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Method Article

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

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DOI:

10.3791/60575

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January 29th, 2020

 , 
1Department of Biological Sciences, Murray State University

In This Article

Summary

Here, we present a protocol for fluorescent antibody-mediated detection of proteins in whole preparations of zebrafish embryos and larvae.

Abstract

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. Although many immunohistochemistry protocols have been optimized for mammalian tissue and tissue sections, these protocols often require modification and optimization for non-mammalian model organisms. Zebrafish are increasingly used as a model system in basic, biomedical, and translational research to investigate the molecular, genetic, and cell biological mechanisms of developmental processes. Zebrafish offer many advantages as a model system but also require modified techniques for optimal protein detection. Here, we provide our protocol for whole-mount fluorescence immunohistochemistry in zebrafish embryos and larvae. This protocol additionally describes several different mounting strategies that can be employed and an overview of the advantages and disadvantages each strategy provides. We also describe modifications to this protocol to allow detection of chromogenic substrates in whole mount tissue and fluorescence detection in sectioned larval tissue. This protocol is broadly applicable to the study of many developmental stages and embryonic structures.

Introduction

The zebrafish (Danio rerio) has emerged as a powerful model for the study of biological processes for several reasons including short generation time, rapid development, and amenability to genetic techniques. As a result, zebrafish are commonly used in high throughput small molecule screens for toxicological research and drug discovery. Zebrafish are also an attractive model for the study of developmental processes given that a single female can routinely produce 50-300 eggs at a time and the optically clear embryos develop externally allowing for efficient visualization of developmental processes. However, early research relied mostly on forward genetic screens using N-ethyl-N-nitrosourea (ENU) or other mutagens due to challenges in establishing reverse genetic techniques. Roughly two decades ago, morpholinos were first used in zebrafish to knockdown targeted genes1. Morpholinos are small antisense oligonucleotides that inhibit translation of target mRNA following microinjection into an embryo at an early developmental stage. A major weakness of morpholinos is that they are diluted as the cells divide and generally lose effectiveness by 72 hours post-fertilization (hpf). While morpholinos remain a powerful tool for zebrafish gene disruption, transcription activator-like effector nucleases (TALENs), zinc-finger nucleases (ZFNs), and clustered regularly interspaced short palindromic repeats (CRISPRs) are more recently being used to directly target the zebrafish genome2,3. These reverse genetic strategies, in combination with forward genetics and high throughput screens, have established the zebrafish as a powerful model to study gene expression and function.

The ability to study gene function generally requires an evaluation of the spatio-temporal distribution of gene or gene product expression. The two most commonly used techniques to visualize such expression patterns during early development are in situ hybridization (ISH) and whole mount immunohistochemistry (IHC). In situ hybridization was first developed in 1969 and relies on the use of labeled antisense RNA probes to detect mRNA expression in an organism4. In contrast, labeled antibodies are used in immunohistochemistry to visualize protein expression. The idea of labeling proteins for detection dates back to the 1930's5 and the first IHC experiment was published in 1941 when FITC-labeled antibodies were used to detect pathogenic bacteria in infected tissues6. ISH and IHC have evolved and improved significantly over the subsequent decades and are now both routinely used in the molecular and diagnostic research laboratory7,8,9,10,11. While both techniques have advantages and disadvantages, IHC offers several benefits over ISH. Practically, IHC is much less time consuming than ISH and is generally less expensive depending on the cost of the primary antibody. In addition, mRNA expression is not always a reliable metric of protein expression as it has been demonstrated in mice and humans that only about a third of protein abundance variation can be explained by mRNA abundance12. For this reason, IHC is an important supplement to confirm ISH data, when possible. Finally, IHC can provide subcellular and co-localization data that cannot be determined by ISH13,14,15. Here, we describe a step-by-step method to reliably detect proteins by immunohistochemistry in whole mount zebrafish embryos and larvae. The goal of this technique is to determine the spatial and temporal expression of a protein of interest in the whole embryo. This technology utilizes antigen-specific primary antibodies and fluorescently tagged secondary antibodies. The protocol is readily adaptable to use on slide-mounted tissue sections and for use with chromogenic substrates in lieu of fluorescence. Using this protocol, we demonstrate that developing zebrafish skeletal muscle expresses ionotropic glutamate receptors, in addition to acetylcholine receptors. NMDA-type glutamate receptor subunits are detectable on the longitudinal muscle at 23 hpf.

