Funding from NIH grant 8P20GM103436 14.

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
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Once eggs are laid, return the adults to home tanks.</li>
	<li>Collect eggs by drawing them up using a transfer pipet or pouring them into a mesh strainer.</li>
	<li>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.</li>
	<li>Remove any eggs that are dead or fail to divide. 
	NOTE: Dead embryos can be easily identified as they become opaque and often appear &#34;cloudy&#34;. If methylene blue is added to embryo medium, the dead embryos take on a dark blue appearance.</li>
	<li>Incubate dishes of eggs at 28.5 &#176;C until they reach the desired stage. For this experiment, raise the embryos until 23 hpf.</li>
	<li>Optional) Transfer the embryos at 24 hpf to 200 &#181;M 1-phenyl 2-thiourea (PTU) in embryo medium to prevent melanogenesis16,17. Alternatively, bleach embryos post-fixation (see optional section 5).</li>
	<li>Change embryo medium or PTU medium daily.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Transfer the embryos to 1.5 mL centrifuge tubes using a plastic or fire-polished pipet.</li>
	<li>Remove embryo medium with a micropipette. Leave only enough liquid to just cover the embryos after each fluid change.</li>
	<li>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.</li>
	<li>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 &#176;C.</li>
	<li>Wash three times in 1x PBS + 1% Triton-X (PBTriton) for 5 min.</li>
	<li>Use the embryos immediately or store at 4 &#176;C for up to 1 week.</li>
	<li>For long term storage, dehydrate the embryos in 100% methanol (MeOH) 2 h or overnight at -20 &#176;C. Store the embryos at -20 &#176;C in MeOH for several months. 
	CAUTION: MeOH is a hazardous material. Wear gloves and dispose of contaminated liquids and solids in designated areas.</li>
</ol>
2. Embryo Preparation
<ol>
	<li>Rehydrate the embryos through serial incubations at room temperature.
	<ol>
		<li>Incubate in 75% MeOH/25% 1x PBS for 5 min, rocking.</li>
		<li>Incubate in 50% MeOH/50% 1x PBS for 5 min, rocking.</li>
		<li>Incubate in 25% MeOH/75% 1x PBS for 5 min, rocking.</li>
		<li>Incubate in 100% PBTriton for 5 min, rocking.</li>
	</ol>
	</li>
	<li>(Optional) Prepare a fresh proteinase K working solution (10 &#181;g/mL in PBTriton) on ice by adding 10 &#181;L of freshly thawed proteinase K stock (10 mg/mL).
	<ol>
		<li>Permeabilize the embryos by digesting up to 30 min in Proteinase K. 
		NOTE: Suggested timing is &#60;24 hpf: no digestion; 24 hpf: 15 min digestion; and 7 days old: 30 min digestion.</li>
		<li>Rinse permeabilized embryos in PBTriton and re-fix in 4% PFA for 20 min at room temperature.</li>
		<li>Wash the embryos three times in PBTriton for 5 min at room temperature with gentle rocking.</li>
	</ol>
	</li>
</ol>
3. Primary Antibody Incubation
<ol>
	<li>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).</li>
	<li>Block the embryos in blocking solution for 1-3 h at room temperature or overnight at 4 &#176;C while rocking.</li>
	<li>Incubate in primary antibody diluted in blocking solution or 1% serum in PBTriton overnight at 4 &#176;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.</li>
	<li>Wash five times in PBTriton for 10 min at room temperature while rocking.</li>
</ol>
4. Secondary Antibody Incubation
<ol>
	<li>Select a secondary antibody based on the host species of the primary antibody and the desired wavelength.</li>
	<li>Incubate in secondary antibody diluted in blocking solution or 1% serum for 2 h at room temperature (or overnight at 4 &#176;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.</li>
	<li>Cover the tubes with aluminum foil or cover with a light-blocking box for this and all subsequent steps.</li>
	<li>Wash three times in PBTriton for 10 min at room temperature while rocking.</li>
	<li>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.</li>
</ol>
Optional Steps
5. Bleaching
<ol>
	<li>Prepare bleach solution in a 1.5 mL tube by adding 810.7 &#181;L of ddH2O, 89.3 &#181;L of 2 M KOH, and 100 &#181;L of 30% H2O2.</li>
	<li>Invert the tube three times to mix.</li>
	<li>Pipette 1 mL of bleach solution direction to the embryos.</li>
	<li>Open the embryo tube cap to allow gas to escape. Gently tap the tube on the bench to dislodge bubbles.</li>
	<li>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).</li>
	<li>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.</li>
	<li>Proceed to documentation or further processing steps below.</li>
</ol>
6. Embryo Dissection and Deyolking
<ol>
	<li>To remove the yolk, transfer a small amount (~200 &#181;L 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.</li>
	<li>Use a plastic transfer pipet to move 1 or more embryos to the PBS droplet.</li>
	<li>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.</li>
	<li>Remove yolk granules and replenish PBS as needed.</li>
	<li>Repeat until embryo is sufficiently free of yolk.</li>
</ol>
