2) TITRE: Make dilutions of phage library in TM10+ gelatin such that 10-100 plaques will form on at least one plate. Mix 100µl of indicator bacteria with an appropriate volume (10-300µl) of phage dilution, incubate for 20 min at 37°C, add 3ml top agar at 48°C, mix by gentle vortexing or inversion and pour evenly on a small (9cm) lambda plate that has been dried well (4-12 hr at 37°C, 2-4 hr at 42°C if fresh). Incubate at 37°C for >8hr and count plaques.
3) PLATING LIBRARY: To 0.3 ml indicator bacteria, add appropriate amount of
phage library stock or dilution (~5-10,000 per plate for genomic, 20-50,000 for
cDNA), mix, incubate 20 min at 37°C, add 8ml top (0.7%) agarose at 48°C, mix
and plate on well dried 15cm plates. Incubate at 37°C for 8-16 hr (cDNAs in
lambda gt10 grow much faster than genomic DNAs in Charon 4).
lifts;
4) LIFTS: Cool plates (>1 hr at 4°C), label nitrocellulose circles with plate names,
layer smoothly on agarose surface, stab holes through filter and agar with
syringe needle (narrow guage for accuracy) dipped in water-insoluble ink and
peel filter off after 30s (first lift), 60s (second lift), 3 min (third lift), 5min
(fourth lift), any time (last lift). Immerse successively in denaturing solution
(5-10 min), neutralising solution (10 min) and 2XSSC (to rinse) with occasional
agitation. Store plates in sealed packages at 4°C. Lay filters on 3mm paper to
air dry for at least 30 min before baking 2 hr at 80°C. Remove moisture from
vacuum oven as required.
hybridisation;
5) HYBRIDISATION: Pre-hybridise for 2-24 hr in aqueous buffer (5 or 2XSSCP,
10X Denhardt's, 250µg/ml salmon sperm DNA, (0.1-0.5% SDS)) at 65°C or in
same buffer with 15-50% de-ionised formamide at 42°C (no SDS when using
formamide) in bags with ~6 filters each. Add denatured probe (nick-translated,
single-stranded, riboprobe, oligonucleotide) to fresh buffer (<106cpm/ml) and
replace pre-hybridisation fluid with ~25ml of this per bag. Incubate at 42°C or
65°C (with shaking if possible) for 8 (nick-translated probe) to 36 hr
(single-stranded probes). Save probe (store at -20°C) for re-screen. Wash filters
in 0.1-0.5XSSC, 0.1% SDS at 65°C (high stringency) or 0.5-6XSSC, 0.1% SDS, at
42°C (low stringency) for a total of 3-4 hr with 3-4 changes. Wrap filters in
Saran wrap (facing a uniform direction for convenience), mark phosphorescent
spots and expose for between 2 and 24 hr.
plaque purification;
6) PLAQUE PURIFICATION Align filters with film and mark syringe needle
positions. Align film with plates and take a small plug (diameter ~2-3mm) of
agarose+agar from around each positive and elute phage in 2ml TM10
(+/-gelatin)+2 drops chloroform for >2hr at room temp with shaking. Dilute
1µl into 2ml TM10 and plate 2-10µl (gt10 libraries) or 20-100µl (Charon 4
libraries) as before in top agarose on dry small or large plates to give 100-1000
plaques. Make lifts and repeat screening procedure using same probe (boiled for
5-10 min before use) and pick isolated positives, or purify a further round if
necessary.
7) RE-USE OF FILTERS: If filters are kept in Saran wrap at 4°C they may be
re-probed after pre-hybridising, probably any time within several weeks (but
check the status of the plates). As the position of positives is already known it
is not really necessary to boil the first probe off.
There exist a number of bacteriophage lambda libraries carrying genomic DNA sequences or cDNA copies of poly A+ RNA isolated from various stages of development or different body parts of D. melanogaster.