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Protocol

The procedures for working with zebrafish breeding adults and embryos described in this protocol were approved by the Institutional Animal Care and Use Committee at Murray State University.

1. Embryo Collection and Fixation

  1. Prepare spawning tanks by placing adult zebrafish mixed sex pairs or groups in tanks with a mesh or slotted liner filled with system water overnight.
  2. At lights on, change the spawning tank water for fresh system water to remove feces. Use a 14 h/10 h light dark cycle with lights coming on at 9 am.
  3. Once eggs are laid, return the adults to home tanks.
  4. Collect eggs by drawing them up using a transfer pipet or pouring them into a mesh strainer.
  5. Transfer the eggs to Petri dishes filled halfway with embryo medium (such as 30% Danieau or E2 embryo medium with 0.5 mg/L methylene blue), limiting the number of embryos per dish to 50.
  6. Remove any eggs that are dead or fail to divide.
    NOTE: Dead embryos can be easily identified as they become opaque and often appear "cloudy". If methylene blue is added to embryo medium, the dead embryos take on a dark blue appearance.
  7. Incubate dishes of eggs at 28.5 °C until they reach the desired stage. For this experiment, raise the embryos until 23 hpf.
  8. Optional) Transfer the embryos at 24 hpf to 200 µM 1-phenyl 2-thiourea (PTU) in embryo medium to prevent melanogenesis16,17. Alternatively, bleach embryos post-fixation (see optional section 5).
  9. Change embryo medium or PTU medium daily.
  10. Dechorionate unhatched embryos using ultra-fine-tip forceps under a stereomicroscope. Alternatively, chemically dechorionate embryos by incubating in 1 mg/mL Pronase in embryo medium for several minutes at room temperature. Remove the embryos from Pronase and wash three times with embryo medium.
  11. Dechorionated embryos will stick to plastic. Keep them in glass or plastic Petri dishes coated with 1-2% agarose dissolved in embryo medium. Move dechorionated embryos using fire-polished Pasteur pipets to minimize damage.
  12. Transfer the embryos to 1.5 mL centrifuge tubes using a plastic or fire-polished pipet.
  13. Remove embryo medium with a micropipette. Leave only enough liquid to just cover the embryos after each fluid change.
  14. Prepare 4% paraformaldehyde (PFA) in 1x phosphate-buffered saline (PBS) in a chemical fume hood.
    CAUTION: PFA is a hazardous material. Wear gloves and dispose of contaminated liquids and solids in designated areas.
  15. Fix the embryos in 4% PFA for 1-2 h with gentle rocking at room temperature. Alternatively, fix the embryos 4 h to overnight at 4 °C.
  16. Wash three times in 1x PBS + 1% Triton-X (PBTriton) for 5 min.
  17. Use the embryos immediately or store at 4 °C for up to 1 week.
  18. For long term storage, dehydrate the embryos in 100% methanol (MeOH) 2 h or overnight at -20 °C. Store the embryos at -20 °C in MeOH for several months.
    CAUTION: MeOH is a hazardous material. Wear gloves and dispose of contaminated liquids and solids in designated areas.

2. Embryo Preparation

  1. Rehydrate the embryos through serial incubations at room temperature.
    1. Incubate in 75% MeOH/25% 1x PBS for 5 min, rocking.
    2. Incubate in 50% MeOH/50% 1x PBS for 5 min, rocking.
    3. Incubate in 25% MeOH/75% 1x PBS for 5 min, rocking.
    4. Incubate in 100% PBTriton for 5 min, rocking.
  2. (Optional) Prepare a fresh proteinase K working solution (10 µg/mL in PBTriton) on ice by adding 10 µL of freshly thawed proteinase K stock (10 mg/mL).
    1. Permeabilize the embryos by digesting up to 30 min in Proteinase K.
      NOTE: Suggested timing is <24 hpf: no digestion; 24 hpf: 15 min digestion; and 7 days old: 30 min digestion.
    2. Rinse permeabilized embryos in PBTriton and re-fix in 4% PFA for 20 min at room temperature.
    3. Wash the embryos three times in PBTriton for 5 min at room temperature with gentle rocking.

3. Primary Antibody Incubation

  1. Select a commercial blocking solution or serum matching the secondary antibody host species (ex. 10% goat serum in PBTriton) with or without 2 mg/mL Bovine Serum Albumin (BSA).
  2. Block the embryos in blocking solution for 1-3 h at room temperature or overnight at 4 °C while rocking.
  3. Incubate in primary antibody diluted in blocking solution or 1% serum in PBTriton overnight at 4 °C while rocking. In this experiment, the primary antibodies used were anti-NMDAR1, anti-pan-AMPA receptor, and anti-phospho-Histone H3, each diluted to a final concentration of 1:500 in 1% goat serum in PBTriton.
  4. Wash five times in PBTriton for 10 min at room temperature while rocking.