7. Flat Mounting on Slides
<ol>
	<li>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 &#181;L micropipette tip or insect pin.</li>
	<li>Wick away excess PBS with a Kim wipe or paper towel.</li>
	<li>Add 2-3 drops of mounting media to the slide and coverslip.</li>
	<li>Air dry for approximately 5 to 10 min.</li>
	<li>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.</li>
	<li>Allow to dry approximately 10 min before imaging.</li>
</ol>
8. Mounting in Agarose
<ol>
	<li>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.</li>
	<li>Heat in a microwave, swirling every 30 s, until agarose is completely dissolved.</li>
	<li>Make 1 mL aliquots in 1.5 mL centrifuge tubes. Store the aliquots at room temperature.</li>
	<li>Cover tube caps with cap locks before heating.</li>
	<li>Place agarose tubes in a floating tube holder in a beaker half-filled with water.</li>
	<li>Microwave the beaker with floating tubes for 2-3 min, or until agarose is completely melted.</li>
	<li>Transfer an embryo to the bridged slide with a plastic pipette or a 200 &#181;L micropipette with a trimmed tip (to reduce shear stress).</li>
	<li>Position the embryo on a rectangular coverslip using insect pins and add approximately 20 &#181;L melted agarose directly to the embryo.</li>
	<li>Quickly orient the region of interest closest to the coverslip using 00 insect pins. 
	NOTE: This is an upside-down mount.</li>
	<li>Return the agarose tube to the hot water tube float between each use and microwave as needed.</li>
	<li>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.</li>
</ol>
9. Mounting on Bridged Slides
<ol>
	<li>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.</li>
	<li>Transfer 1-2 deyolked embryos to the bridged slide with a plastic pipette or a 200 &#181;L micropipette with a trimmed tip (to reduce shear stress).</li>
	<li>Wick away excess fluid with a Kim wipe or paper towel.</li>
	<li>Add a drop of &#8805;80% glycerol directly to the embryo.</li>
	<li>Cover with a rectangular cover glass. The droplet of glycerol should touch the cover glass.</li>
	<li>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.</li>
	<li>Slide the rectangular cover glass gently to roll the embryo into position for imaging.</li>
</ol>
10. DAB Staining
NOTE: This section begins after step 4.2 above and replaces the rest of step 4.
<ol>
	<li>Incubate the embryos in a blocking solution with a peroxidase-conjugated secondary antibody for 2 h at room temperature or overnight at 4 &#176;C while rocking.</li>
	<li>Wash three times in PBTriton for 10 min at room temperature.</li>
	<li>Transfer the embryos to a culture plate or depression slide with a transfer pipette.</li>
	<li>Mix 50 &#181;L of 1% DAB (3,3&#39;-diaminobenzidine) dissolved in ddH2O and 50 &#181;L 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.</li>
	<li>Cover HRP-stained embryos with the DAB solution prepared above and monitor for color development (typically 1-5 min) under a microscope.</li>
	<li>After reaching the desired level of color development, rinse the embryos briefly in PBS.</li>
	<li>Transfer the embryos back to a 1.5 mL tube before fixation.</li>
	<li>Re-fix the embryos for 15-20 min in 4% PFA at room temperature.</li>
	<li>Wash the embryos three times in PBTriton for 5 min.</li>
	<li>Proceed to documentation.</li>
</ol>
11. Modified Protocol for Staining Sectioned Tissue That is Mounted on Slides
<ol>
	<li>Encircle tissue to be stained with a pap pen.</li>
	<li>Transfer the slides to a humid chamber.</li>
	<li>Add 1 mL ofPBS directly to the slide.</li>
	<li>Incubate 7 min at room temperature to remove embedding medium.</li>
	<li>Pour off PBS by inverting slide.</li>
	<li>Rehydrate 1 min in up to 1 mL of TNT buffer (100 mM Tris pH 8.0, 150 mM NaCl, 0.1% Tween20).</li>
	<li>Block in up to 1 mL of blocking solution (commercial or 10% serum + 2% BSA) for 1 h at room temperature.</li>
	<li>Incubate overnight in primary antibody diluted in 1% serum or blocking solution at 4 &#176;C.</li>
	<li>Wash five times in up to 1 mL of TNT buffer at room temperature.</li>
	<li>Incubate in secondary antibody for 2 h at room temperature or overnight at 4 &#176;C. Cover the chamber with foil or use a dark lid.</li>
	<li>Wash five times in TNT at room temperature. Pour off last wash.</li>
	<li>Mount with 2-3 drops of mounting medium and coverslip. Let sit 5-10 min.</li>
	<li>Seal the cover glass onto the slide using clear nail polish. Allow to dry completely before imaging.</li>
</ol>
12. Documentation
<ol>
	<li>Record the full procedure and any deviations in a lab notebook.</li>
	<li>Record the concentration, name, catalog number, manufacturer, and lot number of the primary antibody.</li>
	<li>Place appropriately mounted sample on the microscope stage. Locate the region of interest.</li>
	<li>Select a relatively bright example. Set camera exposure and gain so that signal is sufficiently bright without saturating.</li>
	<li>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.</li>
</ol>