Considerations in Choice of Library:
(1) REPRESENTATION
The Maniatis (Maniatis et al., 1978) and EMBL (for vector see Frischauf et al., 1983; Oregon R library, Pirrotta et al., 1983; Canlon S library, V. Pirrotta, unpublished) libraries were made by sonication and partial Sau3A digestion, respectively, of high molecular weight DNA. The DNA content of each library (6x105 x16 kb ; Maniatis) far exceeds (1.65x105 kb) that of the Drosophila genome. Theoretically, therefore, the genome should be fully represented by each of these libraries, with the possible exception of DNA that is not readily extracted, DNA that has an unusual structure (e.g. heterochromatic) and DNA that contains sequences which cannot be propagated in rec A+ hosts due to recombination or "poisoning". Practical experience has confirmed these expectations. It has been possible not only to clone a large number of specific genes but also to establish an extensive series of overlapping phage in chromosomal "walks" in either of these two phage libraries. Difficulties in cloning have been encountered for sequences that include a long inverted repeat (head to head defective P elements at the sn locus). Less severe differences in the ability of recombinant phages to grow have been inferred from under- and overrepresentation of particular characterised phage. This effect is rarely important but may be minimised by the use of unamplified libraries.
The lambda EMBL vectors tolerate a large insert (23 kb) and recombinants apparently grow slightly more vigorously than Charon 4 recombinants. These benefits are to be weighed against the greater body of gene cloning history with the Maniatis library and the more random nature of the latter library (from sheared DNA as opposed to partial Mbo I digestion).
The representation of particular sequences in cDNA libraries is subject to many more variables than pertain to genomic libraries. Amongst these are the number of phage with inserts of reasonable size, the abundance of the specific RNA in the source material for the library, the possible absence of a polyA tract, the possible presence of strong stop sequences for reverse transcriptase and the under-representation of sequences derived from the 5' end of the RNA. It is therefore strongly recommended that the initial isolation of any particular gene should be attempted using genomic rather than cDNA libraries even if the proportional representation of the sequence is expected to be higher in the latter library. (Of course this does not apply when the use of specific cDNA libraries is a crucial step in the protocol used to identify a particular gene).
The expected frequency of occurrence of a specific cDNA in a library can be roughly predicted from the abundance of the corresponding RNA in the source for the library. As most cDNA libraries include at least 105-106 and up to 5x107 independent recombinants it should be possible to isolate cDNAs for RNAs of abundance down to about 0.0001% (assuming that the majority of packaged phage include bona fide inserts of reasonable length).
(2) LENGTH OF INSERTS
For genomic DNA libraries the constraints of packaging limit the size of inserts to between 15 and 23 kb.
For cDNA libraries the size of inserts depends upon a large number of experimental variables and is thus best judged a posteriori. However, it should be noted that the method of construction can influence the quality of inserts in a predictable fashion. (i) The use of hairpin formation to prime the second cDNA strand rather than G-C tailing makes the presence of full-length or nearly intact 5' ends very unlikely (G-C tailing is now generally used). (ii) Most cDNA libraries include EcoRI linkers at each end. Thus, methylation of internal EcoRI sites is required before addition of linkers to prevent internal cleavage. Although treated with EcoRI methylase during construction, most libraries nevertheless contain some (~20%) recombinants that have been cleaved at internal EcoRI sites. A large proportion of cDNAs that have been examined have additional or 3' derived sequences of 20-30 bp at their 5' ends. It is not clear if this is a general problem or whether it is specific to those libraries that have been examined.
PLATING OUT BACTERIOPHAGE LAMBDA LIBRARIES
1. Number of plaques to screen
a. Genomic library
The number of recombinants of average insert length 16.5 kb that must be screened
to detect a particular sequence in a Drosophila genomic library; is given in the
following Table:
| Number of recombinants screened | Probability of finding at least one phaage | Probable number of phage containing target sequence |
| 5,000 | 39% | 0.5 |
| 10,000 | 63% | 1 |
| 20,000 | 86% | 2 |
| 40,000 | 98% | 4 |
| 80,000 | 99% | 8 |
| 120,000 | 100 - 6 x 10-4% | 12 |
Thus, a genomic library is generally screened by plating out 6-12 15 cm plates, each with 5-10,000 phage.
b. cDNA library
As the number of cDNAs in a library for a given mRNA species cannot be accurately predicted, it is best to screen as large a number of recombinants as reasonable. The maximum number of phage that can be plated on one 15 cm plate and still give a reasonable hybridisation signal is in the order of 20-50,000. Theoretically, any number of plates can be screened simultaneously. However, for an amplified library, the number of new (as opposed to clonally related) recombinants being screened is greatly diminished as the number of plaques being screened approaches the complexity of the library. Hence it is usually better to screen several libraries up to a fraction (1/3 or 1/2) of their complexity than to screen the equivalent number of plaques from just one library.
