ALTERNATIVAS DE EXPANSION URBANA

The technique is relatively simple: DNA or RNA probes are first labelled with reporter molecules. The probe and the target chromosomes or nuclei are denatured. Complementary sequences in the probe and target are then allowed to reanneal. After washing and incubation in fluorescently labelled affinity reagents, a discrete fluorescent signal is visible at the site o f probe hybridisation.

3.1.3.1 C hrom atin preparation and dénaturation

Cells are hypotonically swollen and fixed on slides by standard methanol:acetic acid fixation, which provides excellent preservation of cell and chromosome morphology without altering the target nucleic acid, whilst still allowing access and hybridisation of the probe. Slides are then treated with RNAase to remove endogenous RNA which might act as a competitor for the labelled probe and result in an increased unspecific background signal. Careful washing after the RNAase treatment ensures that no enzyme remains to degrade the hybridisation probe.

Base, acid or high temperatures can be used to denature the chromosomes. The form er two methods tend to be less efficient, but give good results if the DNA sequences of interest are repeated (e.g. ribosomal DNAs, satellite DNAs). Highly efficient dénaturation methods are needed for unique sequence localisation; in such cases slides are incubated briefly at 70°C in a solution containing 70% formamide to dissociate the DNA into single strands, and then fixed by dehydration in cold ethanol to quench the dénaturation reaction and thereby reduce strand reannealing before the addition of probe. Such heat treatment followed by ethanol dehydration also reduces the loss of morphology that occurs when part of a chromosome does not adhere tightly to the slide. Formamide alters the dielectric constant of a solution and thereby lowers the melting point of DNA approximately 0.7°C per % of formamide (McConnaughy et al 1969). This decrease in the temperature required to melt the chromosomal DNA helps to maintain good chromosome cytology. After dénaturation, slides should be used for hybridisation within no more than 12 hours, because the efficiency of hybridisation decreases rapidly as a function of storage time after dénaturation.

3.1.3.2 Probe labelling

A wide variety of nucleic acid probes, probe labelling reactions, and labels have been used for FISH. Total genomic DNA can be used as a species-specific label, as the repetitive sequences it contains show little evolutionary conservation and it is these sequences that reanneal more rapidly than the more highly conserved unique sequences because of their high frequency in probe and target. Chromosome-specific repeats can be used to tag particular chromosomes and are described in more detail in section 3.2.1.1. More recently, chromosome ‘painting’ probes have been developed (reviewed in Lichter et al 1991). These consist of collections of DNA sequences derived from a single human chromosome (e.g. from somatic cell hybrids carrying the desired chromosome as its only human material). Prehybridisation with unlabelled genomic DNA suppresses hybridisation of repetitive elements in the collection that are common to many chromosome types, so that only the chromosome of interest is highlighted. These probes have proved invaluable for the detection of cytogenetic abnormalities. Single-copy sequences can be mapped using FISH, although the efficiency of hybridisation site detection decreases with decreasing probe size. The efficiency of labelling large insert probes (such as cosmids or YACs) is > 90% under suppression conditions.

Although probes may be directly conjugated with fluorescent molecules (Koch et al 1992), the signals obtained are relatively weak, limiting the technique to the detection of repeated DNA sequences or abundant mRNA species. Alternative labelling techniques have been developed in an effort to produce more sensitive or convenient technology. The most widespread approach is to label probes with reporter molecules which, after hybridisation, bind fluorescent affinity reagents. Typical reporter molecules include biotin, digoxygenin, dinitrophenyl (DNP), aminoacetylfluorene (AAF), mercury and sulphonate (reviewed in Bauman et al 1990). For the first three of these reporter molecules, enzymatic incorporation of nucleotides modified with biotin, digoxygenin or DNP is usually preferred over chemical labelling techniques employing photoreactive compounds (photobiotin, photodigoxygenin, or photoDNP) because of a higher labelling efficiency (Lichter et al 1991). However, chemical modification schemes using the other reporter molecules AAF, mercury or sulphonate, have been used successfully for sensitive nonradioactive detection of hybridised nucleic acids (Landegent et al 1987). Alternatively, probes can be labelled by PCR amplification between known priming sequences (primer-induced sequence-specific labelling or PRINS, Koch et al 1992) or by RNA transcription from appropriate vectors, in the presence of labelled nucleotides.