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L., Gallina, D., Kahana, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phenothiourea+sensitizes+zebrafish+cranial+neural+crest+and+extraocular+muscle+development+to+changes+in+retinoic+acid+and+IGF+signaling.">Phenothiourea sensitizes zebrafish cranial neural crest and extraocular muscle development to changes in retinoic acid and IGF signaling.</a> PloS one. 6 (8), e22991 (2011).</li><li>Inoue, D., Wittbrodt, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=One+for+all--a+highly+efficient+and+versatile+method+for+fluorescent+immunostaining+in+fish+embryos.">One for all--a highly efficient and versatile method for fluorescent immunostaining in fish embryos.</a> PLoS One. 6 (5), e19713 (2011).</li></ol>

The authors have no information to disclose.

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...

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&#39;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.

<table><tbody><tr><td>Agarose</td><td>Fisher Scientific</td><td>BP160-100</td><td></td></tr><tr><td>Aluminum foil, heavy duty</td><td>Kirkland</td><td></td><td>Any brand may be substituted</td></tr><tr><td>Anti-NMDA antibody</td><td>Millipore Sigma</td><td>MAB363</td><td></td></tr><tr><td>Anti-phospho-Histone H3 (Ser10), clone RR002</td><td>Millipore Sigma</td><td>05-598</td><td></td></tr><tr><td>Anti-pan-AMPA receptor (GluR1-4)</td><td>Millipore Sigma</td><td>MABN832</td><td></td></tr><tr><td>Bovine serum albumin (BSA)</td><td>Fisher Scientific</td><td>BP1600-100</td><td></td></tr><tr><td>Calcium Nitrate [Ca(NO&lt;sub&gt;3&lt;/sub&gt;)&lt;sub&gt;2&lt;/sub&gt;]</td><td>Sigma Aldrich</td><td>C4955</td><td></td></tr><tr><td>Centrifuge tubes, 1.5 mL</td><td>Axygen</td><td>MCT150C</td><td></td></tr><tr><td>Clear nail polish</td><td>Sally Hanson</td><td></td><td>Any nail polish or hardener may be subsituted</td></tr><tr><td>Depression (concavity) slide</td><td>Electron Miscroscopy Sciences</td><td>71878-01</td><td></td></tr><tr><td>Diaminobenzidine</td><td>Thermo Scientific</td><td>1855920</td><td></td></tr><tr><td>Embryo medium, Danieau, 30%</td><td></td><td></td><td>17.4 mM NaCl, 0.21 mM KCl, 0.12 mM MgS04, 0.18 mM Ca(NO&lt;sub&gt;3&lt;/sub&gt;)&lt;sub&gt;2&lt;/sub&gt;, 1.5 mM HEPES in ultrapure water.</td></tr><tr><td>Embryo medium, E2</td><td></td><td></td><td>7.5 mM NaCl, 0.25 mM KCl, 0.5 mM MgSO&lt;sub&gt;4&lt;/sub&gt;, 75 &amp;mu;M KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, 25 uM Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;, 0.5 mM CaCl&lt;sub&gt;2&lt;/sub&gt;, 0.35 mM NaHCO&lt;sub&gt;3&lt;/sub&gt;, 0.5 mg/L methylene blue</td></tr><tr><td>Floating tube holder</td><td>Thermo Scientific</td><td>59744015</td><td></td></tr><tr><td>Fluorescence compound microscope</td><td>Leica Biosystems</td><td>DMi8</td><td></td></tr><tr><td>Fluorescence stereomicroscope</td><td>Leica Biosystems</td><td>M165-FC</td><td></td></tr><tr><td>Glass coverslips 18 mm x 18 mm</td><td>Corning</td><td>284518</td><td></td></tr><tr><td>Glass coverslips 22 mm x 60 mm</td><td>Thermo Scientific</td><td>22-050-222</td><td></td></tr><tr><td>Glass slides</td><td>Fisher Scientific</td><td>12-544-4</td><td></td></tr><tr><td>Glycerol</td><td>Fisher Scientific</td><td>BP229-1</td><td></td></tr><tr><td>Goat anti-mouse IgG Alexa 488</td><td>Invitrogen</td><td>A11001</td><td></td></tr><tr><td>HEPES solution</td><td>Sigma Aldrich</td><td>H0887</td><td></td></tr><tr><td>Humid chamber with lid</td><td>Simport</td><td>M920-2</td><td></td></tr><tr><td>Hydrogen peroxide, 30%</td><td>Fisher