Histology FISH Accessory Kit - Agilent · P04260US_02/K579911-5 p. 3/26 Intended Use For in vitro...

P04260US_02/K579911-5 p. 1/26

Histology FISH Accessory Kit Code K5799

2nd edition

For fluorescence in situ hybridization (FISH) on formalin-fixed, paraffin-embedded tissue sections.

The kit contains reagents sufficient for 20 tests.

P04260US_02/K579911-5 p. 2/26

Table of content/ Table des matières/ Inhalt

ENGLISH

Intended Use ................................................................................................................................ 3

Introduction ................................................................................................................................... 3

Reagents ...................................................................................................................................... 3

Materials provided .................................................................................................................... 3

Materials required but not provided .......................................................................................... 4

Precautions .................................................................................................................................. 5

Storage ......................................................................................................................................... 8

Specimen Preparation .................................................................................................................. 8

Paraffin-embedded sections ..................................................................................................... 8

INSTRUCTIONS FOR USE .......................................................................................................... 9

A. Reagent Preparation ........................................................................................................... 9

A.1 Pre-Treatment Solution ................................................................................................ 9

A.2 Stringent Wash Buffer .................................................................................................. 9

A.3 Wash Buffer ................................................................................................................. 9

A.4 Ethanol series .............................................................................................................. 9

A.5 Pepsin solution ............................................................................................................ 9

B. Staining Procedure ............................................................................................................ 10

B.1 Procedural notes ....................................................................................................... 10

B.2 Treatment of tissues prior to staining ......................................................................... 10

B.3 Staining protocol ........................................................................................................ 11

B.3.a. Staining protocol for FISH probes diluted in ethylene carbonate-based hybridization buffer ................................................................................................................. 11

B.3.b. Staining protocol for FISH probes diluted in formamide-based hybridization buffer ...................................................................................................................................... 14

Quality Control ............................................................................................................................ 17

Interpretation of Results .............................................................................................................. 18

Troubleshooting .......................................................................................................................... 19

Appendix 1 ................................................................................................................................. 23

Appendix 2 ................................................................................................................................. 23

References ................................................................................................................................. 25

Explanation of symbols ............................................................................................................... 26

P04260US_02/K579911-5 p. 3/26

Intended Use

For in vitro diagnostic use.

Histology FISH Accessory Kit is intended for use in the fluorescence in situ hybridization (FISH) technique on formalin-fixed, paraffin-embedded tissue section specimens.

Reagents provided in this Histology FISH Accessory Kit can also be used together with HER2/CEN-17 IQISH Probe Mix (Code K5731, Vial 3). If reagents from Dako Histology FISH Accessory Kit, Code K5799, are used together with reagents from Code K5731, the procedure outlined in the package insert for Code K5731 must be followed.

Introduction Histology FISH Accessory Kit contains all key reagents, except for the probe, required to complete a FISH procedure for formalin-fixed, paraffin-embedded tissue section specimens. After deparaffinization and rehydration, specimens are heated in Pre-Treatment Solution followed by proteolytic digestion using Pepsin. After the heating and proteolytic pre-treatment steps, user-provided FISH probes are applied and specimens are sealed with Coverslip Sealant before denaturation and hybridization. Following hybridization a stringent wash is performed, which ensures removal of unbound and unspecifically bound FISH probes before dehydration with ethanol. Finally, the specimens are mounted with Fluorescence Mounting Medium containing a blue fluorescence counterstain. Results are interpreted using a fluorescence microscope equipped with appropriate filters (see Appendix 1).

Reagents

Materials provided

The materials listed below are sufficient for 20 tests (a test is defined as one 22 mm x 22 mm target area). The kit provides materials sufficient for up to 10 individual FISH runs (four separate runs, when using the pepsin immersing method).

Histology FISH Accessory Kit is shipped on dry ice. To ensure that kit components have not been exposed to high temperatures during transport, dry ice should still be present upon receipt. Note that some kit components may remain unfrozen, this will not affect the performance of the Histology FISH Accessory Kit.

Vial 1

Vial 2A

Vial 2B

Pre-Treatment Solution (20x) 150 mL, 20x concentrated MES (2-[N-morpholino]ethanesulphonic acid) buffer.

Pepsin 4 x 6.0 mL, ready-to-use Pepsin solution, pH 2.0; contains stabilizer and an antimicrobial agent.

Pepsin Diluent (10x) 24 mL, concentrated 10x Dilution buffer, pH 2.0; contains an antimicrobial agent.

Vial 4 Stringent Wash Buffer (20x) 150 mL, 20x concentrated SSC (saline-sodium citrate) buffer with detergent Tween-20.

P04260US_02/K579911-5 p. 4/26

Vial 5

Vial 6

Fluorescence Mounting Medium 0.4 mL Ready-to-use fluorescence mounting medium with 500 μg/L DAPI (4’,6-diamidine-2-phenylindole)..

Wash Buffer (20x) 500 mL, 20x concentrated Tris/HCl buffer.

Item 7 Coverslip Sealant 1 tube Ready-to-use solution for removable sealing of coverslip.

NOTE: All reagents are interchangeable with the corresponding reagents in Dako HER2 IQFISH pharmDx (Code K5731). If reagents from Dako Histology FISH Accessory Kit, Code K5799, are used together with reagents from Code K5731, the procedure outlined in the package insert for Code K5731 must be followed.

Materials required but not provided

Laboratory reagents

Distilled or deionized water

Ethanol, 96%

Xylene or xylene substitutes

Laboratory equipment

Absorbent wipes

Adjustable pipettes

Calibrated partial immersion thermometer (range 37–100 C)

Coverslips (18 mm x 18 mm or 22 mm x 22 mm and 24 mm x 50 mm or 24 mm x 60 mm)

Forceps

Fume hood

Dako Hybridizer (Code S2450/S2451)*

Heating block or hybridization oven*

Humid hybridization chamber*

Microcentrifuge

Slides, Dako Silanized Slides, Code S3003, poly-L-lysine-coated slides, or superfrost Plus slides

Staining jars or baths

Metal or plastic cradles

Timer (capable of 0–60 minute intervals)

Water bath with lid (capable of maintaining 37 (±2) C, 63 (±2) C, 65 (±2) C and from 95 C to

99 C

Microwave oven with sensing capability if pre-treatment is performed using microwave oven** (see Step 1 in Section B.3.a or B.3.b. Staining protocol, Method B). Microwave proof container, lid must contain holes with a diameter of e.g. one cm

* Heating block for digestion (37 (±2) C), heating block or hybridization oven for denaturation (66 (±2) C (B.3.a.)

or 82 (±2) C) (B.3.b.) and humid hybridization chamber for hybridization (45 (±2) C) can be used, but we recommend Dako Hybridizer, Code S2450/S2451.

** Water bath with lid (capable of maintaining 95-99 C) or a microprocessor-controlled pressure chamber such as Dako Pascal, Code S2800 can be used instead of a microwave oven.

P04260US_02/K579911-5 p. 5/26

Microscope equipment and accessories

Filters for fluorescence microscope: DAPI filter and suitable filters for the used fluorochrome, e.g. FITC/Texas Red double filter, FITC and Texas Red mono filters for Dako FISH Probes – see Appendix 1 for details.

Fluorescence microscope with a 100 watt mercury lamp as light source should be used for Dako FISH Probes. Other light sources are not recommended with these filters.

Microscope slide folder (cardboard tray for 20 slides with hinged cover or similar).

Precautions 1. For in vitro diagnostic use.

2. For professional users.

3. Vial 1, Pre-Treatment Solution (20x), does not require hazard labeling. Safety Data Sheet (SDS) is available for professional users on request.

