Fluorescence in situ hybridization for the detection of transgene fragments integrated in plant genomes experiments
Over the past 30 years, fluorescent chromosome analysis techniques have facilitated the understanding of genome construction and deepened knowledge of the organization of DNA and chromosome evolution. Fluorescent staining allows the identification of chromosomal aneuploidy and polyploidy in species, selections and monocultures, including chromosomal variation resulting from hybridization or histoculture and genetic transformation. The source of this experiment is "Experimental Guide to Transgenic Technology and Field Identification of Wheat Crops" [English] H.D. Jones P.R . Hewley, eds.
Operation method
Detection of transgenic fragments integrated in plant genomes by fluorescence in situ hybridization
Materials and Instruments
Nucleotides Move The basic method of in situ hybridization (Fig. 14.2 ) has evolved from Southern hybridization: the substrate is the chromosome and nucleus on the slide, and the probe is a labeled sequence of the DNA to be tested [i.e., in this book, the transgene and the control, Fig. 14.3(a)]. The probes and chromosomes are denatured separately or simultaneously to produce hybridized molecules (Fig. 14.2). Most experimental protocols take hybridization overnight, which is important for low-copy FISH. For detection of repetitive DNA sequences, a hybridization time of a few hours is sufficient. As with Southern hybridization, several washes are required to remove unbound probes and to characterize the tightness of the hybridization, i.e., to characterize the similarity between the probe and the target DNA sequence, which is necessary to maintain stable hybridization in a double-stranded DNA helix. To prepare the chromosome, a double helix destabilizer such as formamide can also modulate hybridization tightness, in addition to conditions such as hybridization temperature, sodium ion concentration, and washing solution. In direct FISH, the chromosomes can be restained immediately, and DAPI restaining is commonly used; in non-direct FISH using fluorescein and digitonin labeling, immunocytochemistry is required for detection. For more product details, please visit Aladdin Scientific website.
Ice water 8-hydroxyquinoline colchicine Fixative Enzymatic buffer
Slides Glass coverslips Dissecting needles and forceps
Chromosomes are fluorescently stained [ Fig. 14.1 (a ) ], and the ploidy level of transgenic strains and individuals is detected according to the chromosome fixation and preparation procedure (see Sections 3.1 and 3.2), followed by restaining, sealing, and analyzing steps (see Sections 3.7 and 3.8).
1. Material fixation
Preparation of mid-division chromosomes containing a large number of clean and spread out chromosomes is an important factor in ensuring the success of FISH and chromosome staining. Plant material used for fixation and chromosome preparation must be healthy, disease-free and actively growing. It can be any meristematic tissue, but what material is used for the assay during the growth of the transgenic plant is determined by the timing of the experiment, which can be the root tip of a seedling, the tip of a newly grown root on an old plant at the edge of a pot, or the tip of a hydroponically grown plant. For some species, cells of the meristematic tissue of shoots, leaves or new shoots may also be used.
In most cases, obtaining the maximum number of chromosomes in mid-division is important for the success of the experiment. In order to obtain more cells in mid-division, it is often necessary to pre-treat the material so that chromosome condensation occurs and spindle microtubules are disrupted to obtain sufficiently dispersed chromosomes. Spindle microtubule inhibitors such as colchicine (widely used for chromosome counting) can create highly condensed chromosomes, while other chromosome condensers, such as ice water or 8-hydroxyquinoline, produce more stretched chromosomes that are more suitable for FISH analysis.
( 1 ) All steps are performed in a clean container using clean forceps. Avoid exposing the material to fixatives and chemicals during the collection and pretreatment phases (steps 2~4) (see Note 11).
( 2 ) Select suitable plant material to obtain dividing cells. Confirm that the material is grown under suitable conditions. 
( 3 ) Select a cell division blocker to add to the test tube.
( 4 ) Transfer roots (1~2 cm long) or shoots to 3~5 ml of division blocker (see Note 12), using 5 times as much reagent as the material, and then tighten the cap.
