Fluorescein in situ hybridization for detection of chromosomal aberrations and aneuploidies caused by environmental toxins

Summary

Many chemical agents and ionic radiation of different nature can cause damage to chromosome integrity, resulting in abnormal chromosome distribution in mitosis and meiosis. Fluorescein in situ hybridization has been used to detect and quantify specific structural and numerical aberrations in mid-mitotic and interphase cells with a variety of probes complementary to different D N A sequences. Probes targeting specific chromosomal intact sequences have allowed the detection of stable alignment changes in mid-division cells that are closely associated with cell transformation and tumor development; probes that recognize sequences around the periphery of all the chromosomal mitotic grains of a species have been used to differentiate between micronuclei with and without mitotic grains, and, based on this, to further deduce whether they are caused by chromosome loss or breakage. In turn, probes that recognize sequences around the periphery of specific chromosomal mitophagy can be used to count interphase divisions by counting the

the number of chromosomes in the nucleus and discovering hyperploid cells that may have arisen due to chromosome nondisintegration. Ultimately, probes that hybridize to telomere sequences common to all chromosomes can be used to label telomeres and quantify their number and average length, cellular parameters that have recently been suggested to correlate with genetic instability. All of these techniques are based on the basic principles of synaptophysin hybridization , with some specific adjustments.

Written by Martin, this experiment is from "Environmental Genomics Lab Guide".

Operation method

Fluorescein in situ hybridization for detection of chromosomal aberrations and aneuploidies caused by environmental toxins

Move

I. Materials 1 Slide preparation and material fixation

(1) Clean slides.

(2) Methanol - Acetic acid solution (3 : 1)

2 Pre-treatment of material on the slide
3 Denaturation of targets and probes

(1) Denaturation solution (70% formamide-2X SSC): Prepare a 50 mL solution by adding 35 mL of deionized formamide (Kodak, Rochester, NY), 5 mL of evaporated water, and 5 mL of 20X SSC. Adjust p H to 7.0 with HCl.

(2) Hot plate.

4 In situ hybridization

(1) Probe.

(2) Wet box. A wet box can be prepared from a clean plastic box with a lid. Soak several paper towels in water and lay them on the bottom of the box. Cover and place the box in a 37°C incubator.

(3) Glass coverslips.

(4) Glass coverslip sealant (rubberized glue).

5 Rinsing after hybridization

(1) Rinsing solution: 50 % formamide (Fluka, Brooks, SG, Switzerland) in 2 X SSC. Prepare a 100 mL solution by adding 50 mL of formamide, 10 mL of 20 X SSC and 40 mL of distilled water. Adjust p H to 7.0.

(2) 4 X S S C + 0 . (2) 4 X SSC + 0.1 % Tween-20 solution: Add 2000 mL of 20X SSC and 0.5 mL of Tween-20 to a total volume of 500 mL of distilled water, mix well on a magnetic stirrer, and store at room temperature (RT) for up to one month.

6 Detection and Amplification
7 Re-dyeing
II. Methodology 1 Slide preparation and material fixation

Intermediate chromosome smears or interphase nuclear glass microscope slides are prepared according to standard cytogenetic procedures readily available in the laboratory (see Notes 2 and 3). More details on the preparation of intermediate and interphase cells for FISH can be found in references 18 and 19.

2 Slide aging

The purpose of aging is to fix the biological material to the glass surface and to make the chromosome structure resistant to the subsequent denaturation step of DNA. Too fresh a preparation, if not aged, can result in the loss of most nuclei, chromosomes or distortion of chromosome shape during denaturation. Aging too long reduces the efficiency of hybridization (see Note 4). Aging slides can be done by keeping the slides at room temperature for 2 to 3 days, or overnight at 65°C, or by heating at 90°C for 30 min.

3 Pre-treatment of material on slides (see Note 5)

(6) Place the slide in 1 % formaldehyde dissolved in P B S + M g C V solution and incubate for lOmin at room temperature.

(7) Rinse the slide with PBS for 5 min at room temperature.

(8) The slides are rinsed and dehydrated in a series of 70 %, 85 % and 100% cold ethanol solutions. All rinses are carried out in a staining vat, 2 mhi at a time, starting at a concentration of 70% (see Note 6).