1) CELLS : Make a saturated culture of the appropriate bacterial strain ( C600 or Q358 ) in O/N broth + 0.2% maltose. The culture may be grown from single colonies but can conveniently be grown in only a few hours from (0.1-1 ml) aliquots of frozen cells. The saturated culture can be stored at 4°C for at least a week but is best stored after pelleting the cells and resuspending in an approximately equal volume of TM 10 + gelatin. This avoids any potential problem of phage destabilisation during absorption due to absence of Mg2+ ( O/N broth contains no Mg2+ )
2) PHAGE : Dilute the bacteriophage lambda stock in TM10 (+/- gelatin) so that the required number of phage per plate (5-50,000) can be measured accurately (10 µl- 0.5 ml). The titre of the phage should first be accurately measured by making dilutions in TM10+gelatin and plating as below. Titres are generally stable for phage stored at 4°C inTM10 and chloroform.
3) Mix 2-500 µl of the indicator bacteria in TM10 (~4-20x108 cells; 1 OD600 = 8 X 108 cells/ml) with the appropriate volume of phage dilution. When plating phage at very high density it is best to use slightly more cells (500 µl of 3x concentrated cells) so that small plaques form, allowing a more even representation of phage.
4) Incubate for 15-20 min at 37°C
5) Add 8 ml top agarose at 48°C to the mixture of cells and phage. Mix by inversion or gentle vortexing and pour evenly on a 15 cm lambda agarose plate. Even spreading of top agarose is facilitated if plates are warm (37°C, 42°C). Allow ~10 mins. to set.
6) Incubate inverted at 37°C for 8-16 hrs.
IMPORTANT POINTS
(i) Check for contamination of solutions or bacteria with phage by plating one plate
as for the others but with no added phage.
(ii) All solutions and media in which phage are stored, mixed or incubated must contain Mg2+ (~10 mM) otherwise the bacteriophage particles fall apart.This may be seen as a reduction in titre (possibly down to zero) and can give rise to plaques of variable size.
(iii) Top agarose (at the right concentration) must be used for plating (as opposed to agar) to minimise the chance of peeling this layer off when taking lifts.
(iv) Top agarose should not be too hot (>50°C) otherwise small plaques in reduced titre result.
(v) Plates must be very dry (no gloss and preferably with ridges) to prevent problems when taking lifts.
MAKING LIFTS FROM PLATED-OUT LIBRARIES
Multiple lifts; from one plate :
By leaving the nitrocellulose filters on the plate for successively increasing periods
of time it is possible to make several lifts from one plate that contain reasonably
similar amounts of phage. Duplicates are to be recommended whenever the number of
filters to be screened is not already excessive and particularly in screens at low
stringency where spurious signals (not due to DNA-DNA hybridisation) may appear.
For the times listed below for making four lifts, the first two lifts usually remove
considerably more phage than the third and fourth. It is not possible to reduce the
contact time for the first lift if the plates were originally very dry as the filters require
some time to become evenly wetted.
1) Cool the plates down to 4°C (>1 hr, exposed in the cold room) so that top
agarose surface is as hard as possible.
2) Label nitrocellulose circles according to plate designations in non water-soluble ink (most ball-points are OK).
3) Layer individual filters smoothly on the surface of the plates making contact from the centre outwards. If the plates were very dry, a gloved hand will be required to ensure that the entire filter wets in a reasonable time.
4) With a syringe needle dipped in water-insoluble ink mark at least three (five is
better) asymmmetrically disposed spots on the filter by stabbing vertically
through to the agar.The spots should be easily visible but fine enough to allow
accurate alignment of filter and plate (25G5/8 needle is good). Peel the filter off
from one edge and immerse, plaque side up in denaturing solution. When making
multiple lifts peel the filter off after fixed periods of time (after making
alignment spots):
Lift 1 - 30 sec
Lift 2 - 1 min
Lift 3 - 3 min
Lift 4 - 5 min
To maximise signal strength, leave the filter on the top agarose for as long as possible for the last lift (some phage will always remain).
5) Agitate denaturing solution occasionally to ensure filters are being wetted.
6) Transfer filters in same orientation after 5-10 mins. (excess is better than too little time) to neutralising solution. Leave for 10 mins. with some agitation to allow mixing of solutions at the surface of the filters.
7) Transfer to 2xSSC to rinse off high salt solutions (2-3 min with agitation or longer if convenient).
8) Lay filters, DNA side up on 3 mm paper to air dry for at least 30 min.
