The work was supported by the PRESTO program of the Japan Science and Technology Agency&#160;(JPMJPR11AA)&#160; and a National Institutes of Health grant (R01GM107733).

All fish-related procedures were carried out with the approval of the Institutional Animal Care and Use Committee (IACUC) at Harvard Medical School.
1. Tool and Reagent Preparation
<ol>
	<li>Make a wire loop to chop embryos
	<ol>
		<li>Take 20 cm of stainless steel wire that is stiff and non-corrosive with a diameter of 40 &#956;m. Loop the wire through into glass capillary (1.0 mm outer diameter, 0.5 mm inner diameter, no filament), making a small loop at the top (loop length is 1.0 mm)</li>
		<li>Put a little droplet of clear nail polish onto the tip of the glass capillary between the wire loop to hold it in place. Let dry. Make sure not to get any nail polish onto the loop portion, as it may damage the embryos.</li>
		<li>Attach the glass capillary with the loop onto a wooden chopstick (9&#8221; bamboo disposable chopsticks broken in half) using lab tape. Leave about 2.5 cm of the glass capillary extending beyond the chopstick, so that the chopstick part of the tool does not dip into the water. Adjust this length to preference.</li>
	</ol>
	</li>
	<li>Alternatively, make a glass needle to chop embryos
	<ol>
		<li>Pinch one end of glass Pasteur pipette with forceps, while holding the other side with a hand. Heat the thin part of the pipette over a spirit lamp or Bunsen burner.</li>
		<li>Hand-pull the glass pipette. The ideal pipette as a diameter of approximately 30 &#956;m and a gentle curve (radius of curvature = approximately 5 mm). Proper diameter and curvature is obtained with practice and some chance.</li>
	</ol>
	</li>
	<li>Prepare methyl cellulose
	<ol>
		<li>Methyl cellulose is used to hold the embryos while chopping. Make 2% methyl cellulose solution in &#8531;x Ringer&#8217;s solution (116 mM NaCl, 2.9 mM KCl, 1.8 mM CaCl2, and 5 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES; pH 7.2)). Shake methyl cellulose powder in &#8531;x Ringer&#8217;s solution at 4 &#176;C overnight.</li>
		<li>Optionally, add ~1.5 mL phenol red (0.5% in DPBS stock solution) to 10 mL 2% methyl cellulose solution, until red, to make the solution visible. Shake it until the color becomes uniform.</li>
	</ol>
	</li>
</ol>
2. Preparation of Zebrafish Embryos for Surgical Size Reduction
<ol>
	<li>Collect embryos
	<ol>
		<li>Place one or two male and one or two female zebrafish in a mating chamber filled with a lot of water. Separate males and females using a plastic divider. Use the AB line for somite imaging (section 4) and a transgenic reporter line of neural tube patterning (nkx2.2:mem-gfp, dbx1b:gfp, or Olig2:dsred) for neural tube imaging. Leave them in the chamber overnight. 
		NOTE: The choice of adults is important for obtaining eggs that survive surgery well. Typically, young healthy females produce healthy eggs.</li>
		<li>On the next morning, transfer the chamber into shallow water and place the chamber with slight tilt. Remove the divider so the fish can mate.</li>
		<li>Collect the eggs by pouring into a tea strainer. Transfer the eggs into a Petri dish with egg water (for 20x egg water, 6 g instant sea salt, 1.5 g CaSO4 and 1 L H2O; use at 1x). For better staging, collect the eggs right after spawning. Place the embryos in a 28.5 &#176;C incubator.</li>
		<li>If necessary, inject morpholino, mRNA, etc., at 1&#8211;4 cell stages following a common protocol7,8. Inject mRNA for fluorescent membrane label (mem-mCherry, mem-mBFP1, or mem-mCitrine) for neural tube imaging (section 5).</li>
	</ol>
	</li>
	<li>Dechorionate using pronase
	<ol>
		<li>Around the 128-cell to 256-cell stage, transfer healthy embryos to a 35 mm glass dish filled with egg water. Remove as much egg water as possible from the dish.</li>
		<li>Add 1 mL of 20 mg/mL pronase. Gently swirl embryos around by moving the plate and gently pipette the solution up and down to help dechorionation using a glass pipette.</li>
		<li>When the chorions start to lose elasticity (usually 1-4 min after addition of pronase, depending on the amount of eggs and water), add as much egg water as possible to dilute pronase. Assess elasticity loss by gently touching the chorion with forceps; the chorion should &#160;hold the dent without immediately bouncing back. Transfer the eggs to another dish with egg water using a glass pipette (at this point, most of the chorions are not broken).
		<ol>
			<li>Alternatively, gently pour the embryos from the dish into a large 400 mL glass beaker filled with egg water without exposing embryos to air. Then, let the embryos completely settle. Tilt the beaker to let embryos fall to one side. Collect these embryos using a glass pipette and transfer to a new 35 mm glass dish filled with &#8531;x Ringer&#8217;s solution.</li>
		</ol>
		</li>