(9) Air-dry the slides in air or under an air nozzle.

4 Denaturation of targets and probes

(1) Preheat the dish to 70°C.

(2) Prepare IOOjuL denaturing solution per slide in I.5m L microfuge tubes. Denaturing solution must be preheated to 70°C .

(3) Add IOOfX L of Denaturing Solution to each slide and cover with a glass coverslip (see Note 8).

(4) Denature the slides (not more than 3 slides at a time) in a dish at 70°C for 2 min (see Note 9).

(5) Immediately remove the coverslips by immersing the slides in a vat of 70 % cold ethanol and rinse for 2 m in. Remove the coverslips from the vat with forceps and repeat the rinsing with 85 % and 100 % cold ethanol solutions.

(6) Air-dry the slides in air or under an air nozzle.

(7) Preheat the probe at 37°C for 5 min (see Note 10).

(8) Gently vortex the mixture and centrifuge for 2~3s to collect the contents from the bottom of the tube.

(9) Use 30/^L commercial probe per slide or IOm L probe per 22m m X 22m m area (see Note 11). Place the probe in a 0.5 m L m i cr ○ fUge tube (see Note 12) and denature for 5 min at 70°C in a water bath.

(10) Immediately place the tube on ice until use.

5 In Situ Hybridization

(1) Each slide is covered with a 20m m X 50m m glass coverslip using a 30 ML probe (see Notes 8 and 11).

(2) Seal the coverslip with glass coverslip sealant around the perimeter of the coverslip.

(3) Incubate in a wet box at 37°C for 12-16 h. Incubate at 37°C for 2 hours.

6 Rinsing after hybridization

(1) Carefully remove the sealant with tweezers without removing the coverslip. The coverslip will come off during the rinsing process; use forceps to remove the coverslip from the staining vat. According to the probe data sheet (see Note 14), different probes require different levels of hybridization and rinsing conditions. Therefore, there are two post-hybridization rinsing protocols: a low stringency protocol and a high stringency protocol. Low stringency post-hybridization rinses (all solutions below should be preheated in a water bath)

(2) Rinse the slides three times in a glass staining vat containing 50 mL of rinsing solution at 37° C for 5 min each time with intermittent shaking.

(3) At 42°C, rinse the slides three times in 50mL of 2X SCC at pH 7 for 5 min each time with intermittent shaking.

(4) At room temperature, rinse the slides in a staining vat with 4 X S S C + 0.1 % T w e e n -20 solution. (4) Rinse the slide with 4 X SCC + 0.1 % Tween-20 solution in a staining bath at room temperature. Do not let the slides dry out (see Note 16).

Rinse after high stringency hybridization (preheat in a water bath for all the following solutions)

2). Rinse the slides three times at 42°C in a glass staining vat containing 50 mL of rinsing solution for 5 min each time with intermittent agitation.

3』. At 60°C, rinse the slides three times in 50 m L of 0. I X S S C with a p H of 7 for 5 m i n each time with intermittent shaking.

4. At room temperature, the slides were stained in a staining vat with 4 X SSC + 0 . 1 % Tween-20 solution in a staining bath. Do not let the slides dry out (see Note 16).

If the DNA probe is labeled with fluorescein-d U T P, re-stain immediately (see 4. 9). If the probe is labeled with hemi-antigen-d U T P, continue with the following steps (Probing).

7 Probing (see Note 12)

(1) Remove the slide from 4X SSC+ 0 . 1 % Tween-20 solution and quickly blot the excess liquid from the edges.

(2) Add IOOfxL of sealing solution (see Notes 17 and 18) to each slide, cover with a plastic coverslip, and incubate in a wet box at 37T: 3Omin.

(3) Carefully remove the coverslip with forceps and tilt the slide to allow the liquid to flow down briefly. The probe can be probed with a single antibody or with a conjugated antibody. Combined antibodies allow probes to be probed with two or more different half-antigen-d U T P labels.

(4) Add IOOuL of primary (one-time) antibody [fluorescein-labeled ovalbumin and/or fluorescein-mouse digoxin antibody anti-DIG] to each slide and re-cover with a plastic coverslip (see Notes 1 9 and 20). Incubate in a wet box at 37°C for 30-60 min.