Scientific</td><td>H325-500</td><td></td></tr><tr><td>Immunedge pap pen</td><td>Vector labs</td><td>H-4000</td><td></td></tr><tr><td>Insect pins, size 00</td><td>Stoelting</td><td>5213323</td><td></td></tr><tr><td>Magnesium Sulfate (MgSO&lt;sub&gt;4&lt;/sub&gt; &amp;middot; 7H&lt;sub&gt;2&lt;/sub&gt;O)</td><td>Sigma Aldrich</td><td>63138</td><td></td></tr><tr><td>Mesh strainer</td><td>Oneida</td><td></td><td>Any brand may be substituted</td></tr><tr><td>Methanol</td><td>Sigma Aldrich</td><td>34860</td><td></td></tr><tr><td>Methylene blue</td><td>Sigma Aldrich</td><td>M9140</td><td></td></tr><tr><td>Micro-tube cap lock</td><td>Research Products International</td><td>145062</td><td></td></tr><tr><td>Microwave oven</td><td>Toastmaster</td><td></td><td></td></tr><tr><td>Mouse IgG</td><td>Sigma Aldrich</td><td>I8765</td><td></td></tr><tr><td>Normal goat serum</td><td>Millipore Sigma</td><td>S02L1ML</td><td></td></tr><tr><td>Nutating mixer</td><td>Fisher Scientific</td><td>88-861-044</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Fisher Scientific</td><td>04042-500</td><td></td></tr><tr><td>Pasteur pipettes</td><td>Fisher Scientific</td><td>13-678-20C</td><td></td></tr><tr><td>PBTriton</td><td></td><td></td><td>1% TritonX-100 in 1x PBS</td></tr><tr><td>Permount mounting medium</td><td>Fisher Chemical</td><td>SP15-500</td><td></td></tr><tr><td>Petri dish (glass)</td><td>Pyrex</td><td>3160100</td><td></td></tr><tr><td>Petri dish (plastic)</td><td>Fisher Scientific</td><td>FB0875713</td><td></td></tr><tr><td>1-phenyl 2-thiourea</td><td>Acros Organics</td><td>207250250</td><td></td></tr><tr><td>Phosphate buffered saline (PBS), 10x, pH 7.4</td><td>Gibco</td><td>70011-044</td><td></td></tr><tr><td>Phosphate buffered saline (PBS), 1x</td><td></td><td></td><td>1x made from 10x stock diluted in dH&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>Potassium Chloride (KCl)</td><td>Sigma Aldrich</td><td>P9333</td><td></td></tr><tr><td>Potassium Hydroxide (KOH)</td><td>Fisher</td><td>P250-500</td><td></td></tr><tr><td>Potassium Phosphate Monobasic (KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma Aldrich</td><td>P5655</td><td></td></tr><tr><td>Pronase</td><td>Sigma Aldrich</td><td>10165921001</td><td></td></tr><tr><td>Proteinase K</td><td>Invitrogen</td><td>AM2544</td><td></td></tr><tr><td>Sodium Chloride (NaCl)</td><td>Sigma Aldrich</td><td>S7653</td><td></td></tr><tr><td>Sodium Phosphate Dibasic (Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma Aldrich</td><td>S7907</td><td></td></tr><tr><td>Spawning tank with lid and insert</td><td>Aquaneering</td><td>ZHCT100</td><td></td></tr><tr><td>SuperBlock PBS</td><td>Thermo Scientific</td><td>37515</td><td></td></tr><tr><td>Superfrost + slides</td><td>Fisher Scientific</td><td>12-550-15</td><td></td></tr><tr><td>Superglue gel</td><td>3M Scotch</td><td></td><td></td></tr><tr><td>TNT</td><td></td><td></td><td>100 mM Tris, pH 8.0; 150 mM NaCl; 0.1% Tween20; made in dH&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>Transfer pipette</td><td>Fisher</td><td>13-711-7M</td><td></td></tr><tr><td>Trichloracetic Acid (Cl&lt;sub&gt;3&lt;/sub&gt;CCOOH)</td><td>Sigma Aldrich</td><td>T6399</td><td></td></tr><tr><td>Tris Base</td><td>Fisher Scientific</td><td>S374-500</td><td></td></tr><tr><td>TritonX-100</td><td>Sigma Aldrich</td><td>T9284</td><td></td></tr><tr><td>Tween20</td><td>Fisher Scientific</td><td>BP337-500</td><td></td></tr><tr><td>Ultrafine forceps</td><td>Fisher Scientific</td><td>16-100-121</td><td></td></tr><tr><td>Water, ultrapure/double distilled</td><td>Fisher Scientific</td><td>W2-20</td><td></td></tr></tbody></table>