		<li>Wait for several minutes until most of the chorions lose elasticity completely. Remove the remaining chorions from embryos by gently pipetting the eggs. Remove damaged embryos and incubate the embryos in a 28.5 &#176;C incubator.</li>
	</ol>
	</li>
</ol>
3. Surgical Size Reduction and Recovery
<ol>
	<li>Prepare one clean 35 mm glass dish. Spread approximately 0.5 mL of 2% methyl cellulose (in &#8531;x Ringer&#8217;s solution, with phenol red to help visualize) near the center of the bottom of the larger dish using a plastic spatula. Thinly and evenly spread the methyl cellulose to a thickness of approximately 0.5 mm.</li>
	<li>Pour approximately 30 mL of &#8531;x Ringer&#8217;s solution to the side of the dish and allow to spread onto the rest of the dish, covering the methyl cellulose.</li>
	<li>Place dechorionated embryos at the 256-cell-1k-cell stage on 2% methyl cellulose. Adjust the orientation of the embryos onto the side to visualize both the cells and the yolk. (Figure 1)</li>
	<li>Chop approximately 30%-40% of cells from the blastoderm near the animal pole by cutting perpendicular to the animal-vegetal axis using the wire loop (or the glass needle) (Figure 1, top panels). After removing the cells, gently tap the ends together to help the remaining cells stick back. Within minutes, the dead cells should slough off when the embryo starts to heal.</li>
	<li>Make a small wound to the yolk near the vegetal pole instead of &#8220;chopping&#8221; the yolk (Figure 1 middle panels). Wound the yolk by nicking the egg membrane with the mounted wire. Yolk will ooze out for few minutes after wounding, and then the wound will heal (Figure 1 bottom panels).</li>
	<li>When the yolk stops oozing out, move the embryo using a pipet outside of the methyl cellulose in the same dish to allow them to better recover.</li>
	<li>Repeat steps 3.4&#173;-3.6 for all embryos. Leave the dish stable for 30 min while the embryos recover.</li>
	<li>Transfer the embryos to a new dish with fresh &#8531;x Ringer&#8217;s solution and put them in 28.5 &#176;C incubator to allow them to completely recover.
	<ol>
		<li>(Optional) If the stage of interest is the early somite stage, embryos can be incubated at 20 &#176;C after being incubated at 28.5 &#176;C until the shield stage, to adjust timing for experimentation and imaging.</li>
	</ol>
	</li>
</ol>
4. Live Imaging of Zebrafish Somitogenesis
<ol>
	<li>Prepare 100 mL of 1% agarose solution by adding 1 g of agarose to 100 mL of egg water, heating until agarose is fully dissolved. Let cool for 5-10 min to a temperature of 62-72 &#176;C.</li>
	<li>Prepare a mount for imaging. 
	NOTE: This guide was developed for use with an inverted wide-field microscope.
	<ol>
		<li>Pour ~15 mL of 1% agarose in to a 100 mm x 15 mm plastic Petri dish.</li>
		<li>Gently place a Dorsal Mount V1 mold template5 onto the bottom of the Petri dish. Keep the mold on the bottom by using a weight, such as 15 mL tube with water. 
		NOTE: This is so the embryos are as close to the bottom of the Petri dish as possible, when using an inverted microscope. Although the embryos are mounted laterally for somite imaging, here Dorsal Mount V15 is used since it holds the embryos better at early somite stages.</li>
		<li>Let cool until agarose is fully solidified. Pour in ~30 mL of egg water with 0.01% tricaine, and carefully remove mold by gently prying off with forceps.</li>
	</ol>
	</li>
	<li>Mount embryos for somite imaging
	<ol>
		<li>Prepare 1% Low Melting Agarose in &#8531;x Ringer&#8217;s solution (or egg water) and keep it at 42 &#176;C. Wait until the temperature comes down to 42 &#176;C.</li>
		<li>Under a dissection microscope, place one embryo per well. Pipette approximately 1 &#956;L of Low Melting Agarose in the well to finely adjust the well size to the size of individual embryos that vary in size, especially between those with and without size reduction.</li>
		<li>Quickly orient the embryos before the Low Melting Agarose is solidified so the embryos face completely laterally to the dish. After mounting embryos, gently place the cover slip in the agarose mount to hold the embryos in place. The orientation is particularly important for the long-term somite imaging because the embryos have to be exactly laterally mounted for all the somite boundaries to be clearly imaged at later stages.</li>
		<li>Submerge away from the mounting area and manipulate a 25 mm x 25 mm cover glass such that it is 45 degrees offset from the orientation of the square indent made by the mold. Slide the coverslip over the mold in this orientation until each corner is resting of a different side of the mold. 
		NOTE: Although this protocol is for an inverted microscope, placing a cover glass on top of the mold is still important so that the embryos do not move while transferring to the microscope.</li>
	</ol>
	</li>
	<li>Imaging somite formation process