(5) At 37°C, rinse the slides three times for 5 min each with 50 mL of 4 X SSC + 0.l% Tween-20 solution with intermittent shaking.

8 Magnification

(1) Add IOOuL of secondary antibody [biotinylated ovalbumin and/or fluorescein-mouse digoxin antibody (anti-DIG)] to each slide and re-cover with a plastic coverslip (see Notes 19 and 20). Incubate in a wet box at 37°C for 30 to 60 m i n .

(2) Incubate at 37°C in 50 m L of 4 X S S C + 0 . 1 % Tween-20 solution 3 times for 5 min each time, with intermittent shaking.

(3) Add IOO yL of tertiary (three times) antibody [fluorescein-labeled ovalbumin and/or fluorescein-labeled mouse digoxin antibody] to each slide and re-cover with a plastic cover slip (see Notes 19 and 20). Incubate in a wet box at 37°C for 30 to 60 min.

(4) Rinse the slides three times at 37°C in 50 mL of 4 X SSC +0, 1 % Tween-20 solution for 5 min each time with agitation (see Note 21).

9 Restaining

Propidium iodide

(1) Add 30 juLPI/anti-melatonin (2fig;mL) to each slide and cover with a glass coverslip.

(2) Seal the edges of the coverslips with nail polish (see Note 22). Store at 4° C for several months to maintain good signal.

Use D A P I

(1) Incubate slides for lOmin in staining solution.

(2) Rinse slides in 2X SSC for 10 min to elute residual dye.

(3) Rinse the slide in distilled water for a few seconds to remove salt stains.

(4) Air-dry the slide at room temperature and add 30uL of Anti-fade Solution.

(5) Cover the glass coverslip and seal the edge of the slide with nail polish (see Note 22). Store at 4°C for several months

signal well.

1 0 Slide analysis

1 0.1 Chromosome coloring

Chromosome mapping (Fig. 2a, see plate) was developed primarily for the purpose of biological radioactivity determination to detect stable exchange-type aberrations such as reciprocal translocations (5). Indeed, the analysis of double-stranded chromosomes (nonstable aberrations) on solid-stained chromosome preparations is very reliable for the assessment of recent acute radiation exposures, but is not applicable to the assessment of chronic or past exposures because of the decrease in the production of double-stranded chromosomes with time after radiation (20).
For the proper analysis of mid-mitotic aberrations in color painting (and solid staining as well), the discovery of mitotic grains may

may be crucial, especially in the case of mouse proximal mitotic chromosomes: either by double-staining with DAPI, which itself produces a bright mitotic heterochromatin signal after alkaline "thermal denaturation", or by staining with mitotic F LSH.

At the mid-mitotic stage of solid staining, conventional nomenclature classifies each simple translocation as a separate event (e.g., a double-stranded plus an anaphase fragment) and distinguishes between complete (if no nonrecombining breaks are detected) and incomplete exchanges (e.g., a double-stranded chromosome without an anaphase fragment). The application of coloration techniques has led to the creation of new nomenclature systems. These new nomenclature systems are mainly based on the identification of color switches between fluorescein-binding probes and repainting dyes. According to the draft terminology for aberration identification and nomenclature (P A I N T ) (21) each non-normally stained chromosome or fragment is described by the letters 'A' or 'a', respectively, to indicate the chromosomal material of the diplotene dye. The letters 'B' and 'b' are used to indicate the colored material, with upper case letters indicating regions containing a mitophore and lower case letters indicating regions without a mitophore. Thus, a typical P A I N T classification of aberrations is t (A b ), die (A B ), or ace (ab), where t stands for translocation, die for double-attachment, and ace for an anaphase fragment. Another independently developed nomenclature system
Another independently developed system of nomenclature is the Savage and Simpson (S & S) system (22 ), which chooses to interpret aberrations in terms of the mechanics of their formation.