9) Sandwich the filters between 3mm or other (Blot block) paper and bake in the vacuum oven for 2 hr at 80°C. No matter how long the filters were air dried they will retain a considerable amount of moisture. For the sake of the vacuum pump, seal off the evacuated chamber and wipe off condensation on the door intermittently.
10) The filters can now be stored for several days or weeks or used immediately. The plates from which the lifts were taken should be stored at 4°C, sealed in a plastic bag and labelled. When using nitrocellulose filters straight from the pack (not sterile) it is common to see some contaminant bacteria on the plates after a few days. However, such contaminants are generally lost rather than amplified on subsequent re-screening and amplification of phage.
IMPORTANT POINTS
1) Filters should peel off easily from the top agarose without removing any
agarose. If some very small pieces of agarose stick they should be removed while
in the denaturing solution. If a lot of the top agarose peels off, stop. Even after
extensive cleaning it is likely that there will be a high hybridisation background
and no guarantee of an even signal. Repeat plating out, checking that plates are
very dry, top agarOSE is of correct composition (should set at ~45°C), plates
are chilled before lifts are made (and that nitrocellulose wets easily).
2) If filters are not neutralised for long enough they will turn yellow and become brittle upon baking. Whether this affects anything other than the ease of handling the filters is debatable.
HYBRIDISATION:
hybridisation;
PROBES
1. Nick-translated or random primed plasmid DNA or fragments
2. Single-stranded probes from ss phage or cDNAs
3. RNA probes
4. Oligonucleotide probes
The choice of probe is sometimes dictated by the nature of the experiment, e.g.
screening for genes that are differentially expressed (cDNA) or finding genes from
amino acid sequences (oligonucleotides). However, when there is a choice the following
advantages and disadvantages of each type of probe should be considered.
1a) Nick-translated or random primed plasmid DNA: Easiest probe to make
reproducibly sufficiently hot for library screening. Do not use if libraries are
contaminated with plasmid sequences.
1b) Nick-translated or random primed fragments: Although the Drosophila genomic libraries do not contain sequences highly related to commonly used plasmids there is some benefit to using fragments as opposed to plasmids to screen at low stringency. Background hybridisation is generally lower (perhaps due to impurities in plasmid DNA preps.) and the possibility that a strong signal is due to plasmid cross-homology is immediately eliminated. Perhaps surprisingly, the use of fragments purified only once (twice is OK) on an agarose gel does not eliminate strong hybridisation to plasmid-containing phage in contaminated libraries.
2) Single-stranded probes: Presumably because all radioactive material added to
the hybridisation is as a defined DNA molecule, the use of M13 derived
single-stranded probes gives the cleanest signals. Such probes can also be made
of higher specific activity than nick-translated probes. However, it is necessary
to start off with a lot of radioactive dNTPs to incorporate the same amount of
radioactivity into single-stranded as nick-translated probes (because in the
nucleotide excess that is used to generate probes of maximal specific activity, a
lot of hot dNTPs are incorporated into vector rather than probe DNA). Hence,
starting with the same amount of radioactive dNTPs the final signal achieved
(under equivalent conditions) is generally lower (up to 5-fold) using
single-stranded probes but much cleaner. The use of single-stranded probes is
therefore highly recommended for low stringency hybridisations (sensitivity can
be increased by scaling up from the standard protocol and by hybridising for
longer than for nick-translated probes).
M13-derived "double-stranded" probes can be made as for
single-stranded probes but omitting the fragment purification step and can be
used as a substitute for nick-translated probes. However, (i) if anything, the
cold strand will be in excess over the hot strand; (ii) the probe will be long
(which in some people's view is not desirable); and (iii) the probe cannot be
used at low stringency (certainly for the Maniatis library) as ~100 signals per
5000 plaques are seen, presumably as a result of sequences cross-homologous to
M13.
3) RNA probes: RNA probes have not been widely used for screening libraries. It is straightforward to make very hot probes and backgrounds appear to be lower than for nick-translated probes. However, the increased stability of RNA/RNA and RNA/DNA hybrids relative to DNA/DNA hybrids can lead to two problems. First, some regions (or all) of the probe may be unavailable for hybridisation, particularly at low stringency, and second, capricious cross-hybridisation may (and has) been seen, requiring hybridisation at several stringencies to assess the significance of such signals.
4) The highest theoretical specific activity of a terminally labelled oligonucleotide
(~104 Ci/mmol) is only about 3-fold lower than for an average nick-translated
probe of average length 400 bp (assuming 50% of probe is newly synthesised
from a mixture with 2 hot nucleotides at 300 Ci/mmol).