( 5 ) Incubate.
( a ) Ice water: 24 h
( b ) 8-hydrocarbon-based quinoline: 30~45 min at the temperature required for growth, then 4°C, 30~45 min.
( c ) Colchicine: 2~4 h at room temperature or 12~16 h at 4°C.
( 6 ) Absorb the material quickly and place it in fresh fixative. To ensure rapid penetration, make sure the fixative is not contaminated with moisture. In addition to marking the tubes with an (inked) felt pen, use a pencil to make a good mark on a post-it note or a small piece of paper and place the piece of paper into the fixative together.
( 7 ) Leave at room temperature for 1 h, then store at 4°C or -20°C (see Note 13 ).
3.2 Chromosome Preparation
Chromosome preparations for both anaphase and interphase, either by fluorescence staining or FISH, are performed on carrier sheets.
It is recommended to pretreat the material with protein hydrolase to remove cell walls and cytoplasm before placing the samples on slides, dropping acetic acid, and then covering the slides for pressing. All operations were performed at room temperature unless otherwise indicated. Small petri dishes were used for washing and material preparation as described in section 2. 2.
( 1 ) Wash the fixative on the plant material twice with enzyme buffer for 10 min each time.
( 2 ) Transfer plant material to 1 to 2 ml of enzymatic solution and incubate at 37°C for 30 min to 2 h, depending on the material (see Note 2).
( 3 ) Transfer the plant material from the enzyme digest to the enzyme buffer.
( 4 ) Transfer 2 or 3 portions of material to 0.5 ml of 45% acetic acid and leave for a few minutes. This step can be omitted if the material is too soft.
( 5 ) Soak the slide in 96% ethanol and dry it with a lint-free cloth. Place a drop of 60% acetic acid on the slide.
Then place a small piece of root tip or other material into the acetic acid. Replace the acetic acid as needed.
( 6 ) Under a dissecting microscope, tear the tissue or tap the tissue with a glass rod and remove the particulate matter. Cells in the dividing phase are more likely to float.
( 7 ) Wipe the coverslip with hand paper and place it on a slide for phase contrast microscopy. To obtain the optimal slice, press the slide with your hand (thumb), applying as much or as little pressure as necessary. The coverslip may also be tapped.
( 8 ) Observe cell density and mid-division index under the microscope. Place a good sample on a metal plate in dry ice for 5-10 min ( not too long ). The coverslips were gently picked apart with the tip of a stiletto knife and air-dried.
( 9 ) Browse the unsealed slides under a phase contrast microscope. Look for well-dispersed nuclei that are not aggregated in piles, and the slide should be free of surface membranes and impurities. Mid-disintegration chromosomes should be well dispersed and free of cytoplasmic and impurity debris, etc. Record each slide to optimize the next preparation. Small chromosomes can be detected by DAPI staining (see Section 3.7).
( 10 ) Mark the position on the slide where the sample is located by drawing a line on the slide, as the sample position will not be visible on a moist slide in subsequent steps.
( 11 ) Store slides dry at 4°C for several days or at -20°C for several months (see Note 13).
3.3 Probe labeling
Since instructions based on random priming or nick forward principles are available in commercial kits, the experimental protocols for labeling DNA are not described in this chapter. Direct PCR labeling is also commonly used, which involves the introduction of a labeled molecule in combination with the amplification of a specific fragment of the clone, or of a specific fragment of total genomic DNA.
For FISH to be successful, a suitable proportion of labeled molecules must be introduced into the probe to facilitate detection. For more efficient polymerase action, labeled nucleotides (dUTP or dCTP ) are mixed 1:2 with unlabeled TTP or CTP. Reagent manufacturers often suggest recommended concentrations and dilutions of labeled nucleotides for optimal integration efficiency. This is the amount required as a starting point for standard experiments, although some very expensive labeled nucleotides, especially freshly opened reagents that have not been freeze-thawed many times, can be reduced to 50% to 70% of the recommended amount. Another important factor is the length of the probe after labeling. The probe should not be too long, usually 200~600 bp, so as not to penetrate into the DNA of the chromosome. Random priming and nick forward methods produce probes that are just the right length for FISH, although sometimes it may be necessary to adjust the DNA enzyme component of the nick forward enzyme system. Labeling larger insert clones by PCR may not result in good probes.