4. Secondary Antibody Incubation

  1. Select a secondary antibody based on the host species of the primary antibody and the desired wavelength.
  2. Incubate in secondary antibody diluted in blocking solution or 1% serum for 2 h at room temperature (or overnight at 4 °C) while rocking.
    NOTE: Fluorescent secondary antibodies are light sensitive. We used 1:500 goat-anti-mouse Alexa488 diluted in 1% goat serum in PBTriton.
  3. Cover the tubes with aluminum foil or cover with a light-blocking box for this and all subsequent steps.
  4. Wash three times in PBTriton for 10 min at room temperature while rocking.
  5. Transfer the embryos to a 50% glycerol solution in PBS over a bed of 2% agarose in embryo medium and proceed to documentation or proceed to further processing steps below.

Optional Steps

5. Bleaching

  1. Prepare bleach solution in a 1.5 mL tube by adding 810.7 µL of ddH2O, 89.3 µL of 2 M KOH, and 100 µL of 30% H2O2.
  2. Invert the tube three times to mix.
  3. Pipette 1 mL of bleach solution direction to the embryos.
  4. Open the embryo tube cap to allow gas to escape. Gently tap the tube on the bench to dislodge bubbles.
  5. Monitor the bleaching process (use a microscope if necessary) and stop the reaction when pigment is sufficiently removed (approximately 5 min for 24 hpf or 10 min for 72 hpf).
  6. Carefully remove the bleaching solution with a micropipette and rinse embryos three times in 1 mL of PBTriton.
    NOTE: Embryos are sticky in this step.
  7. Proceed to documentation or further processing steps below.

6. Embryo Dissection and Deyolking

  1. To remove the yolk, transfer a small amount (~200 µL or enough to completely cover the embryo but restrictive enough to limit where it can float) of 1x PBS to a depression slide or a plain glass slide.
  2. Use a plastic transfer pipet to move 1 or more embryos to the PBS droplet.
  3. Use ultra-fine forceps and 00 insect pins to break apart the yolk and very gently scrape yolk granules from the ventral surface of the embryo (see also Cheng et al., 2014)18.
  4. Remove yolk granules and replenish PBS as needed.
  5. Repeat until embryo is sufficiently free of yolk.

7. Flat Mounting on Slides

  1. Transfer deyolked embryos to a charged glass slide with a plastic pipette or a 1 mL micropipette with a trimmed tip (to reduce shear stress). Orient as desired with a 200 µL micropipette tip or insect pin.
  2. Wick away excess PBS with a Kim wipe or paper towel.
  3. Add 2-3 drops of mounting media to the slide and coverslip.
  4. Air dry for approximately 5 to 10 min.
  5. Seal the cover glass onto the slide with clear nail polish.
    NOTE: The edges of the cover glass must be completely covered with a thin, continuous layer of nail polish.
  6. Allow to dry approximately 10 min before imaging.

8. Mounting in Agarose

  1. Prepare 1% agarose in embryo medium by adding 0.5 g of agarose to 50 mL of embryo medium in a microwave-safe flask or beaker of at least 3x greater volume than desired.
  2. Heat in a microwave, swirling every 30 s, until agarose is completely dissolved.
  3. Make 1 mL aliquots in 1.5 mL centrifuge tubes. Store the aliquots at room temperature.
  4. Cover tube caps with cap locks before heating.
  5. Place agarose tubes in a floating tube holder in a beaker half-filled with water.
  6. Microwave the beaker with floating tubes for 2-3 min, or until agarose is completely melted.
  7. Transfer an embryo to the bridged slide with a plastic pipette or a 200 µL micropipette with a trimmed tip (to reduce shear stress).
  8. Position the embryo on a rectangular coverslip using insect pins and add approximately 20 µL melted agarose directly to the embryo.
  9. Quickly orient the region of interest closest to the coverslip using 00 insect pins.
    NOTE: This is an upside-down mount.
  10. Return the agarose tube to the hot water tube float between each use and microwave as needed.
  11. Image using a microscope when the agarose hardens. Keep the mounted embryo upside-down for use on an inverted microscope. Flip the coverslip over (so the agarose is under the coverslip) for use on upright microscopes.