4. Vial 2A, Pepsin, 5-10% propan-2-ol, 0.1-1% pepsin A, and <0.1% 3(2H)-Isothiazolone, 5-chloro-2-methyl-, mixt. with 2-methyl-3(2H)-isothiazolone. Vial 2a is labeled:

Danger

H314 Causes severe skin burns and eye damage. H334 May cause allergy or asthma symptoms or breathing difficulties if

inhaled. H317 May cause an allergic skin reaction. H373 May cause damage to organs through prolonged or repeated

exposure. P280 Wear protective gloves. Wear eye or face protection. Wear

protective clothing. P285 In case of inadequate ventilation wear respiratory protection. P260 Do not breathe vapor. P264 Wash hands thoroughly after handling. P272 Contaminated work clothing should not be allowed out of the

workplace. P314 Get medical attention if you feel unwell. P304 + P340 + P310 IF INHALED: Remove victim to fresh air and keep at rest in a

position comfortable for breathing. Immediately call a POISON CENTER or physician.

P342 + P311 If experiencing respiratory symptoms: Call a POISON CENTER or physician.

P301 + P310 + P330 + P331

IF SWALLOWED: Immediately call a POISON CENTER or physician. Rinse mouth. Do NOT induce vomiting.

P303 + P361 + P353 + P363 + P310

IF ON SKIN (or hair): Take off immediately all contaminated clothing. Rinse skin with water or shower. Wash contaminated clothing before reuse. Immediately call a POISON CENTER or physician.

P302 + P352 IF ON SKIN: Wash with plenty of soap and water. P333 + P313 If skin irritation or rash occurs: Get medical attention. P305 + P351 + P338 + P310

IF IN EYES: Rinse cautiously with water for several minutes. Remove contact lenses, if present and easy to do. Continue rinsing. Immediately call a POISON CENTER or physician.

P405 Store locked up. P501 Dispose of contents and container in accordance with all local,

regional, national and international regulations.

P04260US_02/K579911-5 p. 6/26

5. Vial 2B, Pepsin Diluent (10x), contains 30-60% propan-2-ol, and 5-10% 2-amino-2-(hydroxymethyl) propane-1,3-diol hydrochloride. Vial 2B is labeled:

Danger

H225 Highly flammable liquid and vapour. H319 Causes serious eye irritation. H335 May cause respiratory irritation. H336 May cause drowsiness or dizziness. H373 May cause damage to organs through prolonged or repeated

exposure. P280 Wear protective gloves. Wear eye or face protection. P210 Keep away from heat, hot surfaces, sparks, open flames and other

ignition sources. No smoking. P241 Use explosion-proof electrical, ventilating, lighting and all material-

handling equipment. P242 Use only non-sparking tools. P243 Take precautionary measures against static discharge. P233 Keep container tightly closed. P271 Use only outdoors or in a well-ventilated area. P260 Do not breathe vapor. P264 Wash hands thoroughly after handling. P314 Get medical attention if you feel unwell. P304 + P340 + P312 IF INHALED: Remove victim to fresh air and keep at rest in a

position comfortable for breathing. Call a POISON CENTER or physician if you feel unwell.

P303 + P361 + P353 IF ON SKIN (or hair): Take off immediately all contaminated clothing. Rinse skin with water or shower.

P305 + P351 + P338 IF IN EYES: Rinse cautiously with water for several minutes. Remove contact lenses, if present and easy to do. Continue rinsing.

P337 + P313 If eye irritation persists: Get medical attention. P405 Store locked up. P403 Store in a well-ventilated place. P235 Keep cool. P501 Dispose of contents and container in accordance with all local,


6. Vial 4, Stringent Wash Buffer (20x), contains 10-30% sodium chloride, and 10-30% 2-amino-2-(hydroxymethyl) propane-1,3-diol hydrochloride. Vial 4 is labeled:

Warning

H319 Causes serious eye irritation. H315 Causes skin irritation. P280 Wear protective gloves. Wear eye or face protection. P264 Wash hands thoroughly after handling. P302 + P352 + P362-2 + P363

IF ON SKIN: Wash with plenty of soap and water. Take off contaminated clothing. Wash contaminated clothing before reuse.

P332 + P313 If skin irritation occurs: Get medical attention. P305 + P351 + P338 IF IN EYES: Rinse cautiously with water for several minutes.

Remove contact lenses, if present and easy to do. Continue rinsing.

P337 + P313 If eye irritation persists: Get medical attention.

P04260US_02/K579911-5 p. 7/26

7. Vial 6, Wash Buffer (20x), contains 10-30% sodium chloride, and 10-30% trometamol. Vial 6 is labeled:

Warning

H319 Causes serious eye irritation. H315 Causes skin irritation. P280 Wear protective gloves. Wear eye or face protection. P264 Wash hands thoroughly after handling. P302 + P352 + P362-2 + P363

IF ON SKIN: Wash with plenty of soap and water. Take off contaminated clothing. Wash contaminated clothing before reuse.

P332 + P313 If skin irritation occurs: Get medical attention. P305 + P351 + P338 IF IN EYES: Rinse cautiously with water for several minutes.

Remove contact lenses, if present and easy to do. Continue rinsing.

P337 + P313 If eye irritation persists: Get medical attention.

8. Coverslip Sealant contains 60-100% naphtha (petroleum), hydrotreated light, and is labeled:

Danger

H225 Highly flammable liquid and vapour. H304 May be fatal if swallowed and enters airways. P280 Wear protective gloves. Wear eye or face protection. P210 Keep away from heat, hot surfaces, sparks, open flames and other

ignition sources. No smoking. P241 Use explosion-proof electrical, ventilating, lighting and all material-

handling equipment. P242 Use only non-sparking tools. P243 Take precautionary measures against static discharge. P233 Keep container tightly closed. P301 + P310 + P331 IF SWALLOWED: Immediately call a POISON CENTER or

physician. Do NOT induce vomiting. P303 + P361 + P353 IF ON SKIN (or hair): Take off immediately all contaminated

clothing. Rinse skin with water or shower. P405 Store locked up. P403 Store in a well-ventilated place. P235 Keep cool. P501 Dispose of contents and container in accordance with all local,


9. Please refer to the Safety Data Sheet (SDS) for additional information.

10. Tissue fixation methods and thickness of specimen other than those specified may affect tissue morphology and/or signal intensity.

11. Incubation times and temperatures, or methods other than those specified, may give poor results.

12. Reagents have been optimally diluted. Further dilution may result in poor performance.

13. Only clean staining jars should be used for the pepsin immersion method (Step 2. Method C).

P04260US_02/K579911-5 p. 8/26

Storage Store in the dark at 2-8 °C. Alternatively, all reagents may be stored frozen.

The Pepsin (Vial 2A) may be adversely affected if exposed to heat. Do not leave this component at room temperature.

Fluorescence Mounting Medium (Vial 5) may be adversely affected if exposed to excessive light levels. Do not store this component in light.

Do not use the kit after the expiry date stamped on the kit box. If reagents are stored under conditions other than those specified, the user must validate reagent performance (1).

There are no obvious signs indicating instability of this product. Therefore, it is important to evaluate normal tissue previously shown to “FISH well” as control of the assay run. If an unexpected fluorescence pattern is observed which cannot be explained by variations in laboratory procedures, and a problem with the Histology FISH Accessory Kit is suspected, contact Dako Technical Services.

Specimen Preparation

Specimens from biopsies, excisions or resections must be handled to preserve the tissue for FISH analysis. Follow the recommended method of tissue processing for immunocytochemical staining described below. For further information see reference (2). The use of tissue exposed to acid decalcification for FISH is not recommended (3-6). EDTA as decalcifier has been reported to preserve the DNA better (6) for (F)ISH (3, 4, 7) techniques.