	<ol>
		<li>Prewarm the microscope to 28 &#176;C in an incubator built with foamcore boards and a cabinet heater.</li>
		<li>Place the Petri dish with the mounted embryos onto the microscope stage. Find the embryo mounted in the top-left well of the agarose mount with lower magnification lens, then switch the objective to 10x.</li>
		<li>Set up image acquisition. 
		NOTE: This instruction is for bright field.</li>
		<li>Find conditions for the light power, exposure time, and condenser so that the somite boundary can be clearly seen. Using z-stack settings set the lowest and highest desired imaging plane. Set the total time and time interval. Find and register the xy positions of all the embryos mounted so multiple embryos can be timelapse imaged at once.</li>
	</ol>
	</li>
	<li>Measure the lengths of PSM and somites using Fiji16.</li>
</ol>
5. Imaging of Neural Tube Patterning
<ol>
	<li>Prepare 100 mL of 1% agarose solution by adding 1 g of agarose to 100 mL of egg water, heating until agarose is fully dissolved. Let cool for 5-10 min to 62-72 &#176;C. Adjust tricaine to 0.01%.</li>
	<li>Prepare a dorsal mount for imaging. 
	NOTE: This guide is developed for use with an upright fluorescence microscope. For inverted microscopes coverslip bottom dishes are necessary and mounting orientation in step 3 is flipped.
	<ol>
		<li>Pour in ~15 mL of 1% agarose in to a 100 mm x 15 mm plastic Petri dish. Gently float a Dorsal Mount V1 mold on the surface of the agarose to avoid introducing air bubbles. Let cool until agarose is fully solidified.</li>
		<li>Carefully remove the mold with forceps or a razor blade. Fill the dish with egg water with 0.01% tricaine and cover until use.</li>
	</ol>
	</li>
	<li>Mount embryos for neural tube imaging.
	<ol>
		<li>Allow transgenic or injected fluorescent size-reduced and control embryos to develop until the 20-25 somite stage (roughly 18-22 h after fertilization). Embryos expressing a fluorescent membrane label (mem-mCherry, mem-mBFP1, or mem-mCitrine) and a transgenic reporter of neural tube patterning (nkx2.2:mem-gfp, dbx1b:gfp, or Olig2:dsred) are used for imaging.</li>
		<li>Replace egg water medium in prepared dorsal mount with 0.01% tricaine working solution. Gently pipette in embryos to be imaged in to wells of the dorsal mount.</li>
		<li>Manipulate embryos into the correct orientation such that the head is facing forward in the mount and the tail is pointing towards the rear with the dorsal portion facing towards the water surface.</li>
		<li>Orient the embryo such that the tail is lying flat and not pointed downward towards the bottom of the dish. It is sometimes helpful to rest the posterior portion of the tail on the side ledge of the mounting well to prevent the tail from sinking and inadvertently imaging the hindbrain. Repeat for each embryo. Make certain with size reduced embryos that they do not fall to deeply into the wells to be imaged.</li>
		<li>Submerge away from the mounting area and manipulate a 25 mm x 25 mm cover glass such that it is 45&#176; offset from the orientation of the square indent made by the mold. Slide the coverslip over the mold in this orientation until each corner is resting of a different side of the mold.</li>
		<li>Slowly rotate the coverglass until it gently falls on to the mounting area. Check to be sure embryos are still mounted in the correct positions and orientations. If too much movement is caused by the falling of the coverglass, remove gently and repeat mounting from step C.</li>
	</ol>
	</li>
	<li>Image in 3-D the developing spinal cord.
	<ol>
		<li>Gently transport the dish containing mounted embryos to the confocal microscope. For long-term live imaging, be sure to use an incubated microscope. Otherwise, low temperatures are tolerable.</li>
		<li>Under brightfield illumination, navigate to the embryo to image and center the field of view on the preferred anterior-posterior position. To enable a detailed comparison between embryos, this position on each embryo must be consistent.</li>
		<li>Set the imaging parameters to excite and capture signal from the fluorescent proteins being imaged. Parameters can be tuned by eye to create a desired image on the initial sample. To enable measurement consistency, apply these settings to all embryos in the dataset. If many z-stacks are required optimize settings for speed.</li>
		<li>Using z-stack settings, set the lowest and highest desired imaging plane. For optimal image quality in the z-stack, set the z-resolution to the highest that is optimal for the aperture setting. Good images can be generated with 1 &#956;m z-spacing.</li>
	</ol>
	</li>
	<li>Analyze imaging data by any preferred method. 
	NOTE: In this case, analysis was performed with custom MATLAB scripts that enable simple segmentation of the neural portions of an image and quantification of imaging signal.</li>
</ol>

