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 margin18,19,20,21. Because one cannot tell the position of the organizer from the morphology of embryos, randomly cutting the embryo along the animal-vegetal axis produces dorsalized or ventralized embryos. Second, unlike frog embryos, zebrafish embryos go through a process called epiboly, where cells move towards the vegetal pole around a separated yolk until it is completely surrounded by cells. If only a portion of blastoderm is removed, fewer cells remain to engulf a yolk of relatively larger volume, and as a result, morphology appears affected after epiboly. Therefore, we employ two-step chopping in which we chop blastulae near the animal pole, to avoid cutting off the organizer, and wound the yolk membrane, to make the yolk size proportional to the blastula.
In addition to two-step chopping, we found the medium in which the size reduction surgery is performed is critical for recovery of embryos following the surgery. Among several media we tried (Egg water, Egg water + Albumin, Danieau buffer, L15, L15 + FBS, &#8531;x Ringer, 1x Ringer), only &#8531;x Ringer and 1x Ringer yielded high survival rates; in other media, embryos failed to recover from the wounds.
An important troubleshooting tip for low survival rate is to use healthy embryos from healthy and young parental fish. We noted that even when the control non-size reduced embryos show almost 100% survival rate, when size reduced, embryos from older fish tend to show lower survival rate. Also, note that the survival rate tends to decrease when the size reduction is combined with additional perturbations, such as morpholino injection.
The simplicity of the size reduction technique described here allows researchers to apply this technique without specialized equipment or intensive training. Further, since the size reduced embryos remain smaller until later stages of development (once they start eating, their size seems to catch up with the control fish), this technique can be applied to study scaling of many tissues and organs. Therefore, this technique makes it possible to combine size reduction and quantitative in vivo live imaging to study scaling and size control of various systems.

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 compared to the control (dechorionation only) embryos, other than size difference, throughout the developmental stages (see top and middle panels in Figure 2). On the other hand, blastula-only chopped embryos showed a peculiar morphology, especially at earlier stages. During epiboly, the embryos had a constricted and indented appearance (See bottom panel for 70% epiboly in Figure 2). At the following somite stage, midline structures were found to be flattened (i.e. DV length is relatively shorter than ML length) at many axial levels (See bottom panels for 8 and 20 somite in Figure 2). At later stages, the body structures adjacent to the yolk, such as the mid- and hindbrain, and the first ~10 somites, still showed a relatively flattened shape, possibly due to increased tension from the relatively larger yolk.
Somite size reduction in size reduced embryos 
Somites are segmental structures that appear transiently during embryogenesis and give rise to vertebrae and skeletal muscle. From presomitic mesoderm (PSM), somites are formed one by one from the anterior to posterior direction in a periodic manner (e.g. 25 min for zebrafish, 2 h for mice) (Figure 3A). We performed time lapse imaging of somite formation both for control and chopped embryos and measured the size of most newly formed somites (Figure 3B). In both control and chopped embryos, the sizes of somites that were formed at later stages were found to be smaller compared to the ones from earlier stages. Also, throughout the somite formation stages, chopped embryos had smaller somites than the ones in control embryos (Figure 3C).
Neural tube heights are reduced following size reduction 
To see the effect of embryo size reduction on neural tube size, we performed our two-step chopping technique on mem-mCherry injected embryos and imaged their spinal cords at 20 hpf using our confocal imaging system (Figure 4A,B). In this dataset, neural tube heights were reduced following size reduction by 12.4% &#177; 3.2%, as measured manually using custom image analysis code (Figure 4C). Taken together, these data show that size reduction reduces neural tube height. This technique can be used to measure the effects of size reduction on neural patterning.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59434/59434fig1.jpg" /> 
Figure 1: Size reduction technique. Approximately 30%-40% of the cells were cut from the animal pole (top panels). The membrane surrounding the yolk was carefully wounded so that the yolk oozed out (middle panels). For the following few minutes, yolk oozed out and then the wounds on both blastoderm and yolk healed up (bottom panels). Scale bar = 200 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59434/59434fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59434/59434fig2.jpg" /> 
Figure 2: Comparison between two methods of size reduction. Control embryos (top panels, top embryos for 24 hpf and 30 hpf), size reduced embryos with two-step chopping (blastula and yolk, middle panels, middle embryos for 24 hpf and 30 hpf) and size reduced embryos with blastula-only chopping (bottom panels, bottom embryos for 24 hpf and 30 hpf) are compared along developmental stages. Note that in blastula-only chopped embryos, the blastoderm volume is much smaller compared to the yolk (at 70% epiboly). As a result, the embryo has a disproportionately flattened shape at somite stages (i.e., DV axis is relatively shorter compared to AP axis in blastula-only chopped embryos, compared to the control or a two-step chopped one). Scale ba = 200 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59434/59434fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59434/59434fig3.jpg" /> 
Figure 3: Size reduction reduces the length of somites. (A) Schematic illustration of somite formation. (B) Bright field images of control and chopped embryos over time. Yellow arrowheads indicate the most newly formed somite at each somite stage. (C) Somite length (in anterior-posterior axis) measurements over time for both control and chopped embryos. Error bars represent standard deviation. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59434/59434fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59434/59434fig4.jpg" /> 
Figure 4: Size reduction reduces the height of the neural tube. (A-B) Example images of normal sized (A) and size reduced (B) tg(ptch2:kaede) embryos which were injected at the single cell stage with mem-mCherry mRNA. Scale bar = 20 &#956;m. (C) Neural tube heights extracted from manual segmentation of the neural tube in each z-stack. Statistically significant differences are observed in average neural height when values are compared using an unpaired t-test (p = 0.0397). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59434/59434fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Supplemental Movie 1" src="/files/ftp_upload/59434/59434movie1.jpg" /> 
Supplemental Movie 1: Comparison between two-step chopping versus blastula-only chopping. Top row = control embryos, middle row = size reduced embryos with two step chopping, bottom row = size reduced embryos with blastula only chopping. Movies were taken every 3 min for 12 h. Scale bar = 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59434/Supplemental_movie.avi" target="_blank">Please click here to view this video. (Right-click to download.)</a>