NOTE: Dako Histology FISH Accessory Kit performance has not been validated on decalcified tissue.

Paraffin-embedded sections Only tissue preserved in neutral-buffered formalin and paraffin-embedded are suitable for use. Specimens should e.g. be blocked into a thickness of 3 or 4 mm and fixed 18-24 hours in neutral- buffered formalin. The tissues are then dehydrated in a graded series of ethanol and xylene, followed

by infiltration by melted paraffin held at no more than 60 C. Properly fixed and embedded tissues

will keep indefinitely prior to sectioning and slide mounting if stored correctly (15-25 C) (2, 8). Other fixatives are not suitable. Extended fixation time might increase the incubation time required in the Pepsin digestion step.

Tissue specimens should be cut into sections of 2-6µm, collected on slides from a water bath and then air-dried. The optimal thickness of tissue sections is 2-3 µm for Dako Split Signal FISH Probes. Sections of 4–6 µm may be used for other FISH applications.

The paraffin should be melted at 60 C for 30-60 minutes. The sections should then be cooled to

room temperature (20-25 C) and stored at 2-8 C.

It is recommended that tissue sections are mounted on Dako Silanized Slides, Code S3003, or poly-L-lysine-coated or Superfrost Plus slides. In general, specimens should be analyzed within

6 months of sectioning when stored at 2-8 C. If tissue sections are stored under other conditions than those specified, the user must verify the conditions.

P04260US_02/K579911-5 p. 9/26

INSTRUCTIONS FOR USE

A. Reagent Preparation It is convenient to prepare the following reagents prior to staining:

A.1 Pre-Treatment Solution Crystals may occur in Vial 1, but they will dissolve at room temperature. Ensure that no crystals are present before preparation of reagent. Dilute a sufficient quantity of Vial 1 (Pre-Treatment Solution 20x) by diluting the concentrate 1:20 in distilled or deionized water. Unused diluted solution may be stored at 2-8 °C for one month. Discard diluted solution if cloudy in appearance.

A.2 Stringent Wash Buffer Dilute a sufficient quantity of Vial 4 (Stringent Wash Buffer 20x) by diluting the concentrate 1:20 in distilled or deionized water. Unused diluted buffer may be stored at 2-8 °C for one month. Discard diluted buffer if cloudy in appearance.

A.3 Wash Buffer Dilute a sufficient quantity of Vial 6 (Wash Buffer 20x) by diluting the concentrate 1:20 in distilled or deionized water. Unused diluted buffer may be stored at 2-8 °C for one month. Discard diluted buffer if cloudy in appearance.

A.4 Ethanol series From a 96% ethanol solution, prepare 3 jars containing 70%, 85%, and 96% ethanol. Store covered jars at room temperature or at 2-8 °C, and use for a maximum of 200 slides. Discard solutions if cloudy in appearance.

A.5 Pepsin solution A pepsin solution is only needed when using the pepsin immersing method (B.3.a. and B.3.b Step 2 Method C). Prepare pepsin solution as follows;

For a six slide capacity container prepare 60 mL pepsin solution:

Add 48 mL of room temperature (20-25 C) distilled or deoinized water to the container.

Add 6 mL of cold (2-8 C) Pepsin Diluent (10x) (Vial 2B) to the container.

Add 6 mL of cold (2-8 C) Pepsin (Vial 2A) to the container.

Put lid on the container and equilibrate the pepsin solution to 37 (±2) C in a water bath. For a 24 slide capacity container prepare 240 mL pepsin solution:

Add 192 mL of room temperature (20-25 C) distilled or deoinized water to the container.

Add 24 mL of cold (2-8 C) Pepsin Diluent (10x) (Vial 2B) to the container.

Add 24 mL of cold (2-8 C) Pepsin (Vial 2A) to the container.

Put lid on the container and equilibrate the pepsin solution to 37 (±2) C in a water bath. Equilibrated pepsin solution should be used within 5 hours.

P04260US_02/K579911-5 p. 10/26

B. Staining Procedure

B.1 Procedural notes The user should read these instructions carefully and become familiar with all components prior to use (see Precautions).

If kit components are stored frozen, it is recommended to transfer the reagents to 2–8 C the day before performing the analysis to allow proper temperature equilibration. All reagents should be equilibrated to the relevant temperature prior to use.

Vial 1: The diluted Pre-Treatment Solution should be equilibrated to 95-99 C if water bath is used (Section B.3.a or B.3.b Staining protocol, Step 1: Pre-Treatment, Method A). If microwave oven with sensing capability is used for pre-treatment (Section B.3.a. or B.3.b. Staining protocol, Step 1: Pre-Treatment, Method B) the diluted Pre-Treatment Solution should be equilibrated to room temperature 20-25 °C.

Vial 2A: Pepsin should be applied at 2-8 C (Section B.3.a. or B.3.b. Staining protocol, Step 2: Method A and B) and kept cold continuously.

Vial 2B: Pepsin Diluent (10x) should be applied at 2-8 C (Section B.3.a. or B.3.b. Staining protocol, Step 2: Method C)

Vial 4: One jar of the diluted Stringent Wash Buffer should be equilibrated to room temperature

and another jar should be equilibrated to 63 (±2) C (Section B.3.a.) or 65 (±2) C (Section B.3.b) , prior to use.

Vial 5: Fluorescence Mounting Medium may be applied at any temperature from 2-25 C.

Vial 6: The diluted Wash Buffer should be equilibrated to room temperature (20-25 C).

Item 7: Coverslip Sealant may be applied at room temperature (20-25 °C). All steps must be performed at the outlined temperature.

The pre-staining procedure includes dehydrations followed by drying of the specimens. Ensure that the specimens are completely dry before proceeding to the next step. Do not allow specimens to dry during the other procedural steps.

If the staining procedure has to be interrupted, slides may be kept in Wash Buffer after the

deparaffinization step for up to 1 hour at room temperature (20-25 C).

B.2 Treatment of tissues prior to staining

Deparaffinization and rehydration: Prior to performing the procedure, tissue slides must be deparaffinized to remove paraffin and rehydrated. Avoid incomplete removal of paraffin. Residual paraffin will result in increased non-specific staining. This step should be performed at room temperature (20-25 °C).

1. Place slides in a xylene bath and incubate for 5 (±1) minutes. Change bath and repeat once.

2. Tap off excess liquid and place slides in 96% ethanol for 2 (±1) minutes. Change bath and repeat once.

3. Tap off excess liquid and place slides in 70% ethanol for 2 (±1) minutes. Change bath and repeat once.

4. Tap off excess liquid and place slides in diluted Wash Buffer (see INSTRUCTIONS FOR USE, Section A.2) for a minimum of 2 minutes. Commence staining procedure as outlined in Section B.3.a. or B.3.b., Step 1, Pre-Treatment.

Fresh xylene and alcohol solutions should be prepared after every 200 slides have been processed.

Xylene substitutes may be used.

NOTE: The reagents and instructions supplied in this kit have been designed for optimal performance. Further dilution of the reagents or alteration of incubation temperatures may give erroneous or discordant results. Differences in tissue processing and technical procedures in the user’s laboratory may invalidate the assay results.

P04260US_02/K579911-5 p. 11/26

Optional maturing of specimens: Before deparaffinization and rehydration, incubate slides at

60 C for e.g. 1 hour on a heating block, a block in a hybridization oven or on the Dako Hybridizer.

Alternatively, incubate slides at 37 C for 1-2 days followed by 1 hour at 60 C. Continue with deparaffinization and rehydration.

NOTE: This step is optional. Matured specimens may reduce potential background noise.

