A single plaque contains enough DNA for detectable hybridization to a labeled probe. Plaque hybridization thus allows the detection of particular plasmid or bacteriophage clones among a large number in a genomic or in a cDNA library to facilitate their selection. This technique is complementary to phenotype selection, immunoscreening or hybrid selection of mRNA and translation. If on the other hand a partial sequence is known (e.g., deduced from N-terminal sequence from a protein), then it is straightforward to synthesize a biotin-labeled oligomer and to select the appropriate mRNA from the cells to generate cDNA clones from the selected mRNA or even to amplify directly through a PCR method.
Plaque hybridization follow the same essential steps:
(i) distribution and growing of bacteria, phage, etc., on plates (‘master plates’) and replica plating on membranes; (ii) processing of the replica membranes by lysis of the bacteria or other cells or phages in situ and hydrolysis by RNase; (iii) fixation of DNA and hybridization; (iv) location of a positive signal and picking a corresponding clone on the master plate.
For the uninitiated, it is useful to mix specific recombinant clones (e.g., 100 colony-forming units) with, e.g., 1000 colony-forming units of nonrecombinant bacteria (or wildtype phage with recombinant phage, 50 plaque-forming units each, etc.) and gain experience before using precious material
Phage libraries are attractive because inserts can be large, the packaging and infection is efficient (10-50 more efficient than plasmids and thus more representative of the target sequences), phage libraries can be stored for long periods in bacteria-free, chloroform saturated buffers in glass and the titers can be well established.
Standard plaque hybridization methods
A library can be either amplified or screened directly. Amplification will yield a library that can be a source for many experiments. However, amplification may also change the composition of the library due to a different reproduction rate of the individual phages. Primary libraries are therefore screened directly if enough material is available or when a limited screening will be required. Otherwise, only a fraction of the primary library should be amplified. It is useful to keep in mind that to maintain the integrity of lambda phage particles Mg2+ is necessary and that maltose induces the expression of the lambda phage receptor. The inclusion of MgSO4, and 0.2% maltose in the media for the bacteria and MgSO4, in the SM buffer, used for the dilution of phage, is recommended but not obligatory once phage has injected the DNA into the cells (during the 15 min incubation at 37°C). Moreover, the presence of nonviable cells should be minimized since they are ‘dead ends’ for phage and decrease the efficiency of infection.
The number of plaques that is required to obtain the desired clone for 200000 plaques whereas for genomic libraries this can be 100 times larger. Titration allows the desired number of plaques per plate to be obtained. When plates are too wet, plaques tend to smear over the plate. If the plaques remain small, the plates may have been too dry or the cell density too high (a multiplicity of infection of 0.01-0.05 phage particles per cell is optimal). The low background of nitrocellulose membranes is sometimes sacrificed for more durable nylon membranes. This choice is also determined by the abundance of the target. At low abundance, it becomes more important to have lower background levels and nitrocellulose is recommended. Nylon membranes do not offer a higher detectability of the target than nitrocellulose due to the high target concentration.
Modified plaque hybridization procedures
Several replicas can be made from a master plate but it is often useful to amplify plaques in situ to increase the signal/noise ratio particularly if oligomers are used as probes. In the method of Woo (1979) and Vogeli and Kaytes (19871, a membrane is dipped in a bacterial suspension before the plaque lift and then incubated on fresh agar plates. The transferred phages infect the growing bacteria and the signal may be increased five-fold. In detail, plaques are allowed in the master plate until = 0.5 mm (if not too dense) and then incubated at 4°C for 1 h to harden the agarose. Although both Woo and Vogeli and Kaytes dipped the numbered membrane into a suspension of bacteria in fresh LB medium with 10 mM MgSO4, this is not essential since the lawn provides enough cells during transfer. The membranes are placed on the lawn with plaques and are marked by stabbing with waterproof black drawing ink and incubated for a few min at 4°C. Several replicas are then placed, phage side up, on fresh LB + Mg2+ plates and incubated until visible plaques appear (10 h; bacterial growth on nylon is slower).
Hybridization to colony or plaque nucleic acids
Detection of pertinent clones requires three steps: (i) prehybridization to block nonspecific sites on the membranes; (ii) hybridization; (iii) washing and detection of positive signals. The large majority of plaque or colony hybridizations have been achieved with radioactive probes. Biotinylated probes often give background signals with bacterial material and require special steps to reduce these. However, DIG probes have become an attractive alternative with very little background, lack of nonspecific hot spots on the film and very good resolution. Moreover, it is fast and safe. As with dot blot hybridization, it is possible to choose between formamide-containing and aqueous hybridization solutions. For mamide, requiring lower hybridization temperatures, prolongs the useful life of the nitrocellulose membranes. However, aqueous conditions tend to give superior results (typically 30 min prehybridization and 6 h hybridization at 65°C). Bacterial debris can be removed before hybridization, but after DNA fixation, by washing for 60 min in 5 X SSC and 0.5% SDS at 65°C and scraping the debris off gently with gloved hands or with Kimwipes soaked in the same buffer.
