Preparation of Amniocytes for Interphase Fluorescence In Situ Hybridization (FISH)
UNIT 8.9
Stuart Schwartz1 1
R Cytogenetics Laboratory, Laboratory Corporation of America Holdings, Research Triangle Park, North Carolina
FISH has been used to detect and clarify deletions and/or other structural rearrangements, and also has applications in interphase analysis. This unit describes preparation of uncultured amniotic fluid cells for FISH analysis. Cells are swollen, and then slides are prepared by standard methods. The cells are then fixed and permeabilized for subsequent FISH. An alternate protocol describes attachment of amniocytes to a glass or plastic surface followed by hypotonic swelling, fixation, and permeabilization for subsequent FISH. Interphase FISH C 2015 by John Wiley & Sons, analysis of amniotic fluid cells is also described. Inc. Keywords: prenatal diagnosis r FISH r fluorescence in situ hybridization r amniotic fluid r aneuploidy
How to cite this article: Schwartz, S. 2015. Preparation of amniocytes for interphase fluorescence in situ hybridization (FISH). Curr. Protoc. Hum. Genet. 85:8.9.1-8.9.16. doi: 10.1002/0471142905.hg0809s85
INTRODUCTION Use of fluorescence in situ hybridization (FISH) has increased by several orders of magnitude over the past two decades. In a clinical setting, the technique is used most commonly as an adjunct to standard cytogenetic analysis, but can be a stand-alone procedure. Although FISH has been used to detect or clarify gains, losses, and/or structural rearrangements, considerable work has focused on its application in interphase analysis. Most of this work is currently associated with the analysis of leukemia and solid tumors where karyotyping has been difficult or no material (e.g., paraffin sections) is available for karyotype analysis. Interphase analysis has now been utilized extensively for prenatal diagnosis. Although initial controversy surrounded the use of FISH as an investigational procedure for this purpose, there are both recommendations (e.g., from the American College of Medical Genetics) for its utilization and available FDA-approved probes. The types of information that FISH provides, combined with easy utilization and rapid turnaround time, make FISH an attractive adjunct to cytogenetics. Whereas classical cytogenetic methods yield a diagnosis from amniotic fluid cells in 7 to 12 days, interphase analysis by FISH permits detection of specific aneuploidies in 48 hr. Basic Protocol 1 describes preparation of uncultured amniotic fluid cells for FISH analysis. Cells are swollen, and then slides are prepared using standard methods. These are then fixed and permeabilized for subsequent FISH. The Alternate Protocol describes Clinical Cytogenetics Current Protocols in Human Genetics 8.9.1-8.9.16, April 2015 Published online April 2015 in Wiley Online Library (wileyonlinelibrary.com). doi: 10.1002/0471142905.hg0809s85 C 2015 John Wiley & Sons, Inc. Copyright
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attachment of amniocytes to a glass or plastic surface followed by hypotonic swelling, fixation, and permeabilization for subsequent FISH. In contrast to routine cytogenetic procedures for culture of amniotic fluid cells (UNIT 8.4; Minehart Miron, 2012), cells are incubated just long enough for them to attach to the surface, a much quicker procedure. The purpose is to obtain interphase cells attached to plastic coverslips (or glass slides); no metaphase spreads are expected. This procedure is more time-consuming than Basic Protocol 1, but may allow for increased hybridization efficiency because dead cells will not attach to the slides and because changes in the morphology of the cells when they attach make them more permeable to probes; however, it is rarely used today. Two protocols have been given but many exist, and it is incumbent on each laboratory to determine what works best in their laboratory. Basic Protocol 2 for FISH can be used for interphase cells obtained by any of the protocols. The protocol can be utilized with the FDA-approved probe set, or with any repetitiveDNA probes, unique probes, or single-copy probes (e.g., commercially available or from BAC libraries). The FISH protocol described can be used with directly labeled DNA probes (see UNIT 4.3; Knoll and Lichter, 2005, for a more detailed discussion of FISH). CAUTION: Radioactive, biological, and chemical substances require special handling; see APPENDIX 2 for guidelines. BASIC PROTOCOL 1
PREPARATION OF UNCULTURED AMNIOCYTES FOR INTERPHASE FISH ANALYSIS In this procedure, amniotic fluid cells are obtained, washed, swollen in hypotonic solution, and spread onto slides by standard methods. After fixation the slides are used in FISH (see Basic Protocol 2 and UNIT 4.3; Knoll and Lichter, 2005) and analyzed by fluorescence microscopy (UNIT 4.4; McNamara et al., 2005). Unlike the methods in UNIT 8.4 (Minehart Miron, 2012), this protocol does not involve culturing of cells, but can be done on cultured cells if needed. This protocol describes the FDA-approved approach, and as such, the steps presented are as given in the Abbott/Vysis AneuVysion product insert; see https://www.abbott molecular.com/static/cms_workspace/pdfs/US/Package_Insert_04_02_2012.pdf.
Materials Whole amniotic fluid specimen 1× trypsin/EDTA (0.05% trypsin/0.53 mM EDTA·4Na in Hanks’ Balanced Salt Solution without or Mg) Hypotonic solution: 0.56% (w/v) KCl Fixative: 3:1 (v/v) methanol:glacial acetic acid 2× SSC, pH 5.3 (APPENDIX 2D) Pepsin working solution: 2.5 mg pepsin added to 50 ml of 0.01 N HCl Phosphate-buffered saline (PBS; APPENDIX 2D) Post-fixation solution (see recipe) 70%, 85%, and 100% ethanol (see recipe for ethanol wash solutions) 15-ml screw-cap conical centrifuge tubes (e.g., BD Falcon) Clinical tabletop centrifuge (e.g., IEC Centra-HN) Glass microscope slides, pre-cooled Cotton-plugged Pasteur pipets Preparation of Amniocytes for Interphase FISH
Additional reagents and equipment for chromosome slide preparation (UNIT 4.1; Bangs and Donlon, 2005)
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Slide preparation from uncultured amniotic fluid 1. Centrifuge 2 to 5 ml of whole amniotic fluid specimen in a 15-ml centrifuge tube, for 5 min at 200 × g, room temperature. The specimen should not appear bloody or brown. Samples should normally be processed the day the amniotic fluid is received, as rapidity is the essence of this procedure. If necessary, however, samples can be successfully prepared several days after the fluid has been obtained. If not used immediately, amniotic fluid should be stored at room temperature in the dark. Because no cell growth is involved, samples that have been refrigerated or exposed to heat can still be used successfully. The amount of fluid to be used for this test depends on the gestational age at which amniocentesis is performed and the amount of fluid received by the cytogenetics laboratory. See Critical Parameters and Troubleshooting for guidelines.
2. Resuspend the pellet in 2 to 5 ml of 1× trypsin/EDTA and incubate in a 37 ± 1°C water bath for at least 15 min. 3. Centrifuge the suspension for 5 min at 200 × g, room temperature. 4. Resuspend the pellet in 2 to 5 ml of 0.56% KCl and incubate for 20 min in a 37 ± 1°C water bath. 5. Add 0.8 to 2 ml of fixative to the cells/hypotonic solution and vortex gently. 6. Centrifuge the suspension for 5 min at 200 × g, room temperature, and resuspend the pellet in 1 ml fresh fixative. Store fixed specimens at 4°C for at least 30 min or until ready to perform FISH. For long-term storage, keep fixed specimens at −20°C (±10°C) in fixative. 7. Before placing cells on slides, adjust volume of cell suspension according to size of cell pellet by adding additional fixative. If needed, particularly after a prolonged storage (>1 month), wash pellets with fixative by repeating step 6 before slide preparation. 8. To prepare slides for FISH, drop the cell suspension directly onto one or two cold glass slides, making two hybridization areas (15 to 25 μl of cell suspension per area). See UNIT 4.1 (Bangs and Donlon, 2005), Support Protocol, for details of slide making. Achieving the proper concentration for slide making requires practice and can only be arrived at through trial and error. Slides should contain enough nuclei for analysis. In general, one to three slides can be made from each sample, depending on cell concentration.
9. Proceed to steps 10 to 18, if desired. If the optional pretreatment procedure will not be used, age the specimen slides in 2× SSC for 30 min to 1 hr at 37°C or at room temperature for 24 hr, with the slide box uncovered, before hybridization or storage. Slides may be used immediately for FISH analysis (see Basic Protocol 2 and UNIT 4.3; Knoll and Lichter, 2005), or stored at room temperature until the next day. They may be stored for longer periods at 4° or –20°C.
Pretreatment for uncultured amniotic fluid cells (optional) The following method is recommended in order to achieve optimum FISH results, especially for late-gestational-age specimens and specimens to be hybridized immediately following slide preparation. 10. Place slide(s) prepared from uncultured amniocytes, in 2× SSC for 1 hr at 37 ± 1°C.