Thus oligonucleotide hybridisation is potentially of similar sensitivity to standard
screens with nick-translated probes. This should allow for the use of degenerate
oligonucleotides.
The appropriate conditions to use for the hybridisation of oligonucleotides; can be
worked out from the following formula (quoted in Meinkoth & Wahl 1984)
Td = 4(G+C) + 2(A+T)
where Td is the temperature at which 50% of duplexes dissociate and A,G,C,T are the numbers of the corresponding bases in the oligonucleotide. Any mis-match leads to a reduction of about 5°C in Td. Hybridisation is normally conducted at 5°C below Td.
In practice oligonucleotides of 30 bases with 2-4 mis-matches have successfully
been used by hybridising in 7XSSCP at 37°C and washing in 7XSSCP at 42°C.
PRE-HYBRIDISATION
Filters should be pre-hybridised for as long as possible but 2-4 hrs. is usually
sufficient. As the conditions for low stringency hybridisation allow more non-specific
attachment of probe to the filters, 12-24 hr pre-hybridisation is suggested in this
instance. It is possible to pre-hybridise in one solution and hybridise in another but
logic would suggest that it is best to use the same composition of solution in the two
cases. Thus, filters should be pre-hybridised in aqueous solution (see below) at 65°C
or, for low stringency in formamide solutions at 42°C in the same bags to be used for
hybridisation. Six filters per bag is certainly not excessive. Large volumes (~50 ml per
bag with 6 filters) should be used for pre-hybridisation.
HYBRIDISATION
a. High stringency
Hybridisation will only take place with perfect or near perfect homologues at 65°C in the following solution
2X or 5X SSCP
10X Denhardt's
250 µg/ml herring/salmon sperm DNA (should be boiled immediately prior to use
to denature)
(0.1-0.5% SDS) - optional
Using 250 ng nick-translated probe (~10-25 X 106 cpm) in 25 ml in each bag with 6 filters, 8hr is probably sufficient (12-16 hr is generally used) for hybridisation to approach completion. (Theoretically, a probe of 2 kb should be 50% renatured by 8 hr). Under these conditions a signal should be observable in 4-12 hr (at room temp.-Kodak XAR-5). When using single-stranded probes (generallly at lower concentration than nick-translated probes) longer hybridisation times (24-36 hr) should improve the signal but are not necessary.
b. Low stringency
Although it is possible to screen libraries simultaneously at several stringencies (but take care that all lifts have roughly equal amounts of DNA), it is best either to ascertain the optimal stringency for screening by performing genomic Southerns at a variety of stringencies or to use the lowest possible stringency under which signals stand out relative to background. Another strategy, that of hybridising at very low stringency and washing at successively increasing stringencies is not recommended as (i) it takes a long time and uses a lot of film; (ii) sometimes probe becomes irreversibly stuck to filters even though they are being kept ostensibly wet throughout; and (iii) this procedure does not allow for optimal conditions of hybridisation.
In 5 X SSCP
10 X Denhardt's
250 µg/ml salmon sperm DNA
25% formamide at 42°C, background can be consistently as low as for high
stringency hybridisation; 20% formamide is generally also clean but in 15% formamide
at 42°C (and appropriate washing conditions) it is very hard to get clean filters.
Under the latter conditions, probes that recognise essentially unique sequences in the
genome will illuminate a smear of DNA on a genomic Southern that in sum exceeds
specific hybridisation.
Each 1% formamide lowers the Tm of a hetero- (or homo-) duplex by between 0.6 and 0.7°C (McConaughy et al., 1969; Meinboth and Wahl, 1983). Thus a range of 15-50% formamide at 42°C covers a Tm range of at least 21°C, the upper limit representing standard conditions for high stringency hybridisation. The Tm of imperfectly matched duplexes is apparently decreased by about 1°C per 1% in the number of mis-matched base pairs (Bonner et al., 1973).
The plaques containing DNA with the best homology to the probe will always give the strongest signals. However, this signal will decline (and background will increase) if the stringency becomes too low. Thus, it is best to screen at the highest stringency at which a signal can be clearly seen on a genomic Southern. The best general conditions for low stringency hybridisation are to use a single-stranded probe and to hybridise for 20-36 hr. Although on theoretical grounds it may seem best to substitute salmon sperm DNA with e.g. E.coli DNA when using probes derived from (for example) a mammalian source at low stringency, this has not been necessary in practice.