Well-separated colonies contain large amounts of target DNA and can be detected with probe at about (1-5) X lo5 cpm/ml whereas dense screens require about 5 x lo6 cpm/ml. The probe is alkali-, microwave- or heat-denatured and added to fresh hybridization solutions. After washing, wet or damp membranes are placed on used X-ray film covered with Saran wrap and then covered with Saran wrap before placing on unexposed X-ray film. The length of autoradiography depends on the amount of label but can be as short as a few minutes but is usually 1-24 h. Plaque hybridization may require up to ten times longer autoradiography.
Probes can also be stripped in order to rehybridize with a different probe although the same information can be obtained from replicas. Since only a very small amount of the probe will hybridize, it is possible to recover the probe, denature and reuse again. Denaturation in aqueous solutions should be at = 100°C for 15 min, whereas for solutions containing formamide, 15 min at 70°C should suffice.
Despite the wide use of radioprobes in colony or plaque hybridization assays, nonradioactive probes can be advantageous. The use of biotinylated probes, initially the most common among nonradioactive detection systems, is limited since biotin-streptavidin systems tend to give high background levels with bacterial material unless specific measures are taken. The main restriction is that monoclonal antibodies (commercially available) should be used since polyclonal antisera often contain antibodies against bacteria. The main drawback of nonradioactive probes is the ability to reprobe the same membrane. It is possible, however, to strip a membrane of its probe after a colorimetric detection and to perform a chemiluminescent detection or vice versa.
Biotinylated probes have been applied in colony hybridization and plaque hybridization. They require proteinase K and chloroform treatments to reduce background. After lysis and neutralization the membranes are incubated in 30 ml of 1 X SSC, containing 200 pg/ml proteinase K, for 1 h at 37°C. After rinsing twice for 2 min in 30 ml 90% ethanol (wt/wt, not v/v otherwise it exceeds the tolerance level of some batches of nitrocellulose), the membranes are dried and 100 ml of chloroform are passed through each membrane using the filtration device, air-dried and incubated for 5 min in 0.3 M NaCl (30 ml/membrane; optional) and baked in vacuo at 80°C. (Pre) hybridization is in the presence of 45% formamide at 42°C and 100-200 ng/ml biotinylated probe is used. Hybridization solutions can be reused at least 10 times over a time span of at least 5 months if stored at -20°C. Background levels increase with the density of the bacterial colonies but positive colonies can still be distinguished at very high densities. Note that the chloroform treatment tends to shrink the membranes and makes them brittle.
Colony and plaque hybridization with the DIG system is very useful. Following standard transfers to nitrocellulose membranes and baking, membranes are prehybridized in 5 X SSC (containing 0.5% BM blocking reagent (0.1% sarkosyl and 0.02% SDS) for 1-8 h at 68°C. Fresh buffer with 5-200 ng/ml DIG-probe is used for the overnight hybridization at 68°C. Post hybridization washes include two rinses for 5 min at room temperature in 2 X SSC and 0.1% SDS and stringency washes at 65-68°C with 0.1% SDS in 0.1-0.5 x SDS (depending on the degree of stringency required). For immunological detection, membranes are washed for 1 min in wash buffer (0.1 M Tris-HC1, pH 7.5, containing 0.15 M NaCI) and then blocked with BM blocking buffer (for colorimetric detection 0.5% blocking reagent and for chemiluminescent detection 2% blocking reagent in wash buffer). After blocking, anti-DIG Fab fragments conjugated with alkaline phosphatase, diluted 1/5000 in blocking buffer, are allowed to react for 30 min. Enzyme is detected. Colonies can be detected in 10-60 min and background staining is minimal although it increases with time. In the case of nylon membranes, 7% SDS in 50 mM phosphate buffer is used during (prelhybridization and for the blocking buffer increasing the concentration of blocking reagent to 1% and adding 50 kg/ml carrier DNA. It is then very important to rinse the membranes well with the Tris buffer between hybridization and immunological detection since phosphate is a potent competitive inhibitor of alkaline phosphatase.
