bacteriophage l expression libraries
Protocol created by Amanda Liggins & Alison Banham - LRF Haemato-oncology Group, University of Oxford
- The library must be constructed from mRNA extracted from a cell line or tissue that expresses the protein of interest, preferably in reasonable abundance.
- The library must be cloned in a vector suitable for expression in E. coli e.g. λgt11 or λZAP. There are a number of commercially available libraries (e.g. Clontech, Stratagene) that have been used successfully to identify a number of antigens.
- The library should have a complexity of at least 106 as in theory (not in practice as internal initiation occurs) only 1 in 6 inserts will be in the correct frame and orientation for expression as a ß-galactosidase fusion protein.
- There are two types of library. Those generated by oligo-dT priming, which will favour C-terminal epitopes or those generated by random priming, which should be better for N-terminal epitopes. In practice a mixture could be used.
- Ideal = monoclonal or polyclonal antibodies, absolutely specific for conformation- independent epitopes displayed on both native and denatured forms of the protein, high titre and IgG.
- Antisera that fail to detect the protein by Western blotting are probably specific for native epitopes and may be unsuitable.
- The antibody must recognise unmodified epitopes e.g. not oligosaccharides or phosphorylated epitopes, as these will not be present on prokaryotically- expressed proteins.
- Test a range of dilutions of the antibody against non-recombinant plaques, a sample of the library and a series of dilutions of native and denatured antigen. Suggested dilution ranges are 1:200-1:5000 for polyclonals or 1:10- 1:100 for hybridoma supernatants. The optimal dilution is that which displays a high reactivity with the antigen but does not react non-specifically with prokaryotic proteins. Ideally the antibody should be able to detect as little as 50-100pg of protein in the size of a small bacterial colony.
- The best antibodies are either pooled monoclonals that tend to have low background signal or high-titre polyclonal antibodies that don't react with the host components. Background from polyclonals can always be reduced by preclearing or affinity purifying the antiserum.
- The amount of antigen expressed varies widely and is dependent on:
The toxicity of the fusion protein
Its rate of degradation
Its physical state
Antibodies capable of recognising 50-100pg of denatured antigen should allow all but the most labile fusion proteins to be recognised.
Titration of the cDNA Library
- 2 days before they are required pour LB plates in 90mm petri dishes (500ml medium makes about 20 plates).
- 1 day before required prepare a plating culture: Just before going home inoculate a single bacterial colony from a fresh plate into 100ml LB medium supplemented with 0.2% maltose and 10mM MgSO4. λgt11 use E.coli Y109hsdR and include 50 µg/ml ampicillin. λZAP use E.coli XL1 Blue MRF’ and include 12.5 µg/ml tetracycline.
- Incubate culture overnight at 30°C and 225rpm.
- Pellet cells at 3000rpm (approx. 4000g) for 10 minutes at R.T.
- Gently resuspend the cells in 40ml of 10mM MgSO4. Read OD600 and dilute to OD600= 0.5 with 10mM MgSO4. These plating bacteria are now ready for use and can be stored at 4°C for up to two weeks but are best when used fresh.
- Late afternoon: Dry the plates well 1-2h at 37°C with the lids slightly open. Melt top agar (0.7% agarose in LB) in microwave and then allow to equilibrate to 45°C in a waterbath.
- Prepare serial dilutions of the library in SM buffer aiming to have 10-100's of colonies / 0.1ml. Library titers are usually 109-1012 pfu/ml.
- Aliquot 0.1ml of each of the last 4-5 dilutions into sterile 5ml+ tubes. Add 0.1ml of plating bacteria per tube (OD 0.5) and vortex gently. Incubate without shaking at 37°C for 20 min.
- One sample at a time, add 3ml molten top agar to the tube of bacteria, mix and immediately pour onto a warm LB plate, swirling to evenly coat the plate.
- Allow the top agar to set then invert the plates and incubate O/N at 37°C.
- Next day count the plaques and calculate the original library titer. This is worth doing as library titers do drop on storage.
- 2 days before they are required pour large LB plates (140mm diameter; require approx. 45ml medium/plate).
- Also require plating bacteria prepared as described previously.
- Calculate the number of plates required to screen the library using 5 x 104 plaques per 140mm plate. Aim to screen 10-20 plates in the primary screen.
- Dry the plates well at 37°C 1-2h with lids ajar. Keep warm at 37°C. Melt top agar (microwave) and equilibrate to 45°C.
- Dilute the library in SM buffer to obtain 5 x 104 plaques per 0.2ml.
- Aliquot diluted library into the appropriate number of 15ml tubes for the number of plates poured. Add 0.6ml of plating cells, vortex and leave unshaken at 37°C for 20 min.