Another factor affecting the success of labeling is the purity of the template DNA and the length of the DNA template sequence used for labeling. When using cloned fragments, make sure that the DNA prepared in small quantities is pure and free of bacterial contamination so that the insert can be cleanly and completely digested. For smaller insert fragments (100 bp to 2 kb), PCR amplification with M13 sequencing universal primers is recommended to obtain very pure template DNA, and it is advisable to purify the template DNA by electrophoresis of the PCR product followed by excision and recovery of the target bands prior to labeling with random primers or cut-preferences. For longer products, small amounts of extracted DNA can be labeled directly after plasmid linearization or digestion of the target fragment.
However, it has been found that large DNA molecules often do not make good templates, possibly because the enzyme is ineffective if the polymerase does not stop at the end of the DNA template molecule. Therefore, it is better to label large DNA molecules after they have been sonicated, heated, or enzymatically cleaved (see [ 4 ], [ 19 ]).
The biotin labeling method [ Fig. 14.3 (a ) ] is recommended for the detection of transgenic fragments because biotin is the smallest half-antigen, is usually the easiest to incorporate, and specialized biotin kits are commercially available. Secondly, many different affinogens, streptavidin, or anti-biotin antibodies that bind tightly and stably to fluorescent dyes are commercially available (e.g., Alexa dyes and Molecular Probes, Inc.). As a second control or indicator probe, which can be labeled with digoxigenin or direct fluorescent dyes, 5S rDNA is recommended because it is widely available and has both primary and secondary fluorescent sites in many species [Fig. 14.3 ( a )], and provides the best control for FISH experiments. 45S rDNA can be used, but often has strongly fluorescent sites that outperform the fluorescent signal of the transgene. hybridization signal of the transgene. Another very good use is repetitive DNA sequences, which can help identify chromosomes.
3.4 Preparation of slide material
Before starting in situ hybridization, the slide material should be pretreated to enhance penetration of the probe into the target site, e.g., by treating the surface proteins with pepsin or proteolytic hydrolase, and to reduce binding of non-specific probes to detection reagents, e.g., RNAase and pepsin/proteolytic hydrolase treatments. Afterwards the samples are remobilized with paraformaldehyde and ethanol so that the samples remain stable after several washes.
( 1 ) All steps are performed in a staining vat at room temperature unless otherwise indicated.
( 2 ) 200 μl of RNAase was added to each slide at the scratch mark, covered with a plastic coverslip, and incubated for 1 h in a humidor.
( 3 ) Remove the coverslips and wash the slides twice in 2x SSC for 5 min each time.
( 4 ) Shake the slides in 10 mmol/L HCl for about 2 min, quickly add 200 μl of pepsin solution to each slide, cover with a plastic coverslip, and incubate at 37℃ for 10 min (see Note 6).
( 5 ) Elute the coverslips in distilled water, and then wash the slides with 2X SSC twice for 5 min each time.
( 6 ) Place the slide in a staining vat with paraformaldehyde fixative in a fume hood for 10 min.
( 7 ) The slides were washed twice with 2X SSC for 5 min each time.
( 8 ) The slides were dehydrated with graded ethanol (70%, 90% and 96% for 2 min each) and dried in the air.
3.5 Hybridization mix denaturation and hybridization
( 1 ) Prepare the hybridization solution in centrifuge tubes, see Table 14.1, and mix thoroughly. 
( 2 ) Add 34 μl of hybridization solution, 2 μl of transgene probe, 1 μl of indication probe or control probe into the centrifuge tube, and make up to 40 μl with distilled water as probe mix.
( 3 ) Denature the probe, put it into a 70°C water bath for 10 min, and then put it on ice for 5 min.
