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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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(NO3)2]</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(NO3)2, 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 MgSO4, 75 uM KH2PO4, 25 uM Na2HPO4, 0.5 mM CaCl2, 0.35 mM NaHCO3, 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 x 18</td>
			<td>Corning</td>
			<td>284518</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass coverslips 22 x 60</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 (MgSO4 &#183; 7H2O)</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 dH2O</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 (KH2PO4)</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 (Na2HPO4)</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 dH2O</td>
		</tr>
		<tr>
			<td>Transfer pipette</td>
			<td>Fisher</td>
			<td>13-711-7M</td>
			<td></td>
		</tr>
		<tr>
			<td>Trichloracetic Acid (Cl3CCOOH)</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

Research

JoVE Journal

Developmental Biology

19.2K 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

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Multi-target Chromogenic Whole-mount In Situ Hybridization for Comparing Gene Expression Domains in Drosophila Embryos

To analyze gene regulatory networks active during embryonic development and organogenesis it is essential to precisely define how the different genes are expressed in spatial relation to each other in situ. Multi-target chromogenic whole-mount in situ hybridization (MC-WISH) greatly facilitates the instant comparison of gene expression patterns, as it allows distinctive visualization of different mRNA species in contrasting colors in the same sample specimen. This provides the possibility to relate gene expression domains topographically to each other with high accuracy and to define unique and overlapping expression sites. In the presented protocol, we describe a MC-WISH procedure for comparing mRNA expression patterns of different genes in Drosophila embryos. Up to three RNA probes, each specific for another gene and labeled by a different hapten, are simultaneously hybridized to the embryo samples and subsequently detected by alkaline phosphatase-based colorimetric immunohistochemistry. The described procedure is detailed here for Drosophila, but works equally well with zebrafish embryos.

Multi-target Chromogenic Whole-mount In Situ Hybridization for Comparing Gene Expression Domains in Drosophila Embryos

We describe a multi-target chromogenic whole-mount in situ hybridization (MC-WISH) procedure in intact Drosophila embryos allowing simultaneous and specific detection of three different mRNA distribution patterns by contrasting color precipitates.

To analyze gene regulatory networks active during embryonic development and organogenesis it is essential to precisely define how the different genes are ...

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize mammalian adaptive immunity. Drosophila and mammalian hematopoiesis occur in spatially and temporally distinct phases to produce several blood cell lineages. Both systems maintain reservoirs of blood cell progenitors with which to expand or replace mature lineages. The hematopoietic system allows Drosophila and mammals to respond to and to adapt to immune challenges. Importantly, the transcriptional regulators and signaling pathways that control the generation, maintenance, and function of the hematopoietic system are conserved from flies to mammals. These similarities allow Drosophila to be used to genetically model hematopoietic development and disease.
Here we detail assays to examine the hematopoietic system of Drosophila larvae. In particular, we outline methods to measure blood cell numbers and concentration, visualize a specific mature lineage in vivo, and perform immunohistochemistry on blood cells in circulation and in the hematopoietic organ. These assays can reveal changes in gene expression and cellular processes including signaling, survival, proliferation, and differentiation and can be used to investigate a variety of questions concerning hematopoiesis. Combined with the genetic tools available in Drosophila, these assays can be used to evaluate the hematopoietic system upon defined genetic alterations. While not specifically outlined here, these assays can also be used to examine the effect of environmental alterations, such as infection or diet, on the hematopoietic system.

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Drosophila and mammalian hematopoietic systems share many common features, making Drosophila an attractive genetic model to study hematopoiesis. Here we demonstrate dissection and mounting of the major larval hematopoietic organ for immunohistochemistry. We also describe methods to assay various larval hematopoietic compartments including circulating hemocytes and sessile crystal cells.

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize ...

Biosensing Motor Neuron Membrane Potential in Live Zebrafish Embryos

The protocols described here are designed to allow researchers to study cell communication without altering the integrity of the environment in which the cells are located. Specifically, they have been developed to analyze the electrical activity of excitable cells, such as spinal neurons. In such a scenario, it is crucial to preserve the integrity of the spinal cell, but it is also important to preserve the anatomy and physiological shape of the systems involved. Indeed, the comprehension of the manner in which the nervous system-and other complex systems-works must be based on a systemic approach. For this reason, the live zebrafish embryo was chosen as a model system, and the spinal neuron membrane voltage changes were evaluated without interfering with the physiological conditions of the embryos. 
Here, an approach combining the employment of zebrafish embryos with a FRET-based biosensor is described. Zebrafish embryos are characterized by a very simplified nervous system and are particularly suited for imaging applications thanks to their transparency, allowing for the employment of fluorescence-based voltage indicators at the plasma membrane during zebrafish development. The synergy between these two components makes it possible to analyze the electrical activity of the cells in intact living organisms, without perturbing the physiological state. Finally, this non-invasive approach can co-exist with other analyses (e.g., spontaneous movement recordings, as shown here).

Protocols described here allow for the study of the electrical properties of excitable cells in the most non-invasive physiological conditions by employing zebrafish embryos in an in vivo system together with a fluorescence resonance energy transfer (FRET)-based genetically encoded voltage indicator (GEVI) selectively expressed in the cell type of interest.