- Do one tube at a time. Add 10-11ml of top agar, mix gently by inversion and pour onto a warm plate, swirling to evenly coat the plate.
- When all the plates are set, transfer to 42°C for 3-4h until plaques are just visible as pinpricks.
- Number filters with a soft pencil and also number matching plates. Soak one nitrocellulose filter per plate in 10mM IPTG and pat dry. Work quickly to stop plates cooling or filters drying out. Place a filter on a plate (don't lift and reposition) and transfer to 37°C incubator.
- Incubate for at least 2h, but usually 4h. If not doing duplicates, filters can be left O/N. Apparently removing the lid for the last 20 minutes prior to removing the filter helps to prevent the top agar from sticking to the filter. If real problems occur with agar sticking, or there are no duplicate filter lifts, put plates at 4°C for 30 minutes or 5 minutes at -20°C before attempting to remove the filters.
- Mark the orientation of the filter with a needle and then remove the filter using flat-edged forceps and place in TBS-T (0.05% Tween 20).
- If doing duplicate filters overlay a 2nd IPTG soaked filter and leave at 37°C for 3- 6h.
- Then repeat step 11. Store plates at 4°C, as they will be needed to pick any positive plaques.
- Wash filters for 5-10 min to remove any agar and reduce background. Transfer to fresh TBS-T and wash gently for 30 min at R.T. After this treat filters in batches of 5 per petri dish.
- Block O/N at 4°C or 30 min R.T. in TBS-T + 5% dried low-fat milk.
- Transfer to primary antibody solution (antibody diluted in block; 50ml per petri dish) and incubate 30 min -1h at R.T.
- Rinse filters with TBS-T and then wash filters 1x in TBS-T for 10 min.
- Add secondary antibody against the host species of the primary antibody (horse- radish peroxidase (HRP)-conjugated or alkaline phosphatase (AP)-conjugated) at recommended dilution in TBS-T and incubate at R.T. for 30 min.
- Wash filters 4 x 15 min in TBS-T.
- Add DAB (if used HRP-conjugated secondary) or AP substrate, allow colour to develop and then stop reaction with water.
- This chromatographic detection method should identify positive plaques.
Isolation of Positive Plaques
- Positive plaques should be identified by a colour reaction, these can be checked if duplicate filter lifts were made. Pick both strong and weak positives as these may represent different clones or sizes of cDNA.
- Align the filter and petri dish using needle holes on a light box. Core the plaque of interest and 3-4 surrounding negative plaques from the plate using the wide end of a sterile pasteur pipette. Transfer this to a sterile eppendorf tube containing 0.5ml SM buffer. Vortex the tube and incubate 1-2h at R/T or O/N at 4°C to elute the phage. Add 20µl or 1 drop of chloroform, vortex and microfuge at 13000rpm for 1 min to separate the phases. This phage stock is stable for 1 year at 4°C.
Second Round Screening of Positive Plaques
- Titre phage on 90mm petri dishes plate 100, 10 or 1µl using 200µl of plating bacteria (see titration of library).
- Rescreen the appropriate dilution on 90mm dishes for individual positives, as described above.
- Following identification of individual positives, pick with the fine end of a pasteur pipette, transfer to 0.5ml SM and prepare a phage stock as above.
In Vivo Excision of the pBK-CMV Phagemid
- Grow O/N 100ml cultures of E.coli XL1-Blue MRF' and E.coli XLOLR strains supplemented with 0.2% maltose and 10mM MgSO4 cells in NZY broth at 30°C.
- Pellet cells at 3000rpm (approx 4000g) for 10 min and resuspend to an OD600 of 1.0 in 10mM MgSO4.
- In a 15ml Falcon tube combine 200µl XL1-Blue MRF' cells, 250µl phage stock and 1µl of ExAssist (Stratagene) helper phage.
- Incubate unshaken at 37°C for 15 min.
- Add 3 ml NZY broth and incubate at 37°C and 300rpm for 3 hours (doesn't always go cloudy).
- Heat at 65-70°C for 20 min to lyse cells, pellet debris by centrifugation at 3000rpm (approx 4000g) and 4°C for 15 min.
- Decant the supernatant into a sterile Falcon tube. These can be stored at 4°C for 1-2 months.
- To plate excised phagemids set up 2 microtubes per sample containing 200µl XLOLR cells. Add either 100µl or 10µl from step 7.
- Incubate at 37°C unshaken for 15 min.
- Add 300µl NZY broth and incubate at 37°C and 300rpm for 45 min.
- Plate 200µl from each tube on LB-kanamycin (50 µg/ml) plates and incubate O/N at 37°C.
- Bacterial colonies can be grown up in LB-kanamycin and DNA can be prepared using standard plasmid kits.