Name: surgical size reduction of zebrafish for the study of embryonic pattern scaling
Uploaded: 2019-05-03T14:30:00.000+00:00
Duration: 6 min 31 s
Description: Watch this Scientific Journal Video about Surgical Size Reduction of Zebrafish for the Study of Embryonic Pattern Scaling at JoVE.com

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><tbody><tr><td>60 mm PYREX Petri dish</td><td>CORNING</td><td>&amp;nbsp;3160-60</td><td></td></tr><tr><td>Agarose</td><td>affymetrix</td><td>75817</td><td>For making a mount for live imaging</td></tr><tr><td>Agarose, low gelling temperature Type VII-A</td><td>SIGMA-ALDRICH</td><td>A0701-25G</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>EMD</td><td>CX0130-1</td><td>For 1/3 Ringer&#039;s solution</td></tr><tr><td>CaSO&lt;sub&gt;4&lt;/sub&gt;</td><td></td><td></td><td>For egg water</td></tr><tr><td>Cover slip (25 mm x 25 mm, Thickness 1)</td><td>CORNING</td><td>2845-25</td><td></td></tr><tr><td>Disposable Spatula</td><td>VWR&amp;nbsp;</td><td>80081-188</td><td></td></tr><tr><td>Foam board</td><td>ELMER&#039;S</td><td>951300</td><td>For microscope incubator</td></tr><tr><td>Forcept (No 55)</td><td>FST</td><td>11255-20</td><td></td></tr><tr><td>Glass pipette</td><td>VWR</td><td>14673-043</td><td></td></tr><tr><td>HEPES</td><td>SIGMA Life Science</td><td>H4034</td><td>For 1/3 Ringer&#039;s solution</td></tr><tr><td>INCUKIT XL for Cabinet Incubators</td><td>INCUBATOR Warehouse.com</td><td></td><td>For microscope incubator</td></tr><tr><td>Instant sea salt</td><td>Instant Ocean</td><td>138510</td><td>For egg water</td></tr><tr><td>KCl</td><td>SIGMA-ALDRICH</td><td>P4504</td><td>For 1/3 Ringer&#039;s solution</td></tr><tr><td>Methyl cellulose</td><td>SIGMA-ALDRICH</td><td>M0387-100G</td><td></td></tr><tr><td>NaCl</td><td>SIGMA-ALDRICH</td><td>S7653</td><td>For 1/3 Ringer&#039;s solution</td></tr><tr><td>Petri dish</td><td>Falcon</td><td>351029</td><td>For making a mount for live imaging</td></tr><tr><td>Phenol red</td><td>SIGMA Life Science</td><td>P0290</td><td></td></tr><tr><td>Pipette pump</td><td>BEL-ART PRODUCTS</td><td>F37898</td><td></td></tr><tr><td>Pronase</td><td>EMD Millipore Corp</td><td>53702-250KU</td><td></td></tr><tr><td>Tricaine-S (MS222)</td><td>WESTERN CHEMICAL INC</td><td>NC0135573</td><td></td></tr><tr><td>Ultra thin bright annealed 316L dia. 0.035 mm Stainless Steel Weaving Wires</td><td>Sandra</td><td></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

Universidad San Sebastian

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

Microbead Implantation in the Zebrafish Embryo

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic conservation, as well as similarities in cellular, molecular, and physiological processes, with other vertebrates including humans. During early ontogeny, zebrafish embryos are optically transparent, allowing researchers to visualize the dynamics of organogenesis using a simple stereomicroscope. Microbead implantation is a method that enables tissue manipulation through the alteration of factors in local environments. This allows researchers to assay the effects of any number of signaling molecules of interest, such as secreted peptides, at specific spatial and temporal points within the developing embryo. Here, we detail a protocol for how to manipulate and implant beads during early zebrafish development.

The zebrafish is an excellent model system for genetic and developmental studies. Bead implantation is a valuable tissue manipulation technique that can be used to interrogate developmental mechanisms by introducing alterations in local cellular environments. This protocol describes how to perform microbead implantation in the zebrafish embryo.

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic ...

Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning fundamental developmental biology through to large-scale genetic and chemical screens. However, many experiments, especially those related to infection and xenograft models, rely on microinjection and imaging of embryos and larvae, which are laborious techniques that require skill and expertise. To improve the precision and throughput of current microinjection techniques, we developed a series of microstructured devices to orient and stabilize zebrafish embryos at 2 days post fertilization (dpf) in ventral, dorsal, or lateral orientation prior to the procedure. To aid in the imaging of embryos, we also designed a simple device with channels that orient 4 zebrafish laterally in parallel against a glass cover slip. Together, the tools that we present here demonstrate the effectiveness of photolithographic approaches to generate useful devices for the optimization of zebrafish techniques.

