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Method Article

DNA-FISH Metaphase Spreads to Distinguish Extrachromosomal DNA from Homogeneously Staining Regions in Human Cancer Cell Lines

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DOI:

10.3791/71970

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August 4th, 2026

 ,  , 
1Department of Molecular and Human Genetics, Baylor College of Medicine

In This Article

Summary

We report a DNA-FISH metaphase spread protocol that visually detects locus copy number and location within the genome. This approach enables single-cell resolution of amplification states, specifically in cancer cell lines containing extrachromosomal DNA and homogeneously staining regions.

Abstract

Whole-genome sequencing identifies focal DNA amplifications with base-pair resolution but cannot determine whether amplified sequences reside on extrachromosomal DNA (ecDNA, also known as double minutes) or within chromosomally integrated homogeneously staining regions (HSRs). DNA fluorescence in situ hybridization (DNA-FISH) metaphase spreads remain the gold standard for distinguishing these amplification states at single-cell resolution. Here, we present a detailed protocol for DNA-FISH metaphase spreads using human cancer cell lines, encompassing cell culture, metaphase arrest, hypotonic treatment, fixation, chromosome spreading, fluorescent probe hybridization, and fluorescence imaging. The protocol incorporates intermediate quality-control steps to verify successful chromosome dispersion and optimize metaphase spread quality, making the workflow accessible to laboratories without specialized cytogenetics expertise. Results demonstrate clear visualization of ecDNA and HSR amplification states using locus-specific probes and illustrate common technical artifacts that can affect interpretation. This protocol provides a robust and reproducible approach for studying the structural organization of oncogene amplification in cancer cells.

Introduction

The human genome is organized not only by its sequence, but also by its structure, and changes in either sequence or structure can lead to cancer1,2,3,4. Oncogene amplification is present in approximately 40% of all tumors, where it contributes to tumor initiation, growth, metastasis, and drug resistance5. In some cases, amplified oncogenes are carried on extrachromosomal DNAs (ecDNAs, also called double minutes), which are circular, self-replicating, double-stranded DNA elements that reside outside of the linear chromosomes6,7,8,9. ecDNAs are hundreds of kilobases to megabases in size10,11. Due to their lack of centromeres, they segregate asymmetrically during mitosis, leading to as many as 100+ copies in a single nucleus12,13,14,15,16,17,18. This uneven segregation generates copy number variability across daughter cells, which drives rapid tumor evolution and poor patient outcome17,19.

ecDNAs exist in a dynamic relationship with homogeneously staining regions (HSRs) (Figure 1A)8,18. HSRs are focal, intrachromosomal amplifications that can arise through the reintegration of ecDNAs into a single locus in the genome8,10,18,20,21,22,23,24,25,26,27,28,29,30,31. These two states, ecDNAs and HSRs, can interconvert, revealing that oncogene amplification is structurally plastic24,25,32,33. This interconversion has therapeutic relevance because ecDNAs can transition to the HSR state in the presence of drugs32,33,34,35,36,37,38. Cells that undergo this transition to the HSR state then become resistant to the drug under therapeutic pressure33. HSRs also appear to be more frequent in malignant tumors than in benign tumors39. Despite their clinical relevance, no FDA-approved strategies target the HSR state, in part because methods to distinguish these amplification structures remain underutilized.

Although ecDNA and HSR states have distinct clinical consequences, current sequencing approaches cannot reliably distinguish them. Whole-genome sequencing (WGS) now identifies amplified regions with base-pair resolution40. WGS has been valuable in defining the sequence composition of ecDNAs and HSRs, but at the expense of spatial resolution. Unlike traditional cytogenetic approaches, WGS alone cannot determine whether an amplified sequence resides on ecDNAs or is integrated into a chromosome as an HSR41,42,43. As a result, methods that preserve the spatial context of amplification remain essential for understanding the biology of these distinct states.

For the past 40 years, DNA fluorescence in situ hybridization (DNA-FISH) metaphase spreads have served as the gold standard for identifying and distinguishing ecDNA and HSR amplification states42,44,45,46. In fact, ecDNAs and HSRs were first identified and characterized through metaphase-based cytogenetic analyses7,47. Before the widespread use of fluorescent probes, in situ hybridization was performed using radiolabeled DNA probes, which have also been used to visualize ecDNAs and HSRs48,49. As WGS gained in popularity, DNA-FISH metaphase spreads became less common. Many aspects of the workflow are scattered across the cytogenetic literature, and optimized protocols are typically retained within individual research groups or cytogenetics labs that treat the method as proprietary50,51. Here, we describe a DNA-FISH metaphase spread protocol to visually resolve amplification state at the single-cell level.