WASHING
After hybridisation, cut a corner off the bag, pour off the fluid into a 15/50 ml
plastic tube (to be used for the re-screen) and immediately cut the bag open and
immerse filters in wash solution.The conditions of washing have to be adjusted to the
hybridisation conditions, taking into account that hybridisation is optimal at 25°C
below Tm whereas hybrids, once formed, are stable under conditions much
closer to the Tm. Thus the following set of conditions may be used :
| Hybridization | Washing |
| % FORM | XXSCP | Th /°C | Tm /°C | Tm-Th -25/°C | Tm= %mis | XSSC | Tw /°C | Tm /°C | Tm-Tw -10°C |
| 0 | 2 | 65 | 91.3 | 1.3 | 2.6 | 0.2 | 65 | 72.8 | -2.2 |
| 0 | 2 | 65 | 91.3 | 1.3 | 2.6 | 0.5 | 65 | 80.1 | 5.1 |
| 0 | 5 | 65 | 98.6 | 8.6 | 17.2 | 2 | 65 | 91.3 | 16.3 |
| 40 | 5 | 42 | 69.8 | 2.8 | 5.6 | 1 | 65 | 85.7 | 10.7 |
| 30 | 5 | 42 | 77.0 | 10.0 | 20.0 | 0.2 | 42 | 72.8 | 20.8 |
| 25 | 5 | 42 | 80.6 | 13.6 | 27.2 | 0.5 | 42 | 80.1 | 28.1 |
| 20 | 5 | 42 | 84.2 | 17.2 | 34.4 | 2 | 42 | 91.3 | 39.3 |
| 15 | 5 | 42 | 87.8 | 20.8 | 41.6 | 5 | 42 | 98.6 | 46.6 |
| 0 | 5 | 42 | 98.6 | 31.6 | 63.2 | 5 | 20 | 98.6 | 68.6 |
Tm = 69.3 + 0.41(%G+C) Marmur & Doty (1962)
For most vertebrates and Drosophila the G-C content of the genome is about 40%, so that the average Tm at 1XSSC is 85.7°C.
(ii) The dependence of Tm on ionic strength, µ is given by
(Tm)µ2 - (Tm)µ1 = 18.5 log10 (µ2/µ1)
(iii) The Tm of duplexes is lowered by 0.72°C for every 1% of formamide in solution
(Bonner et al. 1973).
(iv) The optimal temperature for hybridisation is 25°C below Tm. Hence the fifth
column should be close to zero for perfect duplexes. It has been found (Bonner
et al. 1973) that optimal rates of reassociation for two imperfectly matched
strands occurs at a temperature that is lower than for a perfect duplex by
approximately half the difference between the two Tms. Thus conditions that
give a (Tm-Th-25)=n are ideal for a heteroduplex in which the Tm is 2n lower
than for a homoduplex (Tm= 2 X (Tm-Th-25)).
(v) The Tm of a heteroduplex that contains n% mis-matches is lowered roughly n°C relative to a homoduplex (Bonner et al. 1973).
(vi) The washing conditions given are such that the Tm of a perfect duplex is roughly 10°C higher than the washing temperature (Tw) and correspondingly higher for imperfectly matched heteroduplexes (columns 6 and 10 should be roughly the same)
(vii) All figures given are derived from solution hybridisation data. These conditions have been succesfully used in practice (actually, with wash conditions shifted up by one position on the table).
All washing solutions normally include 0.1-0.5% SDS
After the last wash, replace with a little fresh washing solution; take filters from this clean solution and wrap between sheets of Saran wrap while still moist. This allows subsequent washing at higher stringency or removal of probe prior to re-use of filters. Put phosphorescent markers on Saran wrap for alignment and expose.
Important Points:
1) Formamide used in hybridisations should be of good quality: freshly de-ionised
or stored frozen.
2) As it can be a relatively long time before the reason for a hybridisation signal is elucidated, it is worthwhile clarifying some of the possibilities in the initial screen by
(a) using a probe that does not include any vector sequence.
(b) including a genomic Southern in the hybridisation (particularly at low
stringency, so that the number of different DNAs being illuminated can be
judged).
(c) using two or more different probes either to circumvent possible vector
contamination or to define more precisely the nature of the hybridising DNA.
3) Background
The inclusion of a genomic Southern should help to distinguish the source of the
problem.