Microinjection of zebrafish embryos and larvae is a crucial but challenging technique used in many zebrafish models. Here, we present a range of microscale tools to aid in the stabilization and orientation of zebrafish for both microinjection and imaging.

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning ...

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

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

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

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

Neuroscience

Spinal Cord Transection in the Larval Zebrafish

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability to reinitiate spinal neurogenesis. However, some anamniotes including the zebrafish Danio rerio exhibit both sensory and functional recovery even after complete transection of the spinal cord. The adult zebrafish is an established model organism for studying regeneration following spinal cord injury, with sensory and motor recovery by 6&#160;weeks post-injury. To take advantage of in vivo analysis of the regenerative process available in the transparent larval zebrafish as well as genetic tools not accessible in the adult, we use the larval zebrafish to study regeneration after spinal cord transection. Here we demonstrate a method for reproducibly and verifiably transecting the larval spinal cord. After transection, our data shows sensory recovery beginning at 2&#160;days post-injury (dpi), with the C-bend movement detectable by 3 dpi and resumption of free swimming by 5 dpi. Thus we propose the larval zebrafish as a companion tool to the adult zebrafish for the study of recovery after spinal cord injury.

After spinal transection, adult zebrafish have functional recovery by six weeks post-injury. To take advantage of larval transparency and faster recovery, we present a method for transecting the larval spinal cord. After transection, we observe sensory recovery beginning at 2&#160;days post-injury, and C-bend movement by 3 days post-injury.

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability ...

Quantifying Liver Size in Larval Zebrafish Using Brightfield Microscopy

In several transgenic zebrafish models of hepatocellular carcinoma (HCC), hepatomegaly can be observed during early larval stages. Quantifying larval liver size in zebrafish HCC models provides a means to rapidly assess the effects of drugs and other manipulations on an oncogene-related phenotype. Here we show how to fix zebrafish larvae, dissect the tissues surrounding the liver, photograph livers using bright-field microscopy, measure liver area, and analyze results. This protocol enables rapid, precise quantification of liver size. As this method involves measuring liver area, it may underestimate differences in liver volume, and complementary methodologies are required to differentiate between changes in cell size and changes in cell number. The dissection technique described herein is an excellent tool to visualize the liver, gut, and pancreas in their natural positions for myriad downstream applications including immunofluorescence staining and in situ hybridization. The described strategy for quantifying larval liver size is applicable to many aspects of liver development and regeneration.

Here we demonstrate a method for quantifying liver size in larval zebrafish, providing a way to assess the effects of genetic and pharmacologic manipulations on liver growth and development.

In several transgenic zebrafish models of hepatocellular carcinoma (HCC), hepatomegaly can be observed during early larval stages. Quantifying larval ...

A 3-D Tail Explant Culture to Study Vertebrate Segmentation in Zebrafish

Vertebrate embryos pattern their major body axis as repetitive somites, the precursors of vertebrae, muscle, and skin. Somites progressively segment from the presomitic mesoderm (PSM) as the tail end of the embryo elongates posteriorly. Somites form with regular periodicity and scale in size. Zebrafish is a popular model organism as it is genetically tractable and has transparent embryos that allow for live imaging. Nevertheless, during somitogenesis, fish embryos are wrapped around a large, rounding yolk. This geometry limits live imaging of PSM tissue in zebrafish embryos, particularly at higher resolutions that require a close objective working distance. Here, we present a flattened 3-D tissue culture method for live imaging of zebrafish tail explants. Tail explants mimic intact embryos by displaying a proportional slowdown of axis elongation and shortening of rostrocaudal somite lengths. We are further able to stall axis elongation speed through explant culture. This, for the first time, enables us to untangle the chemical input of signaling gradients from the mechanistic input of axial elongation. In future studies, this method can be combined with a microfluidic setup to allow time-controlled pharmaceutical perturbations or screening of vertebrate segmentation without any drug penetration concerns.

Here, we present the protocol for 3-D tissue culture of the zebrafish posterior body axis, enabling live study of vertebrate segmentation. This explant model provides control over axis elongation, alteration of morphogen sources, and subcellular resolution tissue-level live imaging.

Vertebrate embryos pattern their major body axis as repetitive somites, the precursors of vertebrae, muscle, and skin. Somites progressively segment from ...