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Protocol

All human cell culture procedures were performed in compliance with institutional guidelines.

1. Arrest cells in metaphase using Colcemid (N-methyl-N-deacetyl-colchicine)

  1. Begin with approximately 2.0 × 106–2.0 × 107 cancer cells or other cultured cells. Too few cells will yield fewer metaphase spreads, whereas too many cells may result in insufficient fixation.
  2. Determine the appropriate Colcemid incubation time for the experimental system. Incubation time affects the number of metaphase spreads obtained and chromosome length, and optimal conditions vary among cell types.
    NOTE: Shorter incubation times result in fewer cells arrested in metaphase at the time of collection. Longer incubation times produce more condensed chromosomes with shorter arms, making analysis more difficult. For cancer cell lines with an approximate doubling time of 24 h, incubation periods ranging from 45 min to overnight produce suitable metaphase spreads. Optimize the incubation time for cells with slower doubling times. A 2 h incubation is generally sufficient to produce an adequate number of metaphase spreads with chromosome lengths suitable for analysis.
  3. Calculate the amount of Colcemid required to achieve a final concentration of 10 µL/mL in the appropriate cell culture medium.
    NOTE: Maintain cells in logarithmic growth by passaging or seeding them at least 24 h before metaphase arrest. Treat cells with Colcemid when cultures reach 70%–90% confluence.
  4. Dilute the Colcemid stock solution 1:1 with sterile ultrapure water. Add the appropriate volume to the culture and incubate for 45 min to overnight in a cell culture incubator.

2. Harvest cells

  1. Prepare fresh Carnoy's fixative (3:1 methanol: acetic acid). Keep the fixative on ice throughout the fixation procedure.
  2. Prepare a fresh 0.075 M KCl solution and warm it in a 37 °C water bath.
  3. Collect the cells as a pellet and record the cell count. Use this value to calculate the final resuspension volume.
    NOTE: Detach adherent cells by trypsinization. Transfer suspension cells directly into 15 mL centrifuge tubes. Dispose of Colcemid and Colcemid-containing media according to institutional hazardous waste guidelines.
  4. Wash the cells with PBS.
  5. Gently resuspend the cells in 10 mL of warm 0.075 M KCl and mix by gentle inversion. This hypotonic treatment swells the cells, allowing them to rupture and disperse chromosomes during slide preparation.
  6. Incubate the cells in a 37 °C water bath for 30 min.
  7. Slowly add 2 mL of ice-cold Carnoy's fixative while gently mixing the suspension by inversion.
  8. Allow the cells to stand for 5 min.
  9. Centrifuge at 500 × g for 5 min at room temperature (RT).
  10. Carefully aspirate the supernatant or remove it with a pipette.
  11. Slowly add 10 mL of ice-cold Carnoy's fixative in 0.5 mL increments while gently mixing after each addition.
  12. Allow the cells to stand for 10 min at RT.
  13. Centrifuge at 650 × g for 5 min at RT.
  14. Repeat steps 2.10–2.13 twice
  15. During the final wash, leave the cells suspended in Carnoy's fixative.
    PAUSE: Store the fixed cell suspension at −20 °C or proceed immediately to Section 3.
    NOTE: For samples containing fewer than approximately 3 × 106 cells, perform fixation in 1.5 mL microcentrifuge tubes instead of larger conical tubes. Reduce the volumes of 0.075 M KCl and Carnoy's fixative 10-fold while maintaining all incubation times, temperatures, and centrifugation conditions.

3. Tabletop dropping

  1. Retrieve the sample from −20 °C and thaw it to RT.
  2. Centrifuge at 800 × g for 7 min at RT.
  3. Label the glass slides and place them on ice.
  4. Using the cell count obtained before fixation, calculate the volume of Carnoy's fixative required to resuspend the pellet at a final concentration of approximately 5 × 106 cells/mL. After centrifugation, carefully remove the supernatant with a pipette, leaving the calculated volume of fixative above the pellet.
  5. Resuspend the fixed cells in Carnoy's fixative using a P1000 pipette.
  6. Transfer approximately 300 µL of the fixed-cell suspension into a 1.5 mL microcentrifuge tube.
  7. Thoroughly resuspend the sample using a P200 pipette by pipetting up and down several times.
  8. Aspirate 60 µL of the resuspended cells using a P200 pipette.
  9. Hold the pipette tip approximately 1 m above the glass slide on ice and aim for the center of the slide.
  10. Slowly dispense one drop of the cell suspension onto the slide.
    NOTE: Mark the drop location on the edge of the slide with an ethanol-resistant marker to facilitate identification during subsequent steps.
  11. Allow the cells to dry on ice for 10 min.
  12. Place the slides on a slide incubator at 80 °C for 10 min.
  13. Remove the slides with tongs and allow them to cool to the touch.
  14. Examine the slides using a phase-contrast microscope with a 10× or 20× objective. Confirm the presence of multiple well-dispersed metaphase spreads without excessive overlap.

