Dec 11, 2017
Paraffin embedding, microtomy and fluorescence in situ hybridization (FISH) of whole adult Drosophila
- Carolyn Elya1,
- Ciera Martinez1
- 1Department of Molecular and Cell Biology, University of California, Berkeley, CA
External link: https://doi.org/10.7554/eLife.34414
Protocol Citation: Carolyn Elya, Ciera Martinez 2017. Paraffin embedding, microtomy and fluorescence in situ hybridization (FISH) of whole adult Drosophila. protocols.io https://dx.doi.org/10.17504/protocols.io.k5ecy3e
Manuscript citation:
Elya C, Lok TC, Spencer QE, McCausland H, Martinez CC, Eisen M, Robust manipulation of the behavior of by a fungal pathogen in the laboratory. eLife doi: 10.7554/eLife.34414
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We currently use this protocol and it is working.
Created: December 04, 2017
Last Modified: March 11, 2018
Protocol Integer ID: 9094
Abstract
Paraffin embedding and sectioning is a classic technique used in a variety of biological disciplines. Though the fruit fly Drosophila melanogaster are one of the most commonly-employed model organisms, the immense Drosophila literature is lacking in current protocols for sectioning whole, adult flies. Here, we adopted a plant histology protocol to embed entire adult flies for thin sectioning and present our protocol for performing FISH on the subsequent sections. In the interest of reproducibility, we have tried to include details that are normally omitted from published protocols.
Guidelines
You will need the following reagents, consumables and equipment for each of the three steps as follows:
I. Paraffin Infiltration of Adult Flies
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II. Sectioning embedded flies via microtome
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III. FISH on paraffin sectionsCold room or refrigerator
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*We’ve successfully used an 18mer DNA probe tagged with 5’ Alexfluor-647 (Thermo Fisher) that targets a ~11,000-fold copied sequence in genome.
#Consists of wet paper towels and something to prop up slides above towels in a sealable plastic container. Construction described in protocol.
Materials
MATERIALS
Chloroform
Acetic Acid (Glacial)Bio Basic Inc.Catalog #AC1001.SIZE.1L
Ethanol, 200 proof
double distilled water (ddH2O)
HistoclearCatalog #HS2001GLL
ParaplastCatalog #39501006
Glass scintillation vialsFisher ScientificCatalog #FS74500-20
Glass seralogical pipettes
Glass Pasteur pipettes with rubber bulbs
Graduated plastic transfer pipettes Fisher ScientificCatalog #13-711-9BM
Embedding ringsElectron Microscopy SciencesCatalog #62350W
Embedding base moldsFisher ScientificCatalog #22-363-553
Razor bladesFisher ScientificCatalog #12-640
Polysine slidesFisher ScientificCatalog #12-545-78
ProLong Gold with DAPICatalog #P36931
Hydrochloric acidMerck MilliporeSigma (Sigma-Aldrich)
Tris-HCl
Sodium chloride
Sodium dodecyl sulphate
Formamide
Potassium chloride
Disodium phosphate
Monopotassium phosphate
Triton X-100
818 high profile microtome blades
Safety warnings
- Histoclear is very toxic to marine life. Do not dispose of Histoclear down the drain.
- All chemicals should be used according to SDS recommendations.
I. Paraffin Infiltration of Adult Flies: Day 1 - Fixation
I. Paraffin Infiltration of Adult Flies: Day 1 - Fixation
Prepare fresh Carnoy’s solution (6:3:1 ethanol:chloroform:acetic acid) and chill to 4C before use.
For each sample, anesthetize adult flies and transfer to appropriate glass scintillation vial containing ice cold Carnoy’s.
Incubate samples in Carnoy’s solution at 4C at least overnight and no longer than 36 hours.
I. Paraffin Infiltration of Adult Flies: Day 2 - Dehydration
I. Paraffin Infiltration of Adult Flies: Day 2 - Dehydration
Wash each sample three times in 70% ethanol as follows:
- Aspirate solution from each scintillation vial with glass pipette and discard.
- Add 5-10 mL 70% EtOH and incubate at RT 5 min.