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The authors declare no competing or financial interests.

Historically, among vertebrate animals, size reduction has been mainly performed using amphibian embryos, by bisecting the embryos along animal-vegetal axis at a blastula stage12. However, there are mainly two differences between frog and zebrafish embryos when we bisect embryos. First, at the stage when zebrafish embryos become tolerant of bisecting (blastula stage), the organizer is located in a restricted area of blastula margin18,...

Scientists have long known that embryos have a remarkable ability to form constant body proportions although embryo size can vary greatly both under natural and experimental conditions1,2,3. Despite decades of theoretical and experimental studies, this robustness to size variation, termed scaling, and its underlying mechanisms remain unknown in many tissues and organs. In order to directly capture the dynamics of the developing system, we established a reproducible and simple size reduction technique in zebrafish4, which has the great advantage in in vivo live imaging5.
Zebrafish has served as a model vertebrate animal to study multiple disciplines of biology, including developmental biology. In particular, zebrafish is ideal for in vivo live imaging6 because 1) development can proceed normally outside the mother and the egg shell, and 2) the embryos are transparent. In addition, the embryos can withstand some temperature and environmental fluctuations, which allows them to be studied in laboratory conditions. Also, in addition to conventional gene expression perturbation by morpholino and mRNA injection7,8, recent advances in CRISPR/Cas9 technology has made reverse genetics in zebrafish highly efficient9. Furthermore, many classical techniques in embryology, such as cell transplantation or tissue surgery can be applied4,10,11.
Size reduction techniques were originally developed in amphibian and other non-vertebrate animals12. For example, in Xenopus laevis, another popular vertebrate animal model, bisection along the animal-vegetal axis at blastula stage can produce size-reduced embryos12,13. However, in our hands this one-step approach results in dorsalized or ventralized embryos in zebrafish, presumably because dorsal determinants are distributed unevenly and one cannot know their localization from the morphology of embryos. Here we demonstrate an alternative two-step chopping technique for zebrafish that produces normally developing but smaller embryos. With this technique, cells are first removed from the animal pole, a region of na&#239;ve cells lacking in organizer activity. To balance the amount of yolk and cells, which is important for epiboly and subsequent morphogenesis, yolk is then removed. Here, we detail this protocol and provide two examples of size invariance in pattern formation; somite formation and ventral neural tube patterning. Combined with quantitative imaging, we utilized the size reduction technique to examine the how the sizes of somites and neural tube are affected in size reduced embryos.