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11. Place slide(s) in freshly made pepsin working solution for 13 min at 37 ± 1°C. 12. Rinse slide(s) in PBS at room temperature for 5 min. 13. Place slide(s) in post-fixation solution for 5 min. 14. Rinse slide(s) in PBS at room temperature for 5 min. 15. Dry slide(s). To speed drying, use a cotton-plugged Pasteur pipet to direct compressed air flow onto the slide. 16. Immerse slide(s) in 70% ethanol at room temperature. Allow the slide(s) to stand in the ethanol wash for 1 min. 17. Remove the slide(s) from 70% ethanol. Repeat step 7 with 85% ethanol, followed by 100% ethanol. 18. Proceed to denaturation of slides(s) as described in Basic Protocol 2. ALTERNATE PROTOCOL
PREPARATION OF AMNIOCYTES ATTACHED TO A SURFACE FOR INTERPHASE FISH ANALYSIS This protocol describes preparation of amniocytes attached to coverslips or slides. These cells are swelled in hypotonic medium and fixed for subsequent FISH analysis. This method is in some cases preferable to that of Basic Protocol 1 in that dead cells will not attach to a coverslip (or slide), so that overall efficiency of the procedure is increased. When cells prepared by this methodology are analyzed by FISH, better hybridization efficiency between probes and interphase chromosomes is observed. This may be due to an increase in surface area available for hybridization, an increase in permeability of membranes, or other factors. Because some time may be required for attachment of cells to the coverslip and, in some cases, for initial growth, turnaround time may be longer with this protocol than with Basic Protocol 1. However, this protocol is rarely used today. A number of variations to this method are given in annotations to the steps. Regardless of variations, the major factor in this protocol is reliance on cellular attachment factors to increase the number of living cells that will attach to the coverslip or slides. NOTE: Depending on the protocol to be used in subsequent FISH analysis, cells can be initially attached either to slides or coverslips. For the sake of brevity, only coverslips are mentioned in the protocol steps.
Additional Materials (also see Basic Protocol 1) Chang in situ medium (Irvine Scientific) supplemented with penicillin/streptomycin (100 U/ml penicillin/100 μg/ml streptomycin added from 100× stock; BioWhittaker) or Amniomax complete medium (Life Technologies) Additional reagents and equipment for in situ preparation, harvest, and culture of amniotic fluid samples (UNIT 8.4; Minehart Miron, 2012) NOTE: All incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. 1. Prepare amniotic fluid for culturing by performing steps 1 to 9 of the Basic Protocol in UNIT 8.4 (Minehart Miron, 2012), using either Chang in situ medium or Amniomax medium at step where plates/coverslips are set up. Note that neither medium contains FBS. Preparation of Amniocytes for Interphase FISH
Amniotic fluid obtained for routine chromosomal studies and established for in situ cultures can also be utilized for FISH analysis. There are several procedures for attaching amniotic
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fluid cells to coverslips and establishing in situ cultures; see UNIT 8.4 (Minehart Miron, 2012) for one method.
2. Place tissue culture plates with coverslips in incubator. Leave undisturbed for 24 hr to allow cells to attach firmly to the coverslips. 3. After 24 hr of incubation, gently add an additional 1.5 ml of the same medium (Chang or Amniomax) that is already in the culture. Continue incubation. 4. After an additional 24 to 48 hr of incubation, carefully remove plates from incubator. Gently remove medium from plate without tipping and add 2 ml room temperature hypotonic solution around edge of plate. Let stand 20 min at room temperature. 5. Gently add 2 ml of room temperature fixative to the hypotonic solution, adding dropwise around edge of plate. Let stand 5 min at room temperature, then gently aspirate and discard solution. 6. Gently add 2 ml of fresh room temperature fixative as in step 5. Let stand 20 min at room temperature, then gently remove and discard solution. Repeat this procedure two more times. 7. Gently add 2 ml of fresh room temperature fixative to the first plate and put a lid on it. Repeat for each plate. 8. Dry coverslips as in the Basic Protocol of UNIT 8.4 (Minehart Miron, 2012), steps 21 to 23. Coverslip drying time depends on the laboratory environment, especially humidity. Drying time is not as critical in analysis of interphase nuclei as in producing metaphases for banding. Coverslips may be used immediately for FISH analysis (see Basic Protocol 2 in this unit; also see UNIT 4.3; Knoll and Lichter, 2005) or stored at room temperature until the next day. They may be stored for longer periods of time at 4° or –20°C.
INTERPHASE FISH ANALYSIS OF AMNIOTIC FLUID CELLS Detailed protocols for fluorescence in situ hybridization are provided in UNIT 4.3 (Knoll and Lichter, 2005). A brief protocol is given here, especially for situations where FISH is used in a laboratory only for analysis of amniotic fluid cells.
BASIC PROTOCOL 2
FISH can be accomplished using commercial probes or probes labeled in the laboratory. The method outlined here is for DNA probes received from a commercial supplier that have already been directly labeled. Such probes may recognize α-satellite DNA repetitive sequences or unique sequences. A variety of probes can be utilized with only minor variations in conditions for hybridization, washing, and detection. For guidelines to determine the number of cells to be analyzed, see Critical Parameters and Troubleshooting. This protocol describes the FDA-approved approach, and as such, the steps presented are as given in the Abbott/Vysis AneuVysion product insert: see https://www .abbottmolecular.com/static/cms_workspace/pdfs/US/Package_Insert_04_02_2012.pdf.
Materials Denaturing solution (see recipe), 73°C70%, 85%, and 100% ethanol (see recipe for ethanol wash solutions), ice cold Slide or coverslip containing amniotic fluid cells (see Basic Protocol 1 or see Alternate Protocol)
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Probes (Vysis/Abbot): CEP 18 (alpha-satellite DNA probe) CEP X (alpha-satellite DNA probe) CEP Y (alpha-satellite DNA probe) LSI 13 LSI 21 Rubber cement 0.4× SSC/0.3% NP-40 solution (see recipe) 2× SSC/0.1% NP-40 solution (see recipe) 2 × SSC (APPENDIX 2D), 37°C Moist chamber (UNIT 4.3) Coplin jars pH meter or pH paper 73°C water bath Forceps 45° to 50°C slide warmer 22 × 22–mm glass coverslips 5-ml syringe DAPI II counterstain Fluorescence microscope with epi-illumination and filter set appropriate for fluorochrome used [(UNIT 4.3 (Knoll and Lichter, 2005) and UNIT 4.4 (McNamara et al., 2005)] Denaturation of specimen DNA 1. Prewarm the humidified hybridization chamber (moist chamber; an airtight container with a piece of damp blotting paper or paper towel approximately 1 in. × 3 in. taped to the side of the container) to 37 ± 2°C by placing it in the 37 ± 2°C incubator prior to slide preparation. 2. Add denaturing solution to Coplin jar and place in a 73 ± 1°C water bath for at least 30 min. Verify the solution temperature before use. This step is critical. The temperature of the denaturing solution must be 72° to 74°C; therefore, the temperature of the water bath must be raised 1°C for every additional slide placed in the Coplin jar.
3. Verify that the pH of the denaturing solution is 7.0 to 8.0 before each use. Denature the specimen DNA by immersing the prepared slides in the denaturing solution at 73 ± 1°C for 5 min. Do not denature more than four slides at one time per Coplin jar. 4. Using forceps, remove the slide(s) from the denaturing solution and immediately place into a 70% ethanol wash solution at room temperature. Agitate the slide to remove the formamide (a component of the denaturing solution). Allow the slide(s) to stand in the ethanol wash for 1 min. 5. Remove the slide(s) from 70% ethanol. Repeat step 4 with 85% ethanol, followed by 100% ethanol. 6. Drain the excess ethanol from the slide by touching the bottom edge of the slide to a blotter, and wipe the underside of the slide dry with a laboratory wipe. 7. Place the slide(s) on a 45 to 50°C slide warmer no more than 2 min before you are ready to apply the probe solution. Preparation of Amniocytes for Interphase FISH
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IMPORTANT NOTE: If the timing of the hybridization is such that the slide is ready more than 2 min before the probe is ready, the slide should remain in the jar of 100% ethanol. Do not air dry slide before placing it on the slide warmer.
Probe preparation 8. Allow the probe to warm to room temperature, thus decreasing the viscosity and allowing for accurate pipetting. 9. Vortex to mix. Spin the tube briefly (1 to 3 sec) in microcentrifuge to bring the contents to the bottom of the tube. Gently vortex again to mix. The commercially available probe from Vysis/Abbot is pre-denatured and ready to apply to the denatured target area on the specimen slide. The following steps are for DNA probes obtained from commercial sources (Vysis/Abbot). Refer to the product data sheet for any modifications of this procedure that are specific to a particular probe. See UNIT 4.3 (Knoll and Lichter, 2005) for details about working with different probes and protocols for labeling probes in the laboratory.
Hybridization The five probes listed in the materials list are applied in two mixes. Mix 1 consists of the centromere for chromosomes 18 (CEP 18), X (CEP X), and Y (CEP Y). Mix 2 consists of sequence specific probes on chromosomes 13 (LSI 13) and 21 (LSI 21). 10. Apply 10 μl of CEP 18/X/Y probe mix to one hybridization area on the slide(s) and 10 μl of LSI 13/21 probe mix to the other hybridization area on the slide(s). Immediately place a 22 mm × 22 mm glass coverslip over the probe solution and allow the solution to spread evenly under the coverslip. Air bubbles will interfere with hybridization and should be avoided. IMPORTANT NOTE: Do not pipet either probe solution onto multiple target areas before applying the coverslips. Each probe solution goes on once; neither should be put on multiple times.
11. Seal coverslip with rubber cement as follows. Draw the rubber cement into a 5-ml syringe, then eject a small amount of rubber cement around the periphery of the coverslip overlapping the coverslip and the slide, forming a seal around the coverslip. 12. Place the slide into the pre-warmed 37 ± 2°C hybridization chamber, cover the chamber with a tight lid, and incubate at 37 ± 2°C for 6 to 24 hr. Moist chambers in many laboratory are generally plastic containers (e.g., Tupperware or Rubbermaid) with two 1-ml pipets taped horizontally to the bottom. This allows the slide to be kept off the bottom of the container, which is lined with paper towels moistened with water or 2× SSC to maintain humid conditions. A typical moist chamber is illustrated in UNIT 4.3 (Knoll and Lichter, 2005); however, a variety of different containers can be utilized. 22 × 22–mm coverslips can be used effectively if the investigator wishes to hybridize two different probes on one slide. This makes it possible to use a greater number of probes with a minimum number of slides, but fewer nuclei will be available for analysis with each probe. Plastic coverslips are preferable to glass for hybridization, detection, and amplification steps, because they can be easily removed and replaced on the slide repeatedly. Glass coverslips are required for mounting. Although the procedure calls for overnight incubation, the time of hybridization can be as brief as 2 to 4 hr when repetitive DNA probes are utilized. Clinical Cytogenetics
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Post-hybridization washes 13. Add 0.4× SSC/0.3% NP-40 to a Coplin jar. Prewarm the 0.4× SSC/0.3% NP-40 solution by placing the Coplin jar in the 73 ± 1°C water bath for at least 30 min or until the solution temperature has reached 73 ± 1°C. The temperature of the wash solution must return to 73 ± 1°C before washing each batch.