B.3 Staining protocol Follow one of the two staining protocol variants, B.3.a or B3.b. Do not interchange steps between the two procedures:

Staining protocol B.3.a describes the half-day procedure that should be applied for FISH probes diluted in an ethylene carbonate-based hybridization buffer.

Staining protocol B.3.b describes the two-day procedure that should be applied for FISH probes diluted in a formamide-based hybridization buffer.

B.3.a. Staining protocol for FISH probes diluted in ethylene carbonate-based hybridization buffer

Step 1: Pre-Treatment (microwave oven, water bath, Pascal)

Pre-treatment can be performed either by using water bath as described in method A), by use of microwave oven with sensing capability as described in method B) or by use of Pascal pressure cooker as described in method C).

Method A: Pre-treatment using water bath Fill staining jars, e.g. Coplin jars, with the diluted Pre-Treatment Solution (see INSTRUCTIONS FOR USE, Section A.1). Place staining jars containing diluted Pre-Treatment Solution in water

bath. Heat water bath and the Pre-Treatment Solution to 95-99 C. Measure temperature inside jar with a calibrated thermometer to ensure correct temperature. Cover jars with lids in order to stabilize the temperature and avoid evaporation.

Immerse the room temperature deparaffinized sections into the preheated Pre-Treatment Solution in

the staining jars. Re-check temperature and incubate for 10 (±1) minutes at 95-99 C.

Remove the entire jar with slides from the water bath. Remove lid and allow the slides to cool in the Pre-Treatment Solution for 15 minutes at room temperature.

Transfer the slides to a jar with diluted Wash Buffer (see INSTRUCTIONS FOR USE, Section A.3)

for 3 minutes at room temperature (20-25 C).

Replace Wash Buffer and soak sections for another 3 minutes.

NOTE: The Pre-Treatment Solution is designed for single use application only. Do not re-use.

Method B: Pre-treatment using microwave oven with sensing capability (Recommended)

Fill a plastic jar with diluted room temperature (20-25 C) Pre-Treatment Solution. Immerse the deparaffinized sections in Pre-Treatment Solution, cover the jar with a punctured lid and place it in the microwave oven. Select the boiling sensor function and a program that runs for 10 minutes after boiling temperature has been reached*.

Following the 10 minutes incubation take the jar with slides out of the oven, remove the lid and cool for 15 minutes at room temperature. Transfer the slides to a jar with diluted Wash Buffer and soak

for 3 minutes at room temperature (20-25 C). Replace Wash Buffer and soak sections for another 3 minutes.

* The use of a microwave oven with a sensing capability means that the oven must include a sensor and programs which initially heat the Pre-Treatment Solution to the boiling point and subsequently maintain the required pre-treatment temperature (above 95 ºC) while counting down the preset time (10 (±1) minutes). Some microwave oven models with sensing capability may not include the possibility to freely set a count-down time. If the model only includes pre-set programs, be sure to select a program which maintain the required pre-treatment temperature (above 95 ºC) for at least 10 (±1) minutes and manually stop the program after 10 (±1) minutes.

NOTE: The Pre-Treatment Solution is designed for a single use application only. Do not re-use.

Method C: Pre-treatment using Pascal pressure chamber

P04260US_02/K579911-5 p. 12/26

Place 500 mL of deionized water inside the Pascal pan. Fill a plastic jar with 250 mL Pre-Treatment Solution. Immerse the deparaffinized sections in Pre-Treatment Solution and place the jar inside the Pascal pan. Incubate at 121 °C for 1 minute. Release pressure after cooling to 90 °C. Carefully read the Pascal Handbook for information on proper handling of the Pascal pressure chamber. Remove the container after the pressure is released. Remove lids and allow the slides to cool in the Pre-Treatment Solution for another 15 minutes at room temperature. Transfer the slides to a jar containing diluted Wash Buffer (see INSTRUCTIONS FOR USE, Section A.2). Soak sections for 3 minutes at room temperature.

Replace Wash Buffer and soak sections for another 3 minutes. Continue to Step 2.

NOTE: The Pre-Treatment Solution is designed for a single use only. Do not re-use. Do not use a sealed, airtight container when performing pre-treatment in a microwave oven as the pressure inside the container will increase and may explode or cause injury when opening. It is possible to use a metal slide holder in the microwave oven if the metal holder is submerged in pre-treatment solution during the whole microwave treatment. Do not otherwise use metal in a microwave oven. Follow the microwave manufacturer’s instructions. Avoid continuous boiling of the Pre-Treatment Solution during the 10 minutes incubation. Perform regular temperature control using a calibrated thermometer to verify correct temperature conditions. Step 2: Pepsin, ready-to-use (RTU) or pepsin solution

Pepsin incubation can be performed by direct application of RTU pepsin drops to the slides either

at room temperature (20-25 C) (Method A) or at 37 C (Method B). Alternatively, slides can be

immersed into a pepsin solution and incubated at 37 (±2) C (Method C).

Method A and Method B: Tap off excess buffer. Using lintless tissue (such as an absorbent wipe or gauze pad), carefully wipe around the specimen to remove any remaining liquid and to keep reagents within the prescribed area.

Apply 5-8 drops (250 µL) of cold (2-8 C) Pepsin (Vial 2A) to cover specimen. Always store Pepsin

at 2-8 C.

Method A: Pepsin, RTU - Incubation at 20-25 C

Incubate for 5-15 minutes at room temperature (20-25 C). An incubation time of 5-15 minutes will be adequate for most specimens, but the optimal incubation time may depend on tissue fixation and/or thickness of specimen and should be determined by the user.

Tap off excess Pepsin and soak sections in the diluted Wash Buffer (see INSTRUCTIONS FOR

USE, Section A.3) for 3 minutes at room temperature (20-25 C).

Replace diluted Wash Buffer and soak sections for another 3 minutes. Continue to dehydration.

Method B: Pepsin, RTU - Incubation at 37 C

Place specimen with Pepsin on a heating block at 37 C – e.g. Dako Hybridizer – and incubate for 3-5 minutes. An incubation time of 3-5 minutes will be adequate for most specimens, but the optimal incubation time may depend on tissue fixation and/or thickness of specimen and should be determined by the user.

Tap off excess Pepsin and soak sections in diluted Wash Buffer for 3 minutes at room temperature

(20–25 C).

Replace Wash Buffer and soak sections for another 3 minutes. Continue to dehydration.

Dehydrate tissue sections through a graded series of ethanol: 2 minutes in 70% ethanol, 2 minutes in 85% ethanol, and 2 minutes in 96% ethanol.

Allow tissue sections to air dry completely.

Method C: Pepsin solution - Immersion of slides into 37 C pepsin solution The kit contains reagents sufficient for four separate runs (60 mL pepsin solution, small container for six slides) or a single run (240 mL pepsin solution, large container for 24 slides). Prepare the pepsin solution as described in section A.5.

P04260US_02/K579911-5 p. 13/26

Put lid on the container and equilibrate the pepsin solution to 37 (±2) C in a water bath. Ensure that the temperature has stabilized. Measure temperature inside the container with a calibrated thermometer to ensure correct temperature.

Tap of excess wash buffer. Immerse slides into the 37 (±2) C pepsin solution and incubate for 20-30 minutes. An incubation time of 20-30 minutes will be adequate for most specimens, but the optimal incubation time may depend on tissue fixation and/or thickness of specimen and should be determined by the user.

Tap off exces pepsin solution and soak sections in diluted Wash Buffer for 3 minutes at room

temperature (20-25 C).

Replace Wash Buffer and soak sections for another 3 minutes. Continue to dehydration.