4. Pretreat the slide

  1. Prepare Coplin jars for the wash steps
    1. Prepare each reagent fresh: 70% acetic acid (40 mL), PBS (40 mL), PBS (45 mL), 70% ethanol (40 mL), 85% ethanol (40 mL), and 100% ethanol (40 mL).
  2. Place the glass slides in 70% acetic acid for 2 min.
  3. Place the glass slides in 40 mL of PBS for 30 s.
  4. Place the glass slides in 45 mL of PBS for 5 min.
  5. Dehydrate the glass slides sequentially in 70%, 85%, and 100% ethanol for 1 min each.
  6. Air-dry the slides at RT for 10 min.

5. Prepare the probe

NOTE: This protocol uses commercially available locus-specific DNA-FISH probes that hybridize to a genomic region of interest.

  1. Select a probe targeting the amplified locus to be examined. Thaw the probe and hybridization buffer according to the manufacturer's instructions. Allow the probe to reach RT before use.
  2. Prepare a probe mixture containing 1 µL of probe and 4 µL of hybridization buffer for each glass slide.
    NOTE: The optimal probe concentration may depend on the manufacturer and the probe. The concentration specified here has been optimized as per the probes used in this study. However, optimize the probe concentration for each new locus.
  3. Apply 5 µL of the probe/hybridization buffer mixture to the center of each drop location on the glass slide.
    NOTE: Probes are light-sensitive. Minimize light exposure throughout the remaining procedure by storing the slides in a dark drawer between steps and while processing other slides.
  4. Carefully place a coverslip over the sample.
  5. Seal the edges of the coverslip with rubber cement.
  6. Place the glass slides in a humid chamber.
  7. Hybridize the probe according to the manufacturer's protocol. Allow hybridization to proceed overnight (12–18 h).

6. Post-hybridization washes, counterstaining, mounting, and imaging

  1. Prepare Coplin jars for the wash steps
    1. Prepare each reagent fresh: 2× SSC (40 mL), 0.4× SSC (40 mL), 70% ethanol (40 mL), 85% ethanol (40 mL), 100% ethanol (40 mL), and PBS (45 mL).
    2. Prepare and store SSC as a 20× stock solution (3 M NaCl, 0.3 M sodium citrate, pH 7.0). To prepare 1 L, dissolve 175.3 g NaCl and 88.2 g sodium citrate in 1 L of ultrapure water. Dilute the stock to prepare 2× SSC and 0.4× SSC working solutions as needed.
  2. Warm the 0.4× SSC solution to 72 °C in a water bath.
  3. Remove the glass slides from the humid chamber and carefully remove the rubber cement.
  4. Place the glass slides in 2× SSC for 2 min at RT. Carefully remove the coverslips.
  5. Place the glass slides in prewarmed 0.4× SSC at 72 °C for 2 min.
  6. Place the glass slides in 2× SSC for 1 min at RT.
  7. Dehydrate the glass slides sequentially in 70%, 85%, and 100% ethanol for 1 min each.
  8. Air-dry the slides at RT for 10 min in the dark.
  9. Dilute 1 mg/mL DAPI 1:1000 in PBS to a final volume of 1 mL.
    NOTE: This aliquot can be stored at 4 °C for up to 1 week.
  10. Pipette 10 µL of the diluted DAPI onto each glass slide at the marked drop location.
  11. Carefully place a coverslip over the sample to ensure the DAPI solution spreads evenly.
  12. Incubate the slides in the dark for 5–10 min.
  13. Place the glass slides in PBS for 30 s.
  14. Remove the coverslips.
  15. Place the glass slides in PBS for 2 min.
  16. Dehydrate the glass slides by briefly dipping them in 70%, 85%, and 100% ethanol for approximately 10 s each.
  17. Air-dry the slides at RT for 10 min in the dark.
  18. Add 15 µL of mounting medium to each glass slide, then place a coverslip on top.
  19. Allow the mounting medium to spread evenly, then gently blot away any excess.
  20. Seal the edges of the coverslip with nail polish.
  21. Allow the nail polish to dry for 15 min at RT in the dark.
    PAUSE: Store the slides at 4 °C for long-term storage.
  22. Image the slides.
    NOTE: Images were acquired using an inverted fluorescence microscope with a 60× oil-immersion objective.
  23. Collect multiple z-stacks for each fluorescence channel to capture signal throughout the full depth of the metaphase spread.
  24. Deconvolve the image stacks using a maximum-likelihood estimation algorithm to enhance signal clarity and reduce out-of-focus fluorescence.
  25. Process and analyze the final images in ImageJ, and select the optimal focal plane for each metaphase spread before downstream analysis.