whole mount immunohistochemistry in zebrafish embryos and larvae

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.

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...

Watch this Scientific Journal Video about Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae at JoVE.com

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

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

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. ...

whole-mount-immunohistochemistry-in-zebrafish-embryos-and-larvae

Universidad San Sebastian

Research

JoVE Journal

Developmental Biology

19.6K Views.  Murray State University. Here, we present a protocol for fluorescent antibody-mediated detection of proteins in whole preparations of zebrafish embryos and larvae.

Video: Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

Immunostaining Phospho-epitopes in Ciliated Organs of Whole Mount Zebrafish Embryos

The rapid proliferation of cells, the tissue-specific expression of genes and the emergence of signaling networks characterize early embryonic development of all vertebrates. The kinetics and location of signals - even within single cells - in the developing embryo complements the identification of important developmental genes. Immunostaining techniques are described that have been shown to define the kinetics of intracellular and whole animal signals in structures as small as primary cilia. The techniques for fixing, imaging and processing images using a laser-scanning confocal compound microscope can be completed in as few as 36 hr.
Zebrafish (Danio rerio) is a desirable organism for investigators who seek to conduct studies in a vertebrate species that is affordable and relevant to human disease. Genetic knockouts or knockdowns must be confirmed by the loss of the actual protein product. Such confirmation of protein loss can be achieved using the techniques described here. Clues into signaling pathways can also be deciphered by using antibodies that are reactive with proteins that have been post-translationally modified by phosphorylation. Preserving and optimizing the phosphorylated state of an epitope is therefore critical to this determination and is accomplished by this protocol.
This study describes techniques to fix embryos during the first 72 hr of development and co-localize a variety of relevant epitopes with cilia in the Kupffer&#39;s Vesicle (KV), the kidney and the inner ear. These techniques are straightforward, do not require dissection and can be completed in a relatively short period of time. Projecting confocal image stacks into a single image is a useful means of presenting these data.

Techniques are described to immunostain phospho-epitopes in whole zebrafish embryos and then conduct two-color fluorescent confocal localization in cellular structures as small as primary cilia. The techniques for fixing and imaging can define the location and kinetics of the appearance or activation of specific proteins.

The rapid proliferation of cells, the tissue-specific expression of genes and the emergence of signaling networks characterize early embryonic development ...