(a) Filters: Any agarose sticking to the filters while making lifts will give rise to
background. Batches of filters can vary but this has not often proved to be a
problem. It is also not clear that yellow filters (due to incomplete
neutralisation) lead to high background; there is a consensus that rinsing the
filters before baking is important.
(b) Probe: Nick-translated probes used at high radioactive concentration (106cpm/ml-do not exceed this) will always give some (uniform) background, but signals are also strong and very easily visible. The parameter most strongly favoured for adjustment to optimise signal/noise is the DNase I concentration used in nick-translation. Single-stranded probes in general do not give any significant background (when made under standard conditions); rather the only problem here is the strength of the signal. If the signal is very low (or zero) make sure the probe really was single-stranded.
(c) Hybridisation conditions: As the kinetics of the hybridisation are generally
not a problem it is best to use large hybridisation volumes to allow easy flow
over the (multiple) filters and to keep the concentration of counts below
106/ml. Also, use plenty of pre-hybridisation fluid as there is a large surface
area to coat and pre-hybridise for as long as possible, particularly at low
stringency.
PICKING POSITIVES & PLAQUE PURIFYING
plaque purifying;
1) Align autoradiographs with filters and mark the position of the syringe needle
holes on the film.
2) Mark positives with unique labels, so that after plaque purification it is possible to return to any plaques that did not check out as expected. To distinguish positives from background due to cassette contamination, cosmic rays and bubbles on the original plate the following criteria can be used :
(i) Shape: the source of radioactivity should be diffuse (as opposed to a point source) and should be roughly circular. Sometimes plaques will streak when lifts are taken giving a comet (a circular spot and a tail, not necessarily connected); this is a clear indication of a real positive. Duplicates help to identify the original position of the plaque but, irrespective of relative intensity, the circular region of a comet signal is the place to pick.
(ii) Intensity: Partial overlap between probe and recombinant phage DNA as well as variable plaque size means that wide variation in signal intensity is possible. Nonetheless, intensity is the best available guide to distinguish positives from background and also for low stringency hybridisation to distinguish between the quality of homology between the probe and various phage.
(iii) Clustering of positives is a clear indication of artifactual signals.
(iv) Number: Expected numbers of positives can be worked out for genomic libraries and, less reliably, for cDNA libraries. If there are an excessive number of positives, consider the possibilities that (i) the probe contains some repetitive sequences or (ii) the probe includes some "vector sequences" that are also present in the library (pUC, pBR, polylinker, ß-galactosidase sequences).
3) Align syringe needle marks on plates with their positions on the films and take a plug of agarose (down to the bottom of the plate) that overlies each positive. Because of the inaccuracy in lining up the orientation marks and because the filters generally contract a little during hybridisation and washing (if not pre-autoclaved) it is best to pick a small area around the positive such that you are confident that the positive must have been picked even given a maximal error in alignment. This can be done by using the wider end of a Pasteur pipette but is better done with multiple stabs of the other end to pick a (roughly) circular area of about 1/3-1/4 the size of the large end.
4) Elute the phage from the agar/agarose plug in 2ml of TM10 (+/-gelatin) with 1-2 drops of chloroform for at least 2 hr. Elution for less than 2 hr will lead to uneven titres. The titre of phage in the eluate is somewhat variable, depending on plating conditions, the number of lifts taken and (reproducibly) the type of bacteriophage lambda.
For the Maniatis library each plaque should yield a titre of 106-107 pfu. For cDNA libraries in gt10 the titre is 5-10 fold higher. The method of plaque picking described above removes 10-20 plaques from plates with 10-20,000 plaques. Thus, the expected number of infectious phage in the eluate will be in the range 1x107-1x108 (Maniatis) and 1x108-1x109 (most cDNA libraries). It should only be necessary to plate out about 100 phage to find several positives; however, because of variability in the growth rate of phage and the possibility that the required plaque was initially comparitively small on a crowded plate it is often worth plating out a large number (500-1000) to be certain of not losing a positive. In the latter case a large plate must be used. Thus, the following dilutions are recommended for plaque purification with an error margin that may lead to overcrowding rather than a loss of a positive signal. Dilutions are made in TM10.
Maniatis:
[Plug in 2ml] 1µlÆ[2ml] Plate 50-100µl Æ ~500 plaques
gt10 libraries:
[Plug in 2ml] 1µlÆ [2ml] Plate 5-10µl Æ ~500 plaques
Phage are plated out as for the initial plating but using (1/3) fewer bacteria and 3 ml top agarose if using small rather than large plates. Should the number of plaques be too great or too small return either to the original eluate or dilution, both of which should be of stable titre if kept at 4°C (or room temp. for a day). However, if the titre was initially lower than expected due to incomplete elution, it will now be higher than "expected" as elution will now be complete.