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

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

6.7K Views.  Harvard Medical School. Hoewel schaalvergroting een van de universele en fundamentele kenmerken van ontwikkelingssystemen is, zijn de onderliggende mechanismen niet begrepen in veel organen en weefsels. De groottereductietechniek van zebravissen die in deze video wordt gepresenteerd, zal uw begrip van mechanismen die ten grondslag liggen aan dynamische schaalprocessen bevorderen. Voordelen van het werken met zebravissen zijn dat het gemakkelijk is om live beeldvorming, genetica en kwantitatieve analyse uit te voeren...

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

A 3-dimensional (3D)-printed Template for High Throughput Zebrafish Embryo Arraying

The zebrafish is a globally recognized fresh water organism frequently used in developmental biology, environmental toxicology, and human disease related research fields. Thanks to its unique features, including large fecundity, embryo translucency, rapid and simultaneous development, etc., zebrafish embryos are often used for large scale toxicity assessment of chemicals and drug/compound screening. A typical screening procedure involves adult zebrafish spawning, embryos selection, and arraying the embryos into multi-well plates. From there, embryos are subjected to exposure and the toxicity of chemical, or the effectiveness of the drugs/compounds can be evaluated relatively quickly based on phenotypic observations. Among these processes, embryos arraying is one of the most time-consuming and labor-intensive steps that limits the throughput level. In this protocol, we present an innovative approach that makes use of a 3D-printed arraying template coupled with vacuum manipulation to speed up this laborious step. The protocol herein describes the overall design of the arraying template, a detailed experimental setup and step-by-step procedure, followed by representative results. When implemented, this approach should prove beneficial in a variety of research applications using zebrafish embryos as testing subjects.

A 3-dimensional 3D-printed Template for High Throughput Zebrafish Embryo Arraying

Here, we present a protocol to design and fabricate a zebrafish embryo arraying template, followed by a detailed procedure on the use of such template for high throughput zebrafish embryo arraying into a 96-well plate.

The zebrafish is a globally recognized fresh water organism frequently used in developmental biology, environmental toxicology, and human disease related ...

Environment

Dissection of Larval Zebrafish Gonadal Tissue

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex determination system, where several genes distributed throughout the genome collectively determine the sex identities of individual fish. Currently, the genes involved in regulating gonad development and how they work remain elusive. Normally, isolating gonadal tissue is the first step to examine the sex developmental processes. Here, we present a procedure to isolate gonadal tissue from 17 dpf (days post fertilization) and 25 dpf zebrafish larvae. The isolated gonadal tissue may be subsequently examined by morphology and gene expression profiling.

Here, we present a protocol for isolating gonadal tissue of larval zebrafish, which will facilitate investigations of zebrafish sex differentiation and maintenance.

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex ...

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 ...

Real Time and Repeated Measurement of Skeletal Muscle Growth in Individual Live Zebrafish Subjected to Altered Electrical Activity

A number of methods can be used to visualize individual cells throughout the body of live embryonic, larval or juvenile zebrafish. We show that live fish with fluorescently-marked plasma membranes can be scanned in a confocal laser scanning microscope in order to determine the volume of muscle tissue and the number of muscle fibers present. Efficient approaches for the measurement of cell number and size in live animals over time are described and validated against more arduous segmentation methods. Methods are described that permit the control of muscle electrical, and thus contractile, activity. Loss of skeletal muscle contractile activity greatly reduced muscle growth. In larvae, a protocol is described that allows reintroduction of patterned electrical-evoked contractile activity. The described methods minimize the effect of inter-individual variability and will permit analysis of the effect of electrical, genetic, drug, or environmental stimuli on a variety of cellular and physiological growth parameters in the context of the living organism. Long-term follow-up of the measured effects of a defined early-life intervention on individuals can subsequently be performed.

Optical clarity is a major advantage for cell biological and physiological work in zebrafish. Robust methods for measurement of cell growth in individual animals are described that permit novel insights into how growth of skeletal muscle and neighboring tissues are integrated with whole body growth.

A number of methods can be used to visualize individual cells throughout the body of live embryonic, larval or juvenile zebrafish. We show that live fish ...

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 ...

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

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