When the FISH technique is used for the analysis of aberrations caused by chemical mutagens (6, 23 ) - a new type of aberration not seen in radiation studies - it must be taken into account in the correct determination of the effects caused by chemical mutagens: i.e., the presence of one or more extra holochromosomes in the mid-stage cell, accompanied by a species-specific number of haploid chromosomes. These extra chromosomes originate from chromosome haplotype exchanges in the peripheral region of the mitophagy and appear only in irradiated G2 phase cells. Unless all chromosomes are hybridized with probes that specifically bind different fluorescent dyes to them, and a computerized image analysis system is used to assign a different color to each of them, usually only a portion of the entire karyotype is stained, and aberrations involving stained chromosomes thus represent only a portion of the total number of resulting aberrations. It may be important to predict the fate of exposed cells or individuals by assessing the full number of aberrations. Assuming that aberrations are randomly distributed across chromosomes, in order to convert the observed number of aberrations into a ratio of the total number of aberrations to the total genome, it is necessary to know the proportion of the total genome that is covered by each color probe (or cocktail of probes). If the household represents the proportion of the genome that is stained by a given color, 9 is the proportion of the genome that is stained by a second color, and r is the fraction of the genome that is stained, the fraction of detectable exchanges is computed as S = 2p 9+ 2 p r+ 2 gr. Multiplying the number of measured intermediate phases by S yields the average value in each cell (genome). The ratio of the total number of observed aberrations to the mean in each cell gives the estimated frequency of aberrations/genome. Similarly, it is assumed that TZ is when the whole genome can be used to
When the whole genome is available for aberration scanning, the number of measurable midphases is known, and n/S denotes the proportion of colored midphases to the total number of measurable midphases, which is equivalent to the information that can be provided by the n all-analyzed midphases (24).

IO.2 F ISH Staining of the Ropes

Mitotic F I S H staining allows the detection of potential aneuploidy mutagens by measuring the frequency of chromosome loss and chromosome nondisjunction.

To assess chromosome loss, probes specifically targeting all chromosomal mitotic sequences were used to detect the presence of mitophagy in micronuclei. Probe-labeled micronuclei were mitosis-positive (C + ) and were assumed to contain one or more chromosomes attributed to chromosome loss during the previous late mitotic phase. To assess chromosome nondisjunction, probes for detecting the mitotic (peripheral) regions of specific chromosomes were used to rate the number of chromosomes present in each sister interphase nucleus in a binucleate cell. Songocystin B-blocked cells were used in the experiments, thus retaining the two sister nuclei of a single mitotic division in the same cytoplasm. In an aneuploid nucleus, Xf any autosomes, there should be two spots for each probe. Undisjunction, which occurred in the last mitosis, can be detected by probing for signals of the 2 + 2 form that should be present in the sister nuclei, such as the appearance of 3 + 1 or 4 + 0 combinations; nondisjunction of the X and Y chromosomes in the male cell produces signals of the 2 + 0 form (Fig. 2b, see plate). Only cells with the correct total number of hybridization signals and similar fluorescence intensities were included in the analysis. It has been shown that chromosome nondisjunction may be a more sensitive indicator of aneuploidy activity than chromosome loss (25). This suggests that molecular cytogenetic analysis of interphase cells is of great importance as a rapid and reliable method of assessing spontaneous and induced chromosome nondisjunction.

10.3 Quantitative F I S H for telomere length estimation

1 0.3.1 Quantitative image analysis

Telomeres protect the ends of chromosomes from end-to-end fusion. Shortening of telomere length beyond a critical length drives cells into replicative aging or is the cause of chromosomal instability. Therefore, telomere length measurement is an effective way to assess cellular longevity or to study chromosomal instability caused by physical or chemical factors. Recently, a new in situ technique, quantitative fluorescence in situ hybridization (Q-HSH ), has been developed. This technique, which can be applied to mid-mitotic and interphase nuclei, is based on the use of a peptide nucleic acid (P N A ) telomeric oligonucleic acid probe, which is comparable to the standard D N A oligonucleic acid.
probe that produces stronger and more specific signals than the standard D N A oligonucleic acid probe (17). Currently, Q -F IS H is capable of detecting individual length differences in telomeres smaller than Ikb ( 2 6 ) . It has been hypothesized that telomere length is directly correlated with its integrated fluorescence intensity value (IFI) because the fluorescent probes used may quantitatively hybridize to telomeric repeats, allowing detection of small telomere length differences.