Step: 3 FISH probe diluted in ethylene carbonate-based hybridization buffer (provided by the user)

NOTE: FISH probe mixes diluted in ethylene carbonate-based hybridization buffer must be stored at -18 °C between uses. Storage at -18 °C results in separation of the buffer into two phases. Prior to use ensure that only one phase is present by equilibrating the probe mix to room temperature

(20-25 C) followed by mixing. Thaw the FISH probe mix at room temperature (20-25 °C) for a maximum of 30 minutes (protect from strong light), then thoroughly whirl the vial for 15 seconds at 2500 rpm using a vortex mixer.

Apply an appropriate amount of probe mix, e.g. 10 µL, to the center of the tissue section. Immediately place a 22 mm x 22 mm glass coverslip over the probe mix and allow it to spread evenly under the coverslip. Avoid air bubbles. If air bubbles are observed, gently tap them away from the tissue using forceps.

Seal coverslip with Coverslip Sealant by ejecting the Sealant around the periphery of the coverslip. Allow the Coverslip Sealant to overlap the coverslip and the slide, thereby forming a seal around the coverslip. Make sure that the Coverslip Sealant covers the entire edge of the coverslip.

Prepare Dako Hybridizer* (Code S2450 or S2451) for a hybridization run. Make sure that Humidity Control Strips (Code S2452) are saturated and optimal for use. Start the Hybridizer and choose a program that will:

Denature at 66 C for 10 minutes followed by hybridization at 45 C for 60-120 minutes.

Place slides in the Hybridizer, make sure the lid is properly closed and start program. Please refer to Dako Hybridizer Instruction Manual for details.

* Instrumentation that allows for conditions identical to the ones described above may be used for denaturation and hybridization:

Place slides on a flat metal or stone surface (heating block or on a block in a hybridization oven) preheated to 66 (±1) C. Denature for exactly 10 minutes. Place slides in a preheated humidified hybridization chamber. Cover the chamber with a lid

and incubate at 45 (±2) C for 60-120 minutes.

Step 4: Stringent Wash

Fill two staining jars, e.g. Coplin jars, with the diluted Stringent Wash Buffer (see INSTRUCTIONS FOR USE, Section A.2). A minimum volume of 100 mL or 15 mL per slide in each jar is recommended.

Place one of the staining jars containing diluted Stringent Wash Buffer at room temperature in a fume hood and the other in a water bath. Heat water bath and the diluted Stringent Wash Buffer to

63 (±2) C. Ensure that the temperature has stabilized. Cover jar with lid in order to stabilize the temperature and avoid evaporation. Measure temperature inside the water bath jar with a calibrated thermometer to ensure correct temperature. The Stringent Wash Buffer contains detergent and may become turbid at 63 °C; this will not affect performance.

Using forceps or gloves take slides from the hybridization chamber and gently remove Coverslip Sealant as well as coverslip and place slides in the room temperature pre-wash jar, one at a time.

P04260US_02/K579911-5 p. 14/26

As soon as all coverslips have been removed, transfer slides from the room temperature, pre-wash

jar to the 63 (±2) C jar in the water bath.

Immediately after transferring the slides into the 63 (±2) C diluted Stringent Wash Buffer in the water bath, the timer should be started. Perform stringent wash for exactly 10 minutes.

Remove slides from the diluted Stringent Wash Buffer, and soak sections in diluted Wash Buffer

for 3 minutes at room temperature (20-25 C).

Change diluted Wash Buffer and soak sections for another 3 minutes.


Allow tissue sections to dry completely. Step 5: Mounting

Apply 15 µL of Fluorescence Mounting Medium containing DAPI (Vial 5) to the target area of the slide and apply a glass coverslip.

NOTE: Slides may be read after 15 minutes or within 7 days after mounting. However, fading occurs if slides are exposed to light or high temperatures. To minimize fading, store slides in the

dark at -18-8 C.

B.3.b. Staining protocol for FISH probes diluted in formamide-based hybridization buffer

DAY 1

Step 1: Pre-Treatment (microwave oven, water bath, Pascal)

Pre-treatment can be performed either by using water bath as described in method A), by use of microwave oven with sensing capability as described in method B) or by use of Pascal pressure chamber as described in method C).

Method A: Pre-treatment using water bath Fill staining jars with diluted Pre-Treatment Solution (see INSTRUCTIONS FOR USE, Section A.1). Place staining jars containing diluted Pre-Treatment Solution in water bath. Heat water bath and

the Pre-Treatment Solution to 95-99 C. Measure temperature inside jar with a calibrated thermometer to ensure correct temperature. Cover jars with lids (no holes) in order to stabilize the temperature and avoid evaporation.

Immerse the deparaffinized sections into the preheated Pre-Treatment Solution.

Re-check temperature, close the water bath’s lid and incubate for 10 (±1) minutes at 95-99 C.

Remove the entire jar with slides from the water bath. Remove lid (do not remove slides). Allow the slides to cool in the Pre-Treatment Solution for 15 minutes at room temperature. Transfer the slides to a jar containing diluted Wash Buffer (see INSTRUCTIONS FOR USE, Section A.2) for 3 minutes at room temperature.


Method B: Pre-treatment using microwave oven with sensing capability (Recommended)

Fill a container with diluted room temperature (20-25 C) Pre-Treatment Solution. Immerse the deparaffinized sections into the solution and close the lid. The lid must contain holes allowing pressure to escape.

Place the container with slides into the microwave oven and heat. Before heating, make sure the outside of the container and microwave oven cavity are dry.

Incubate just below the boiling point (not less than 95 °C) for 10 minutes.

After incubation remove lid (do not remove slides).

Allow the slides to cool in the Pre-Treatment Solution for 15 minutes at room temperature (20-25 °C).

The slides must be covered with buffer during the whole pre-treatment procedure.

P04260US_02/K579911-5 p. 15/26

Transfer the slides to a jar containing diluted Wash Buffer (see INSTRUCTIONS FOR USE, Section A.2) for 3 minutes at room temperature.


Method C: Pre-treatment using Pascal pressure chamber Place 500 mL of deionized water inside the Pascal pan. Fill a plastic jar with 250 mL Pre-Treatment Solution. Immerse the deparaffinized sections in Pre-Treatment Solution and place the jar inside the Pascal pan. Incubate at 121 °C for 1 minute. Release pressure after cooling to 90 °C. Carefully read the Pascal Handbook for information on proper handling of the Pascal pressure chamber. Remove the container after the pressure is released. Remove lids and allow the slides to cool in the Pre-Treatment Solution for another 15 minutes at room temperature. Transfer the slides to a jar containing diluted Wash Buffer (see INSTRUCTIONS FOR USE, Section A.2). Soak sections for 3 minutes at room temperature.


NOTE: The Pre-Treatment Solution is designed for a single use only. Do not re-use. Do not use a sealed, airtight container when performing pre-treatment in a microwave oven as the pressure inside the container will increase and may explode or cause injury when opening. It is possible to use a metal slide holder in the microwave oven if the metal holder is submerged in pre-treatment solution during the whole microwave treatment. Do not otherwise use metal in a microwave oven. Follow the microwave manufacturer’s instructions. Avoid continuous boiling of the Pre-Treatment Solution during the 10 minutes incubation. Perform regular temperature control using a calibrated thermometer to verify correct temperature conditions.

Step 2: Pepsin, ready-to-use (RTU) or pepsin solution Pepsin incubation can be performed by direct application of RTU pepsin drops to the slides either

at room temperature (20-25 C) (Method A) or at 37 C (Method B). Alternatively, slides can be

immersed into a pepsin solution and incubated at 37 (±2) C (Method C)

Method A and method B: Tap off excess buffer. Using lintless tissue (such as an absorbent wipe or gauze pad), carefully wipe around the specimen to remove any remaining liquid and to keep reagents within the required area.