A Simple and Effective Transplantation Device for Zebrafish Embryos

Classical embryological manipulations, such as removing cells and transplanting cells within or between embryos, are powerful techniques to study complex developmental processes. Zebrafish embryos are ideally suited for these manipulations since they are easily accessible, relatively large in size, and transparent. However, previously developed devices for cell removal and transplantation are cumbersome to use or expensive to purchase. In contrast, the transplantation device presented here is economical, easy to assemble, and simple to use. In this protocol, we first introduce the handling of the transplantation device as well as its assembly from commercially and widely available parts. We then present three applications for its use: generation of ectopic clones to study signal dispersal from localized sources, extirpation of cells to produce size-reduced embryos, and germline transplantation to generate maternal-zygotic mutants. Finally, we show that the tool can also be used for embryological manipulations in other species such as the Japanese rice fish medaka.

Embryological manipulations such as extirpation and transplantation of cells are important tools to study early development. This protocol describes a simple and effective transplantation device to perform these manipulations in zebrafish embryos.

Classical embryological manipulations, such as removing cells and transplanting cells within or between embryos, are powerful techniques to study complex ...

Robust Zebrafish Cell Transplantation for Regeneration and Development Research

High-resolution Cell Transplantation in Embryonic and Larval Zebrafish

Development and regeneration occur by a process of genetically encoded spatiotemporally dynamic cellular interactions. The use of cell transplantation between animals to track cell fate and to induce mismatches in the genetic, spatial, or temporal properties of donor and host cells is a powerful means of examining the nature of these interactions. Organisms such as chick and amphibians have made crucial contributions to our understanding of development and regeneration, respectively, in large part because of their amenability to transplantation. The power of these models, however, has been limited by low genetic tractability. Likewise, the major genetic model organisms have lower amenability to transplantation. 
The zebrafish is a major genetic model for development and regeneration, and while cell transplantation is common in zebrafish, it is generally limited to the transfer of undifferentiated cells at the early blastula and gastrula stages of development. In this article, we present a simple and robust method that extends the zebrafish transplantation window to any embryonic or larval stage between at least 1 and 7 days post fertilization. The precision of this approach allows for the transplantation of as little as one cell with near-perfect spatial and temporal resolution in both donor and host animals. While we highlight here the transplantation of embryonic and larval neurons for the study of nerve development and regeneration, respectively, this approach is applicable to a wide range of progenitor and differentiated cell types and research questions.

Author Spotlight: Mapping Cellular Connectivity in the Zebrafish Nervous System During Development and Regeneration

Here we present a protocol to transplant cells with high spatial and temporal resolution in zebrafish embryos and larvae at any stage between at least 1 and 7 days post fertilization.

Development and regeneration occur by a process of genetically encoded spatiotemporally dynamic cellular interactions. The use of cell transplantation ...

Removing Zebrafish Scales for Culturing and Bone Biology Studies

In Vitro Culturing Technique for Studying Cellular Dynamics in Zebrafish Scales

Zebrafish scales offer a variety of advantages for use in standard laboratories for teaching and research purposes. Scales are easily collected without the need for euthanasia, regenerate within a couple of weeks, and are translucent and small, allowing them to be viewed using a standard microscope. Zebrafish scales are especially useful in educational environments, as they provide a unique opportunity for students to engage in hands-on learning experiences, particularly in understanding cellular dynamics and in vitro culturing methods. The main objective of this protocol is to describe a method for collecting and maintaining zebrafish scales in culture for use in a variety of biological studies using basic laboratory equipment. Additionally, the protocol details their use in understanding bone homeostasis by examining the activity of bone cells involved in bone resorption and deposition. It also includes additional protocols for general techniques, such as the visualization of nuclei and apoptotic cells. The in vitro culturing protocol produces reliable results with minimal reagents and equipment. This article discusses the benefits of using in vitro cultures of zebrafish scales to foster scientific inquiry and outlines the resources needed to support their integration into educational settings.

Author Spotlight: Advancing Bone Cell Activity Research with Zebrafish Scales 

This protocol offers a method to study cellular dynamics using a simple in vitro culture technique. It provides an opportunity for zebrafish researchers and educators to study cellular processes, such as those related to bone homeostasis and basic cell biology, by visualizing fluorescent nuclei and apoptotic cells within the scales.

Zebrafish scales offer a variety of advantages for use in standard laboratories for teaching and research purposes. Scales are easily collected without ...

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

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

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

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

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

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

Summary

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Chapters in this video

Microbead Implantation in the Zebrafish Embryo

Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae

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

Spinal Cord Transection in the Larval Zebrafish

Quantifying Liver Size in Larval Zebrafish Using Brightfield Microscopy

A 3-D Tail Explant Culture to Study Vertebrate Segmentation in Zebrafish

A Simple and Effective Transplantation Device for Zebrafish Embryos

High-resolution Cell Transplantation in Embryonic and Larval Zebrafish

In Vitro Culturing Technique for Studying Cellular Dynamics in Zebrafish Scales

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