Biotin and digoxigenin labelling combined with fluorescent detection are currently the most widely used because of the high sensitivity of detection and the commercial availability of the reagents. Typically employed are dUTP or UTP analogs that contain a biotin molecule covalently attached to the C-5 position of the pyrimidine ring through an allylamine linker arm (Langer et al 1981). Such modified nucleotides can function as suitable substrates for DNA or RNA polymerases in vitro, and can thus be introduced into DNA or RNA molecules

to form efficient labelled probes without significantly altering their hybridisation characteristics. Both biotin- and digoxygenin-labelled probes are also stable for years.

The size of the probe is critical for in situ hybridisation and should be between 100 and 500 nucleotides, a size that maximises specific hybridisation as it is ideal for the formation of the probe networks required to efficiently detect unique sequences, whilst decreasing background hybridisation (Lawrence and Singer 1985). The most convenient method for introducing label into a double-stranded probe is nick translation (Rigby et al 1977), because probes of the desired size range can be obtained readily by adjusting the DNAase concentration. DNA probes can also be prepared using short random DNA primers and the Klenow fragment of E. Coli DNA pol (Feinberg and Vogelstein 1983). However, this tends to yield probes of a shorter length, limiting the formation of probe networks (Kamakari 1994) although suitable for the detection of tandemly repeated probes. In the case of probes labelled by chemical modification, sonication can be used to achieve fragments of the required size.

3.1.3.3 H ybridisation

Labelled probes are mixed in a hybridisation buffer containing formamide, salt, dextran sulphate and unlabelled carrier DNA (e.g salmon sperm DNA) or E.Coli t-RNA. Early studies used high salt buffers (5xSSC) and elevated temperatures (60-68°C) to promote accurate base- pairing and therefore efficient hybridisation. Such conditions often adversely affected the cytology, making chromosome identification difficult after hybridisation. The presence of formamide and moderate salt concentrations in the hybridisation mixture lowers the temperature required for accurate nucleic acid annealing, permitting hybridisation at temperatures of 37-45®C (McConnaughy et al 1969) which helps to preserve chromosome m orphology.

Dextran sulphate increases the rate of reassociation of DNA as described in section 3.1.1, whilst unlabelled carrier DNA or E. Coli tRNA are included to reduce non-specific binding of the probe to chromatin and glass.

For repetitive probes, the mixture is denatured and applied directly to slides after chromosome dénaturation, or is directly applied to slides and denatured simultaneously with the target DNA. For unique sequence contained in large-insert probes, the probe is preannealed with excess unlabelled genomic or Cotl DNA to reduce repetitive sequence binding to the target before applying to the slides. Hybridisation between target and probe occurs during overnight incubation (16-18 hrs) of the slides at 37°C.

3.1.3.4 P ost-hybridisation w ashes

After hybridisation, excess unhybridised probe must be removed by extensive washing of the slides. In the case of DNA-DNA hybrids, non-specifically bound DNA probe can be removed by washes in salt solutions of varying ionic strengths, or by incubation in low salt at high temperatures. Temperatures of between 55-70°C are well below the melting point of 50% GC content DNA and will ensure fidelity of the hybrids of interest whilst providing a

stringency sufficient to remove non-specifically bound probes. However, incubations at high temperatures for prolonged periods destroys the architecture of the chromosomes. Standard post-hybridisation washes are thus performed with 50% formamide in 2xSSC followed by 2xSSC at a temperature 5-7°C higher than the hybridisation temperature. The number and duration of washes may vary according to the probe size.

3.1.3.5 Fluorescent detection

Hybridised probes are detected by incubating the slides in immunofluorescent reagents to produce a fluorescent signal at the hybridisation site. The extraordinary affinity of biotin for the glycoprotein avidin (Langer et al 1981) is frequently exploited for the detection of biotin-labelled probes by using fluorochrome-conjugated avidin. Anti-biotin antibodies can also be used. To overcome the potential problems associated with the endogenous presence of biotin in cytological material which may cause ‘autofluorescence’ and contribute to undesirable background signals, detection of biotin-labelled probes is performed in a ‘blocking buffer’ containing high salt concentration (typically 4 x SSC) which reduces the non-specific binding of biotin affinity reagents (Lawrence et al 1990). Probes labelled with digoxygenin, DNP, AAF or sulphonate are detected with specific antibodies followed by fluorescently labelled anti-immunoglobulins.

