Name: Dissection of Zebrafish Craniofacial Tissues Upon Staining with Alcian Blue (Video) | JoVE Journal
Uploaded: 2025-09-12T13:50:00.000+00:00
Duration: 5 min 57 s
Description: Learn to dissect and stain craniofacial cartilages in 5-day-old zebrafish larvae using Alcian blue. This protocol enables detailed characterization of palate morphology at tissue and cellular levels.

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5 minutes

We acknowledge Dr. Kalidas Kohale and his team for the maintenance of the fish facility. SRN acknowledges financial support from the Department of Atomic Energy (DAE), Govt. of India (Project Identification no. RTI4003, DAE OM no. 1303/2/2019/R\&#38;D-II/DAE/2079 dated 11.02.2020), the Max Planck Society Partner Group program (M.PG.A MOZG0010) and the Science and Engineering Research Board Start-up Research Grant (SRG/2023/001716). We thank Dr. Shweta Verma from the lab for insightful comments. We thank Swetha Nagarajan and Upal Chatterjee for help with making figures for the manuscript.

The zebrafish maintenance and experimental procedures used in this study were approved by the institutional animal ethics committee, vide Reference TIFR/IAEC/2023-1.
1. Reagent preparation
<ol>
	<li>E3 media:
	<ol>
		<li>Prepare 50x stock 1 by mixing 3.25 g of sodium phosphate dibasic (0.458 mM Na2HPO4), 0.29 g of potassium dihydrogen phosphate (0.042 mM KH2PO4), 11.93 g of sodium chloride (4.084 mM NaCl), and 0.48 g of potassium chloride (0.128 mM KCl) in 1 L of distilled water.&#160;</li>
		<li>Prepare 50x stock 2 by mixing 2.43 g of calcium chloride (0.33 mM CaCl2) and 4.07 g of magnesium sulfate (0.33 mM MgSO4) in 1 L of distilled water.</li>
		<li>To prepare 500 mL of 1x E3 media, to 480 mL of distilled water, add 10 mL each of stocks 1 and 2 (pH = 7.4). 
		NOTE: Wear gloves during the preparation of these solutions.</li>
	</ol>
	</li>
	<li>4% paraformaldehyde (PFA) in 1x PBS (Phosphate-buffered saline):
	<ol>
		<li>Prewarm 100 mL of 1x PBS at 68 &#176;C. Add 4 g of PFA and incubate the solution at 68 &#176;C with periodic mixing until the PFA is completely dissolved. 
		NOTE: Wear gloves and a mask while preparing this reagent, preferably under a fume hood as PFA is highly toxic.</li>
	</ol>
	</li>
	<li>0.05% MS-222 (commonly known as tricaine) solution:
	<ol>
		<li>Make 4% tricaine stock in E3 and store at 4 &#176;C. To make 10 mL of tricaine stock, add 0.40 g of MS-222 in 7 mL of E3. Shake the tube containing the solution gently to mix and make up the volume to 10 mL. To make 50 mL of 0.05% tricaine solution, add 625 &#181;L of the 4% stock to 49.375 mL of E3. 
		NOTE: Wear gloves and a mask while preparing this reagent.</li>
	</ol>
	</li>
	<li>0.4% Alcian blue stock solution in 70% ethanol
	<ol>
		<li>To prepare 2 mL of the stock solution, dissolve 8 mg of Alcian blue in 1 mL of 50% ethanol. Heat the solution at 37 &#176;C for 15-20 min with periodic mixing to dissolve. Once the stain dissolves, make up the solution to 70% ethanol by adding 900 &#181;L of 100% ethanol and 100 &#181;L of distilled water. 
		NOTE: The stock solution can be stored at room temperature for up to 2 weeks.</li>
	</ol>
	</li>
	<li>Stain solution -- 0.02% Alcian blue, 200 mM magnesium chloride (MgCl2), and 70% ethanol
	<ol>
		<li>To prepare 5 mL of Alcian blue stain solution, add 1 mL of 1 M MgCl2, 3.5 mL of 100% ethanol, and 250 &#181;L of 0.4% Alcian blue stock (8 mg of Alcian blue in 2 mL of 70% ethanol) to 250 &#181;L of water and mix well. 
		NOTE: Prepare the stain solution fresh for best staining results.</li>
	</ol>
	</li>
	<li>Bleach solution:
	<ol>
		<li>Mix equal volumes of 3% hydrogen peroxide (H2O2) and 2% potassium hydroxide (KOH). To make 2 mL of bleach solution, mix 1 mL of 3% hydrogen peroxide and 1 mL of 2% potassium hydroxide. 
		NOTE: Prepare this solution fresh before beginning this step. Since H2O2 is light-sensitive, protect it from being exposed to light as much as possible. Wear gloves while handling H2O2 and KOH.</li>
	</ol>
	</li>
	<li>Solution I: 20% glycerol + 0.25% KOH 
	NOTE: As glycerol is highly viscous, pipette with care. Wear gloves while preparing these solutions.
	<ol>
		<li>To prepare 5 mL of solution I, add 2 mL of 50% glycerol and 625 &#181;L of 2% KOH to 2.375 mL of water and mix well.</li>
	</ol>
	</li>
	<li>Solution II: 50% glycerol + 0.25% KOH
	<ol>