( 4 ) Drop the probe mixture onto the slide where the chromosomes are located, cover with a plastic coverslip, take care to remove all air bubbles, if there are any air bubbles, it is necessary to uncover the coverslip and re-cover it.
( 5 ) Place the slide on a heating block and raise the temperature to the desired denaturation temperature, usually 75°C for 8 min. Depending on the species, preparation method and storage time of the material, the denaturation temperature ranges from 70 to 95°C for 6 to 12 min (see Note 14).
( 6 ) Cool down to 37°C and place at 37°C for 10-20 min, keeping the temperature at 37°C for hybridization.
( 7 ) If the heating plate can be humidified, keep the slide on the heating plate, if not, transfer the slide to an oven humidor or water bath for hybridization. Be careful not to let the sample dry out or allow condensation to accumulate on the sample. Overnight (16 h ).
3.6 Rigorous washing and hybridization site detection
( 1 ) Prepare the wash solution and assay buffer, see section 2 . 6.
( 2 ) Remove the slide from the hybridization and check that the sample is not evaporated dry or wetted by condensation.
( 3 ) Float the coverslips away at 42°C, 2X SSC, taking care not to scrape between slides.
( 4 ) Wash the slide at 42°C, 2X SSC for 2 min.
( 5 ) The slides were incubated twice at 42℃ in rigorous washing solution for 5 min each time, and the temperature of the washing solution was measured and recorded accurately. Use 0.1% x SSC high rigor wash or 2X SSC low rigor wash (see Note 8).
( 6 ) The slides are washed three times in 2x SSC at 42°C.
( 7 ) Remove the slide from the water bath and allow it to cool naturally for 10-15 min.
( 8 ) Transfer the slide to assay buffer and leave for at least 5 min.
( 9 ) Add 200 μl of BSA to the mark of each slide, cover with a plastic coverslip and leave for 5~30 min.
( 10 ) Prepare the assay solution according to the probe marker (see section 2. 6), remove the BSA, aspirate 50 μl of assay solution onto the slide, cover with a plastic coverslip, and incubate for 60 min at 37°C in a warming oven (see Note 9).
( 11 ) Remove the coverslip and wash with assay buffer three times for 5 min each time.
3.7 Re-staining and sealing
Most of the samples need to be double-stained with DNA to visualize the chromosomes. The dye color must be different from the probe color and the signal must not be so bright that it blurs the hybridization signal. Therefore, it is recommended to stain before blocking. Some experiments combine staining and blocking in one step, which saves a step but may lead to overstaining and high background values. The first fluorescent FISH experiment uses propidium iodide staining, and the chromosomes appear red when blue or green light is used as excitation light.
In dual-target FISH experiments, when fluorescent probes are labeled with red and green, it is appropriate to use DAPI with blue light under UV excitation (Fig. 14.3) because it often shows heterochromatin banding patterns at the same time. DAPI is also a good dye for detecting the quality and number of chromosomes during chromosome preparation, and it is especially useful when the chromosomes are very small [Fig. 14.1 (a)].
( 1 ) Apply 50-100 μI of the staining solution to the slide, cover with a plastic coverslip, and leave for 10 min at room temperature in the dark (see Note 5).
( 2 ) Remove the coverslip and wash briefly with assay buffer (see 4 . 2 .). 2 . 6 section).
( 3 ) Aspirate the buffer, add a drop of anti-fluorescence quenching sealer solution to the coverslip, and cover the glass coverslip. In conjunction with the filter paper sheet squeeze the slide by hand to remove excess sealing solution (see Note 15).
( 4 ) Slide Observation. It may take several hours for the quencher to penetrate into the sample, so it may be left overnight before observation and photographs are taken.
3.8 Observation and Photography
Observe probe hybridization sites and chromosome staining using a fallout fluorescence microscope. This chapter is not a description of the use of the microscope, but only the following suggestions for some common problems and solutions.