9. Mounting on Bridged Slides

  1. To make bridged slides, glue square coverslips to the glass slide using a small dot of superglue.
    NOTE: There should be a trough at least 5 mm wide between the coverslips. Two #1 coverslips high is typically appropriate for 24-48 hpf embryos while three coverslips high may be necessary for 72 hpf.
  2. Transfer 1-2 deyolked embryos to the bridged slide with a plastic pipette or a 200 µL micropipette with a trimmed tip (to reduce shear stress).
  3. Wick away excess fluid with a Kim wipe or paper towel.
  4. Add a drop of ≥80% glycerol directly to the embryo.
  5. Cover with a rectangular cover glass. The droplet of glycerol should touch the cover glass.
  6. Add more glycerol to the space between the cover glass and slide as needed to completely cover the embryo with a margin of glycerol on the sides of the embryo.
  7. Slide the rectangular cover glass gently to roll the embryo into position for imaging.

10. DAB Staining

NOTE: This section begins after step 4.2 above and replaces the rest of step 4.

  1. Incubate the embryos in a blocking solution with a peroxidase-conjugated secondary antibody for 2 h at room temperature or overnight at 4 °C while rocking.
  2. Wash three times in PBTriton for 10 min at room temperature.
  3. Transfer the embryos to a culture plate or depression slide with a transfer pipette.
  4. Mix 50 µL of 1% DAB (3,3'-diaminobenzidine) dissolved in ddH2O and 50 µL of 0.3% hydrogen peroxide and bring to 1 mL with PBS.
    CAUTION: DAB is a hazardous material. Wear gloves and dispose of DAB contaminated liquids and solids in designated areas.
  5. Cover HRP-stained embryos with the DAB solution prepared above and monitor for color development (typically 1-5 min) under a microscope.
  6. After reaching the desired level of color development, rinse the embryos briefly in PBS.
  7. Transfer the embryos back to a 1.5 mL tube before fixation.
  8. Re-fix the embryos for 15-20 min in 4% PFA at room temperature.
  9. Wash the embryos three times in PBTriton for 5 min.
  10. Proceed to documentation.

11. Modified Protocol for Staining Sectioned Tissue That is Mounted on Slides

  1. Encircle tissue to be stained with a pap pen.
  2. Transfer the slides to a humid chamber.
  3. Add 1 mL ofPBS directly to the slide.
  4. Incubate 7 min at room temperature to remove embedding medium.
  5. Pour off PBS by inverting slide.
  6. Rehydrate 1 min in up to 1 mL of TNT buffer (100 mM Tris pH 8.0, 150 mM NaCl, 0.1% Tween20).
  7. Block in up to 1 mL of blocking solution (commercial or 10% serum + 2% BSA) for 1 h at room temperature.
  8. Incubate overnight in primary antibody diluted in 1% serum or blocking solution at 4 °C.
  9. Wash five times in up to 1 mL of TNT buffer at room temperature.
  10. Incubate in secondary antibody for 2 h at room temperature or overnight at 4 °C. Cover the chamber with foil or use a dark lid.
  11. Wash five times in TNT at room temperature. Pour off last wash.
  12. Mount with 2-3 drops of mounting medium and coverslip. Let sit 5-10 min.
  13. Seal the cover glass onto the slide using clear nail polish. Allow to dry completely before imaging.

12. Documentation

  1. Record the full procedure and any deviations in a lab notebook.
  2. Record the concentration, name, catalog number, manufacturer, and lot number of the primary antibody.
  3. Place appropriately mounted sample on the microscope stage. Locate the region of interest.
  4. Select a relatively bright example. Set camera exposure and gain so that signal is sufficiently bright without saturating.
  5. Compare staining intensity of the same region of interest using the same exposure settings when comparing between experimental antibody-labeled embryos and control antibody (ex. IgG) embryos.

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Results

Whole mount immunohistochemistry uses antibodies to detect the spatial pattern of protein expression in the intact animal. The basic workflow of immunohistochemistry (depicted in Figure 1) involves breeding zebrafish, raising and preparing embryos, blocking non-specific antigens, using an antigen-specific primary antibody to target the protein of interest, detecting that primary antibody with a labeled secondary antibody, mounting the specimen, and documentin...

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Discussion

Immunohistochemistry is a versatile tool that can be used to characterize the spatio-temporal expression of virtually any protein of interest in an organism. Immunohistochemistry is used on a wide variety of tissues and model organisms. This protocol has been optimized for use in zebrafish. Immunohistochemistry in different species may require different fixation and handling techniques, blocking solutions depending on species and the presence of endogenous peroxidases, and incubation times due to the thickness and compos...