		<li>To prepare 5 mL of solution II, add 3.570 mL of 70% glycerol and 625 &#181;L of 2% KOH to 805 &#181;L of water and mix well.</li>
	</ol>
	</li>
	<li>Storage solution: 50% glycerol + 0.1% KOH
	<ol>
		<li>To prepare 5 mL of storage solution, add 3.570 mL of 70% glycerol and 250 &#181;L of 2% KOH to 1.180 mL of water and mix well.</li>
	</ol>
	</li>
	<li>3% agarose
	<ol>
		<li>Dissolve 3 g of agarose in 100 mL of distilled water. Microwave till the solution becomes clear.</li>
	</ol>
	</li>
	<li>0.1% PBST (0.1% Tween 20 in 1x PBS)
	<ol>
		<li>To make 1 L of 1x PBS, add 8 g of NaCl, 0.20 g of KCl, 1.44 g of Na2HPO4, and 0.24 g of KH2PO4 in 1,000 mL of distilled water. Mix the solution and adjust the pH to 7.4. To make 500 mL of 0.1% PBST, add 5 mL of 10% Tween 20 to 495 mL of 1x PBS.</li>
	</ol>
	</li>
	<li>0.50 &#181;g/mL DAPI solution
	<ol>
		<li>Make 10 mg/mL DAPI stock in 1x PBS. To make 10 mL of 0.50 &#181;g/mL DAPI solution, add 0.50 &#181;L of the stock to 10 mL of 1x PBS and shake vigorously to ensure that DAPI dissolves completely. 
		NOTE: Wear gloves while preparing these solutions. Protect DAPI from light as it is light-sensitive. The DAPI stock can be stored at 4 &#176;C for more than a year.</li>
	</ol>
	</li>
</ol>
2. Embryo collection
<ol>
	<li>Set up pairs of male and female zebrafish in breeding tanks the evening before embryos are required, with the male and female fish separated by barriers.</li>
	<li>Remove the barriers the next morning and collect embryos within 15-20 min after the removal of the barriers. 
	NOTE: The timing of barrier removal is helpful in the staging of zebrafish embryos and larvae.</li>
	<li>Raise the embryos at 28.5 &#176;C until they reach 5 dpf. 
	NOTE: Be sure to replace the E3 media with a fresh batch of preheated E3 at 28.5 &#176;C once every 2 days.</li>
</ol>
3. Alcian blue staining
NOTE: Wear gloves while performing this experiment
<ol>
	<li>Anesthetize 20-25 5-day-old zebrafish larvae using 0.05% tricaine in a 2 mL microcentrifuge tube (MCT).</li>
	<li>After the larvae stop moving, which typically takes ~4-5 min, remove the anesthetic and fix them in 1.5 mL of 4% PFA at room temperature (RT) on a rocker at 60 rpm for 2 h.</li>
	<li>Remove as much of the fixative as possible using a pipette and add 1.5 mL of 50% ethanol to the sample. Incubate for 10 min while rocking at 60 rpm at RT.</li>
	<li>Remove as much of the ethanol as possible using a pipette and add 1.5 mL of the Alcian blue stain solution.</li>
	<li>Incubate the fixed larvae in the stain solution for 18-20 h on a rocker at 60 rpm at RT. 
	NOTE: Incubating for a shorter or longer duration of time can lead to certain regions of the cartilage to be understained or overstained, respectively. Here, 18-20 h of staining seemed to work best for clearly visualizing the neuro and the viscerocranium of a 5-day-old larva.</li>
	<li>Remove the stain solution, add 1.5 mL of distilled water, and incubate for 1-2 min on a rocker at RT. 
	NOTE: Staining can be stopped at this step to visualize cell boundaries in the dissected cartilages. However, a disadvantage of stopping the staining protocol at this step without performing subsequent clearing steps is that the boundaries of the neuro and the viscerocranium will still remain slightly blurred, and it becomes more challenging to perform the dissections precisely.</li>
	<li>Add 1 mL of the bleach solution to the sample and incubate for 20 min at RT in the open without any cover over the plate or wells. 
	NOTE: Do not cover the MCTs during this step as the reaction between hydrogen peroxide and potassium hydroxide produces oxygen gas, which could cause the cover to pop off if closed.</li>
	<li>Remove the bleach solution and perform the following tissue clearing steps.
	<ol>
		<li>Add 1.5 mL of solution I to the sample and keep on a rocker at 60 rpm for 40 min at RT.</li>
		<li>Remove solution I and add 1.5 mL of solution II to the sample and keep on a rocker at 60 rpm for 2 h at RT. 
		NOTE: The clearing steps help remove the excess stain, improving the optical clarity of the samples and thus aiding in performing the dissections efficiently. Handle samples with care while changing solutions, as any damage can affect the future analysis of their morphology.</li>
	</ol>
	</li>
	<li>Remove solution II and add 1.5-2 mL of the storage solution. Because of the presence of glycerol, stained samples gradually sink to the bottom within 10 min. 