A variety of fluorescent labels are available for use in FISH, differing in their absorption and emission spectra, that is, the range of light wavelengths required for excitation and those re-emitted by that substance. The most commonly used fluorochromes are fluorescein isothyocyanate (FITC) which is excited by blue light of 495nm and emits in the green-yellow region at 515nm, rhodamine and Texas Red which are excited at 550nm and 595nm and both emit in the red region at 575nm and 615nm, respectively, and 7-amino-4-methyl coum arin-3- acetic acid (AMCA) which is excited by UV light (350nm) and emits in the blue region at 450nm. The sensitivity of the fluorescence detection can be increased using multiple layers o f affinity reagents or antibodies e.g. for biotinylated probes, avidin may be used for primary detection, followed by alternating layers of anti-avidin antibody and avidin (sandwich amplification).

3.1.3.6 C hrom osom e counterstaining and banding

The most commonly used fluorescent dye for chromosome staining when banding is not required is propidium iodide which excites at 340nm and emits in the red region at 600- 610nm. Chromosomes can be banded after hybridisation, and although the bands are crude they are sufficient to identify chromosomes and regionally localise probes. The assignment o f a sequence to an individual band may also provide information on the general functional significance of that region since light G-bands are early replicating, rich in CpG islands, contain many housekeeping genes, and large numbers of ALU repeats, whilst dark G-bands are late replicating with fewer but tissue-specific genes and more LI repeats (Bickmore and Sumner 1989). Banding methods that can be applied in parallel to fluorescent probe detection

and allow simultaneous visualisation of bands and fluorescent signal include the use of actinomycin/diamidinophenylindole (DAPI; emission in the blue region) staining to highlight G-bands (where band contrast may be improved by growth in 5-bromodeoxyuridine (BrdU), Hoescht staining and UV irradiation for R-bands, or ALU or LINE repetitive sequences can be added to the hybridisation mix to depict R- or G-bands respectively. Quinacrine (emits in the yellow region) can also be used to generate a Q-banding pattern, as can DAPI used alone.

3.1.3.7 Signal visu alisation

Most conventional fluorescence microscopes are suitable for viewing the signals from painted whole chromosomes, stained subchromosomal regions or localised repetitive or single­ copy probes, provided that they are fitted with appropriate objectives and filter sets for visualisation of the relevant fluorochromes. Excitation filters select the optimum wavelength o f light to excite the fluorochrome, whilst barrier filters both suppress the excess exciting light and select out the emission wavelength of the fluorochrome to render it visible.

A filter set can be chosen specifically for a given fluorochrome (single band-pass filter sets), or for the simultaneous detection of two or more fluorochromes having different wavelength specificities, such as FITC for signal detection and propidium iodide for chromosome staining (double band-pass filter sets), or similarly for viewing three distinct fluorochromes (triple band-pass filter sets). Such multiple band-pass filter sets have greatly simplified multiple colour FISH mapping experiments by minimising the image shift that can occur when filters are changed. The consequent displaced registration of images detected on multiple exposure of photographic films can introduce errors where analysis involves fine measurements or localisation of probes that map very close to one another.

Images of hybridised cells can be collected with high resolution on high-speed colour film or into computer memory via a static CCD (charge coupled device) camera or a confocal laser scanning microscope equipped with a photomultiplier. CCD cameras and confocal laser scanning microscopes allow the generation of digitised images and thus facilitate the collection and analysis of hybridisation data by enhancing contrast and improving the signal-to-noise ratio by averaging images to allow detection of small signals. The former system is the most sensitive to date, by virtue of its high efficiency in counting emitted photons over a broad spectral range of wavelengths, enabling the recording of signals which are not visible to the observer’s eye. The second system is especially suitable for three-dimensional microscopy, where a series of optical sections through a labelled specimen is obtained. 3D reconstruction is accomplished by applying appropriate computer software to a stack of digitised images. Laser scanning confocal microscopy is designed to eliminate most of the out-of-focus fluorescence obtained in each section by conventional image devices. The generation of high quality optical sections, greatly reduces the amount of mathematical operations required to reconstruct the object.