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Disclosures

The authors have no information to disclose.

Acknowledgements

Funding from NIH grant 8P20GM103436 14.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
AgaroseFisher ScientificBP160-100
Aluminum foil, heavy dutyKirklandAny brand may be substituted
Anti-NMDA antibodyMillipore SigmaMAB363
Anti-phospho-Histone H3 (Ser10), clone RR002Millipore Sigma05-598
Anti-pan-AMPA receptor (GluR1-4)Millipore SigmaMABN832
Bovine serum albumin (BSA)Fisher ScientificBP1600-100
Calcium Nitrate [Ca(NO3)2]Sigma AldrichC4955
Centrifuge tubes, 1.5 mLAxygenMCT150C
Clear nail polishSally HansonAny nail polish or hardener may be subsituted
Depression (concavity) slideElectron Miscroscopy Sciences71878-01
DiaminobenzidineThermo Scientific1855920
Embryo medium, Danieau, 30%17.4 mM NaCl, 0.21 mM KCl, 0.12 mM MgS04, 0.18 mM Ca(NO3)2, 1.5 mM HEPES in ultrapure water.
Embryo medium, E27.5 mM NaCl, 0.25 mM KCl, 0.5 mM MgSO4, 75 μM KH2PO4, 25 uM Na2HPO4, 0.5 mM CaCl2, 0.35 mM NaHCO3, 0.5 mg/L methylene blue
Floating tube holderThermo Scientific59744015
Fluorescence compound microscopeLeica BiosystemsDMi8
Fluorescence stereomicroscopeLeica BiosystemsM165-FC
Glass coverslips 18 mm x 18 mmCorning284518
Glass coverslips 22 mm x 60 mmThermo Scientific22-050-222
Glass slidesFisher Scientific12-544-4
GlycerolFisher ScientificBP229-1
Goat anti-mouse IgG Alexa 488InvitrogenA11001
HEPES solutionSigma AldrichH0887
Humid chamber with lidSimportM920-2
Hydrogen peroxide, 30%Fisher ScientificH325-500
Immunedge pap penVector labsH-4000
Insect pins, size 00Stoelting5213323
Magnesium Sulfate (MgSO4 · 7H2O)Sigma Aldrich63138
Mesh strainerOneidaAny brand may be substituted
MethanolSigma Aldrich34860
Methylene blueSigma AldrichM9140
Micro-tube cap lockResearch Products International145062
Microwave ovenToastmaster
Mouse IgGSigma AldrichI8765
Normal goat serumMillipore SigmaS02L1ML
Nutating mixerFisher Scientific88-861-044
ParaformaldehydeFisher Scientific04042-500
Pasteur pipettesFisher Scientific13-678-20C
PBTriton1% TritonX-100 in 1x PBS
Permount mounting mediumFisher ChemicalSP15-500
Petri dish (glass)Pyrex3160100
Petri dish (plastic)Fisher ScientificFB0875713
1-phenyl 2-thioureaAcros Organics207250250
Phosphate buffered saline (PBS), 10x, pH 7.4Gibco70011-044
Phosphate buffered saline (PBS), 1x1x made from 10x stock diluted in dH2O
Potassium Chloride (KCl)Sigma AldrichP9333
Potassium Hydroxide (KOH)FisherP250-500
Potassium Phosphate Monobasic (KH2PO4)Sigma AldrichP5655
PronaseSigma Aldrich10165921001
Proteinase KInvitrogenAM2544
Sodium Chloride (NaCl)Sigma AldrichS7653
Sodium Phosphate Dibasic (Na2HPO4)Sigma AldrichS7907
Spawning tank with lid and insertAquaneeringZHCT100
SuperBlock PBSThermo Scientific37515
Superfrost + slidesFisher Scientific12-550-15
Superglue gel3M Scotch
TNT100 mM Tris, pH 8.0; 150 mM NaCl; 0.1% Tween20; made in dH2O
Transfer pipetteFisher13-711-7M
Trichloracetic Acid (Cl3CCOOH)Sigma AldrichT6399
Tris BaseFisher ScientificS374-500
TritonX-100Sigma AldrichT9284
Tween20Fisher ScientificBP337-500
Ultrafine forcepsFisher Scientific16-100-121
Water, ultrapure/double distilledFisher ScientificW2-20

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