14. Add 2× SSC/0.1% NP-40 to a second Coplin jar and place at room temperature. Discard both wash solutions after 1 day of use. 15. Remove rubber cement seal and coverslips. Immediately place the slide into the Coplin jar containing 0.4× SSC/0.3% NP-40 at 73 ± 1°C. Agitate the slide for approximately 3 sec. Repeat for the other slides, then incubate 2 min. Do not place more than four slides in the wash at one time. If more than four slides are to be washed, verify that the temperature of the wash solution is 73 ± 1°C before each use. 16. Remove each slide from the wash bath and place in the jar of 2× SSC/0.1% NP-40 at room temperature for 5 to 60 sec, agitating for 1 to 3 sec as the slides are placed in the bath. 17. Allow the slide to air-dry in the dark (a closed drawer or a shelf inside a closed cabinet is sufficient.) 18. Apply 10 μl of DAPI II counterstain to each target area of the slide and apply a glass coverslip. Store the slide(s) in the dark prior to signal enumeration. Store hybridized slides (with coverslips) at −20°C (±10°C) in the dark. Under these conditions, the slides can be stored for up to 12 months without significant loss in fluorescence signal intensity. For long-term storage, the coverslips may be sealed to prevent desiccation and the slides stored at −20°C (±10°C). BASIC PROTOCOL 3
SIGNAL ENUMERATION Assessing slide adequacy Evaluate slide adequacy using the following criteria: Probe signal intensity: The signal should be bright, distinct, and easily evaluable. Signals should be in either bright, compact, oval shapes or stringy, diffuse, oval shapes. Background: The background should appear dark or black and free of fluorescence particles or haziness. Cross-hybridization/target specificity: The probe should hybridize to and illuminate only the corresponding target DNA on the chromosome. On cultured specimens, metaphase spreads may be evaluated to identify any cross-hybridization to non-target sequences. At least 98% of cells should show one or more signals for acceptable hybridization (see guidelines for signal enumeration below). If any of the above features are unsatisfactory, troubleshoot and process a fresh slide.
Preparation of Amniocytes for Interphase FISH
Selection of best viewing area and evaluable nuclei Use a 25× objective to scan the hybridized area and examine the specimen distribution. Select an area where the specimen is distributed sparsely, few interphase nuclei or metaphase spreads are overlapping, and several interphase nuclei can be scanned within a viewing field. Avoid areas where the distribution of cells is dense, cells are overlapped, or the nuclear border of individual nuclei is unidentifiable. Avoid areas containing clumps of cells. Enumerate only those cells with discrete signals.
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Figure 8.9.1 Optimal and suboptimal amniocyte slides. Preparations from 17-gestational-week amniotic fluid stained with Giemsa stain to permit better nuclear visualization, showing: (A) too few cells; (B) good number of cells; (C) too many cells for optimal FISH analysis.
Enumeration scan Using a 40× or 63× objective, begin analysis in the upper left quadrant of the selected area and, scanning from left to right, count the number of signals in each evaluable metaphase spread or within the nuclear boundary of each evaluable interphase cell. Areas on the slide with a high cell density should be randomly skipped in order to scan the entire target area. Continue the scanning until 50 nuclei are enumerated and analyzed for each target. When mosaicism (10% to 60% aneuploid cells) is expected, 200 nuclei per target should be enumerated. A target with less than 50 evaluable nuclei should be either supplemented with an additional slide or considered an uninformative case.
Interphase enumeration Enumerate the fluorescent signals in each evaluable interphase nucleus using a 40× or 63× objective. Follow the signal-counting guidelines in Figures 8.9.1 and 8.9.2. Objectives with higher magnification (e.g., 63× or 100×) should be used to verify or resolve questions about split or diffused signals.
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Figure 8.9.2 Identification of aneuploidy by FISH. Patient was referred because of ultrasound indication of holoprosencephaly and abnormal triple-marker screen. (A) Hybridization with a chromosome 13/21 α-satellite repetitive probe revealed five signals in a majority of cells, indicating presence of an extra chromosome (either 13 or 21). Arrow indicates signal out of plane of focus. (B) Hybridization with chromosome 21 cosmid probe revealed two signals in a majority of cells, indicating that the fetus does not have trisomy 21. (C) Hybridization with a chromosome 13q telomeric probe revealed three signals in a majority of cells, indicating that fetus is trisomic for chromosome 13. Subsequent metaphase analysis revealed a 47,XX,+13 karyotype, confirming FISH results.
Two signals that are in close proximity and approximately the same sizes but not connected by a visible link are counted as two signals. Count a diffuse signal as one signal if diffusion of the signal is contiguous and within an acceptable boundary. Two small signals connected by a visible link are counted as one signal. For CEP 18, LSI 13, and LSI 21, enumerate the number of nuclei with 0, 1, 2, 3, 4, or >4 signals. Count only those nuclei with one or more FISH signal of any color. If the accuracy of enumeration is in doubt, repeat the enumeration in another area of the slide. Preparation of Amniocytes for Interphase FISH
For CEP X/Y, enumerate the number of nuclei with 0, 1, 2, 3, 4, or >4 signals (for both orange and green signals) and record the counts in a two-way table, and then calculate the percentage of nuclei with X, Y, XY, XX, XXY, XYY, XXX, and others.
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Count only those nuclei with one or more FISH signal of any color. If the accuracy of enumeration is in doubt, repeat the enumeration. REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2D.
Denaturing solution Mix: 49 ml formamide 7 ml 20× SSC pH 5.3 (APPENDIX 2D) 14 ml purified H2 O 70 ml final volume Mix well and place in a glass Coplin jar. Measure pH at room temperature using a pH meter with a glass pH electrode to verify that the pH is between 7.0 to 8.0. Store in a covered container at 2° to 8°C. This solution can be used for up to 1 week. Check pH prior to each use.
Ethanol wash solutions Prepare 70% (v/v), 85% (v/v), and 100% (v/v) ethanol using 100% ethanol and purified water. Add 70 ml of each solution to a Coplin jar and maintain at room temperature. Store unused dilutions in a covered container at room temperature for up to 6 months. Solutions used in the assay may be used for 1 week unless evaporation occurs or the solution becomes diluted due to excessive use.
NP-40, 0.1%, in 2× SSC Mix: 100 ml 20× SSC pH 5.3 (APPENDIX 2D) 849 ml purified H2 O 1 ml NP-40 (IGEPAL CA-630; Sigma-Aldrich, CAS no. 9002-93-1) Add H2 O to 1000 ml Mix thoroughly. Adjust the pH to 7.0 to 7.5 with 1 N NaOH using a pH meter and glass pH electrode to measure the pH. Adjust volume to 1 liter with water. Filter through 0.45-μm filtration unit. Add 70 ml of the solution to a Coplin jar and maintain at room temperature. Store unused solution in a covered container at room temperature for up to 6 months. Discard solution that was used in the assay at the end of each day.
NP-40, 0.3%, in 0.4× SSC Mix: 950 ml purified H2 O 20 ml 20× SSC pH 5.3 (APPENDIX 2D) 3 ml NP-40 (IGEPAL CA-630; Sigma-Aldrich, CAS no. 9002-93-1) Add H2 O to 1000 ml Mix thoroughly. Adjust pH to 7.0 to 7.5 with 1 N NaOH, if necessary, using a pH meter with glass pH electrode. Adjust volume to 1 liter with water. Filter through 0.45-μm filtration unit. Store unused solution in a covered container at room temperature for up to 6 months. Discard solution that was used in the assay at the end of each day.