Incubate samples in the following ethanol concentrations for 1 hour each at RT:
- 70%
- 85%
- 95%
- 100%*
- 100%
01:00:00 70% ethanol
01:00:00 85% ethanol
01:00:00 95% ethanol
01:00:00 100% ethanol
Note
*Samples can be stored after this step overnight at 4C
01:00:00 100% ethanol*
Replace 100% ethanol and incubate at 4C overnight.
I. Paraffin Infiltration of Adult Flies: Day 3 - Tissue infiltration with Histoclear (Xylene alternative)
I. Paraffin Infiltration of Adult Flies: Day 3 - Tissue infiltration with Histoclear (Xylene alternative)
Incubate samples in the following solutions for 2 hours each at RT using glass pipettes and beakers (Histoclear melts plastics):
- 3:1 Ethanol:Histoclear
- 1:1 Ethanol:Histoclear
- 1:3 Ethanol:Histoclear
- Histoclear*
- Histoclear
- Histoclear
02:00:00 3:1 Ethanol:Histoclear
02:00:00 1:1 Ethanol:Histoclear
02:00:00 1:3 Ethanol:Histoclear
02:00:00 Histoclear
02:00:00 Histoclear
Note
*Samples can be stored after this step overnight at 4C
02:00:00 Histoclear*
I. Paraffin Infiltration of Adult Flies: Day 4 - Paraffin infiltration
I. Paraffin Infiltration of Adult Flies: Day 4 - Paraffin infiltration
Incubate samples in new Histoclear for 2 hours at RT.
02:00:00 Histoclear
Add 10-15 paraffin chips to each sample and incubate at least 3 hours at RT with gentle orbital shaking.
Add 15 paraffin chips to each sample and incubate overnight at RT with gentle orbital shaking.
Melt paraffin chips overnight at 60C in Pyrex measuring cup.
I. Paraffin Infiltration of Adult Flies: Day 5 - Paraffin infiltration cont'd
I. Paraffin Infiltration of Adult Flies: Day 5 - Paraffin infiltration cont'd
Unscrew scintillation vial caps by ¼ turn. Incubate samples at 42C until paraffin is dissolved (about 1.5 hours).
01:30:00 Check that paraffin has melted
Aspirate Histoclear/paraffin solution from samples and discard.
Cover samples with ~10 mL molten paraffin and replace cap leaving ¼ turn open. Incubate at least 3 hours at 60C (with optional 50 rpm shaking).
Perform paraffin change as follows:
- Remove samples one at a time from incubator.
- Make sure your flies have sunk to the bottom of the scintillation vial.
- Press bottom of scintillation against gloved palm until the paraffin at the bottom of the vial hardens (it will become opaque), holding your flies in place.
- Carefully decant and discard the molten paraffin.
- Add a new ~10 mL of molten paraffin and replace cap leaving ¼ turn open.
Continue incubating samples at 60C (with optional 50 rpm shaking) overnight. Melt additional paraffin as needed.
I. Paraffin Infiltration of Adult Flies: Day 6 - Paraffin infiltration cont'd
I. Paraffin Infiltration of Adult Flies: Day 6 - Paraffin infiltration cont'd
Perform paraffin change in the morning. Continue incubating samples at 60C for at least 3 hours.
Perform paraffin change in the afternoon. Continue incubating samples at 60C for at least 3 hours.
Perform paraffin change in the late afternoon/evening. Continue incubating samples at 60C overnight. Melt additional paraffin as needed.
I. Paraffin Infiltration of Adult Flies: Day 7 - Paraffin infiltration cont'd
I. Paraffin Infiltration of Adult Flies: Day 7 - Paraffin infiltration cont'd
Perform paraffin change in the morning. Continue incubating samples at 60C for at least 3 hours.
Perform paraffin change in the afternoon. Continue incubating samples at 60C for at least 3 hours.
Perform paraffin change in the late afternoon/evening. Continue incubating samples at 60C overnight. Melt additional paraffin as needed.
I. Paraffin Infiltration of Adult Flies: Day 8 - Paraffin infiltration cont'd
I. Paraffin Infiltration of Adult Flies: Day 8 - Paraffin infiltration cont'd
Perform paraffin change in the morning. Continue incubating samples at 60C for at least 3 hours.
Perform paraffin change in the afternoon. Continue incubating samples at 60C for at least 3 hours.