	NOTE: Stained samples can be stored at 4 &#176;C for more than a year.</li>
</ol>
4. Dissection of the craniofacial skeleton
<ol>
	<li>Melt the 3% agarose solution in a microwave until the solution becomes clear. 
	NOTE: Do not overheat as the solution can bubble out of the flask. Keep a careful check of the solution while performing this step.</li>
	<li>Coat a 90 mm Petri dish with 3% agarose by pouring the molten agarose until one third of the depth of the Petri dish is covered and allowing it to solidify. Dissections will be performed in this Petri dish. 
	NOTE: The Petri dishes with agarose coating can be prepared a week in advance and stored at 4 &#176;C. The agarose coating provides a softer substrate on which the dissections can be performed without damaging the forceps.</li>
	<li>Clean the forceps to be used with 70% ethanol and wipe them with a clean piece of standard lab tissue paper.</li>
	<li>Using a Pasteur pipette, transfer 3-4 stained larvae along with 0.5 mL of storage solution to the agarose-coated petri dish. 
	NOTE: The storage solution ensures more optical clarity for performing the dissections compared to doing the same in E3.</li>
	<li>Orient a stained larva on its lateral side (Figure 1A) and carefully scrape off the yolk from the body of the larva using the forceps 
	NOTE: Figure 1B shows an embryo where the yolk has been removed. It is a good practice to transfer the yolk-scraped embryo to a new Petri dish before proceeding with further dissection steps so that the sticky yolk mass floating around does not interfere with the dissection.</li>
	<li>After the yolk has been removed, hold the larva by its tail region with one forceps and use the other to carefully remove the eyes. 
	NOTE: Figure 1C shows an embryo with the yolk and the eyes removed. Eyes should be removed with caution as their proximity to the craniofacial skeleton can interfere with their dissection.</li>
	<li>Using the forceps, carefully pull and pinch off the brain and other tissues dorsal to the neurocranium. Hold the larva still with one pair of forceps and use the other pair of forceps to make an incision in the middle (Figure 1C, red arrow), followed by pinching off&#160;the tissues (Figure 1D). 
	NOTE: While performing this step, be careful with removing the tissues closer to the anterior-most regions of the neurocranium, as the cartilage might come off with them.</li>
	<li>Separate the head of the larva from the rest of the body. The head region after this step primarily contains the neurocranium and the viscerocranium attached to each other at two points-at the anterior and posterior ends-along with a few small unstained tissues attached to these craniofacial structures (Figure 1E). 
	NOTE: It is difficult to entirely remove the unstained tissues directly attached to the neurocranium and the viscerocranium. Even though some residual tissues will be present, their presence will not affect any downstream shape or size analysis as they are unstained.</li>
	<li>Carefully sever the two connections between the neurocranium and the viscerocranium to separate them (Figure 1F, red dashed lines). See Figure 2 for images of the neurocranium and the viscerocranium both in intact as well as dissected embryos.</li>
	<li>On a clean glass slide, add a drop or two of 100% glycerol. With the help of forceps, carefully transfer the dissected neurocranium or viscerocranium to this slide. Use forceps to gently place a coverslip on these tissues, ensuring no bubbles are formed, and seal the cover slip on all slides with nail polish. 
	NOTE: These slides can be stored at 4 &#176;C for more than a year without losing the stain.</li>
</ol>
5. Quantification of the dimensions of the ethmoid plate
<ol>
	<li>Acquire images of the palate using a camera attached to the stereo microscope (Figure 2D). In the absence of a scientific-grade camera, attach a standard smartphone to the stereo microscope to acquire images.</li>
	<li>For quantification, load the image of a palate into FIJI22 and then, use the line tool to measure the width and the height of the ethmoid plate in microns (see Figure 3A-C) and determine the dimensions using the pixel resolution of the camera.</li>
	<li>Draw an outline around the ethmoid plate and click on Analyze | Set Measurements | area to obtain the area of the ethmoid plate (Figure 3D,E).</li>
</ol>