Chemistry

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

In recent years, the zebrafish embryo has emerged as a popular model to study developmental biology due to traits such as ex utero embryo development and optical transparency. In particular, the zebrafish embryo has become an important organism to study vertebrate kidney organogenesis as well as multiciliated cell (MCC) development. To visualize MCCs in the embryonic zebrafish kidney, we have developed a combined protocol of whole-mount fluorescent in situ hybridization (FISH) and whole mount immunofluorescence (IF) that enables high resolution imaging. This manuscript describes our technique for co-localizing RNA transcripts and protein as a tool to better understand the regulation of developmental programs through the expression of various lineage factors.

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

Cilia development is vital to proper organogenesis. This protocol describes an optimized method to label and visualize ciliated cells of the zebrafish.

In recent years, the zebrafish embryo has emerged as a popular model to study developmental biology due to traits such as ex utero embryo development and ...

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology. Here, we present a high-resolution double fluorescent in situ hybridization protocol for analyzing the precise expression pattern of a single gene and for determining the overlap of the expression domains of two genes. The protocol is a modified version of the standard in situ hybridization using alkaline phosphatase and substrates such as NBT/BCIP and Fast Red 1,2. This protocol utilizes standard digoxygenin and fluorescein labeled probes along with tyramide signal amplification (TSA) 3. The commercially available TSA kits allow flexible experimental design as fluorescence emission from green to far-red can be used in combination with various nuclear stains, such as propidium iodide, or fluorescence immunohistochemistry for proteins. TSA produces a reactive fluorescent substrate that quickly covalently binds to moieties, typically tyrosine residues, in the immediate vicinity of the labeled antisense riboprobe. The resulting staining patterns are high resolution in that subcellular localization of the mRNA can be observed using laser scanning confocal microscopy 3,4. One can observe nascent transcripts at the chromosomal loci, distinguish nuclear and cytoplasmic staining and visualize other patterns such as cortical localization of mRNA. Studies in Drosophila indicate that roughly 70% of mRNAs exhibit specific patterns of subcellular localization that frequently correlate with the function of the encoded protein 5. When combined with computer-aided reconstruction of 3D confocal datasets, our protocol allows the detailed analysis of mRNA distribution with sub-cellular resolution in whole vertebrate embryos.

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology. Here, we present a high-resolution double fluorescent in situ hybridization protocol for analyzing the precise expression pattern of a single gene and for determining the overlap of the expression domains of two genes. We include a propidium iodide nuclear counter-stain to highlight tissue organization.

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology.  Here, we present a high-resolution double ...

Biology

Dissection and Lateral Mounting of Zebrafish Embryos: Analysis of Spinal Cord Development

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene knockdown approaches can be utilized to examine the molecular mechanisms underlying nervous system development. Second, large clutches of developmentally synchronized embryos provide large experimental sample sizes. Third, the optical clarity of the zebrafish embryo permits researchers to visualize progenitor, glial, and neuronal populations. Although zebrafish embryos are transparent, specimen thickness can impede effective microscopic visualization. One reason for this is the tandem development of the spinal cord and overlying somite tissue. Another reason is the large yolk ball, which is still present during periods of early neurogenesis. In this article, we demonstrate microdissection and removal of the yolk in fixed embryos, which allows microscopic visualization while preserving surrounding somite tissue. We also demonstrate semipermanent mounting of zebrafish embryos. This permits observation of neurodevelopment in the dorso-ventral and anterior-posterior axes, as it preserves the three-dimensionality of the tissue.

Developmental processes such as proliferation, patterning, differentiation, and axon guidance can be readily modeled in the zebrafish spinal cord. In this article, we describe a mounting procedure for zebrafish embryos, which optimizes visualization of these events.

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene ...

Neuroscience

A Layered Mounting Method for Extended Time-Lapse Confocal Microscopy of Whole Zebrafish Embryos

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues or cells. A difficulty with imaging whole embryo development is that zebrafish embryos grow substantially in length. When mounted as regularly done in 0.3-1% low melt agarose, the agarose imposes growth restriction, leading to distortions in the soft embryo body. Yet, to perform confocal time-lapse microscopy, the embryo must be immobilized. This article describes a layered mounting method for zebrafish embryos that restrict the motility of the embryos while allowing for the unrestricted growth. The mounting is performed in layers of agarose at different concentrations. To demonstrate the usability of this method, whole embryo vascular, neuronal and muscle development was imaged in transgenic fish for 55 consecutive hours. This mounting method can be used for easy, low-cost imaging of whole zebrafish embryos using inverted microscopes without requirements of molds or special equipment.