<table border="0" cellpadding="0" cellspacing="0" style="width:608px;" width="609">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">60 mm PYREX Petri dish</td>
			<td style="width:107px;">CORNING</td>
			<td style="width:119px;">&#160;3160-60</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Agarose</td>
			<td style="width:107px;">affymetrix</td>
			<td align="right" style="width:119px;">75817</td>
			<td>For making a mount for live imaging</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Agarose, low gelling temperature Type VII-A</td>
			<td style="width:107px;">SIGMA-ALDRICH</td>
			<td style="width:119px;">A0701-25G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">CaCl2</td>
			<td style="width:107px;">EMD</td>
			<td style="width:119px;">CX0130-1</td>
			<td>For 1/3 Ringer&#39;s solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">CaSO4</td>
			<td style="width:107px;"></td>
			<td style="width:119px;"></td>
			<td>For egg water</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Cover slip (25 mm x 25 mm, Thickness 1)</td>
			<td style="width:107px;">CORNING</td>
			<td style="width:119px;">2845-25</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Disposable Spatula</td>
			<td style="width:107px;">VWR&#160;</td>
			<td style="width:119px;">80081-188</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Foam board</td>
			<td style="width:107px;">ELMER&#39;S</td>
			<td align="right" style="width:119px;">951300</td>
			<td>For microscope incubator</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Forcept (No 55)</td>
			<td style="width:107px;">FST</td>
			<td style="width:119px;">11255-20</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Glass pipette</td>
			<td style="width:107px;">VWR</td>
			<td style="width:119px;">14673-043</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">HEPES</td>
			<td style="width:107px;">SIGMA Life Science</td>
			<td style="width:119px;">H4034</td>
			<td>For 1/3 Ringer&#39;s solution</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:216px;">INCUKIT XL for Cabinet Incubators</td>
			<td style="width:107px;">INCUBATOR Warehouse.com</td>
			<td style="width:119px;"></td>
			<td>For microscope incubator</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Instant sea salt</td>
			<td style="width:107px;">Instant Ocean</td>
			<td align="right" style="width:119px;">138510</td>
			<td>For egg water</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">KCl</td>
			<td style="width:107px;">SIGMA-ALDRICH</td>
			<td style="width:119px;">P4504</td>
			<td>For 1/3 Ringer&#39;s solution</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Methyl cellulose</td>
			<td style="width:107px;">SIGMA-ALDRICH</td>
			<td style="width:119px;">M0387-100G</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">NaCl</td>
			<td style="width:107px;">SIGMA-ALDRICH</td>
			<td style="width:119px;">S7653</td>
			<td>For 1/3 Ringer&#39;s solution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Petri dish</td>
			<td style="width:107px;">Falcon</td>
			<td align="right" style="width:119px;">351029</td>
			<td>For making a mount for live imaging</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Phenol red</td>
			<td style="width:107px;">SIGMA Life Science</td>
			<td style="width:119px;">P0290</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Pipette pump</td>
			<td style="width:107px;">BEL-ART PRODUCTS</td>
			<td style="width:119px;">F37898</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Pronase</td>
			<td style="width:107px;">EMD Millipore Corp</td>
			<td style="width:119px;">53702-250KU</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Tricaine-S (MS222)</td>
			<td style="width:107px;">WESTERN CHEMICAL INC</td>
			<td style="width:119px;">NC0135573</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Ultra thin bright annealed 316L dia. 0.035 mm Stainless Steel Weaving Wires</td>
			<td style="width:107px;">Sandra</td>
			<td style="width:119px;"></td>
			<td>The wire we used was obtained ~20 years ago and we could not find exactly the same one. This product has the same material and diameter as the one we use.</td>
		</tr>
	</tbody>
</table>

surgical size reduction of zebrafish for the study of embryonic pattern scaling

In the developmental process, embryos exhibit a remarkable ability to match their body pattern to their body size; their body proportion is maintained even in embryos that are larger or smaller, within certain limits. Although this phenomenon of scaling has attracted attention for over a century, understanding the underlying mechanisms has been limited, owing in part to a lack of quantitative description of developmental dynamics in embryos of varied sizes. To overcome this limitation, we developed a new technique to surgically reduce the size of zebrafish embryos, which have great advantages for in vivo live imaging. We demonstrate that after balanced removal of cells and yolk at the blastula stage in separate steps, embryos can quickly recover under the right conditions and develop into smaller but otherwise normal embryos. Since this technique does not require special equipment, it is easily adaptable, and can be used to study a wide range of scaling problems, including robustness of morphogen mediated patterning.&#160;

Yolk volume reduction is important for normal morphology 
As recently described in Almuedo-Castillo et al.17, size reduction of embryos can be achieved without reducing yolk volume. To compare with and without yolk volume reduction, we performed both two-step chopping (both blastula and yolk) and blastula-only chopping (Figure 2 and Supplemental Movie 1). Two-step chopped embryos showed seemingly normal overall morphology compare...

Watch this Scientific Journal Video about Surgical Size Reduction of Zebrafish for the Study of Embryonic Pattern Scaling at JoVE.com

Surgical Size Reduction of Zebrafish for the Study of Embryonic Pattern Scaling

Here, we describe a method for reducing the size of zebrafish embryos without disrupting normal developmental processes. This technique enables the study of pattern scaling and developmental robustness against size change.

In the developmental process, embryos exhibit a remarkable ability to match their body pattern to their body size; their body proportion is maintained ...

surgical-size-reduction-zebrafish-for-study-embryonic-pattern

Research

JoVE Journal

Developmental Biology

6.6K Views.  Harvard Medical School. Here, we describe a method for reducing the size of zebrafish embryos without disrupting normal developmental processes. This technique enables the study of pattern scaling and developmental robustness against size change.

Video: Surgical Size Reduction of Zebrafish for the Study of Embryonic Pattern Scaling

Large-scale Zebrafish Embryonic Heart Dissection for Transcriptional Analysis

The zebrafish embryonic heart is composed of only a few hundred cells, representing only a small fraction of the entire embryo. Therefore, to prevent the cardiac transcriptome from being masked by the global embryonic transcriptome, it is necessary to collect sufficient numbers of hearts for further analyses. Furthermore, as zebrafish cardiac development proceeds rapidly, heart collection and RNA extraction methods need to be quick in order to ensure homogeneity of the samples. Here, we present a rapid manual dissection protocol for collecting functional/beating hearts from zebrafish embryos. This is an essential prerequisite for subsequent cardiac-specific RNA extraction to determine cardiac-specific gene expression levels by transcriptome analyses, such as quantitative real-time polymerase chain reaction (RT-qPCR). The method is based on differential adhesive properties of the zebrafish embryonic heart compared with other tissues; this allows for the rapid physical separation of cardiac from extracardiac tissue by a combination of fluidic shear force disruption, stepwise filtration and manual collection of transgenic fluorescently labeled hearts.