The protocols described here are designed to allow researchers to study cell communication without altering the integrity of the environment in which the ...

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 ...

Quantitative Whole-mount Immunofluorescence Analysis of Cardiac Progenitor Populations in Mouse Embryos

The use of ever-advancing imaging techniques has contributed broadly to our increased understanding of embryonic development. Pre-implantation development and organogenesis are two areas of research that have benefitted greatly from these advances, due to the high quality of data that can be obtained directly from imaging pre-implantation embryos or ex vivo organs. While pre-implantation embryos have yielded data with especially high spatial resolution, later stages have been less amenable to three-dimensional reconstruction. Obtaining high-quality 3D or volumetric data for known embryonic structures in combination with fate mapping or genetic lineage tracing will allow for a more comprehensive analysis of the morphogenetic events taking place during embryogenesis.
This protocol describes a whole-mount immunofluorescence approach that allows for the labeling, visualization, and quantification of progenitor cell populations within the developing cardiac crescent, a key structure formed during heart development. The approach is designed in such a way that both cell- and tissue-level information can be obtained. Using confocal microscopy and image processing, this protocol allows for three-dimensional spatial reconstruction of the cardiac crescent, thereby providing the ability to analyze the localization and organization of specific progenitor populations during this critical phase of heart development. Importantly, the use of reference antibodies allows for successive masking of the cardiac crescent and subsequent quantitative measurements of areas within the crescent. This protocol will not only enable a detailed examination of early heart development, but with adaptations should be applicable to most organ systems in the gastrula to early somite stage mouse embryo.

Here, we describe a protocol for whole mount immunofluorescence and image-based quantitative volumetric analysis of early stage mouse embryos. We present this technique as a powerful approach to qualitatively and quantitatively assess cardiac structures during development, and propose that it may be widely adaptable to other organ systems.

The use of ever-advancing imaging techniques has contributed broadly to our increased understanding of embryonic development. Pre-implantation development ...

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...

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 ...

Whole-Mount In Situ Hybridization in Zebrafish Embryos and Tube Formation Assay in iPSC-ECs to Study the Role of Endoglin in Vascular Development

Vascular development is determined by the sequential expression of specific genes, which can be studied by performing in situ hybridization assays in zebrafish during different developmental stages. To investigate the role of endoglin(eng) in vessel formation during the development of hereditary hemorrhagic telangiectasia&#160;(HHT), morpholino-mediated targeted knockdown of eng in zebrafish are used to study its temporal expression and associated functions. Here, whole-mount in situ RNA hybridization (WISH) is employed for the analysis of eng and its downstream genes in zebrafish embryos. Also, tube formation assays are performed in HHT patient-derived induced pluripotent stem cell-differentiated&#160;endothelial cells (iPSC-ECs; with eng mutations). A specific signal amplifying system using the whole amount In Situ Hybridization &#8211; WISH provides higher resolution and lower background results compared to traditional methods. To obtain a better signal, the post-fixation time is adjusted to 30 min after probe hybridization. Because fluorescence staining is not sensitive in zebrafish embryos, it is replaced with diaminobezidine (DAB) staining here. In this protocol, HHT&#160;patient-derived iPSC lines containing an eng mutation are differentiated into endothelial cells. After coating a plate with basement membrane matrix for 30 min at 37 &#176;C, iPSC-ECs are seeded as a monolayer into wells and kept at 37 &#176;C for 3 h. Then, the tube length and number of branches are calculated using microscopic images. Thus, with this improved WISH protocol, it is shown that reduced eng expression affects endothelial progenitor formation in zebrafish embryos. This is further confirmed by tube formation assays using iPSC-ECs derived from a patient with HHT. These assays confirm the role for eng in early vascular development.

Presented here is a protocol for whole-mount in situ RNA hybridization analysis in zebrafish and tube formation assay in patient-derived induced pluripotent stem cell-derived endothelial cells to study the role of endoglin in vascular formation.

Vascular development is determined by the sequential expression of specific genes, which can be studied by performing in situ hybridization assays in ...

Visualization and Analysis of Pharyngeal Arch Arteries using Whole-mount Immunohistochemistry and 3D Reconstruction

Improper formation or remodeling of the pharyngeal arch arteries (PAAs) 3, 4, and 6 contribute to some of the most severe forms of congenital heart disease. To study the formation of PAAs, we developed a protocol using whole-mount immunofluorescence coupled with benzyl alcohol/benzyl benzoate (BABB) tissue clearing, and confocal microscopy. This allows for the visualization of the pharyngeal arch endothelium at a fine cellular resolution as well as the 3D connectivity of the vasculature. Using software, we have established a protocol to quantify the number of endothelial cells (ECs) in PAAs, as well as the number of ECs within the vascular plexus surrounding the PAAs within pharyngeal arches 3, 4, and 6. When applied to the whole embryo, this methodology provides a comprehensive visualization and quantitative analysis of embryonic vasculature.

Here, we describe a protocol to visualize and analyze the pharyngeal arch arteries 3, 4, and 6 of mouse embryos using whole-mount immunofluorescence, tissue clearing, confocal microscopy, and 3D reconstruction.

Improper formation or remodeling of the pharyngeal arch arteries (PAAs) 3, 4, and 6 contribute to some of the most severe forms of congenital heart ...