Reduction of background
Background signals should be identified as much as possible by: (i) use of replica membranes and increasing stringency; (ii) use of negative control and, if available, positive control membranes; (iii) careful preparation of probes; (iv) optimal hybridization conditions. Particularly with radioactive probes, the spontaneous signals (from nonspecific radioactive spots to static electricity-generated dots) can be easily recognized from duplicate membranes. Only those present on both membranes are considered. ‘Static dots’ can be reduced by using a sheet of paper, as found in the X-ray film box, instead of or in addition to the Saran wrap.
Negative controls (clones with irrelevant insert) and positive controls (clones with probe insert) are excellent indicators of the quality of the procedure. Whenever possible, an irrelevant insert should be chosen which has a higher GC content and is longer than the relevant insert. Probes should be carefully prepared. Particularly those with GC tails or GC-rich regions are prone to give high background levels. Radioactive probes should always be filtered before use. Since colony or plaque membranes may contain a considerable amount of bacterial debris, prehybridization solutions should never be used, contrary to slot blot hybridization, as hybridization solutions. Probe should be added to fresh solutions. Oligomer probes tend to adhere nonspecifically to colony proteins on the membranes. Special grade colony/plaque membranes and the use of SDS in the hybridization solution reduces this problem somewhat.
Selection, picking and purification of clones
The size of the signals depends on the size of the area occupied by the recombinant clones but can be as small as pinpoints for high density screens. The positive spots are circled with a red pencil and the autoradiogram, or its mirror image (depending on orientation marks) by reversing the film, is oriented with respect to the master plate on a light box. Positive colonies are picked with a tooth pick if very well separated or all bacteria within the positive area with a sterile wire loop. Several dilutions are made and those with about 25-1000 colonies per 150 mm plate are screened again by colony hybridization until a well-isolated colony is obtained.
Positive plaques are picked with the wide (‘wrong’) or narrow end (depending on the resolution of plaques) of a Pasteur pipette and the agarose plugs are added to SM buffer (100 l of chloroform are added, vortexed and incubated for 30 min. After centrifugation for 3 min at 3000 Xg, phage is plated at different dilutions and the plaque hybridization is repeated for plates containing 25-1000 plaques/l50 mm plate. Well-isolated plaques should then be obtained.
- Plate phage so that about 10000 plaques are obtained per 135 mm plate (or about 5000 per 82 mm plate).
- When plaques have developed, but are not yet confluent, cool plates for 1-2 h at 4°C (not overnight, then place at 4°C and let plaques develop the next day). If plaques develop poorly (poor bacterial growth), then top agar was probably too warm during plating.
- Remove plate covers and let agar air-dry for about 30 min at room temperature. Label each plate and corresponding membrane with a pencil and carefully apply, without trapping air bubbles, to the surface of the top agar (ink-side up). The membrane will wet in about 1 min. Make matching marks by punching the edge of the membrane and the agar with a clean 18-G needle (patterns should be different for the various membranes). Let membrane sit on the agar for about 2-10 min. Peel the membrane off carefully (leave top agar intact). If problems are encountered, longer cooling or higher concentration in top agar (agarose) may be required. Up to five replicas can be made from a plate without regenerating the plaques.
- Alternative: in situ amplification of plaques. Resuspend bacteria in LB+ 10 mM MgSO4. Label the membrane, dip in bacterial suspension and air-dry briefly. Lay the membrane on a plate with plaques (phage will come into contact with bacteria and infect them). Place the membrane, plaque side up, on a fresh LB+Mg2* plate and incubate inverted overnight (considerable amplification of phage will occur). Do not forget to apply matching marks to the membrane and master plate.
- Dry membranes for about 30 min on the bench top and place them on 3MM paper saturated with 0.2 N NaOH+ 1.5 M NaCl (plaque side up) for 2 min. Transfer then to 3MM paper saturated with neutralization solution (1.5 M NaC1+ 0.5 M Tris-HCI, pH 7.4) and with ~ x S S C ,re spectively, 2 min each. Dry membranes and continue with hybridization. Replica membranes can be used to increase contact with each solution to 1 min; however, the NaOH treatment should be kept to less than 5 min for nitrocellulose.
- Pick colonies, corresponding to the positive signal on the film, with a Pasteur pipette and resuspend in 500 p,I of SM buffer (step 4), add a few drops of chloroform and vortex for a few seconds. Spin at 3000 X g for 3 min and collect the supernatants (add a few drops of chloroform).
- Dilute phage stock to = 200 plaques/O.l ml and replate. Repeat plaque hybridization and pick well-isolated plaques. Note: Nylon membranes do not offer a higher detectability than nitrocellulose due to the high concentration of the target in the plaques.