( 1 ) The fluorescent dye and in situ hybridization signals can be very weak and decay rapidly, even when sealed with a good fluorescent antidote, so the microscope should be placed on a fixed benchtop in a fully darkened room and arranged for easy handling. Ensure that the system is properly placed and adjust the light source to be centered.
( 2 ) Ensure that the instrument and immersion oil and UV are fluorescence-specific. The immersion oil should be kept at room temperature in the dark. The heat of the sun in the transportation vehicle, the oil mixture, and the absorption of water by the immersion oil may cause the immersion oil to self-fluoresce or become impervious to UV.
( 3 ) - In general, it is best to use filters specific to a single fluorescence. Multi-channel filters are available, but these slides do not work well due to the varying light intensities of the different fluorescents, and hybrid excitation between the different fluorescents often makes the results difficult to analyze. Make sure that the filters do not change their characteristics over time.
( 4 ) Take photographs using either film or digital cameras. Both are capable of obtaining image resolution that meets publication standards. However, it must be noted that no digital system, no matter how expensive, can compensate for the advantages of direct microscopic observation. When film is used, color film is better than slides because it has a better exposure range and is cheaper to price and print.
( 5 ) Slides should always be placed in an anti-attenuator, which greatly prevents the attenuation of the fluorescent signal.
( 6 ) Special care should be taken before recording the fluorescent signal not to attenuate the chromosome signal, especially the weak FISH hybridization signal. Observation at low magnification generally does not attenuate the signal. However, there is rapid attenuation at 63x and 100x. It is necessary to set up the camera system so that the exposure button can be pressed quickly once the focal plane is focused. Photograph the FISH signal first and do not observe chromosome fluorescence with DAPI until a satisfactory FISH hybridization signal has been found.
( 7 ) When observing a weak fluorescent signal that is barely visible to the naked eye, focusing the image can be difficult. It is possible to alternate between the red and green focal planes, which is not convenient with DAPI.
( 8 ) Whether using a digital camera or regular film, it is easiest to analyze, overlay, and combine various images using image processing software (e.g., Adobe Photoshop, Corel Draw, or other similar packages). However, care must be taken that images are not over-processed. The responsible person must be well aware of the fact that he/she is accountable for the image processing of laboratory members!
3.9 FISH analysis of transgenes
Probably the most difficult part of the experiment is determining whether the observed or recorded signals are real and reliable, especially for low-copy transgenes.
( 1 ) Check that the mid-mitotic phase is intact and that the chromosome morphology is well preserved. Chromosomes can be slightly bloated, but not too hairy. Chromosomes should be unbroken, without holes or distortions.
( 2 ) Make sure the control probe has the expected results.
( 3 ) Watch for background signals and non-specific cross-hybridization signals that may blur out the true FISH signal [ Fig. I 4.3 (b) and Fig. 14.3 (c) ]. Any signal outside the focal plane that produces a clear edge can be removed [ see Fig. 14.3 (a ) ]. A strong signal in the form of a starburst [Fig. 14.3 (d)] often means that the probe or detector has deteriorated.
( 4 ) Counting transgene FISH signals on chromosomes. Analyze a number of mid-mitotic cells and, if possible, identify the chromosomes and the location of the FISH signal. In a normal diploid cell, a diploid locus should produce fluorescent signals at the same sites on both chromatin where the homologous gene is located (see Note 16). When the target band is less than 50 kb, it is unlikely that signals from all loci can be observed in the same mid-division cell because of chromosome helices and extrusions, so do not expect to see all signal sites in any given mid-division cell. However, after observing 5 to 10 or more mid-division cells, it should be possible to determine which is the true signal and which is the background signal. When a signal with high fluorescence intensity and good reliability is observed, it indicates that several transgenes are integrated into the genome in a tandem manner [ [18], [ 19 ] , see Fig. 14 .3 (a ) ].
( 5 ) The most important point is to compare slides obtained from the same probe or from the same batch of experiments as a way to rule out experimental failures, experimental contamination or ineffective labeling of probes, inappropriate antibody dilution, or improper use of other reagents.