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Results

This protocol (Figure 1B) yields clean, well-dispersed, intact metaphase chromosomes suitable for cytogenetic analysis, with chromosomes and DNA-FISH signals clearly visible by fluorescence microscopy. Using a brightfield/phase-contrast microscope at low magnification (10× or 20×) is an important quality-control step to verify that nuclei have burst and chromosomes have dispersed (Figure 2). Successful fixation and tabletop dropping should yield well-dispersed m...

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Discussion

Despite their well-established advantages, DNA-FISH metaphase spreads are underused in the era of WGS. The physical organization of amplified DNA is functionally significant, as it can influence downstream biological behavior and therapeutic response61,62,63,64. Focally amplified oncogenes can exist on ecDNA or within HSRs, and these two amplification states are not functionally equivalent

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Disclosures

The authors declare no competing interests.

Acknowledgements

This work was supported by NIH R00HG011467 and CPRIT RR23004 to G.S.E. Imaging for this project was supported by the Integrated Microscopy Core at Baylor College of Medicine with funding from the NIH (P30 DK056338, P30 CA125123, S10OD030414). The authors thank Dr. Pawel Stankiewicz for his advice early on. The authors also thank Dr. Paul Mischel (Stanford University) for providing the GBM39ec and GBM39hsr cell lines.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
15-mL conical centrifugeany sourceNACentrifugation. 
15-mL conical tubesany sourceNACell collection.
200 proof ethanolany sourceNAWash steps. 
Acetic Acid, glacial, 99.7%Millipore Sigma695092Carnoys fixative and slide wash steps.
Antifade Mounting MediumFisherScientificNC9265087Slide mounting after completed slide preparation
Biological safety cabinet with vaccum outletany sourceNAChemical storage.
Colcemid (N-methyl-N-deacetyl-colchicine)Millipore Sigma10295892001Binds tubulin, arrests cells in metaphase.
Coplin jarsany sourceNAWash steps. 
DAPI (4′,6-diamidino-2-phenylindole)Milipore SigmaD8417-1MGFluorescent DNA stain.
EGFR/CON7 FISH probe Empire GenomicsEGFR-CHR07Locus-specific fluorescent probe. 
Elmer's rubber cementany sourceNAKeep coverslide in place during overnight probe hybridization.
Ethanol resistant permanent markerany sourceNASlide labeling.
Fluorescent microscopeany sourceNASlide imaging. 
Glass Cover Slip, 22 x 22mmThermoFisher50-143-780Probe hybridization on slide.
Glass Slide, precleanedVWR48311-703To drop metaphase spreads and perform DNA-FISH.
Humidified CO2 Incubatorany sourceNALive-cell incubation.
Hybridization BufferEmpire GenomicsHyb-BufferBuffer used with probe. 
Ice bucketany sourceNAKeep Carnoy's fixative cold. 
KCl (2M)ThermoFisherAM9640GSwell cells prior to fixation. 
Methanol, 99.9%, HPLC gradeMillipore Sigma1060351000For Carnoys fixative. 
Microcentrifugeany sourceNACentrifugation. 
Microcentrifuge tubesany sourceNAFor cells during fixation, and for probe preparation. 
MYC / SE 8 ProbeLeicaKI-10106Locus-specific fluorescent probe. 
PBS, pH 7.4Gibco10010-023Wash steps. 
Phase contrast microscopeany sourceNAImaging of metaphase spreads immediately after dropping
Pipette tipsany sourceNAPipetting.
Sodium chlorideMillipore SigmaS9888For SSC buffer.
Sodium citrateMillipore Sigma567446For SSC buffer.
Sterile ultrapure waterany sourceNAWash steps. 
ThermoBrite programmable slide incubatorany sourceNASlide incubation.
Tongsany sourceNAFor slide wash steps. 
Water bathany sourceNAFor KCl incubation. 

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