Post-fixation solution To 48.7 ml of phosphate-buffered saline (PBS; APPENDIX 2D), add: 1.3 ml of 37% formaldehyde (0.95% final) 0.23 g MgCl2 Store up to 1 month at 4°C
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COMMENTARY Background Information
Preparation of Amniocytes for Interphase FISH
Fluorescence in situ hybridization (FISH) with repetitive-element and single-copy DNA probes, as well as with probes made from chromosome-specific libraries, has been utilized for analysis of both metaphase and interphase cells for over two decades now. FISH analysis has made possible studies of alterations of chromosome structure including detection and characterization of duplications, deletions, and/or cryptic translocations (Stumm and T¨onnies, 2008; Zuffardi et al., 2009; Tsuchiya, 2011; Bubendorf and Piaton, 2012; Haferlach, 2012; Jehan et al., 2012; Das and Tan, 2013; Lampert et al., 2013; Riegel, 2014); determination of the chromosomal origin of supernumerary markers (Rodr´ıguez L et al., 2007); and detection of aneuploidy in interphase cells. Detection of aneuploidy by FISH in interphase cells has been applied widely to neoplastic conditions (e.g., the study of alterations in chronic lymphocytic leukemia, myelodyplasia, solid tumors, and acute lymphocytic leukemia, among others; Stumm and T¨onnies, 2008; Tsuchiya, 2011; Bubendorf and Piaton, 2012; Haferlach, 2012; Jehan et al., 2012; Das and Tan, 2013; Lampert et al., 2013). This technology has been utilized to detect aneuploidies in blood smears from newborn infants as well as extensively in amniotic fluid samples and CVS samples (Ward et al., 1993; Tepperberg et al., 2001; Weise and Liehr, 2008). There is one major method and an alternative method of preparing amniotic fluid cells for detection of aneuploidy by FISH. The first method (Basic Protocol 1) uses uncultured cells obtained from amniotic fluid samples. The second method (Alternate Protocol) uses cells that have been placed on a coverslip or slide and allowed a short period of time (up to 48 hr) for attachment. There have been numerous studies published and reviews analyzing the use of the FISH for prenatal diagnosis in interphase cells (Ward et al., 1993; Tepperberg et al., 2001; Weise and Liehr, 2008; Faas et al., 2011). The majority of studies utilized a variation of the basic protocol and analyzed the cells within 2 days of receipt. Results from over 29,000 amniotic fluid samples summarized, after initial use of the protocol, revealed only one false-positive (0.003%) and seven false-negative results (0.024%), demonstrating the efficacy of this technology (Tepperberg et al., 2001). Although there have been isolated instances of false-positive and false-
negative results, this technology has proven to be robust over time. It was initially argued that this technique could be most effectively used for patients with a higher probability of an aneuploid fetus (e.g., with abnormal ultrasonographic findings, maternal age >40 years, or positive maternal serum screen), rather than for every individual who comes to a prenatal diagnostic program. Advantages to analysis based on increased risk of aneuploidy are that if a particular abnormality is suspected, only one probe may be needed. This will save on the amount of specimen needed, as well as on additional analyses to be performed. This in turn will lower the cost of carrying out FISH, which can be both expensive and time-consuming if five different probes must be utilized. However, today the technology is used mostly as an adjunctive procedure for all patients that request testing to facilitate the reporting of results more rapidly, usually within 24 hr. Invariably, five probes are utilized rather than just one.
Critical Parameters and Troubleshooting Cell preparation Preparation of the cells is the most difficult aspect of FISH with amniotic fluid. Preparation of uncultured amniocytes (Basic Protocol 1) is easier, quicker, and much more readily used than preparation of amniocytes attached to coverslips (Alternate Protocol). Over the past decade, this technology has been optimized, and investigators have used different hypotonic solutions, hypotonic treatment times, and fixative solutions than those mentioned here. The optimization of this technology has evolved over the years, leading to fairly standard protocols in most laboratories. However, because technologies are so varied, each laboratory will ultimately determine what works best in their individual laboratory. A successful analysis can be done even in cases where only 1 ml of amniotic fluid is obtained from an 11- to 15-gestational-week pregnancy. In general, however, 2 to 3 ml of amniotic fluid is needed at 12 to 15 weeks; 3 to 5 ml at 14 to 19 weeks; and 5 ml at 20 weeks. These numbers may be slightly conservative, as on the average 3 ml provides successful results. Although more cells are available in amniotic fluid from 26 gestational-week pregnancies, the majority are dead cells. Therefore, additional amniotic fluid may be needed in these cases.
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A possible advantage of the Alternate Protocol is that the morphology of the cells changes after attachment, making them more permeable and thus increasing hybridization efficiency. The procedure described in Klinger et al. (1992), which uses cells attached to slides, does not designate any extra incubation time to allow for cell attachment. However, most laboratories have not been able to reproduce the results of Klinger et al. (1992) using that methodology, but laboratories have had successful hybridization after allowing attachment for 24 hr, and the Alternate Protocol has been designed with that in mind. Several factors affect hybridization of amniotic fluid cells. In general, cells from amniotic fluid received late in gestation (>25 weeks) will hybridize poorly, and the results will be much more difficult to analyze. In such samples, there is usually an elevated number of cells that are dead and will not hybridize, as well as more debris, which makes hybridization less efficient. In most cases, cells from as early as 12 gestational weeks can be used for FISH studies. The difficulty with early-gestation specimens is that there are not enough cells to complete the proper studies. Bloody specimens can also present a problem, which can usually be alleviated by fixation, as the fixative will hemolyze the blood. When uncultured amniotic fluid cells are spun down, red blood cells will be present in the centrifuge tube and possibly on the slide. Because these are anucleate, they will not hybridize with a probe, but their presence may lead to cellular crowding, which results in an overall lower number of cells demonstrating hybridization. Another possible complication in prenatal FISH studies is maternal-cell admixture. Presence of any maternal cells can lead to false conclusions. Therefore, it is important to know whether the sample may be at higher risk for having maternal cells (e.g., whether the specimen was bloody or was obtained through the anterior placenta). It should also be noted in the analysis whether any maternal cells appear to be present. A few studies have evaluated the presence of maternal cells in amniotic fluid samples. Although some of these studies suggest a higher frequency of contamination than others, it is clear that contamination does not frequently cause problems with the use of these protocols (Nub et al., 1994; Rebello et al., 1994). It is very important for every laboratory to determine the exact extent of maternal-cell contamination in their samples based on use of both an X and Y probe in cells from a male fetus; however, the
admixture of maternal cells will vary from one specimen to the next. When there is a hint of the possibility of red cells present, it is advisable to count 75 interphase cells rather than 50 cells as is usually counted. FISH Recommendations from the laboratories utilizing these protocols suggest that the protocol may have to be modified to take into account the hybridization efficiency of the probe and number of cells showing a positive signal, as well as the possibility of maternal cell contamination. Although considerable troubleshooting may be necessary with FISH protocols (see UNIT 4.3; Knoll and Lichter, 2005), the probes used for this analysis work relatively well with little modification. Clinical guidelines The American College of Medical Genetics and the American Society of Human Genetics (ACMG/ASHG) published a position paper in 2000 on the technical and clinical assessment of fluorescence in situ hybridization. This included a section on evaluating the results of FISH on interphase cells in prenatal diagnosis (Test and Technology Transfer Committee, 2000). This position statement includes the following: (1) those requesting interphase FISH testing for prenatal diagnosis should be fully aware of what these focused tests can and cannot do (clinical sensitivity and specificity) including false positive and false negatives; (2) while the speed of diagnosis with this technology is a distinct advantage, the increased cost of FISH must be considered; (3) for management of the fetus, it is reasonable to report positive FISH test results; (4) clinical decision-making should be based on information from two of three of the following— FISH results, confirmatory chromosome analysis, or consistent clinical information; (5) for ultimate management of pregnancies that are FISH positive, these results need to be further characterized by traditional chromosome analysis to determine the mechanism (e.g., aneuploidy or translocation) accounting for the FISH detected abnormality. An overview of the validations and guidelines for the clinical diagnostic use of interphase FISH analysis is given in Section E10 of the American College of Medical Genetics Standards and Guidelines for Clinical Genetics Laboratories (1/2010; https://www.acmg.net/StaticContent/SGs/ Section_E_2011.pdf).
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Figure 8.9.3 Distinct and split signals. Hybridization of cells with chromosome 18 α-satellite probe showing: (A) two signals; (B) two split signals each of which is considered one signal by criteria used in authors’ laboratory.
Preparation of Amniocytes for Interphase FISH
FISH analysis of interphase will always yield higher results based on the ascertainment of patients, specifically in patients with advanced maternal age and specific ultrasound findings. However, once an invasive technique has been done, this technology becomes available to all patients and is often used to provide more rapid results. There are some general rules that should be considered when performing prenatal interphase FISH. Optimally, 50 interphase cells should be analyzed for each probe used (25 counts from two different readers). Because FISH is not usually used in any case as a standalone procedure, analysis of 25 cells is acceptable, though not optimal (and analysis of fewer than 25 is unacceptable). If cell-preparation techniques have been optimized, 70% to 98% of all interphase cells should show hybridization of each probe. Each laboratory should develop its own guidelines regarding acceptable frequency of signals for each probe, but in general 60% of the cells should show an abnormal number of signals for a sample to be considered aneuploid (e.g., three signals for an 18 centromere probe). Usually, detection of an abnormal number of signals in 30% of cells is within experimental error. Cases with 10% but 60% of the cells demonstrating abnormal signals need to be analyzed
further. Controls need not be used for each hybridization if multiple slides are analyzed; however, no conclusions concerning absence of the Y chromosome can be made without a positive signal in another sample. Each time a new lot of probe is utilized, it should be tested on known control slides. Consistency in scoring interphase signals is of the utmost importance; this can be accomplished over time and after multiple evaluations. Because of the three-dimensional nature of interphase cells, not all cells will always be in the same plane of focus. Therefore, in order to detect signals, it is necessary to focus up and down to detect any signals outside the focal plane. Focusing up and down also helps delineate nuclear boundaries in preparations with extensive nuclear overlap. Split signals are in general counted as one signal (Fig. 8.9.3), unless a distance greater than the size of a signal lies between the two spots comprising the putative split signal. In those cases, the signals are counted as two separate signals. Hybridized specimens can be initially scanned (and analyzed in some cases) using a 40× objective. Often, initial analysis is accomplished with either a 63× or 100× objective; when a 40× objective is used, these more powerful objectives can be utilized to clarify any questions that may arise.