Digital images are recorded by a camera on a fluorescence microscope equipped with a multi-filter wheel. Image acquisition must be accomplished using specialized software. Digital images are obtained from Dahl-stained chromosomes and C y3-stained telomeres (Fig. 2c, see plate). A special computer program (TFL-Telo program) was developed for image analysis (27). Briefly, chromosomes and telomeres are identified by segmentation of the DAPI image and the C y 3 image, respectively, and the two images are combined for pixel shift correction. To avoid possible selection bias, the interim phase was selected only on well-prepared chromosome smears.

The T F L -Telo program allows contour lines to be made from separate chromosomes and separate telomere spots, and the two images can also be successfully combined. Ultimately, the fluorescence intensity of telomeres on each chromosome can be expressed as an arbitrary unit of fluorescence (a.u. f. )

10.3.2 Calibration

To ensure a reliable quantitative assessment of telomere length in different samples, two levels of calibration must be applied. First, to correct for differences in daytime light intensity and alignment, images of fluorescent beads (orange beads, size ○.2 Mm; MolecularProbes, Eugene, O R) were obtained and analyzed by a computer program in parallel with the cell samples. Second, to find the relationship between fluorescence intensity and the number of T 2A G 3 repeats, only images with 10 and 80 kb (L Y -S and L Y -S, respectively) were analyzed.
R lymphoma cell lines, respectively)(2 8 ) the length of the defined (T 2A G 3)n could be hybridized and analyzed. Based on the FI value and the slope of the linear relationship between the telomere lengths of L Y -S and L Y -R cells, the telomere lengths of cell samples can be assessed after obtaining their IFl values under the same conditions. In addition to cell lines with defined telomere lengths, plasmids with inserted telomere sequences of known size can optionally be used (27).

1 1 Microscopy Filter Settings

A very important part of FISH is the visualization and recording of results using a fluorescence microscope and camera system. A fluorescence microscope consists of a light source that excites a fluorescent dye and a special filter that transmits the light emitted by the fluorescent dye to a high degree. The filter allows light of a specific wavelength to pass through while blocking other light. Therefore, filters are individually designed for specific fluorescents and must be properly selected. Propidium iodide or D A P I filters should be used for scanning cells or mid-phase smears, while filters specific for probe-bound fluorescein are used for target-of-interest imaging. Table 2 gives general guidelines for the selection of filters from microscope manufacturers.