Apply 5–8 drops (250 µL) of cold (2–8 C) Pepsin (Vial 2A) ensuring that the specimen is covered.

Always store Pepsin at 2-8 C.

Method A: Pepsin, RTU – Incubation at 20-25 C

Incubate for 5-50 minutes at room temperature (20-25 C). An incubation time of 10-20 minutes at room temperature will be adequate for most specimens, but optimal incubation time should be determined by the user.

Tap off excess Pepsin and soak sections in the diluted Wash Buffer (see INSTRUCTIONS FOR

USE, Section A.3) for 3 minutes at room temperature (20-25 C).

Replace diluted Wash Buffer and soak sections for another 3 minutes. Continue to Step 3, FISH Probe or see Appendix 2 for optimization of digestion time (optional).

Method B: Pepsin, RTU – Incubation at 37 C

Place specimens with pepsin at 37 C on the Dako Hybridizer or on a preheated heating block.

Incubate for 2–6 minutes at 37 C. This will be adequate for most specimens prepared as described in the Specimen Preparation section. The optimal incubation time depends on tissue type, tissue fixation, thickness of specimen, specimen maturation and should be determined by the user. These factors may increase the required digestion time to 7–15 min or even higher. First time users should identify the optimal digestion time by testing a representative tissue for 1, 3, 6, 12 and 18 minutes.

Tap off Pepsin and soak sections in the diluted Wash Buffer (see INSTRUCTIONS FOR USE,

Section A.3) for 3 minutes at room temperature (20-25 C).

Replace diluted Wash Buffer and soak sections for another 3 minutes. Continue to Step 3, FISH Probe or see Appendix 2 for optimizing of digestion time (optional).

Method C: Pepsin solution - Immersion of slides into 37 C pepsin solution

P04260US_02/K579911-5 p. 16/26

The kit contains reagents sufficient for four seperate runs (60 mL pepsin solution, small container for six slides) or a single run (240 mL pepsin solution, large container for 24 slides). Prepare the pepsin solution as described in section A.5.

Put lid on the container and equilibrate the pepsin solution to 37 (±2) C in a water bath. Ensure that the temperature has stabilized. Measure temperature inside the container with a calibrated thermometer to ensure correct temperature.

Tap of excess wash buffer. Immerse slides into the 37 (±2) C pepsin solution and incubate for 20-30 minutes. An incubation time of 20-30 minutes will be adequate for most specimens, but the optimal incubation time may depend on tissue fixation and/or thickness of specimen and should be determined by the user.

Tap off excess pepsin solution and soak sections in diluted Wash Buffer for 3 minutes at room

temperature (20-25 C).

Replace Wash Buffer and soak sections for another 3 minutes. Continue to Step 3, FISH Probe or see Appendix 2 for optimizing of digestion time (optional).

NOTE: It is recommended that the procedure in Specimen Preparation section is followed. Under-digested tissue might result in no signals or only dizzy signals often with a high level of green cytosolic background.

Step 3: FISH Probe (provided by the user)

NOTE: If reagents from Dako Histology FISH Accessory Kit, Code K5799, are used together with reagents from Code K5731, the procedure outlined in the package insert for Code K5731 must be followed.

The following step should be performed in a fume hood. Fluorochrome-labeled probes may be affected adversely if exposed to excessive light levels. Do not perform the remainder of this procedure in strong light, such as direct sunlight.

Dehydrate tissue sections through a graded series of ethanol: 2 minutes in 70% ethanol, 2 minutes in 85% ethanol, and 2 minutes in 96% ethanol (see INSTRUCTIONS FOR USE, Section A.4).


Apply an appropriate amount of fluorochrome-labeled probe, e.g. 10 µL of a Dako FISH probe to the centre of the tissue section. Immediately place an 18 mm x 18 mm (or e.g. 22 mm x 22 mm) glass coverslip over the probe and allow it to spread evenly under the coverslip. Avoid air bubbles. If air bubbles are observed, gently tap them away using forceps.

Seal coverslip by applying the Coverslip Sealant around the periphery of the coverslip. Allow the Coverslip Sealant to overlap the coverslip and the slide, thereby forming a seal around the coverslip. Make sure that the Coverslip Sealant seals the entire edge of the coverslip.

Place slides in Dako Hybridizer (Code S2450/S2451) and set the denaturation to 82 °C for 5 min and hybridization to 45 °C overnight (14–20 h) when using Dako FISH probes. Continue to Day 2, Step 4, Stringent Wash. See the Hybridizer Handbook for further instructions.

NOTE: A heating block, hybridization oven and hybridization chamber can alternatively be used, see below.

Place slide on a flat metal or stone surface (heating block or on a block in a hybridization oven)

preheated to 82 (±2) C. Denature for 5 minutes ensuring that the temperature of the block does

not drop below 80 C.

Place slides in a preheated humidified hybridization chamber. Cover the chamber with a lid and

incubate overnight (14–20 hours) at 45 (±2) C when using Dako FISH probes.

DAY 2

Step 4: Stringent Wash Fill two staining jars with diluted Stringent Wash Buffer (see INSTRUCTIONS FOR USE, Section A.2). A minimum volume of 100 mL diluted Stringent Wash Buffer is required for 1–6 slides in a jar. For more than 6 slides use at least 15 mL buffer per slide.

Place one of the staining jars containing diluted Stringent Wash Buffer at room temperature in a fume hood and the other in a water bath. Heat water bath and the diluted Stringent Wash Buffer to

P04260US_02/K579911-5 p. 17/26

65 (±2) C. Ensure that the temperature has stabilized. Cover jar with lid in order to stabilize the temperature and avoid evaporation. Measure temperature inside the jar with a calibrated thermometer to ensure correct temperature.

Take slides from the hybridization chamber. Using forceps, gently remove the Coverslip Sealant as well as the coverslip and transfer the slide to the room temperature pre-wash jar.

As soon as all coverslips have been removed, transfer slides from the room temperature, pre-wash

jar to the 65 (±2) C jar in the water bath. Close jar lid and then water bath lid. Perform stringent

wash for exactly 10 minutes at 65 (±2) C.

Remove slides from the diluted Stringent Wash Buffer and soak sections in diluted Wash Buffer for

3 minutes at room temperature (20–25 C).

Change diluted Wash Buffer and soak sections for another 3 minutes.

Dehydrate tissue sections through a graded series of ethanol: 2 minutes in 70% ethanol, 2 minutes in 85% ethanol, and 2 minutes in 96% ethanol (see INSTRUCTIONS FOR USE, Section A.4).

Allow slides to completely air-dry.

Step 5: Mounting Apply 15 µL of Fluorescence Mounting Medium (Vial 5), containing blue fluorescence counterstain, to the target area of the slide and apply a glass coverslip (e.g. 24 mm x 50 mm or 24 mm x 60 mm). Allow the medium to spread evenly over the specimen.

NOTE: Slides may be read after 15 minutes or within 7 days after mounting. Fading occurs if slides

are exposed to light or high temperatures. Slides should be stored in the dark at -18–8 C.

Quality Control 1. Signals must be bright, distinct and easy to evaluate.

2. Normal tissue previously shown to “FISH well” should be used for control of the assay run.

3. Normal tissue cells should have clearly visible fluorescence signals, indicating that the FISH probes have successfully hybridized to the target regions.