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Anticipated Results After establishment of these techniques within the laboratory, analysis should be successful >95% of the time (for both singlecopy and repetitive probes, once the technique has been optimized). However, if samples obtained at a later gestational age are studied, the success rate may be lower because of lower hybridization efficiency. It is crucial for each laboratory to develop its own internal guidelines and conditions for optimal hybridization and analysis. Hybridization efficiencies using samples prepared by either method should be similar once the methods have been optimized for an individual laboratory. To avoid inaccuracies arising from the presence of large numbers of dead cells (e.g., in specimens obtained at advanced maternal age), cells that show no signal are placed in the analysis category “did not hybridize.” These cells are included when determining hybridization efficiency but are not included in the actual analysis. It is possible to get one to three slides with 50 cells each from a 1-ml sample. It is not always advisable to take a specimen from a sample containing
UNIT 8.9
Stuart Schwartz1 1
R Cytogenetics Laboratory, Laboratory Corporation of America Holdings, Research Triangle Park, North Carolina
FISH has been used to detect and clarify deletions and/or other structural rearrangements, and also has applications in interphase analysis. This unit describes preparation of uncultured amniotic fluid cells for FISH analysis. Cells are swollen, and then slides are prepared by standard methods. The cells are then fixed and permeabilized for subsequent FISH. An alternate protocol describes attachment of amniocytes to a glass or plastic surface followed by hypotonic swelling, fixation, and permeabilization for subsequent FISH. Interphase FISH C 2015 by John Wiley & Sons, analysis of amniotic fluid cells is also described. Inc. Keywords: prenatal diagnosis r FISH r fluorescence in situ hybridization r amniotic fluid r aneuploidy
How to cite this article: Schwartz, S. 2015. Preparation of amniocytes for interphase fluorescence in situ hybridization (FISH). Curr. Protoc. Hum. Genet. 85:8.9.1-8.9.16. doi: 10.1002/0471142905.hg0809s85
INTRODUCTION Use of fluorescence in situ hybridization (FISH) has increased by several orders of magnitude over the past two decades. In a clinical setting, the technique is used most commonly as an adjunct to standard cytogenetic analysis, but can be a stand-alone procedure. Although FISH has been used to detect or clarify gains, losses, and/or structural rearrangements, considerable work has focused on its application in interphase analysis. Most of this work is currently associated with the analysis of leukemia and solid tumors where karyotyping has been difficult or no material (e.g., paraffin sections) is available for karyotype analysis. Interphase analysis has now been utilized extensively for prenatal diagnosis. Although initial controversy surrounded the use of FISH as an investigational procedure for this purpose, there are both recommendations (e.g., from the American College of Medical Genetics) for its utilization and available FDA-approved probes. The types of information that FISH provides, combined with easy utilization and rapid turnaround time, make FISH an attractive adjunct to cytogenetics. Whereas classical cytogenetic methods yield a diagnosis from amniotic fluid cells in 7 to 12 days, interphase analysis by FISH permits detection of specific aneuploidies in 48 hr. Basic Protocol 1 describes preparation of uncultured amniotic fluid cells for FISH analysis. Cells are swollen, and then slides are prepared using standard methods. These are then fixed and permeabilized for subsequent FISH. The Alternate Protocol describes Clinical Cytogenetics Current Protocols in Human Genetics 8.9.1-8.9.16, April 2015 Published online April 2015 in Wiley Online Library (wileyonlinelibrary.com). doi: 10.1002/0471142905.hg0809s85 C 2015 John Wiley & Sons, Inc. Copyright
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attachment of amniocytes to a glass or plastic surface followed by hypotonic swelling, fixation, and permeabilization for subsequent FISH. In contrast to routine cytogenetic procedures for culture of amniotic fluid cells (UNIT 8.4; Minehart Miron, 2012), cells are incubated just long enough for them to attach to the surface, a much quicker procedure. The purpose is to obtain interphase cells attached to plastic coverslips (or glass slides); no metaphase spreads are expected. This procedure is more time-consuming than Basic Protocol 1, but may allow for increased hybridization efficiency because dead cells will not attach to the slides and because changes in the morphology of the cells when they attach make them more permeable to probes; however, it is rarely used today. Two protocols have been given but many exist, and it is incumbent on each laboratory to determine what works best in their laboratory. Basic Protocol 2 for FISH can be used for interphase cells obtained by any of the protocols. The protocol can be utilized with the FDA-approved probe set, or with any repetitiveDNA probes, unique probes, or single-copy probes (e.g., commercially available or from BAC libraries). The FISH protocol described can be used with directly labeled DNA probes (see UNIT 4.3; Knoll and Lichter, 2005, for a more detailed discussion of FISH). CAUTION: Radioactive, biological, and chemical substances require special handling; see APPENDIX 2 for guidelines. BASIC PROTOCOL 1
PREPARATION OF UNCULTURED AMNIOCYTES FOR INTERPHASE FISH ANALYSIS In this procedure, amniotic fluid cells are obtained, washed, swollen in hypotonic solution, and spread onto slides by standard methods. After fixation the slides are used in FISH (see Basic Protocol 2 and UNIT 4.3; Knoll and Lichter, 2005) and analyzed by fluorescence microscopy (UNIT 4.4; McNamara et al., 2005). Unlike the methods in UNIT 8.4 (Minehart Miron, 2012), this protocol does not involve culturing of cells, but can be done on cultured cells if needed. This protocol describes the FDA-approved approach, and as such, the steps presented are as given in the Abbott/Vysis AneuVysion product insert; see https://www.abbott molecular.com/static/cms_workspace/pdfs/US/Package_Insert_04_02_2012.pdf.
Materials Whole amniotic fluid specimen 1× trypsin/EDTA (0.05% trypsin/0.53 mM EDTA·4Na in Hanks’ Balanced Salt Solution without or Mg) Hypotonic solution: 0.56% (w/v) KCl Fixative: 3:1 (v/v) methanol:glacial acetic acid 2× SSC, pH 5.3 (APPENDIX 2D) Pepsin working solution: 2.5 mg pepsin added to 50 ml of 0.01 N HCl Phosphate-buffered saline (PBS; APPENDIX 2D) Post-fixation solution (see recipe) 70%, 85%, and 100% ethanol (see recipe for ethanol wash solutions) 15-ml screw-cap conical centrifuge tubes (e.g., BD Falcon) Clinical tabletop centrifuge (e.g., IEC Centra-HN) Glass microscope slides, pre-cooled Cotton-plugged Pasteur pipets Preparation of Amniocytes for Interphase FISH
Additional reagents and equipment for chromosome slide preparation (UNIT 4.1; Bangs and Donlon, 2005)
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Slide preparation from uncultured amniotic fluid 1. Centrifuge 2 to 5 ml of whole amniotic fluid specimen in a 15-ml centrifuge tube, for 5 min at 200 × g, room temperature. The specimen should not appear bloody or brown. Samples should normally be processed the day the amniotic fluid is received, as rapidity is the essence of this procedure. If necessary, however, samples can be successfully prepared several days after the fluid has been obtained. If not used immediately, amniotic fluid should be stored at room temperature in the dark. Because no cell growth is involved, samples that have been refrigerated or exposed to heat can still be used successfully. The amount of fluid to be used for this test depends on the gestational age at which amniocentesis is performed and the amount of fluid received by the cytogenetics laboratory. See Critical Parameters and Troubleshooting for guidelines.
2. Resuspend the pellet in 2 to 5 ml of 1× trypsin/EDTA and incubate in a 37 ± 1°C water bath for at least 15 min. 3. Centrifuge the suspension for 5 min at 200 × g, room temperature. 4. Resuspend the pellet in 2 to 5 ml of 0.56% KCl and incubate for 20 min in a 37 ± 1°C water bath. 5. Add 0.8 to 2 ml of fixative to the cells/hypotonic solution and vortex gently. 6. Centrifuge the suspension for 5 min at 200 × g, room temperature, and resuspend the pellet in 1 ml fresh fixative. Store fixed specimens at 4°C for at least 30 min or until ready to perform FISH. For long-term storage, keep fixed specimens at −20°C (±10°C) in fixative. 7. Before placing cells on slides, adjust volume of cell suspension according to size of cell pellet by adding additional fixative. If needed, particularly after a prolonged storage (>1 month), wash pellets with fixative by repeating step 6 before slide preparation. 8. To prepare slides for FISH, drop the cell suspension directly onto one or two cold glass slides, making two hybridization areas (15 to 25 μl of cell suspension per area). See UNIT 4.1 (Bangs and Donlon, 2005), Support Protocol, for details of slide making. Achieving the proper concentration for slide making requires practice and can only be arrived at through trial and error. Slides should contain enough nuclei for analysis. In general, one to three slides can be made from each sample, depending on cell concentration.
9. Proceed to steps 10 to 18, if desired. If the optional pretreatment procedure will not be used, age the specimen slides in 2× SSC for 30 min to 1 hr at 37°C or at room temperature for 24 hr, with the slide box uncovered, before hybridization or storage. Slides may be used immediately for FISH analysis (see Basic Protocol 2 and UNIT 4.3; Knoll and Lichter, 2005), or stored at room temperature until the next day. They may be stored for longer periods at 4° or –20°C.
Pretreatment for uncultured amniotic fluid cells (optional) The following method is recommended in order to achieve optimum FISH results, especially for late-gestational-age specimens and specimens to be hybridized immediately following slide preparation. 10. Place slide(s) prepared from uncultured amniocytes, in 2× SSC for 1 hr at 37 ± 1°C.
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11. Place slide(s) in freshly made pepsin working solution for 13 min at 37 ± 1°C. 12. Rinse slide(s) in PBS at room temperature for 5 min. 13. Place slide(s) in post-fixation solution for 5 min. 14. Rinse slide(s) in PBS at room temperature for 5 min. 15. Dry slide(s). To speed drying, use a cotton-plugged Pasteur pipet to direct compressed air flow onto the slide. 16. Immerse slide(s) in 70% ethanol at room temperature. Allow the slide(s) to stand in the ethanol wash for 1 min. 17. Remove the slide(s) from 70% ethanol. Repeat step 7 with 85% ethanol, followed by 100% ethanol. 18. Proceed to denaturation of slides(s) as described in Basic Protocol 2. ALTERNATE PROTOCOL
PREPARATION OF AMNIOCYTES ATTACHED TO A SURFACE FOR INTERPHASE FISH ANALYSIS This protocol describes preparation of amniocytes attached to coverslips or slides. These cells are swelled in hypotonic medium and fixed for subsequent FISH analysis. This method is in some cases preferable to that of Basic Protocol 1 in that dead cells will not attach to a coverslip (or slide), so that overall efficiency of the procedure is increased. When cells prepared by this methodology are analyzed by FISH, better hybridization efficiency between probes and interphase chromosomes is observed. This may be due to an increase in surface area available for hybridization, an increase in permeability of membranes, or other factors. Because some time may be required for attachment of cells to the coverslip and, in some cases, for initial growth, turnaround time may be longer with this protocol than with Basic Protocol 1. However, this protocol is rarely used today. A number of variations to this method are given in annotations to the steps. Regardless of variations, the major factor in this protocol is reliance on cellular attachment factors to increase the number of living cells that will attach to the coverslip or slides. NOTE: Depending on the protocol to be used in subsequent FISH analysis, cells can be initially attached either to slides or coverslips. For the sake of brevity, only coverslips are mentioned in the protocol steps.