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The authors have no competing interests to declare.

Craniofacial malformations constitute a majority of all birth defects in humans, often leading to infant mortality. However, the underlying etiology in most craniofacial anomalies remains to be elucidated25. Craniofacial structures are complex in any given vertebrate, and from an undergraduate setting perspective, the first requisite is therefore to have an educational tool that provides students a general overview of distinct structures in the craniofacial region. However, having access to a mamm...

The craniofacial cartilage in vertebrates emerges from a multipotent cell population called the cranial neural crest cells1. These cells are first specified in the dorsal margins of the neural plate in early embryos, which then migrate large distances to reach the craniofacial region and the pharyngeal arches before differentiating2,3. The signaling mechanisms that drive this migration and differentiation are largely conserved among vertebrates4,5. Despite this, the organization, size, and shape of individual cartilage elements are distinct across species, giving rise to diverse craniofacial morphologies6. Zebrafish, in particular, has emerged as a powerful model system to study craniofacial morphogenesis, given the transparent nature of the embryos until larval stages. Using simple staining procedures for marking the cartilage, the morphology of many craniofacial structures in zebrafish has been characterized under different perturbations to signaling pathways7,8,9 as well as when embryos or larvae are exposed to common pollutants present in the environment10. The Alcian blue staining described here for marking the cartilage involves the use of non-acidic conditions standardized by Walker and Kimmel11, which has been subsequently used by many zebrafish labs working on craniofacial development5,12,13,14,15,16,17,18,19, including the description of usage in high-school settings20. Acid-free staining preserves the bony structures intact, allowing for staining cartilage and bones together using Alcian blue and alizarin red, respectively. However, a detailed protocol for staining as well as for performing precise dissection of the craniofacial cartilages, especially for aiding such studies to be undertaken in resource-limited settings, is missing.
Even though the facial skeletal structure in adult zebrafish is fairly complex, consisting of 43 cartilage-derived bones21, the craniofacial cartilage in a larva at 5 days post fertilization (dpf), which has just acquired the ability to feed, is relatively simple and can be broadly divided into a dorsal neurocranium and ventral viscerocranium4. These structures support the brain, contribute to the feeding apparatus and give rise to supporting cartilages for gill tissues. The protocol described here will allow for staining zebrafish cartilages with Alcian blue, followed by a detailed procedure for dissecting the neuro and the viscerocranium into separate structures, which will ultimately enable a careful characterization of the shape and size of various cartilages in these structures. As an example, we measure the dimensions of the anterior region of the palate (roof of the mouth) called the ethmoid plate, which is a part of the neurocranium.