Perform paraffin change in the late afternoon/evening. Continue incubating samples at 60C overnight. Melt additional paraffin as needed.
Note
Embedding should be performed as soon after fly infiltration has been completed. Flies can be safely left in molten agar for up to 3 days following infiltration. Flies can remain in molten agar beyond this, but sample integrity may decline (tissues may soften). How long a sample can remain in molten agar should be determined on case-by-case basis.
I. Paraffin Infiltration of Adult Flies: Day 9 - Embedding flies in paraffin
I. Paraffin Infiltration of Adult Flies: Day 9 - Embedding flies in paraffin
Pre-warm microslide warming table. Pre-warm two transfer pipettes with last “segment” snipped off by placing in 60C incubator.
Note
Experience suggests embedding only one fly per ring/base, though this is left to the user’s discretion. The orientation in which a sample is embedded is also up to the user.
Place embedding mold base onto hottest side of microplate warmer to heat.
Remove one sample from 60C. Using pre-warmed pipette, aspirate up a fly and quickly deposit into base mold, filling base mold to cover fly in paraffin.
Note
Make sure that your fly is touching the bottom of the embedding base and as close to the middle as possible. Samples that are embedded next to the side tend to break or otherwise section poorly.
Expected result
Top base mold with labeled embedding ring.
Expected result
Slide base mold to the middle of the heat gradient. Quickly use tools (forceps or poker) to orient fly in desired position.
Note
Clean wax off of tools periodically using flame to prevent unwanted sticking.
Using pre-warmed pipette, add a mL or so of molten paraffin to begin filling the embedding ring.
Note
Act quickly! You want all of the paraffin to harden together in a continuous mass. If you allow a layer of paraffin to cool before adding the next, your sample will not cut evenly.
Slide sample all the way to the cool end of the gradient.
Continue adding molten paraffin as the previous paraffin has just set but not hardened to continue filling the mold.
Once mold is almost full, allow it to set completely before transferring to the coolest end of the plate for further cooling.
Allow samples to completely set overnight before sectioning.
Expected result
II. Sectioning embedded flies via microtome
II. Sectioning embedded flies via microtome
Tape 5 or 10 mL serological pipette to the heat block.
Heat slide block to 42C.
Clean the area adjacent to the microtome with 70% ethanol.
Place up to 6 slides on the heat block, making sure that polysine side is facing up.
Cover polysine-coated area with ddH2O.
Prepare a sample for sectioning by carefully removing ring from base (do this gently so as not to break your sample off into the plastic base).
Starting from the edge of the paraffin block, use a razor blade to slowly cut away excess paraffin on each side of your sample.
Note
If you try to make a direct cut next to your sample, the paraffin will break and your sample will be lost.
Clamp your sample onto the microtome. Do not over tighten the clamp.
Move the blade cassette away from your sample before installing a fresh blade.
Move the blade cassette toward your sample until you are a few hundred microns away.
In “coarse” mode set to 30 um intervals, progress the blade until you make contact with the paraffin.
Change microtome to “fine” mode and set to 8 um. Make one slice then use your pointy tool to gently hold the slice down.
Make an additional slice, checking that your first slice adheres to the second to start forming a ribbon. If it does not, discard unattached slices and try again.
Continue slicing slowly, taking care to keep the ribbon coming straight away from the blade. As the ribbon grows, you will need to use your teasing needle (or similar tool) to gently pick up and support the growing ribbon.
Expected result
Continue slicing, supporting the ribbon until you have sliced through the entire sample.
Use a new razor blade to cut the ribbon away from the paraffin block.
Using forceps and the teasing needle, carefully lay the ribbon on the bench.
Expected result
Using a slide as reference, cut ribbon ~2 sections shorter than the slide length.
Carefully transfer the ribbon, shiny side facing down, onto a water-coated slide. Arranged the ribbon as straight as you can.
Note
As teasing needle, forceps and razor blade start to retain paraffin, they will grow sticky. Razor blades should be properly disposed of and replaced with a new blade. Paraffin can be removed from forceps and teasing needle by passing the tips through the flame.
Continue adding ribbons in rows to the slide until you cannot safely add additional ribbons.