Additional Materials (also see Basic Protocol 1) Chang in situ medium (Irvine Scientific) supplemented with penicillin/streptomycin (100 U/ml penicillin/100 μg/ml streptomycin added from 100× stock; BioWhittaker) or Amniomax complete medium (Life Technologies) Additional reagents and equipment for in situ preparation, harvest, and culture of amniotic fluid samples (UNIT 8.4; Minehart Miron, 2012) NOTE: All incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. 1. Prepare amniotic fluid for culturing by performing steps 1 to 9 of the Basic Protocol in UNIT 8.4 (Minehart Miron, 2012), using either Chang in situ medium or Amniomax medium at step where plates/coverslips are set up. Note that neither medium contains FBS. Preparation of Amniocytes for Interphase FISH
Amniotic fluid obtained for routine chromosomal studies and established for in situ cultures can also be utilized for FISH analysis. There are several procedures for attaching amniotic
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fluid cells to coverslips and establishing in situ cultures; see UNIT 8.4 (Minehart Miron, 2012) for one method.
2. Place tissue culture plates with coverslips in incubator. Leave undisturbed for 24 hr to allow cells to attach firmly to the coverslips. 3. After 24 hr of incubation, gently add an additional 1.5 ml of the same medium (Chang or Amniomax) that is already in the culture. Continue incubation. 4. After an additional 24 to 48 hr of incubation, carefully remove plates from incubator. Gently remove medium from plate without tipping and add 2 ml room temperature hypotonic solution around edge of plate. Let stand 20 min at room temperature. 5. Gently add 2 ml of room temperature fixative to the hypotonic solution, adding dropwise around edge of plate. Let stand 5 min at room temperature, then gently aspirate and discard solution. 6. Gently add 2 ml of fresh room temperature fixative as in step 5. Let stand 20 min at room temperature, then gently remove and discard solution. Repeat this procedure two more times. 7. Gently add 2 ml of fresh room temperature fixative to the first plate and put a lid on it. Repeat for each plate. 8. Dry coverslips as in the Basic Protocol of UNIT 8.4 (Minehart Miron, 2012), steps 21 to 23. Coverslip drying time depends on the laboratory environment, especially humidity. Drying time is not as critical in analysis of interphase nuclei as in producing metaphases for banding. Coverslips may be used immediately for FISH analysis (see Basic Protocol 2 in this unit; also see UNIT 4.3; Knoll and Lichter, 2005) or stored at room temperature until the next day. They may be stored for longer periods of time at 4° or –20°C.
INTERPHASE FISH ANALYSIS OF AMNIOTIC FLUID CELLS Detailed protocols for fluorescence in situ hybridization are provided in UNIT 4.3 (Knoll and Lichter, 2005). A brief protocol is given here, especially for situations where FISH is used in a laboratory only for analysis of amniotic fluid cells.
BASIC PROTOCOL 2
FISH can be accomplished using commercial probes or probes labeled in the laboratory. The method outlined here is for DNA probes received from a commercial supplier that have already been directly labeled. Such probes may recognize α-satellite DNA repetitive sequences or unique sequences. A variety of probes can be utilized with only minor variations in conditions for hybridization, washing, and detection. For guidelines to determine the number of cells to be analyzed, see Critical Parameters and Troubleshooting. This protocol describes the FDA-approved approach, and as such, the steps presented are as given in the Abbott/Vysis AneuVysion product insert: see https://www .abbottmolecular.com/static/cms_workspace/pdfs/US/Package_Insert_04_02_2012.pdf.
Materials Denaturing solution (see recipe), 73°C70%, 85%, and 100% ethanol (see recipe for ethanol wash solutions), ice cold Slide or coverslip containing amniotic fluid cells (see Basic Protocol 1 or see Alternate Protocol)
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Probes (Vysis/Abbot): CEP 18 (alpha-satellite DNA probe) CEP X (alpha-satellite DNA probe) CEP Y (alpha-satellite DNA probe) LSI 13 LSI 21 Rubber cement 0.4× SSC/0.3% NP-40 solution (see recipe) 2× SSC/0.1% NP-40 solution (see recipe) 2 × SSC (APPENDIX 2D), 37°C Moist chamber (UNIT 4.3) Coplin jars pH meter or pH paper 73°C water bath Forceps 45° to 50°C slide warmer 22 × 22–mm glass coverslips 5-ml syringe DAPI II counterstain Fluorescence microscope with epi-illumination and filter set appropriate for fluorochrome used [(UNIT 4.3 (Knoll and Lichter, 2005) and UNIT 4.4 (McNamara et al., 2005)] Denaturation of specimen DNA 1. Prewarm the humidified hybridization chamber (moist chamber; an airtight container with a piece of damp blotting paper or paper towel approximately 1 in. × 3 in. taped to the side of the container) to 37 ± 2°C by placing it in the 37 ± 2°C incubator prior to slide preparation. 2. Add denaturing solution to Coplin jar and place in a 73 ± 1°C water bath for at least 30 min. Verify the solution temperature before use. This step is critical. The temperature of the denaturing solution must be 72° to 74°C; therefore, the temperature of the water bath must be raised 1°C for every additional slide placed in the Coplin jar.
3. Verify that the pH of the denaturing solution is 7.0 to 8.0 before each use. Denature the specimen DNA by immersing the prepared slides in the denaturing solution at 73 ± 1°C for 5 min. Do not denature more than four slides at one time per Coplin jar. 4. Using forceps, remove the slide(s) from the denaturing solution and immediately place into a 70% ethanol wash solution at room temperature. Agitate the slide to remove the formamide (a component of the denaturing solution). Allow the slide(s) to stand in the ethanol wash for 1 min. 5. Remove the slide(s) from 70% ethanol. Repeat step 4 with 85% ethanol, followed by 100% ethanol. 6. Drain the excess ethanol from the slide by touching the bottom edge of the slide to a blotter, and wipe the underside of the slide dry with a laboratory wipe. 7. Place the slide(s) on a 45 to 50°C slide warmer no more than 2 min before you are ready to apply the probe solution. Preparation of Amniocytes for Interphase FISH
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IMPORTANT NOTE: If the timing of the hybridization is such that the slide is ready more than 2 min before the probe is ready, the slide should remain in the jar of 100% ethanol. Do not air dry slide before placing it on the slide warmer.
Probe preparation 8. Allow the probe to warm to room temperature, thus decreasing the viscosity and allowing for accurate pipetting. 9. Vortex to mix. Spin the tube briefly (1 to 3 sec) in microcentrifuge to bring the contents to the bottom of the tube. Gently vortex again to mix. The commercially available probe from Vysis/Abbot is pre-denatured and ready to apply to the denatured target area on the specimen slide. The following steps are for DNA probes obtained from commercial sources (Vysis/Abbot). Refer to the product data sheet for any modifications of this procedure that are specific to a particular probe. See UNIT 4.3 (Knoll and Lichter, 2005) for details about working with different probes and protocols for labeling probes in the laboratory.
Hybridization The five probes listed in the materials list are applied in two mixes. Mix 1 consists of the centromere for chromosomes 18 (CEP 18), X (CEP X), and Y (CEP Y). Mix 2 consists of sequence specific probes on chromosomes 13 (LSI 13) and 21 (LSI 21). 10. Apply 10 μl of CEP 18/X/Y probe mix to one hybridization area on the slide(s) and 10 μl of LSI 13/21 probe mix to the other hybridization area on the slide(s). Immediately place a 22 mm × 22 mm glass coverslip over the probe solution and allow the solution to spread evenly under the coverslip. Air bubbles will interfere with hybridization and should be avoided. IMPORTANT NOTE: Do not pipet either probe solution onto multiple target areas before applying the coverslips. Each probe solution goes on once; neither should be put on multiple times.
11. Seal coverslip with rubber cement as follows. Draw the rubber cement into a 5-ml syringe, then eject a small amount of rubber cement around the periphery of the coverslip overlapping the coverslip and the slide, forming a seal around the coverslip. 12. Place the slide into the pre-warmed 37 ± 2°C hybridization chamber, cover the chamber with a tight lid, and incubate at 37 ± 2°C for 6 to 24 hr. Moist chambers in many laboratory are generally plastic containers (e.g., Tupperware or Rubbermaid) with two 1-ml pipets taped horizontally to the bottom. This allows the slide to be kept off the bottom of the container, which is lined with paper towels moistened with water or 2× SSC to maintain humid conditions. A typical moist chamber is illustrated in UNIT 4.3 (Knoll and Lichter, 2005); however, a variety of different containers can be utilized. 22 × 22–mm coverslips can be used effectively if the investigator wishes to hybridize two different probes on one slide. This makes it possible to use a greater number of probes with a minimum number of slides, but fewer nuclei will be available for analysis with each probe. Plastic coverslips are preferable to glass for hybridization, detection, and amplification steps, because they can be easily removed and replaced on the slide repeatedly. Glass coverslips are required for mounting. Although the procedure calls for overnight incubation, the time of hybridization can be as brief as 2 to 4 hr when repetitive DNA probes are utilized. Clinical Cytogenetics
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Post-hybridization washes 13. Add 0.4× SSC/0.3% NP-40 to a Coplin jar. Prewarm the 0.4× SSC/0.3% NP-40 solution by placing the Coplin jar in the 73 ± 1°C water bath for at least 30 min or until the solution temperature has reached 73 ± 1°C. The temperature of the wash solution must return to 73 ± 1°C before washing each batch.