This article describes a method to mount fragile zebrafish embryos for extended time-lapse confocal microscopy. This low-cost method is easy to perform using regular glass-bottom microscopy dishes for imaging on any inverted microscope. The mounting is performed in layers of agarose at different concentrations.

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues ...

Eye Removal in Living Zebrafish Larvae to Examine Innervation-dependent Growth and Development of the Visual System

Zebrafish exhibit remarkable life-long growth and regenerative abilities. For example, specialized stem cell niches established during embryogenesis support continuous growth of the entire visual system, both in the eye and the brain. Coordinated growth between the retinae and the optic tectum ensures accurate retinotopic mapping as new neurons are added in the eyes and brain. To address whether retinal axons provide crucial information for regulating tectal stem and progenitor cell behaviors such as survival, proliferation, and/or differentiation, it is necessary to be able to compare innervated and denervated tectal lobes within the same animal and across animals. 
Surgical removal of one eye from living larval zebrafish followed by observation of the optic tectum achieves this goal. The accompanying video demonstrates how to anesthetize larvae, electrolytically sharpen tungsten needles, and use them to remove one eye. It next shows how to dissect brains from fixed zebrafish larvae. Finally, the video provides an overview of the protocol for immunohistochemistry and a demonstration of how to mount stained embryos in low-melting-point agarose for microscopy.

The article explains how to surgically remove eyes from living zebrafish larvae as the first step toward investigating how retinal input influences optic tectum growth and development. In addition, the article provides information about larval anesthetization, fixation, and brain dissection, followed by immunohistochemistry and confocal imaging.

Zebrafish exhibit remarkable life-long growth and regenerative abilities. For example, specialized stem cell niches established during embryogenesis ...

Immunostaining of Dissected Zebrafish Embryonic Heart 

Zebrafish embryo becomes a popular in vivo vertebrate model for studying cardiac development and human heart diseases due to its advantageous embryology and genetics 1,2. About 100-200 embryos are readily available every week from a single pair of adult fish. The transparent embryos that develop ex utero make them ideal for assessing cardiac defects 3. The expression of any gene can be manipulated via morpholino technology or RNA injection 4. Moreover, forward genetic screens have already generated a list of mutants that affect different perspectives of cardiogenesis 5. 
Whole mount immunostaining is an important technique in this animal model to reveal the expression pattern of the targeted protein to a particular tissue 6. However, high resolution images that can reveal cellular or subcellular structures have been difficult, mainly due to the physical location of the heart and the poor penetration of the antibodies. 
 Here, we present a method to address these bottlenecks by dissecting heart first and then conducting the staining process on the surface of a microscope slide. To prevent the loss of small heart samples and to facilitate solution handling, we restricted the heart samples within a circle on the surface of the microscope slides drawn by an immEdge pen. After the staining, the fluorescence signals can be directly observed by a compound microscope. 
Our new method significantly improves the penetration for antibodies, since a heart from an embryonic fish only consists of few cell layers. High quality images from intact hearts can be obtained within a much reduced procession time for zebrafish embryos aged from day 2 to day 6. Our method can be potentially extended to stain other organs dissected from either zebrafish or other small animals.

Immunostaining of Dissected Zebrafish Embryonic Heart

A rapid way to conduct immunostaining of zebrafish embryonic heart is described. Compared to the whole mount immunostaining approach, this method dramatically increases the penetration of the antibodies, which allows obtaining high resolution images that reveal cellular/subcellular structures in the heart within a much reduced processing time.

Zebrafish embryo becomes a popular in vivo vertebrate model for studying cardiac development and human heart diseases due to its advantageous embryology ...

High Resolution Whole Mount In Situ Hybridization within Zebrafish Embryos to Study Gene Expression and Function

This article focuses on whole-mount in situ hybridization (WISH) of zebrafish embryos. The WISH technology facilitates the assessment of gene expression both in terms of tissue distribution and developmental stage. Protocols are described for the use of WISH of zebrafish embryos using antisense RNA probes labeled with digoxigenin. Probes are generated by incorporating digoxigenin-linked nucleotides through in vitro transcription of gene templates that have been cloned and linearized. The chorions of embryos harvested at defined developmental stages are removed before incubation with specific probes. Following a washing procedure to remove excess probe, embryos are incubated with anti-digoxigenin antibody conjugated with alkaline phosphatase. By employing a chromogenic substrate for alkaline phosphatase, specific gene expression can be assessed. Depending on the level of gene expression the entire procedure can be completed within 2-3 days.