4. Failure to detect signals in normal tissue cells indicates assay failure, and results should be considered invalid.

5. Due to tissue sectioning, some normal cells will have less than the expected number of signals.

6. Nuclear morphology must be intact and homogenously stained when evaluated using a DAPI filter. Numerous ghost-like cells, donut cells and a general poor nuclear morphology indicate over-digestion or over-denaturation of the specimen, which can result in loss or fragmentation of signals. Peripheral staining, holes in cells (DAPI filter) and/or strong green cytosolic background staining when observed with a Texas Red/FITC double filter might indicate under-digestion, which can result in loss of signal. Such specimens should be considered invalid.

7. Differences in tissue fixation, processing, and embedding in the user’s laboratory may produce variability in results, necessitating regular evaluation of in-house controls.

P04260US_02/K579911-5 p. 18/26

Interpretation of Results Scan several areas of cells to account for possible heterogeneity. Select an area having good nuclei distribution. Begin analysis in the upper left quadrant of the selected area and, scanning from left to right, count the number of signals within the nuclear boundary of each evaluated nucleus, according to the guidelines below.

▪ Focus up and down to find all of the signals in the individual nucleus.

▪ Do not score nuclei with no signals.

▪ Do not include nuclei that require subjective judgement. Skip nuclei with weak signal intensity and non-specific or high background.

▪ The user must determine the number of nuclei that should be counted for each probe.

▪ Avoid areas where the nuclear borders are ambiguous.

P04260US_02/K579911-5 p. 19/26

Troubleshooting

Problem Probable Cause Suggested Action

1. No signals or weak signals

1a. Inappropriate digestion time.

1a. Ensure that formalin-fixed, paraffin-embedded tissue sections are used.

A prolonged digestion time of e.g. 7–15 minutes or even

higher at 37 C might help. See Section B.3.a or B.3.b Step 2, Troubleshooting point 4d, 4e and Appendix 2.

1b. Kit has been exposed to high temperatures during transport or storage.

1b. Check storage conditions. Ensure that dry ice was present when the consignment was received. Ensure that vials 2A and 5

have been stored at 2–8 C, and that vial 5 has been stored in the dark.

1c. Pre-treatment conditions incorrect.

1c. Ensure that the recommended pre-treatment temperature and time are used. Furthermore that the digestion incubation time has been optimized if required, see Section B.3.a or B.3.b Step 2. The closer the pre-treatment temperature is to the boiling point the stronger signals.

Ensure that the Pepsin is handled at the correct temperature. See Section B.1.

1d. Denaturation conditions incorrect.

1d. Ensure that the recommended denaturation temperature and time are used.

1e. Hybridization temperature incorrect.

1e. Hybridize at 45 (±2) C when using Dako FISH probes.

1f. Evaporation of probe buffer during hybridization.

1f. Ensure sufficient humidity in the hybridization chamber. Use Dako Hybridizer (Code S2450/S2451) and Hybridizer Humidity Control Strips (Code S2452). Use Coverslip Sealant. See Troubleshooting point 5a and 5b.

P04260US_02/K579911-5 p. 20/26

1g. Stringency wash conditions incorrect.

1g. Ensure that the recommended stringency wash volume, temperature and time are used, and that coverslips are removed before performing stringent wash.

1h. Microscope not functioning properly

- Inappropriate filter set - Unsuitable lamp - Mercury lamp too old - Burnout filters - Dirty and/or cracked collector

lenses

- Unsuitable immersion oil

1h. Check the microscope and ensure that the used filters are suitable for use with the fluorochromes, that they are not worn out and that the mercury lamp is suitable and has not been used beyond expected lifetime (See Appendix 1). Use suitable immersion oil for fluorescence. In case of doubt, please contact your local microscope vendor.

1i. Faded signals. 1i. Avoid extended microscopic examination and minimize exposure to strong light sources.

2. Areas without signal

2a. Probe volume too small.

2a. Ensure that the probe volume is large enough to cover the area under the coverslip.

2b. Air bubbles caught during probe application or mounting.

2b. Avoid air bubbles.

If observed, gently tap them away using forceps.

2c. Poor deparaffinization. 2c. Ensure that the paraffin has been complete removed from sections. See section B.2.

3. Excessive back-ground staining

3a. Inappropriate digestion time. Strong green cytosolic background might indicate under-digestion.

3a. Ensure that formalin-fixed, paraffin-embedded tissue sections are used.

A prolonged digestion time of e.g. 7–15 minutes at

37 C might help. See Section B.3.a or B.3.b. Step 2, Troubleshooting point 4e and Appendix 2.

3b. Sections have dried in B.3.

3b. Follow the instructions and avoid drying of slides unless specified.

3c. Paraffin incompletely removed.

3c. Follow the deparaffinization and rehydration procedures outlined in Section B.2

P04260US_02/K579911-5 p. 21/26

3d. Stringency wash temperature too low.

3d. Ensure that the recommended stringency wash temperature is used.

3e. Prolonged exposure of hybridized section to strong light.

3e. Avoid extended microscopic examination and minimize exposure to strong light.

3f. Expired glass slides or non-suitable slides may cause an excessive red “star sky” staining.

3f. Use the recommended slide types and secure that they are not expired. See “Paraffin-embedded sections” section.

3g. Non-fluorescing Immersion oil mixed with the Fluorescence Mounting Medium.

3g. Use large glass coverslips when mounting as recommended. See Section B.3.a or B.3.b, Step 5. This prevents immersion oil from mixing with mounting medium, avoiding auto fluorescence and accidental removal of coverslips by the microscopes objective.

4. Poor nuclei morphology or weak nuclei staining

4a. Incorrect pre-treatment conditions may result in unclear or cloudy appearance.

4a. Ensure that the recommended pre-treatment temperature and time are used. See also Troubleshooting point 4b-e.

4b. Boiling during the pre-treatment may result in morphology damage and lack of signals.

4b. Avoid boiling. See Section B.3, Step 1. See also Troubleshooting point 4d.

4c. Incorrect Pepsin treatment. 4c. Adhere to recommended Pepsin incubation times. See Section B.3, Step 2 and Appendix 2. See also Troubleshooting point 1a, 1c 3a, 4d and 4e.

4d. Too long Pepsin treatment will dissolve the tissue and result in lack of signals. Over-digestion may cause ghost cells or donut nuclei and nuclei with a general poor nuclear morphology to appear.

4d. Shorten the Pepsin incubation time. See Section B.3.a or B.3.b, Step 2 and Appendix 2. Ensure that the section thickness is 2–6 µm. See also Troubleshooting point 1a, 1c, 4b and 4e.

P04260US_02/K579911-5 p. 22/26

4e. Too short Pepsin treatment may result in lack of signals. Under-digested tissue may cause peripheral nuclei stain and holes in nuclei (DAPI filter); high green background in cytosol (Texas Red/FITC double filter). Notice that nuclei in under-digested tissue have nice morphology in contrast to over-digested tissue.

4e. Prolong the Pepsin incubation time to e.g. 2–3 times the used digestion time. See also Troubleshooting point 1a, 1c, 3a, 4a and 4d.

4f. Denaturation temperature too high.

4f. Ensure that the recommended denaturation temperature is used.

4g. Incorrect hybridizations conditions.

4g. See Troubleshooting point 1f and 5b.

5. Coverslip adhere strongly to glass slide after hybridization and thereby hard to remove.

5a. Coverslip inappropriately sealed to glass slide.

5a. Do not force off the coverslip if the coverslip adhere strongly to glass slide. Immerse slide in the jar containing diluted Stringent Wash Buffer at room temperature for five minutes and then remove coverslip. Follow recommendations for coverslip sealing in Section B.3a or B.3.b, Step 3. See also Troubleshooting point 1f and 5b.

5b. Incorrect hybridizations conditions. Might cause reduced or no signals, tissue damage and background staining.