<table><tbody><tr><td>Agarose</td><td>Lonza bioscience</td><td>50004</td><td>For coating petri dishes</td></tr><tr><td>Alcian blue 8G&amp;nbsp;</td><td>Sigma-Aldrich</td><td>A3157</td><td>For staining cartilages</td></tr><tr><td>Calcium Chloride dihydrate</td><td>Sigma-Aldrich</td><td>12022</td><td>For making buffers</td></tr><tr><td>Camera</td><td>Olympus</td><td>DP28</td><td>For acquisition of images</td></tr><tr><td>DAPI</td><td>Roche Life Sciences</td><td>10236276001</td><td>For labelling nuclei</td></tr><tr><td>Ethanol</td><td>Honeywell</td><td>32221</td><td>Solvent for alcian blue stain</td></tr><tr><td>FIJI</td><td></td><td></td><td>Version: ImageJ 1.54f</td></tr><tr><td>Forceps</td><td>Dumont</td><td>Dumont</td><td>For performing dissections</td></tr><tr><td>Glycerol</td><td>MP biomedicals</td><td>193996</td><td>For making washes and mounting samples&amp;nbsp;</td></tr><tr><td>Hydrogen peroxide</td><td>MP biomedicals</td><td>194057</td><td>For making bleach solution</td></tr><tr><td>Magnesium chloride hexahydrate</td><td>SARD Biosciences</td><td>SBRC0217</td><td>For making buffers</td></tr><tr><td>Magnesium sulfate heptahydrate</td><td>Sigma-Aldrich</td><td>M2773</td><td>For making buffers</td></tr><tr><td>MS-222 (Tricaine)</td><td>Sigma-Aldrich</td><td>A5040</td><td>For anaesthetising larvae</td></tr><tr><td>Potassium Chloride</td><td>Sigma-Aldrich</td><td>P9541</td><td>For making buffers</td></tr><tr><td>Potassium hydroxide</td><td>Sigma-Aldrich</td><td>484016</td><td>For making buffers</td></tr><tr><td>Potassium phosphate monobasic</td><td>Sigma-Aldrich</td><td>P0662</td><td>For making buffers</td></tr><tr><td>Sodium Chloride</td><td>Sigma-Aldrich</td><td>S3014</td><td>For making buffers</td></tr><tr><td>Sodium phosphate dibasic</td><td>Sigma-Aldrich</td><td>71640</td><td>For making buffers</td></tr><tr><td>Stereoscope</td><td>Olympus</td><td>SZX61</td><td>For performing dissections</td></tr><tr><td>Tween-20</td><td>HIMEDIA</td><td>TC287</td><td>Detergent, used for permeabilising cells</td></tr></tbody></table>

dissection of zebrafish craniofacial tissues upon staining with alcian blue

Dissection of Zebrafish Craniofacial Tissues Upon Staining with Alcian Blue

Zebrafish, given its optical transparency, is an excellent vertebrate model to study the mechanisms by which craniofacial cartilages form in the embryo. Craniofacial cartilages can be broadly classified into two groups, the neurocranium and the viscerocranium, which in turn can be subdivided into numerous cartilage groups necessary for supporting the brain and constituting the feeding and respiratory apparatus, among others. This protocol will first describe a simple and established staining procedure in zebrafish to visualize these cartilage groups, followed by a methodology to dissect and separate the neuro and the viscerocranium. From these dissections, simple shape and size metrics can be easily obtained, and in this protocol, this is demonstrated for the palate, which is part of the anterior neurocranium, a tissue often affected in most craniofacial disorders in humans. The protocol can be replicated in a straightforward manner even in resource-limited settings, thus providing a valuable educational tool for undergraduate students to get introduced to craniofacial morphology.

The zebrafish craniofacial skeleton is fully cartilaginous by 5 days post fertilization (dpf) and thus, can be stained using Alcian blue, which is a basic dye capable of binding to glycosaminoglycans present abundantly in cartilages23. This results in the cartilage being stained blue (Figure 1) so that it can be distinguished from other tissues present in the larva using a standard stereo microscope available in most undergraduate lab settings. Once the yolk is scrape...