Caveat

(1) Plastic coverslips can be prepared by cutting a piece of parafihn membrane. They are easier to remove from the slide than glass coverslips, so parafilm membranes should be used in preference to plastic, except for those steps that require high temperatures or where sealing equipment is not suitable for plastic.(2) To prepare interim chromosome smears, cultured cells are treated with colchicine amide (lO^g/m L Invitrogen, Paisley, UK) for 3 h. This protocol is applicable to many cell cultures. This protocol is suitable for many cell cultures. For cells with a short cell cycle, e.g. mouse cells, 2 h of colchicine treatment is sufficient. For slow-growing cells, colchicine should be used from 3 h to overnight. The concentration of colchicine should be adapted to the duration of treatment: longer treatments require lower concentrations of colchicine. To obtain the mid-phase in the in vivo assay, use ○-S m L KT 3m 0I/L colchicine per mouse. For the preparation of binucleated cells, human pinocytin B was added to the cells.(3/i g Z m L, Sigma-Aldrich) was added to the culture medium for one "t-cell cycle.(3) This protocol is recommended for cultured cells and cell suspensions obtained from experimental animals or humans, fixed with methanol-acetic acid (3 : 1 ).(4) The conditions under which the slides are aged are always related to the probe used. Therefore, we recommend that slide aging be performed according to the probe data instructions.(5) Many incubations are performed in 50m L glass staining vats. For best results, check the solution temperature by placing a clean thermometer directly into the vat.(6) Store the ethanol solution at 1 20°C before use.(7) When processing multiple slides at the same time, each slide will result in a 1°C decrease in the temperature of the hot plate. Therefore, the temperature of the hot plate must be set i°C higher for each slide to be denatured.(8) Care should be taken to avoid the creation of air bubbles.(9) Time and temperature are important to maintain the morphology of chromosomes and nuclei. Incomplete denaturation of the target D N A leads to loss of signal. The recommended protocol has been shown to be effective for both intermediate and interphase human cells, with higher denaturation temperatures of up to 80°C required for mouse cells.(10) The probes are usually diluted with glycerol or formamide and the solutions are very viscous. We therefore recommend that commercial probes be preheated at 37°C to reduce the viscosity of the solution.(11) For economy, only target areas on slides containing good quality and quantity of intermediate smear or interphase nuclei are hybridized. Selected areas can be marked on the back with a diamond scribe.(12) We recommend wrapping tubes containing fluorescein-labeled probes or fluorescein-labeled antibodies in aluminum foil to protect them from light.(13) If two or more probes are to be used at the same time in the same hybridization, the final volume may exceed 10 to 12 ML for a standard 22 mmX22 mm coverslip. To solve this problem, centrifuge the total volume of the mixed probes at 840(^ for IOmin, discard the supernatant, and air-dry the precipitate under an air nozzle or centrifuge under vacuum at speed with 10 to 12; xL Hybridization Buffer (Hybrisolol). The precipitate was dried under an air nozzle or by speed vacuum centrifugation, resuspended in 10 to 12;xL hybridization buffer (Hybrisol YI, Roche), and the mixture was added to the slide.(14) Temperature, time, and buffer concentration of the hybridization and post-hybridization rinse solutions are important; low stringency results in nonspecific binding of the probe to other sequences, whereas high stringency results in signal loss.(15) This step was initially used to remove nonspecific and/or repetitive DNA from hybridization to cells and chromosomes. there is no standard procedure to refer to, but there are some guidelines that can be helpful. The lower the concentration of the salt solution, the longer the rinse, the higher the temperature, and the greater the stringency, the more unbound or nonspecifically bound probes will be removed. For very short DNA probes (0.5-3 kb) or very complex probes (chromosome painting probes), the rinsing temperature should be lower (up to 45.) and the stringency lower (1-2X SSC). For repetitive probes (e.g. cr satellite repeats), the temperature and stringency should be the highest.(16) If desired, slides may be stored at 4°C in 50 m L of 4 X SSC + 0.1 % Tween-20 solution for up to 2 weeks prior to probing.(17) Unless otherwise specified, the volume of the probing solution is for the entire 22 mmX50 mm slide area. If only an area of 22 mmX22 mm is hybridized, reduce the volume to 1/3.(18) BSA blocks the nonspecific binding of some antibodies to glass, which is important for reducing background, but may also partially block antibody contact with labeled D N A .(19) For purchased fluorescently labeled antibodies, the antibody concentration for in situ hybridization should follow the instructions in the operation manual. As a general guideline, a 1:100 dilution of the 1m g/m L antibody master mix into the probe solution and incubation at 37°C for 30 min should be fine. Each time a new antibody is purchased, it should be tested at different dilutions to find the optimal range of action. As a rule of thumb, we have found that the following antibodies work well for in situ hybridization at the following concentrations:a- For biotin-labeled probes: 1 : 300 for fluorescent ovalbumin D C S (VectorLaboratories) and rhodamine ovalbumin D C S (VectorLaboratories) in the probe solution. 1 : 100 for biotinylated anti-ovalbumin D (Vector Laboratories) in the probe solution.b - For DIG-labeled probes - 1 : 100 to 1 : 200 for mouse anti-DIG (Roche) in the probe solution. 1 : 100 for the DIG antibody (Roche). Anti-DIG rhodamine and anti-DIG fluorescein in the probe solution (Roche) is 1 : 50.(20) If combining antibodies is used, prepare a double concentration of antibody solutions by mixing 5 CVL of each solution on a slide.(21) Rinsing can be done at 37°C or 42°C. The optimal temperature should be determined empirically. The optimal temperature should be determined empirically to remove excess and unboundProbe reactants that are excess and unbound should be empirically determined.(22) Nail polish may improve the shelf life of FISH slides.＜

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