5) Make lifts, pre-hybridise, hybridise and wash as for the initial screen. The same probe can be used for initial and second screen but as most will have renatured during the first hybridisation it should be boiled for 5-10 min immediately prior to second use. (Precipitation of, presumably, BSA occurs if boiling is for too long but this does not affect the results whether additional hybridisation solution is added or not).
6) Signals from positives should be more numerous and stronger than in the initial screen it is therefore not worthwhile making duplicates for a re-screen if only one probe is being used. Positive plaques to be amplified should be well isolated in order to avoid contamination by neighbouring, diffusing phage. The potential problem of contamination is less serious for cDNAs where, in general, a single EcoRI insert will immediately be sub-cloned than for genomic lambda DNAs where mapping or use as an in situ hybridisation probe may be complicated by small amounts of contaminating phage.
7) Stocks of phage can be kept in TM10+gelatin+0.2% chloroform at 4°C or in 7% DMSO, TM10+gelatin at -70°C after snap-freezing. Such stocks can be made from single plaque eluates (106-107pfu/ml) or, more conveniently if they are to be used as a source for large scale growth of phage, as high titre eluates of plate lysates (1010pfu/ml).
ARRAYS
It is sometimes desirable to investigate the nature of a large number of
positively-hybridising phage by further hybridisation before preparing DNA. For this
purpose it is convenient to generate multiple copies of a set of positives. Put 100µl
aliquots (in duplicate if space permits) of each single plaque eluate in a well of a
micro-titre dish. Pour large plates with indicator bacteria (but no phage) in top
agarose. Transfer the phage array to such plates (after the top agarose has set) by
dipping the "clonemaster" (an array of metal prongs) in the micro-titre dish and then
resting it on the surface of the plate. Allow liquid to be absorbed and incubate at
37°C for >8 hr. Make lifts and screen as described above but expect much stronger
signals (and sometimes rings rather than spots; rings are usually observed using phage
in gt10 and can potentially be eliminated by dilution of the phage).
RE-USE OF FILTERS
If moist filters (stored between sheets of Saran wrap) and plates are stored at
4°C, the filters can be re-used several times for reasons of economy and in some cases,
such as chromosome walking, as part of the experimental design.
STRAINS AND MEDIA
C600 (F-, thi-1, htr-1, LeuB6, lacY1, tonA21, supE44, º)
Q358 (hsdR-R, hsdM+R, supF, ø80r)
Lambda plates NZCYM or LBM+ 1.5% agar
Top agarose NZCYM or LBM+ 0.7% agarose
NZCYM per litre 10 g NZamine
5 g NaCl
5 g yeast extract
1 g casamino acids
1.2 g MgSO4
LBM per litre 10 g Bacto-tryptone
5 g Bacto yeast extract
10 g NaCl
1.2 g MgSO4
REFERENCES
Bonner, T, Brenner, D.J., Neufeld, B.R. and Britten, R.J. (1973) Reduction in the rate of
DNA reassociation by sequence divergence
Frischauf, A.M., Lehrach, H., Poustka, A. and Murray, N. (1983) Lambda replacement vectors carrying polylinker sequences. J. Mol. Biol. 170 827-842
Maniatis, T., Hardison, R.C., Lacy, E., Lauer, J., O'Connell, C., Quon, D.,Sim, G.K. and Efstradiadis, A. (1978) The isolation of structural genes from libraries of eukaryotic DNA. Cell 15 687-701
Marmur, J. and Doty, P. (1962) Determination of the base composition of deoxyribonucleic acid from its thermal denaturation temperature. J.Mol.Biol. 5 109-118
Marmur, J. (1961) J.Mol.Biol. 3 208
McConaughy, R.L., Laird, C.D. and McCarthy, B.J.(1969) Nucleic acid reassociation in formamide. Biochem. 8 3289-3295
Meinkoth, J. and Wahl, G. (1984) Hybridization of nucleic acids immobilized on solid supports. Analyt.Bioch. 138 267-284
Pirrotta, V., Hadfield, C. and Pretorius, G.H.J. (1983) Microdissection and cloning of the white locus and 3B1-3C2 region of the Drosophila X chromosome. EMBO J. 2 927-934.