14. Add 2× SSC/0.1% NP-40 to a second Coplin jar and place at room temperature. Discard both wash solutions after 1 day of use. 15. Remove rubber cement seal and coverslips. Immediately place the slide into the Coplin jar containing 0.4× SSC/0.3% NP-40 at 73 ± 1°C. Agitate the slide for approximately 3 sec. Repeat for the other slides, then incubate 2 min. Do not place more than four slides in the wash at one time. If more than four slides are to be washed, verify that the temperature of the wash solution is 73 ± 1°C before each use. 16. Remove each slide from the wash bath and place in the jar of 2× SSC/0.1% NP-40 at room temperature for 5 to 60 sec, agitating for 1 to 3 sec as the slides are placed in the bath. 17. Allow the slide to air-dry in the dark (a closed drawer or a shelf inside a closed cabinet is sufficient.) 18. Apply 10 μl of DAPI II counterstain to each target area of the slide and apply a glass coverslip. Store the slide(s) in the dark prior to signal enumeration. Store hybridized slides (with coverslips) at −20°C (±10°C) in the dark. Under these conditions, the slides can be stored for up to 12 months without significant loss in fluorescence signal intensity. For long-term storage, the coverslips may be sealed to prevent desiccation and the slides stored at −20°C (±10°C). BASIC PROTOCOL 3
SIGNAL ENUMERATION Assessing slide adequacy Evaluate slide adequacy using the following criteria: Probe signal intensity: The signal should be bright, distinct, and easily evaluable. Signals should be in either bright, compact, oval shapes or stringy, diffuse, oval shapes. Background: The background should appear dark or black and free of fluorescence particles or haziness. Cross-hybridization/target specificity: The probe should hybridize to and illuminate only the corresponding target DNA on the chromosome. On cultured specimens, metaphase spreads may be evaluated to identify any cross-hybridization to non-target sequences. At least 98% of cells should show one or more signals for acceptable hybridization (see guidelines for signal enumeration below). If any of the above features are unsatisfactory, troubleshoot and process a fresh slide.
Preparation of Amniocytes for Interphase FISH
Selection of best viewing area and evaluable nuclei Use a 25× objective to scan the hybridized area and examine the specimen distribution. Select an area where the specimen is distributed sparsely, few interphase nuclei or metaphase spreads are overlapping, and several interphase nuclei can be scanned within a viewing field. Avoid areas where the distribution of cells is dense, cells are overlapped, or the nuclear border of individual nuclei is unidentifiable. Avoid areas containing clumps of cells. Enumerate only those cells with discrete signals.
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Figure 8.9.1 Optimal and suboptimal amniocyte slides. Preparations from 17-gestational-week amniotic fluid stained with Giemsa stain to permit better nuclear visualization, showing: (A) too few cells; (B) good number of cells; (C) too many cells for optimal FISH analysis.
Enumeration scan Using a 40× or 63× objective, begin analysis in the upper left quadrant of the selected area and, scanning from left to right, count the number of signals in each evaluable metaphase spread or within the nuclear boundary of each evaluable interphase cell. Areas on the slide with a high cell density should be randomly skipped in order to scan the entire target area. Continue the scanning until 50 nuclei are enumerated and analyzed for each target. When mosaicism (10% to 60% aneuploid cells) is expected, 200 nuclei per target should be enumerated. A target with less than 50 evaluable nuclei should be either supplemented with an additional slide or considered an uninformative case.
Interphase enumeration Enumerate the fluorescent signals in each evaluable interphase nucleus using a 40× or 63× objective. Follow the signal-counting guidelines in Figures 8.9.1 and 8.9.2. Objectives with higher magnification (e.g., 63× or 100×) should be used to verify or resolve questions about split or diffused signals.
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Figure 8.9.2 Identification of aneuploidy by FISH. Patient was referred because of ultrasound indication of holoprosencephaly and abnormal triple-marker screen. (A) Hybridization with a chromosome 13/21 α-satellite repetitive probe revealed five signals in a majority of cells, indicating presence of an extra chromosome (either 13 or 21). Arrow indicates signal out of plane of focus. (B) Hybridization with chromosome 21 cosmid probe revealed two signals in a majority of cells, indicating that the fetus does not have trisomy 21. (C) Hybridization with a chromosome 13q telomeric probe revealed three signals in a majority of cells, indicating that fetus is trisomic for chromosome 13. Subsequent metaphase analysis revealed a 47,XX,+13 karyotype, confirming FISH results.
Two signals that are in close proximity and approximately the same sizes but not connected by a visible link are counted as two signals. Count a diffuse signal as one signal if diffusion of the signal is contiguous and within an acceptable boundary. Two small signals connected by a visible link are counted as one signal. For CEP 18, LSI 13, and LSI 21, enumerate the number of nuclei with 0, 1, 2, 3, 4, or >4 signals. Count only those nuclei with one or more FISH signal of any color. If the accuracy of enumeration is in doubt, repeat the enumeration in another area of the slide. Preparation of Amniocytes for Interphase FISH
For CEP X/Y, enumerate the number of nuclei with 0, 1, 2, 3, 4, or >4 signals (for both orange and green signals) and record the counts in a two-way table, and then calculate the percentage of nuclei with X, Y, XY, XX, XXY, XYY, XXX, and others.
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Count only those nuclei with one or more FISH signal of any color. If the accuracy of enumeration is in doubt, repeat the enumeration. REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2D.
Denaturing solution Mix: 49 ml formamide 7 ml 20× SSC pH 5.3 (APPENDIX 2D) 14 ml purified H2 O 70 ml final volume Mix well and place in a glass Coplin jar. Measure pH at room temperature using a pH meter with a glass pH electrode to verify that the pH is between 7.0 to 8.0. Store in a covered container at 2° to 8°C. This solution can be used for up to 1 week. Check pH prior to each use.
Ethanol wash solutions Prepare 70% (v/v), 85% (v/v), and 100% (v/v) ethanol using 100% ethanol and purified water. Add 70 ml of each solution to a Coplin jar and maintain at room temperature. Store unused dilutions in a covered container at room temperature for up to 6 months. Solutions used in the assay may be used for 1 week unless evaporation occurs or the solution becomes diluted due to excessive use.
NP-40, 0.1%, in 2× SSC Mix: 100 ml 20× SSC pH 5.3 (APPENDIX 2D) 849 ml purified H2 O 1 ml NP-40 (IGEPAL CA-630; Sigma-Aldrich, CAS no. 9002-93-1) Add H2 O to 1000 ml Mix thoroughly. Adjust the pH to 7.0 to 7.5 with 1 N NaOH using a pH meter and glass pH electrode to measure the pH. Adjust volume to 1 liter with water. Filter through 0.45-μm filtration unit. Add 70 ml of the solution to a Coplin jar and maintain at room temperature. Store unused solution in a covered container at room temperature for up to 6 months. Discard solution that was used in the assay at the end of each day.
NP-40, 0.3%, in 0.4× SSC Mix: 950 ml purified H2 O 20 ml 20× SSC pH 5.3 (APPENDIX 2D) 3 ml NP-40 (IGEPAL CA-630; Sigma-Aldrich, CAS no. 9002-93-1) Add H2 O to 1000 ml Mix thoroughly. Adjust pH to 7.0 to 7.5 with 1 N NaOH, if necessary, using a pH meter with glass pH electrode. Adjust volume to 1 liter with water. Filter through 0.45-μm filtration unit. Store unused solution in a covered container at room temperature for up to 6 months. Discard solution that was used in the assay at the end of each day.