To analyse cardiac gene expression profiles during zebrafish heart development, total RNA has to be extracted from isolated hearts. Here, we present a protocol for collecting functional/beating hearts by rapid manual dissection from zebrafish embryos to obtain cardiac-specific mRNA.

The zebrafish embryonic heart is composed of only a few hundred cells, representing only a small fraction of the entire embryo. Therefore, to prevent the ...

Use of the TetON System to Study Molecular Mechanisms of Zebrafish Regeneration

The zebrafish has become a very important model organism for studying vertebrate development, physiology, disease, and tissue regeneration. A thorough understanding of the molecular and cellular mechanisms involved requires experimental tools that allow for inducible, tissue-specific manipulation of gene expression or signaling pathways. Therefore, we and others have recently adapted the TetON system for use in zebrafish. The TetON system facilitates temporally and spatially-controlled gene expression and we have recently used this tool to probe for tissue-specific functions of Wnt/beta&#8211;catenin signaling during zebrafish tail fin regeneration. Here we describe the workflow for using the TetON system to achieve inducible, tissue-specific gene expression in the adult regenerating zebrafish tail fin. This includes the generation of stable transgenic TetActivator and TetResponder lines, transgene induction and techniques for verification of tissue-specific gene expression in the fin regenerate. Thus, this protocol serves as blueprint for setting up a functional TetON system in zebrafish and its subsequent use, in particular for studying fin regeneration.

Here we outline the workflow for using the TetON system to achieve tissue-specific gene expression in the adult regenerating zebrafish tail fin.

The zebrafish has become a very important model organism for studying vertebrate development, physiology, disease, and tissue regeneration. A thorough ...

Hepatocyte-specific Ablation in Zebrafish to Study Biliary-driven Liver Regeneration

The liver has a great capacity to regenerate. Hepatocytes, the parenchymal cells of the liver, can regenerate in one of two ways: hepatocyte- or biliary-driven liver regeneration. In hepatocyte-driven liver regeneration, regenerating hepatocytes are derived from preexisting hepatocytes, whereas, in biliary-driven regeneration, regenerating hepatocytes are derived from biliary epithelial cells (BECs). For hepatocyte-driven liver regeneration, there are excellent rodent models that have significantly contributed to the current understanding of liver regeneration. However, no such rodent model exists for biliary-driven liver regeneration. We recently reported on a zebrafish liver injury model in which BECs extensively give rise to hepatocytes upon severe hepatocyte loss. In this model, hepatocytes are specifically ablated by a pharmacogenetic means. Here we present in detail the methods to ablate hepatocytes and to analyze the BEC-driven liver regeneration process. This hepatocyte-specific ablation model can be further used to discover the underlying molecular and cellular mechanisms of biliary-driven liver regeneration. Moreover, these methods can be applied to chemical screens to identify small molecules that augment or suppress liver regeneration.

To help assess the molecular mechanisms underlying zebrafish biliary-driven liver regeneration, we established a liver injury model in which the nitroreductase-expressing hepatocytes are genetically ablated upon metronidazole treatment. In this protocol, we describe how to adeptly manipulate, monitor and analyze hepatocyte ablation and biliary-driven liver regeneration.

The liver has a great capacity to regenerate. Hepatocytes, the parenchymal cells of the liver, can regenerate in one of two ways: hepatocyte- or ...

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 ...

Development of an Ethanol-induced Fibrotic Liver Model in Zebrafish to Study Progenitor Cell-mediated Hepatocyte Regeneration

Sustained liver fibrosis with continuation of extracellular matrix (ECM) protein build-up results in the loss of cellular competency of the liver, leading to cirrhosis with hepatocellular dysfunction. Among multiple hepatic insults, alcohol abuse can lead to significant health problems including liver failure and hepatocellular carcinoma. Nonetheless, the identity of endogenous cellular sources that regenerate hepatocytes in response to alcohol has not been properly investigated. Moreover, few studies have effectively modeled hepatocyte regeneration upon alcohol-induced injury. We recently reported on establishing an ethanol (EtOH)-induced fibrotic liver model in zebrafish in which hepatic progenitor cells (HPCs) gave rise to hepatocytes upon near-complete hepatocyte loss in the presence of fibrogenic stimulus. Furthermore, through chemical screens using this model, we identified multiple small molecules that enhance hepatocyte regeneration. Here we describe in detail the procedures to develop an EtOH-induced fibrotic liver model and to perform chemical screens using this model in zebrafish. This protocol will be a critical tool to delineate the molecular and cellular mechanisms of how hepatocyte regenerates in the fibrotic liver. Furthermore, these methods will facilitate potential discovery of novel therapeutic strategies for chronic liver disease in vivo.