The protocol described here presents a simple and easy-to-follow procedure for staining and dissecting craniofacial cartilages in a 5-day-old zebrafish larva. It can be used to study the anatomy, shape, and size of these structures under various developmental conditions.

Zebrafish, given its optical transparency, is an excellent vertebrate model to study the mechanisms by which craniofacial cartilages form in the embryo. ...

We are exploring the mechanisms by which zebra fish palate grows over time using fixed samples as well as time lapse imaging of embryos with high spatial temporal resolution using light sheet microscopy. Our protocol is simple and easy to follow. It lays down detailed steps to dissect out, and separate craniofacial cartilages of a five-day old zebrafish larva.This protocol has enabled careful characterization of palate shape at tissue and cellular scales, ultimately allowing us to assess the effect of commentary antigens on palate morphogenesis. To begin, obtain 20 to 25 anesthetized zebra fish larvae, and remove the anesthetic once the larvae have stopped moving. Add 1.5 milliliters of 4%paraform aldehyde to the tube, and place the sample on a rocker set to 60 revolutions per minute at room temperature for two hours.Using a pipette, remove as much of the fixative as possible from the tube. Then add 1.5 milliliters of 50%ethanol to the sample, and place it on a rocker at room temperature set to 60 revolutions per minute for 10 minutes. Now using a pipette, aspirate the ethanol from the tube, and add 1.5 milliliters of alcian blue stain solution to the sample.Incubate the larvae in the stain solution for 18 to 20 hours at room temperature while rocking at 60 revolutions per minute. Next, remove the stain solution using a pipette, and wash the larvae with 1.5 milliliters of distilled water for one to two minutes on a rocker at room temperature. Then add one milliliter of bleach solution to the sample, and incubate at room temperature for 20 minutes without covering the tubes.Using a pipette, remove the bleach solution from the sample, and treat the sample with tissue clearing solutions one and two. After removing solution two, add 1.5 to two milliliters of storage solution to the sample. Due to the glycerol content, observe as the stained samples gradually sink to the bottom within 10 minutes.Place a stained larva on its lateral side in the agarose coated Petri dish. Using forceps, gently scrape off the yolk from the body of the larvae. After yolk removal, transfer the larvae to a fresh Petri dish to avoid interference during dissection, hold the larvae near the posterior of the head using one pair of forceps.With another pair, carefully remove the eyes without damaging the surrounding cranial facial structures. While holding the larva, still with one pair of forceps, make an incision at the center of the tissue dorsal to the neuro cranium. Then pinch and pull off the brain, and associated tissues.Next, separate the head of the larvae from the rest of the body. The resulting head region should include the neuro cranium, and visceral cranium still attached at the anterior and posterior ends. Carefully sever the two connecting points between the neuro cranium, and visceral cranium with forceps to fully separate them.Finally, place one or two drops of 100%glycerol on a clean glass slide, and transfer the dissected neuro cranium or visceral cranium to the slide. Using forceps, gently place a cover slip on top of the tissues ensuring no air bubbles are trapped, and seal the cover slip with nail polish before imaging. The dissected neuro cranium displayed multiple clearly labeled structures, including the ethmoid plate, trabecula, and pericardial cartilage, enabling detailed structural analysis.The visceral cranium included Meckel's cartilage, and additional elements such as the palato quadrate, and basilbranchial cartilages. Alcian blue staining at five days post fertilization enabled clear visualization of the ethmoid plate. The width and length of the ethmoid plate were consistently measurable using image analysis, and the length was significantly greater than the width, and quantified data from 15 samples.The ethmoid plate area was extracted from image data, and confirmed through quantification with clear boundary markings. Distinct cellular arrangements were observed in the ethmoid plate with cuboidal cells in the medial region, and columer cells in the lateral parts. Dappy staining allowed visualization of individual cell nuclei within the ethmoid plate.

dissection-zebrafish-craniofacial-tissues-upon-staining-with-alcian

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JoVE Journal

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Dissection of Zebrafish Craniofacial Tissues Upon Staining with Alcian Blue

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