5b. Ensure sufficient humidity in the hybridization chamber. Use Dako Hybridizer (Code S2450/S2451) and Hybridizer Humidity Control Strips (Code S2452). Follow instructions given in Package Insert for Hybridizer Humidity Control Strips. See recommended hybridization conditions in Section B.3.a or B.3.b, Step 3. See also Troubleshooting point 1f and 5a.

6. High level of green autofluorescence on slide including areas without FFPE tissue

6. Use of expired or unrecommended glass slides

6. Ensure that the coated glass slide (Dako Silanized Slides, Code S3003, or poly-L-lysine-coated slides) have not passed expiry date.

NOTE: If the problem cannot be attributed to any of the above causes, or if the suggested corrective action fails to resolve the problem, please call Dako Technical Services for further assistance.

P04260US_02/K579911-5 p. 23/26

Appendix 1

Histology FISH Accessory Kit, Code K5799

Fluorescence Microscope Specifications

Dako recommends the following equipment for use with the Histology FISH Accessory Kit when using Dako FISH probes:

1. Microscope type

• Epifluorescence microscope.

2. Lamp

• 100 watt mercury lamp (should generally be replaced every 200 hours).

3. Objectives

• For screening of the tissue, fluorescence dry 10x or fluorescence oil immersion 16x objectives are applicable.

• For high power magnification and scoring of signals, only fluorescence oil immersion objectives, e.g. 63x or 100x are recommended.

4. Filters

Filters are individually designed for specific fluorochromes and must be chosen accordingly. Dako recommends the use of a specific DAPI filter in combination with a high quality Texas Red/FITC double filter when using Dako FISH probes.

• DAPI filter.

• Texas Red/FITC double filter.

• Texas Red and FITC single filters can be used for confirmation.

Fluorochrome Excitation Wavelength Emission Wavelength

FITC 495 nm 520 nm

Texas Red 596 nm 615 nm

Filters are specific to each microscope type and the use of appropriate filters is crucial for the interpretation. Further detailed information, can be provided by your microscope manufacturer or your Dako representative or Dako Web site.

5. Oil

• Non-fluorescing immersion oil.

Precautions

Specific fluorochromes require different equipment. For Dako FISH probes a 50 watt mercury lamp is not recommended, rhodamine filters cannot be used, and triple filters are in general not recommended.

A non-optimized microscope may cause problems when reading the fluorescent signals. It is important that the light source has not expired and that it is properly aligned and focused.

Customers should monitor and follow the manufacturer’s recommendations for the mercury lamp and the microscope should be maintained.

An effort should be made to expose the sample to as little of the excitation light as possible in order to minimize fading of the fluorescence.

We recommend that you discuss the set-up of your particular microscope with the manufacturer before starting the fluorescence in situ hybridization, or refer to relevant literature.

Appendix 2

Optional step for optimizing Pepsin digestion time

P04260US_02/K579911-5 p. 24/26

This optional step can be used to evaluate the effect of the pepsin digestion and to help optimizing the digestion time used in Step 2, Pepsin.

To control whether the used pepsin digestion time is sufficient, stain the washed pepsin-digested tissue section with user provided 10 µL propidium iodide (400 ng/mL). Place a 22 mm x 22 mm glass coverslip over the propidium iodide and allow it to spread evenly under the coverslip. Incubate for 1 minute.

Use a fluorescence microscope with Texas Red/FITC double filter (see Appendix 1) to evaluate the red staining of nuclei with propidium iodide. If the tissue digestion is acceptable there should be red nuclei with well-defined perimeters. Remove propidium iodide by soaking sections in diluted Wash

Buffer twice for 3 minutes at room temperature (20-25 C) and continue to Section B3, Staining protocol, Step 3, FISH Probe.

If the nuclei are still green by autofluorescence and not stained red by propidium iodide it is necessary to repeat the digest cycle:

Soak sections in diluted Wash Buffer twice for 3 minutes at room temperature (20-25 C) to remove propidium iodide.

Apply 5–8 drops (250 µL) of cold (2-8 C) Pepsin (Vial 2) to cover specimen. Always store the

Pepsin vial at 2–8 C. Place slides at 37 C on a preheated heating block or on the Dako Hybridizer. Incubate for 10 minutes if no or little digestion has occurred. The 10 minutes is only a guideline; the user should determine the optimal time.

Soak again in diluted Wash Buffer twice for 3 minutes at room temperature (20-25 C) to remove Pepsin.

Apply once more 10 µL propidium iodide (400 ng/mL) for 1 minute. Evaluate staining and soak

sections in diluted Wash Buffer twice for 3 minutes at room temperature (20-25 C). Repeat cycle if necessary or continue to Section B.3.a or B.3.b, Staining protocol, Step 3, FISH Probe.

NOTE: The kit contains reagents sufficient for 20 tests. The use of Wash Buffer and Pepsin in this optional step will reduce the total number of possible tests with the kit.

P04260US_02/K579911-5 p. 25/26

References

1. Clinical Laboratory Improvement Amendments of 1988: Final Rule, 57 CFR 7163, February 28, 1992.

2. Sheehan DC, Hrapchak BB. Theory and practice of histotechnology. St. Louis: CV Mosby Company; 1980.

3. Brown RSD, Edwards J, Bartlet JW, Jones C, Dogan A. Routine acid decalcification of bone marrow samples can preserve DNA for FISH and CGH studies in metastatic prostate cancer. J Histochem Cytochem 2002 50:113-5.

4. Alers JC, Krijtenberg P-J, Visser KJ, van Dekken H. Effect of bone decalcification procedures on DNA in situ hybridization and comparative genomic hybridization: EDTA is highly preferable to a routinely used acid decalcifier. J Histochem Cytochem 1999 47:703-9.

5. Provan AB, Hodges E, Smith AG, Smith JL. Use of paraffin wax embedded bone marrow trepine biopsy specimens as a source of archival DNA. J Clin Pathol 1992 45:763-5.

6. Sarsfield P, Wickham CL, Joyner MV, Ellard S, Jones DB, Wilkins BS. Formic acid decalcification of bone marrow trephines degrades DNA: alternative use of EDTA allows the amplification and sequencing of relatively long PCR products. Mol Pathol 2000 53:336.

7. Reineke T, Jenni B, Abdou MT, Frigerio S, Zubler P, Moch H, Tinguely M. Ultrasonic decalcification offers new perspectives for rapid FISH, DNA, and RT-PCR analysis in bone marrow trephines. Am J Surg Pathol 2006 30: 892-6.

8. Kiernan JA. Histological and histochemical methods: theory and practice. New York: Pergamon Press; 1981.

P04260US_02/K579911-5 p. 26/26

Explanation of symbols

Catalogue number

Temperature limitation

Batch code

In vitro diagnostic medical device

Keep away from sunlight (consult storage section)

Use by

Consult instructions for use

Contains sufficient for <n> tests

Manufacturer

GHS hazard pictogram (consult precautions section)




Revision 2017.08

Manufactured by: Dako Denmark A/S Produktionsvej 42 DK-2600 Glostrup Denmark Tel. +45 44 85 95 00 Fax +45 44 85 95 95 www.agilent.com

Distributed in the US by: Dako North America, Inc. 6392 Via Real Carpinteria, California 93013, USA Tel. 805/566-6655, Toll Free: 800/235-5743 Ordering Information: Tel. 800/235-5763 Fax 805/566-6688 Technical Services: Tel. 800/424-0021

http://www.agilent.com/

Histology FISH Accessory Kit - Agilent · P04260US_02/K579911-5 p. 3/26 Intended Use For in vitro...

Documents

Transcript of Histology FISH Accessory Kit - Agilent · P04260US_02/K579911-5 p. 3/26 Intended Use For in vitro...