Sustained fibrosis with deposition of excessive extracellular matrix proteins leads to cirrhosis. Alcohol abuse is one of the main causes of severe liver disease. We established an ethanol-induced zebrafish fibrotic liver model to study the mechanisms and strategies of promoting hepatocyte regeneration upon alcohol-induced injury.

Sustained liver fibrosis with continuation of extracellular matrix (ECM) protein build-up results in the loss of cellular competency of the liver, leading ...

Generation of Parabiotic Zebrafish Embryos by Surgical Fusion of Developing Blastulae

Surgical parabiosis of two animals of different genetic backgrounds creates a unique scenario to study cell-intrinsic versus cell-extrinsic roles for candidate genes of interest, migratory behaviors of cells, and secreted signals in distinct genetic settings. Because parabiotic animals share a common circulation, any blood or blood-borne factor from one animal will be exchanged with its partner and vice versa. Thus, cells and molecular factors derived from one genetic background can be studied in the context of a second genetic background. Parabiosis of adult mice has been &#160;used extensively to research aging, cancer, diabetes, obesity, and brain development. More recently, parabiosis of zebrafish embryos has been used to study the developmental biology of hematopoiesis. In contrast to mice, the transparent nature of zebrafish embryos permits the direct visualization of cells in the parabiotic context, making it a uniquely powerful method for investigating fundamental cellular and molecular mechanisms. The utility of this technique, however, is limited by a steep learning curve for generating the parabiotic zebrafish embryos. This protocol provides a step-by-step method on how to surgically fuse the blastulae of two zebrafish embryos of different genetic backgrounds to investigate the role of candidate genes of interest. In addition, the parabiotic zebrafish embryos are tolerant to heat shock, making temporal control of gene expression possible. This method does not require a sophisticated set-up and has broad applications for studying cell migration, fate specification, and differentiation in vivo during embryonic development.

This protocol provides step-by-step instruction on how to generate parabiotic zebrafish embryos of different genetic backgrounds. When combined with the unparalleled imaging capabilities of the zebrafish embryo, this method provides a uniquely powerful means to investigate cell-autonomous versus non-cell-autonomous functions for candidate genes of interest.

Surgical parabiosis of two animals of different genetic backgrounds creates a unique scenario to study cell-intrinsic versus cell-extrinsic roles for ...

Culture of Murine Embryonic Metatarsals: A Physiological Model of Endochondral Ossification

The fundamental process of endochondral ossification is under tight regulation in the healthy individual so as to prevent disturbed development and/or longitudinal bone growth. As such, it is imperative that we further our understanding of the underpinning molecular mechanisms involved in such disorders so as to provide advances towards human and animal patient benefit. The mouse metatarsal organ explant culture is a highly physiological ex vivo model for studying endochondral ossification and bone growth as the growth rate of the bones in culture mimic that observed in vivo. Uniquely, the metatarsal organ culture allows the examination of chondrocytes in different phases of chondrogenesis and maintains cell-cell and cell-matrix interactions, therefore providing conditions closer to the in vivo situation than cells in monolayer or 3D culture. This protocol describes in detail the intricate dissection of embryonic metatarsals from the hind limb of E15 murine embryos and the subsequent analyses that can be performed in order to examine endochondral ossification and longitudinal bone growth.

We present a protocol to dissect and culture embryonic day 15 (E15) murine metatarsal bones. This highly physiological ex vivo model of endochondral ossification provides conditions closer to the in vivo situation than cells in monolayer or 3D culture and is a vital tool for investigating bone growth and development.

The fundamental process of endochondral ossification is under tight regulation in the healthy individual so as to prevent disturbed development and/or ...

