Fluorescent in situ hybridization (FISH)

Overview

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Fluorescent in situ hybridization (FISH) is a classic method of research that is used to locate the positions of specific DNA sequences on chromosomes. In FISH, in situ hybridization is coupled with fluorescent labeling where probes are labeled with an F-labeled nucleotide and then targeted to the DNA or gene or interest. In this coupled technology, dsDNA can be particularly identified and targeted and then visualized through a fluorescence microscope or other applicable imaging system.

FISH technology has been used extensively as a macromolecule recognition tool to effectively identify and map chromosomal, and sub-chromosomal, abnormalities during the annotation phase of the Human Genome Project. Now, FISH is used in research more so for its highly sensitive and specific capabilities, which may be used as a diagnostic tool for the identification of genetic and chromosomal disorders, principally directed for clinical diagnosis.

The FISH procedure uses fluorescent labels that are directly complementary to the nucleotide sequence of interest. FISH technology may therefore be used not only to target specific DNA sequences, but mitochondrial RNA (mtRNA), long noncoding RNA (lncRNA), and microRNA (miRNA) in various tissue and cell types, as well.

There are many slight variations in how FISH may be performed, however each procedure follows the same general principles. Firstly, a probe is constructed. The probe is then fluorescently labeled directly by a process called Nick translation. Next, the labeled probe and target DNA/RNA are denatured separately and then sequentially annealed to one another. Locations of the fluorescent probe within the target DNA/RNA can then be identified through various means of analysis. Below is an example FISH protocol using DNA from HeLa cells, using probes constructed through PCR and direct fluorescent labeling methods, along with hybridization and secondary washing steps.

Example Protocol

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1. Probe Creation using dPCR
   
   

   Materials	Specifications
   DNA extraction kit	(Qiagen's FlexiGene DNA kit) used to obtain gDNA from HeLa cells
   Forward primers	5'-TTA GGG TTA GGG TTA GGG TTA GGG TTA GGG-3'
   Reverse primer	5'-CCC TAA CCC TAA CCC TAA CCC TAA CCC TAA-3'
   PCR kit	Complete with enzyme, buffer, or master mixes
   Fluorescent labelers	dUTPs or biotinylated dUTP (at a 1:1 ratio of iF488-dUTP:dTTP)
   dNTPs	Used to provide nucleotides to unzipped strands during PCR
   PCR and gel clean up kit	(Enzo Biosciences) used to purify DNA probes.

   
   
   1. Follow the steps on the DNA extraction kit to extract gDNA from HeLa cells, or use a manual in-house protocol if preferred.
   2. Using a spectrophotometer or absorbance microplate reader, ensure there is at least 100 ng of gDNA starting material.
   3. Combine 100 ng gDNA with appropriate amounts of PCR enzyme and PCR buffer.
      Note: Consequently, a PCR master mix may be used that contains a DNA polymerase, dNTPs, MgCL2 and buffer. If using a PCR master mix, only primers, fluorescent probes, and water (DNase/RNase free, to make up the volume) need to be added to the working sample.
   4. Add 500 uM dNTPs to the working sample.
   5. Add 300 uM of forward and 300 uM of the reverse primers to the working sample.
   6. Add an appropriate amount of dUTPs or biotinylated dUTP.
   7. Perform PCR as per the PCR enzyme's profile
   8. Purify the probes using the PCR and gel clean up kit.
2. FISH protocol
   
   

   Materials	Specifications
   Colcemid	0.1 µg/mL, KaryoMAX Colcemid Solution (Thermo Fisher scientific, Cat; 15212021)
   KCl	0.075 M, prewarmed to 37 ℃
   Fixative1	3:1, methanol:acetic acid, ice cold
   Culture medium	As used for cell culture, prewarmed to 37 ℃
   Transfer tubes	15 mL falcon tube, 1.5 mL Eppendorf tube
   Trypsin	Commercially available
   100% ETOH
   DI H1O
   Poly-L-Lysine
   Glass microscope slide