High Resolution Whole Mount In Situ Hybridization within Zebrafish Embryos to Study Gene Expression and Function

The zebrafish, a small tropical fish, has become a popular model for studying gene function during vertebrate development and disease. The temporal and spatial expression of target genes can be determined by in situ hybridization. Our improved protocol allows for the detection of low abundant transcripts with low non-specific background signal.

This article focuses on whole-mount in situ hybridization (WISH) of zebrafish embryos. The WISH technology facilitates the assessment of gene expression ...

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model congenital abnormalities. During the first day of life, the zebrafish embryo progresses through many developmental stages including fertilization, cleavage, gastrulation, segmentation, and the organogenesis of structures such as the kidney, heart, and central nervous system. The anatomy of a young zebrafish embryo presents several challenges for the visualization and analysis of the tissues involved in many of these events because the embryo develops in association with a round yolk mass. Thus, for accurate analysis and imaging of experimental phenotypes in fixed embryonic specimens between the tailbud and 20 somite stage (10 and 19 hours post fertilization (hpf), respectively), such as those stained using whole mount in situ hybridization (WISH), it is often desirable to remove the embryo from the yolk ball and to position it flat on a glass slide. However, performing a flat mount procedure can be tedious. Therefore, successful and efficient flat mount preparation is greatly facilitated through the visual demonstration of the dissection technique, and also helped by using reagents that assist in optimal tissue handling. Here, we provide our WISH protocol for one or two-color detection of gene expression in the zebrafish embryo, and demonstrate how the flat mounting procedure can be performed on this example of a stained fixed specimen. This flat mounting protocol is broadly applicable to the study of many embryonic structures that emerge during early zebrafish development, and can be implemented in conjunction with other staining methods performed on fixed embryo samples.

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

The zebrafish embryo is an excellent model for developmental biology research. During embryogenesis, zebrafish develop with a yolk mass, which presents three-dimensional challenges for sample observation and analysis. This protocol describes how to create two-dimensional flat mount preparations of whole mount in situ (WISH) stained zebrafish embryo specimens.

The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model ...

In Situ Hybridization Combined with Immunohistochemistry in Cryosectioned Zebrafish Embryos

As a vertebrate, the zebrafish has been widely used in biological studies. Zebrafish and humans share high genetic homology, which allows its use as a model for human diseases. Gene function study is based on the detection of gene expression patterns. Although immunohistochemistry offers a powerful way to assay protein expression, the limited number of commercially available antibodies in zebrafish restricts the application of costaining. In situ hybridization is widely used in zebrafish embryos to detect mRNA expression. This protocol describes how to obtain images by combining in situ hybridization and immunohistochemistry for zebrafish embryo sections. In situ hybridization was performed prior to cryosectioning, followed by antibody staining. Immunohistochemistry and the imaging of a single cryosection were performed after in situ hybridization. The protocol is helpful to unravel the expression pattern of two genes, first by in situ transcript detection and then by immunohistochemistry against a protein in the same section.

In Situ Hybridization Combined with Immunohistochemistry in Cryosectioned Zebrafish Embryos

This protocol describes how to obtain images by combining in situ hybridization and immunohistochemistry of zebrafish embryonic sections. In situ hybridization was performed prior to cryosectioning, followed by antibody staining. It is useful to detect the expression patterns of two genes in zebrafish if there is a paucity of antibodies.

As a vertebrate, the zebrafish has been widely used in biological studies. Zebrafish and humans share high genetic homology, which allows its use as a ...

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

Summary

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Chapters in this video

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A Layered Mounting Method for Extended Time-Lapse Confocal Microscopy of Whole Zebrafish Embryos

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Immunostaining of Dissected Zebrafish Embryonic Heart

High Resolution Whole Mount In Situ Hybridization within Zebrafish Embryos to Study Gene Expression and Function

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

In Situ Hybridization Combined with Immunohistochemistry in Cryosectioned Zebrafish Embryos