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced in vivo throughout the rapid developmental period. Multiple processes can be studied including cell proliferation, gene expression, cell migration and morphogenesis. Importantly, the generation of chimeras through transplantations can be easily performed, allowing mosaic labeling and tracking of individual cells under the influence of the host environment. For example, by combining functional gene manipulations of the host embryo (e.g., through morpholino microinjection) and live imaging, the effects of extrinsic, cell nonautonomous signals (provided by the genetically modified environment) on individual transplanted donor cells can be assessed. Here we demonstrate how this approach is used to compare the onset of fluorescent transgene expression as a proxy for the timing of cell fate determination in different genetic host environments.
In this article, we provide the protocol for microinjecting zebrafish embryos to mark donor cells and to cause gene knockdown in host embryos, a description of the transplantation technique used to generate chimeric embryos, and the protocol for preparing and running in vivo time-lapse confocal imaging of multiple embryos. In particular, performing multiposition imaging is crucial when comparing timing of events such as the onset of gene expression. This requires data collection from multiple control and experimental embryos processed simultaneously. Such an approach can easily be extended for studies of extrinsic influences in any organ or tissue of choice accessible to live imaging, provided that transplantations can be targeted easily according to established embryonic fate maps.

In Vivo Imaging of Transgenic Gene Expression in Individual Retinal Progenitors in Chimeric Zebrafish Embryos to Study Cell Nonautonomous Influences

Live tracking of individual WT retinal progenitors in distinct genetic backgrounds allows for the assessment of the contribution of cell non-autonomous signaling during neurogenesis. Here, a combination of gene knockdown, chimera generation via embryo transplantation and in vivo time-lapse confocal imaging was utilized for this purpose.

The genetic and technical strengths have made the zebrafish vertebrate a key model organism in which the consequences of gene manipulations can be traced ...

A Method for Characterizing Embryogenesis in Arabidopsis

Given the highly predictable nature of their development, Arabidopsis embryos have been used as a model for studies of morphogenesis in plants. However, early stage plant embryos are small and contain few cells, making them difficult to observe and analyze. A method is described here for characterizing pattern formation in plant embryos under a microscope using the model organism Arabidopsis. Following the clearance of fresh ovules using Hoyer's solution, the cell number in and morphology of embryos could be observed, and their developmental stage could be determined by differential interference contrast microscopy using a 100X oil immersion lens. In addition, the expression of specific marker proteins tagged with Green Fluorescent Protein (GFP) was monitored to annotate cell identity specification during embryo patterning by confocal laser scanning microscopy. Thus, this method can be used to observe pattern formation in wild-type plant embryos at the cellular and molecular levels, and to characterize the role of specific genes in embryo patterning by comparing pattern formation in embryos from wild-type plants and embryo-lethal mutants. Therefore, the method can be used to characterize embryogenesis in Arabidopsis.

A Method for Characterizing Embryogenesis in Arabidopsis

This protocol outlines a method for observing embryogenesis in Arabidopsis via ovule clearance followed by the inspection of embryo pattern formation under a microscope.

Given the highly predictable nature of their development, Arabidopsis embryos have been used as a model for studies of morphogenesis in plants. However, ...

Using In Vivo and Tissue and Cell Explant Approaches to Study the Morphogenesis and Pathogenesis of the Embryonic and Perinatal Aorta

The aorta is the largest artery in the body. The aortic wall is composed of an inner layer of endothelial cells, a middle layer of alternating elastic lamellae and smooth muscle cells (SMCs), and an outer layer of fibroblasts and extracellular matrix. In contrast to the widespread study of pathological models (e.g., atherosclerosis) in the adult aorta, much less is known about the embryonic and perinatal aorta. Here, we focus on SMCs and provide protocols for the analysis of the morphogenesis and pathogenesis of embryonic and perinatal aortic SMCs in normal development and disease. Specifically, the four protocols included are: i) in vivo embryonic fate mapping and clonal analysis; ii) explant embryonic aorta culture; iii) SMC isolation from the perinatal aorta; and iv) subcutaneous osmotic mini-pump placement in pregnant (or non-pregnant) mice. Thus, these approaches facilitate the investigation of the origin(s), fate, and clonal architecture of SMCs in the aorta in vivo. They allow for modulating embryonic aorta morphogenesis in utero by continuous exposure to pharmacological agents. In addition, isolated aortic tissue explants or aortic SMCs can be used to gain insights into the role of specific gene targets during fundamental processes such as muscularization, proliferation, and migration. These hypothesis-generating experiments on isolated SMCs and the explanted aorta can then be assessed in the in vivo context through pharmacological and genetic approaches.

Using In Vivo and Tissue and Cell Explant Approaches to Study the Morphogenesis and Pathogenesis of the Embryonic and Perinatal Aorta

Protocols for studying the embryonic and perinatal murine aorta using in vivo clonal analysis and fate mapping, aortic explants, and isolated smooth muscle cells are detailed here. These diverse approaches facilitate the investigation of the morphogenesis of the embryonic and perinatal aorta in normal development and the pathogenesis in disease.

The aorta is the largest artery in the body. The aortic wall is composed of an inner layer of endothelial cells, a middle layer of alternating elastic ...