   1. The fixative must always be freshly prepared.
   
   
   1. Day 1: Cell Treatment With Colcemid
      1. Bring HeLa cells to 70-80% confluency
      2. Harvested cells by trypsinization
         1. Add 1 mL of trypsin to the cells, and softly pipette the sample.
         2. Incubate at 37 ℃ for 3-5 mins.
         3. Add 5 mL of pre-warmed culture medium to the sample and transfer the sample to a 15 mL falcon tube.
      3. Add 0.1 µg/mL of Colcemid and incubate the sample at 37 ℃ for 4 hrs.
      4. After incubation, centrifuge the sample at 1700 rpm for 5 mins.
      5. After centrifugation, carefully remove the supernatant.
      6. Dissolve the pellet in 8 mL of 0.075 M KCl (pre-warm at 37 ℃)
      7. Incubate the sample for 10 mins at 37 ℃.
      8. After incubation, add 1 mL of ice cold fixative to the sample.
      9. Centrifuge the sample at 1700 rpm for 5 mins.
      10. After centrifugation, carefully remove the supernatant.
      11. Add 4 mL of ice cold fixative to the pellet.
      12. Incubate the sample at 4 ℃ for 10 mins.
      13. Inculcate the sample at 37 ℃ for 15 mins.
      14. After incubation, centrifuge the sample at 1700 rpm for 5 mins.
      15. After centrifugation, carefully remove the supernatant.
      16. Add 1 mL of fixative to the sample
      17. Transfer the sample to 1.5 mL Eppendorf tube.
      18. Centrifuged the sample at 2300 rpm for 5 mins. After centrifugation, carefully remove the supernatant.
      19. Add fixative (enough to cover and resuspend cells) to the pellet and store at -20 ℃.
   2. Slide Preparation
      1. Wash slides with 100% ETOH and DI H2O.
      2. Incubate slides with 50 µg/mL of poly-L-Lysine at room temperature.
         Note: Pre-prepared poly-L-Lysine slides are also commercially available.
   3. Day 2: Metaphase Spread
      
      

      Materials	Specifications
      DI H2O	Commercially available
      NP-40 Lysis Buffer	Diluted to 0.1% using 2X SSC Buffer, commercially available
      Acetic acid	45%, Commercially available
      Poly-L-Lysine slides	Previously prepared
      DNA probes
      20X SSC Buffer	To make 100 mL solution: Combine 17.532 g NaCl and 8.823 g Sodium citrate. Dissolved in 100 mL distilled water, and adjust to pH 7.
      ETOH	70% and 100%
      Denaturation buffer	Prewarmed to 80 ℃, 70 % of total volume formamide with 2X SSC, pH 7.
      Sperm DNA	To make single stranded DNA: Boil aliquot amount of stock DNA at 100 ℃ for 5 mins. Put the sample on ice for 5 mins. Store the sample at -20 ℃ until use in the hybridization buffer.
      Hybridization buffer	Includes concentrations by volume: 55 % formamide 2X SSC 11% dextran sulfate 0.11% SDS 220 ug/ml sperm DNA

      
      
      1. The next day, wash the slides with DI H2O.
      2. Wash the slides with 45% acetic acid.
      3. Adhere cells to the slides
         1. Cells in fixative that are stored in -20 ℃ should be centrifuged 1-2 times before
         2. Dropping on slides.
         3. Hold the slides at a 45° angle.
         4. At a 3-5 cm height, apply 1 drop of cells (~15 uL from a 200-300 uL cell suspension) onto the slides.
      4. Allow the slides to air dry for 5 mins.
      5. Incubate the slides at 37 ℃ for 1 hr.
   4. Cell Pretreatment and Denaturation
      1. Incubate prepared slides with enough 0.1 % NP-40 in 2X SSC to cover the sample for 30 mins at 37 ℃.
      2. Add 70% ethanol to prepared slides and allow them to air dry for 3-5 mins.
      3. Add 100% ethanol to prepared slides and allow them to air dry for 3-5 mins.
      4. Add 20-40 uL of pre-warmed denaturation buffer to the prepared slides, and a cover slip to the top.
      5. Incubated the prepared slides at 80 ℃ for 5 mins.
      6. Remove the cover slip by dipping prepared slides in 70% ethanol
      7. Add 100 % ethanol to prepared slides and allow them to air dry for 2 mins.
   5. Probe Preparation
      1. Add 1 uL of the probe, previously made, to 9 uL of hybridization buffer. Vortex well.
      2. Incubated the solution at 37 ℃ for 10 mins.
      3. Denatured the solution at 80 ℃ for 5 mins in a thermal cycler.
      4. Place the solution on ice until applied to the prepared slides.
      5. Hybridization:
         1. Add 10 uL of the solution to the, now denatured, slides. Place a cover slip on top.
         2. Seal the slides with rubber cement. Incubate the slides at 45-50 ℃ for ~5 mins.
         3. Incubated the sealed slides at 37 ℃ until the next day. If necessary, place a wet paper towel under the slides to help maintain moisture.
3. Day-3: Post-Hybridization Wash and DAPI staining:
   