Post-fixation solution To 48.7 ml of phosphate-buffered saline (PBS; APPENDIX 2D), add: 1.3 ml of 37% formaldehyde (0.95% final) 0.23 g MgCl2 Store up to 1 month at 4°C
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COMMENTARY Background Information
Preparation of Amniocytes for Interphase FISH
Fluorescence in situ hybridization (FISH) with repetitive-element and single-copy DNA probes, as well as with probes made from chromosome-specific libraries, has been utilized for analysis of both metaphase and interphase cells for over two decades now. FISH analysis has made possible studies of alterations of chromosome structure including detection and characterization of duplications, deletions, and/or cryptic translocations (Stumm and T¨onnies, 2008; Zuffardi et al., 2009; Tsuchiya, 2011; Bubendorf and Piaton, 2012; Haferlach, 2012; Jehan et al., 2012; Das and Tan, 2013; Lampert et al., 2013; Riegel, 2014); determination of the chromosomal origin of supernumerary markers (Rodr´ıguez L et al., 2007); and detection of aneuploidy in interphase cells. Detection of aneuploidy by FISH in interphase cells has been applied widely to neoplastic conditions (e.g., the study of alterations in chronic lymphocytic leukemia, myelodyplasia, solid tumors, and acute lymphocytic leukemia, among others; Stumm and T¨onnies, 2008; Tsuchiya, 2011; Bubendorf and Piaton, 2012; Haferlach, 2012; Jehan et al., 2012; Das and Tan, 2013; Lampert et al., 2013). This technology has been utilized to detect aneuploidies in blood smears from newborn infants as well as extensively in amniotic fluid samples and CVS samples (Ward et al., 1993; Tepperberg et al., 2001; Weise and Liehr, 2008). There is one major method and an alternative method of preparing amniotic fluid cells for detection of aneuploidy by FISH. The first method (Basic Protocol 1) uses uncultured cells obtained from amniotic fluid samples. The second method (Alternate Protocol) uses cells that have been placed on a coverslip or slide and allowed a short period of time (up to 48 hr) for attachment. There have been numerous studies published and reviews analyzing the use of the FISH for prenatal diagnosis in interphase cells (Ward et al., 1993; Tepperberg et al., 2001; Weise and Liehr, 2008; Faas et al., 2011). The majority of studies utilized a variation of the basic protocol and analyzed the cells within 2 days of receipt. Results from over 29,000 amniotic fluid samples summarized, after initial use of the protocol, revealed only one false-positive (0.003%) and seven false-negative results (0.024%), demonstrating the efficacy of this technology (Tepperberg et al., 2001). Although there have been isolated instances of false-positive and false-
negative results, this technology has proven to be robust over time. It was initially argued that this technique could be most effectively used for patients with a higher probability of an aneuploid fetus (e.g., with abnormal ultrasonographic findings, maternal age >40 years, or positive maternal serum screen), rather than for every individual who comes to a prenatal diagnostic program. Advantages to analysis based on increased risk of aneuploidy are that if a particular abnormality is suspected, only one probe may be needed. This will save on the amount of specimen needed, as well as on additional analyses to be performed. This in turn will lower the cost of carrying out FISH, which can be both expensive and time-consuming if five different probes must be utilized. However, today the technology is used mostly as an adjunctive procedure for all patients that request testing to facilitate the reporting of results more rapidly, usually within 24 hr. Invariably, five probes are utilized rather than just one.
Critical Parameters and Troubleshooting Cell preparation Preparation of the cells is the most difficult aspect of FISH with amniotic fluid. Preparation of uncultured amniocytes (Basic Protocol 1) is easier, quicker, and much more readily used than preparation of amniocytes attached to coverslips (Alternate Protocol). Over the past decade, this technology has been optimized, and investigators have used different hypotonic solutions, hypotonic treatment times, and fixative solutions than those mentioned here. The optimization of this technology has evolved over the years, leading to fairly standard protocols in most laboratories. However, because technologies are so varied, each laboratory will ultimately determine what works best in their individual laboratory. A successful analysis can be done even in cases where only 1 ml of amniotic fluid is obtained from an 11- to 15-gestational-week pregnancy. In general, however, 2 to 3 ml of amniotic fluid is needed at 12 to 15 weeks; 3 to 5 ml at 14 to 19 weeks; and 5 ml at 20 weeks. These numbers may be slightly conservative, as on the average 3 ml provides successful results. Although more cells are available in amniotic fluid from 26 gestational-week pregnancies, the majority are dead cells. Therefore, additional amniotic fluid may be needed in these cases.
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A possible advantage of the Alternate Protocol is that the morphology of the cells changes after attachment, making them more permeable and thus increasing hybridization efficiency. The procedure described in Klinger et al. (1992), which uses cells attached to slides, does not designate any extra incubation time to allow for cell attachment. However, most laboratories have not been able to reproduce the results of Klinger et al. (1992) using that methodology, but laboratories have had successful hybridization after allowing attachment for 24 hr, and the Alternate Protocol has been designed with that in mind. Several factors affect hybridization of amniotic fluid cells. In general, cells from amniotic fluid received late in gestation (>25 weeks) will hybridize poorly, and the results will be much more difficult to analyze. In such samples, there is usually an elevated number of cells that are dead and will not hybridize, as well as more debris, which makes hybridization less efficient. In most cases, cells from as early as 12 gestational weeks can be used for FISH studies. The difficulty with early-gestation specimens is that there are not enough cells to complete the proper studies. Bloody specimens can also present a problem, which can usually be alleviated by fixation, as the fixative will hemolyze the blood. When uncultured amniotic fluid cells are spun down, red blood cells will be present in the centrifuge tube and possibly on the slide. Because these are anucleate, they will not hybridize with a probe, but their presence may lead to cellular crowding, which results in an overall lower number of cells demonstrating hybridization. Another possible complication in prenatal FISH studies is maternal-cell admixture. Presence of any maternal cells can lead to false conclusions. Therefore, it is important to know whether the sample may be at higher risk for having maternal cells (e.g., whether the specimen was bloody or was obtained through the anterior placenta). It should also be noted in the analysis whether any maternal cells appear to be present. A few studies have evaluated the presence of maternal cells in amniotic fluid samples. Although some of these studies suggest a higher frequency of contamination than others, it is clear that contamination does not frequently cause problems with the use of these protocols (Nub et al., 1994; Rebello et al., 1994). It is very important for every laboratory to determine the exact extent of maternal-cell contamination in their samples based on use of both an X and Y probe in cells from a male fetus; however, the
admixture of maternal cells will vary from one specimen to the next. When there is a hint of the possibility of red cells present, it is advisable to count 75 interphase cells rather than 50 cells as is usually counted. FISH Recommendations from the laboratories utilizing these protocols suggest that the protocol may have to be modified to take into account the hybridization efficiency of the probe and number of cells showing a positive signal, as well as the possibility of maternal cell contamination. Although considerable troubleshooting may be necessary with FISH protocols (see UNIT 4.3; Knoll and Lichter, 2005), the probes used for this analysis work relatively well with little modification. Clinical guidelines The American College of Medical Genetics and the American Society of Human Genetics (ACMG/ASHG) published a position paper in 2000 on the technical and clinical assessment of fluorescence in situ hybridization. This included a section on evaluating the results of FISH on interphase cells in prenatal diagnosis (Test and Technology Transfer Committee, 2000). This position statement includes the following: (1) those requesting interphase FISH testing for prenatal diagnosis should be fully aware of what these focused tests can and cannot do (clinical sensitivity and specificity) including false positive and false negatives; (2) while the speed of diagnosis with this technology is a distinct advantage, the increased cost of FISH must be considered; (3) for management of the fetus, it is reasonable to report positive FISH test results; (4) clinical decision-making should be based on information from two of three of the following— FISH results, confirmatory chromosome analysis, or consistent clinical information; (5) for ultimate management of pregnancies that are FISH positive, these results need to be further characterized by traditional chromosome analysis to determine the mechanism (e.g., aneuploidy or translocation) accounting for the FISH detected abnormality. An overview of the validations and guidelines for the clinical diagnostic use of interphase FISH analysis is given in Section E10 of the American College of Medical Genetics Standards and Guidelines for Clinical Genetics Laboratories (1/2010; https://www.acmg.net/StaticContent/SGs/ Section_E_2011.pdf).
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Figure 8.9.3 Distinct and split signals. Hybridization of cells with chromosome 18 α-satellite probe showing: (A) two signals; (B) two split signals each of which is considered one signal by criteria used in authors’ laboratory.
Preparation of Amniocytes for Interphase FISH
FISH analysis of interphase will always yield higher results based on the ascertainment of patients, specifically in patients with advanced maternal age and specific ultrasound findings. However, once an invasive technique has been done, this technology becomes available to all patients and is often used to provide more rapid results. There are some general rules that should be considered when performing prenatal interphase FISH. Optimally, 50 interphase cells should be analyzed for each probe used (25 counts from two different readers). Because FISH is not usually used in any case as a standalone procedure, analysis of 25 cells is acceptable, though not optimal (and analysis of fewer than 25 is unacceptable). If cell-preparation techniques have been optimized, 70% to 98% of all interphase cells should show hybridization of each probe. Each laboratory should develop its own guidelines regarding acceptable frequency of signals for each probe, but in general 60% of the cells should show an abnormal number of signals for a sample to be considered aneuploid (e.g., three signals for an 18 centromere probe). Usually, detection of an abnormal number of signals in 30% of cells is within experimental error. Cases with 10% but 60% of the cells demonstrating abnormal signals need to be analyzed
further. Controls need not be used for each hybridization if multiple slides are analyzed; however, no conclusions concerning absence of the Y chromosome can be made without a positive signal in another sample. Each time a new lot of probe is utilized, it should be tested on known control slides. Consistency in scoring interphase signals is of the utmost importance; this can be accomplished over time and after multiple evaluations. Because of the three-dimensional nature of interphase cells, not all cells will always be in the same plane of focus. Therefore, in order to detect signals, it is necessary to focus up and down to detect any signals outside the focal plane. Focusing up and down also helps delineate nuclear boundaries in preparations with extensive nuclear overlap. Split signals are in general counted as one signal (Fig. 8.9.3), unless a distance greater than the size of a signal lies between the two spots comprising the putative split signal. In those cases, the signals are counted as two separate signals. Hybridized specimens can be initially scanned (and analyzed in some cases) using a 40× objective. Often, initial analysis is accomplished with either a 63× or 100× objective; when a 40× objective is used, these more powerful objectives can be utilized to clarify any questions that may arise.
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Anticipated Results After establishment of these techniques within the laboratory, analysis should be successful >95% of the time (for both singlecopy and repetitive probes, once the technique has been optimized). However, if samples obtained at a later gestational age are studied, the success rate may be lower because of lower hybridization efficiency. It is crucial for each laboratory to develop its own internal guidelines and conditions for optimal hybridization and analysis. Hybridization efficiencies using samples prepared by either method should be similar once the methods have been optimized for an individual laboratory. To avoid inaccuracies arising from the presence of large numbers of dead cells (e.g., in specimens obtained at advanced maternal age), cells that show no signal are placed in the analysis category “did not hybridize.” These cells are included when determining hybridization efficiency but are not included in the actual analysis. It is possible to get one to three slides with 50 cells each from a 1-ml sample. It is not always advisable to take a specimen from a sample containing