Allow a few seconds for all of the ribbons to unwrinkle then use transfer pipette to start removing water from underneath ribbons. Be careful to keep an eye on your ribbons as they will slide when you do this (you can use your teasing needle to hold them in place).
Gently tilt the slide and remove as much water as possible from the slide.
Place slide to dry leaning against the serological pipette taped to the 42C block.
Note
If you place the slide directly on the 42C block, you will damage the tissue.
When you are finished slicing your sample, remove paraffin ring. If you wish to slice additional samples, slide the razor so that you are cutting with a unused portion of the blade.
When you are finished slicing all samples, incubate slides at 42C overnight to dry. After overnight drying, slides can be used for histology, immunohistochemistry or fluorescence in situ hybridization (FISH).
Note
For best results, FISH and immunhistochemistry should be performed as soon after the slides are ready (i.e. the day after sectioning). Note that this protocol has been optimized to work with 8 micron sections.
Expected result
III. FISH on paraffin sections: Day 1
III. FISH on paraffin sections: Day 1
Prepare solutions:
Hybridization solution
20 mM Tris-HCl [pH 8.0]
0.9M NaCl
0.01% sodium dodecyl sulphate
30% formamide
PBSTx
0.8% NaCl [w/v]
0.02% KCl [w/v]
0.115% Na2HPO4 [w/v]
0.02% KH2PO4 [w/v]
0.3% Triton X-100 [v/v]
Warm 0.2 M HCl in staining dish to 37C.
Remove paraffin by incubating slides in 2x changes of Histoclear for 10 min each.
00:10:00 Histoclear 1
00:10:00 Histoclear 2
Rehydrate sections by incubating slides in a graded ethanol series as follows:
- 100% ethanol – 2x 5 min
- 95% ethanol - 2 min
- 85% ethanol - 2 min
- 70% ethanol - 2 min
00:02:00 70% ethanol
00:05:00 100% ethanol, wash 1
00:05:00 100% ethanol, wash 2
00:02:00 95% ethanol
00:02:00 85% ethanol
Unmask antigens by incubating slides in 0.2 M HCl (in water) for 60 min at 37C. Rinse 2-3x in ddH2O to remove acid.
01:00:00 Antigen unmasking
Prepare 80 uL of 100 pmol/µL FISH probe in hybridization buffer for each slide.
Note
100 pmol/µL is the optimized concentration for our probe, but it might not be for yours. The best probe concentration should be determined empirically on a case-by-case basis.
Apply 80 uL of 100 pmol/uL FISH probe to each slide and top with Hybrislip.
Note
Be careful not to introduce any bubbles! If you wind up with bubbles, you can carefully remove Hybrislip and try to lay down again without bubbles.
Put the glass slides in a humid chamber and leave at room temperature, overnight, in the dark.
Note
A humid chamber can be easily constructed using a sealable tupperware container or other appropriate vessel. Line the container with paper towels and apply DI water until completely wet (no standing water). Place spacers (e.g. dowels) on the wet paper towels to prop up your slides. Place the slides on top of the spacers, making sure that they are horizontal, before closing the container.
Note
From this point onward, be sure to keep your slides in the dark as much as possible to avoid bleaching of your probe.
Expected result
III. FISH on paraffin sections: Day 2
III. FISH on paraffin sections: Day 2
The next day, carefully remove the coverslip by gently dunking the slides one-at-a-time in a Coplin jar filled with freshly prepared PBSTx.
Gently dunk the slides 5x in PBSTx to wash off probe.
Note
If you find you have high background fluorescence, you can increase the amount of washing by placing slides in a Coplin jar containing 1x PBSTx and incubating on an orbital shaker at low speed for 5 minutes. Be sure to cover the Coplin jar in foil to prevent your probe from bleaching.
Dunk slides in 1x PBS to remove all detergent (~3 times)
Apply ~80 µl of ProLong Gold (antifade reagent) with DAPI directly onto the slide and cover the slide with a 60 x 24 mm coverslip. Avoid air bubbles.
III. FISH on paraffin sections: Day 2
III. FISH on paraffin sections: Day 2
Allow the slides to dry ~24 hours in the dark at RT before viewing (to cure the ProLong Gold).
Note
Slides should be immediately imaged at 10x-40x with the appropriate excitation and emission filters.
Expected result