   

   Materials	Specifications
   2X SSC buffer	Previously prepared
   Hybridized slides
   NP-40 Lysis Buffer	Diluted to 0.3% in 0.4X SSC Buffer, commercially available
   2X SEC	Previously prepared
   DAPI Mounting Medium	To prepare: Dilute 10mM DAPI in mounting medium until the concentration of 5uM is reached

   
   
   1. Remove the coverslip by placing the slides in a 2X SSC buffer.
   2. Wash the slides with 0.3% NP-40 in 0.4X SSC buffer for 2 mins at 73-75 ℃.
   3. Wash the slides with 2X SSC Buffer for 1 min at room temperature.
   4. Allow the slides to air dry. Mount the cover slip by applying DAPI Mounting Medium.
   5. Images can be acquired using a fluorescence microscope at 20X magnification.

Visualization and Analysis

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The power of FISH can be amplified by the use of multiple fluorescent colors in tandem. Multicolor FISH can be used to identify as many labeled features as there are differing fluorophores used in the hybridization process. Single colors, as well as various combinations of colors, can be simultaneously detected.



Epifluorescent microscopy is commonly used for this multicolor analysis, and similar to other fluorescence microscopes, the excitation light from the light source is passed through a filter before reaching the objective lens and dichroic mirror. This excitation filter allows a single band of a particular wavelength through, dependent on the peak fluorescence excitation wavelength of the fluorophore. The excitation filter can be in the form of filter cubes or a filter wheel with multipass dichroic and barrier filters. The produced signals must be isolated by an emission filter that ensures that only the wavelength emitted from the targeted fluorophore in the sample passes through.

Traditionally, data was collected through digital charged coupled devices (CCDs), high-speed color film, or intensified video cameras, though these methods had issues with color fidelity and there existed problems when processing superimposed images, with different colors, within a single specimen.

Recently, much development has been made to overcome these limitations with the creation of quantitative imaging analysis software suites. These analysis programs are capable of capturing images across multiple wavelengths and/or focal planes, with the ability to retrieve and store large volumes of multicolor image data sets. The software offer the ability to focus at multiple wavelengths, record the slide locations of cells, quantitatively measure probe counts and/or intensity, and determine cell morphology. It may also be able to analyze multicolor metaphase chromosomes, map genes, offer comparative genomic hybridization ratios, as well as determine karyotype generation.

Confocal laser scanning microscopy may also be used for visualization. This technique uses light emitted by a laser, at a specific wavelength to maximally excite the fluorophore, which it will then emit at a somewhat longer wavelength. The difference between these two wavelengths is known as the Stokes shift.

The light runs through a pinhole aperture before hitting the dichromatic mirror directed towards the specimen, where secondary fluorescence is then emitted. Instead of illuminating the entire sample at once, the laser light only illuminates a given spot at a specified depth. Because of this, out-of-focus signals from features located above and/or below the focal plane contribute to the high signal-to-noise ratio of images produced. As the detector pinhole required for focusing images also may reduce signal strength, confocal laser scanning microscopy typically requires higher fluorescence signal yield to be effective.

Considerations and Limitations

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Dozens of variations of FISH have been created to improve the sensitivity, specificity, and image resolution for the associated target samples. Depending on the technique, chemical and physical information can be generated for a number of biological targets. As the target DNA remains intact, (different from other forms of molecular analysis) genetic information about probe positions in relation to chromosome bands or other probes is directly, and efficiently, obtained. FISH has aided in rare disease diagnostics, evaluation of prognosis, and even the chance of remission for diseases like cancer.

In many cases, it may be necessary to multiplex spectral techniques to increase the number of detectable targets. Many experimental steps require empirical optimization specifically for the samples and probes used, which may prove time consuming and costly. Long hybridization times are essential to ensure the appropriate saturation of signals throughout the specimen, so FISH is also an inherently lengthy process and requires experienced personnel, though advances in assay automation may alleviate these limitations in the future.

Though many variations of FISH exist, a major downfall of this technique is that it can only detect known genetic aberrations. FISH can be used only to confirm previously characterized imbalances and abnormalities, and cannot serve as a screening test for novel chromosomal rearrangements.

Product Ordering Information

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