This work was supported by NIH grant DP2OD008470.&#160;

The procedures for working with zebrafish embryos described in this protocol were approved by the Institutional Animal Care and Use Committee at the University of Notre Dame.
1. Ringer&#8217;s Solution Preparation
<ol>
	<li>Make a solution of 116 mM NaCl, 2.9 mM KCl, 3.6 mM CaCl2, and 5 mM Hepes with ultrapure H2O through continual stirring to avoid any salt precipitation.</li>
	<li>After adding salts and while the solution is still stirring, add 100 units/penicillin per mL and 100 &#181;g streptomycin per ml.</li>
	<li>After adding all antibiotics and salts, perform filter sterilization of the solution and store in the sterile container. 
	Note: N-Phenylthiourea (1-Phenyl-2-thiourea) or PTU can be added to Ringer&#8217;s Solution to block the formation of pigment in bead implanted embryos without negative consequences to embryo health. To prepare Ringer&#8217;s/pigmentation blocking solution, use a concentration of 0.003 % PTU. Due to the low concentration of PTU to be added it is advised that a larger volume of Ringer&#8217;s Solution be made, at least 1 L, to avoid adding miniscule amounts of PTU powder. Add PTU during step 1.2, while the solution is stirring.</li>
</ol>
2. Tricaine Solution Preparation
<ol>
	<li>Add 2 g of Ethyl 3-aminobenzoate methanesulfonate salt (Tricaine) per L of solution plus 0.1 M Tris, from a 1 M stock balanced to a pH of 9.5 and made with ultrapure H2O. 
	Note: Tricaine will be used to euthanize the zebrafish embryos at the desired time point to assay bead implantation phenotypes.</li>
</ol>
3. E3 Solution Preparation
<ol>
	<li>Make a solution of 0.25 M NaCl, 10 mM KCl, 12.5 mM CaCl2,and 16.6 mM MgSO4 to 1 L of ultrapure water to create 1 L of E3.</li>
	<li>Add 200 &#181;l of Methylene Blue to the E3 solution to inhibit fungal growth.</li>
</ol>
4. Pulling Needles for Microbead Transfer
<ol>
	<li>Use fire polished borosilicate glass with filament at an O.D.: 1.0 mm, I.D.: 0.5 mm, at a length of 10 cm.</li>
	<li>Place borosilicated glass into a micropipette puller to prepare fine needles. 
	Note: The following four dimensions can be used as a starting point to pull the needles: Heat = 540, Pull = 245, Velocity = 200, and Time = 125.</li>
	<li>After pulling an adequate number of needles, store in covered plastic petri dishes, positioned on strips of modeling clay or adhesive tape to prevent breaking the needle tip, until use.</li>
	<li>To trim the needle tip to the desired bore size, use a pair of fine forceps to score the end of the needle. 
	Note: Needle bore size will vary depending on the microbead size being utilized as well as user handling preferences. A good starting point is to craft bore sizes in a range between 100 and 200 &#181;m if microbeads ranging from 50-100 &#181;m will be utilized.</li>
	<li>Prior to use, insert the trimmed needle into the capillary tube holder to fashion the completed microbead transfer instrument.</li>
</ol>
5. Whisker/Lash Tool Preparation
<ol>
	<li>Obtain a whisker, lash, or other small yet firm piece of natural or synthetic hair filament. Cut near the base of the filament at a 45&#176; angle. Adhere the whisker to a P-1000 micropipette tip using permanent clear adhesive glue.</li>
</ol>
6. Microbead Implantation Tray Preparation
<ol>
	<li>Mix 2 g of agarose with 100 ml of E3 solution to make a 2 % E3 agarose gel.</li>
	<li>Heat the 2 % agarose solution for 1 min and 30 sec in a 250 ml Erlenmeyer flask, swirling the flask every 30 sec in order to avoid the gel from bubbling over.</li>
	<li>Take the lid of a 60 mm x 15 mm petri dish and place it in a larger 150 mm x 15 mm dish with the interior facing upward.</li>
	<li>Pour the hot agarose gel into the 150 mm x 15 mm petri dish, making sure to anchor down the lid of the smaller dish with the index finger. 
	Note: Allow a thin layer of agarose below the lid of the smaller dish to remove condensation that will form when pouring the gel; this will make the beads easier to see under a dissection microscope.</li>
	<li>As the agarose cools, but before it solidifies, place a well mold on the gel. This will allow for easier placement of the embryos.</li>
	<li>Note: In order to avoid bubbles in the wells, place the mold slowly at an angle until it is floating on top of the gel.</li>
	<li>Using a microspatula, carefully remove the well mold once the gel has solidified. Add E3 solution and store at 4 &#176;C.</li>
</ol>
7. Embryo Collection
<ol>
	<li>Prepare mating chambers, separated by dividers, by filling them up with system water, and then placing adult zebrafish male(s) and female(s) pairs in the chambers O/N.</li>
	<li>Collect fertilized eggs using a fine wire mesh strainer (embryonic stages will vary slightly).</li>
	<li>Deposit the fertilized eggs in a clean, 10 cm diameter petri dish by turning the strainer upside down and rinsing it with a fine stream of E3 until there are no eggs left in the strainer, and there approximately 25 ml of E3 solution in the dish.</li>
	<li>After collection, divide the eggs into several dishes (approximately 50-60 eggs per dish) to avoid overcrowding, which might lead to developmental asynchrony within the clutch and make collecting desired times points difficult.</li>
	<li>Incubate embryos in E3 solution at 28.5 &#176;C to enable developmental assessment according to the standard zebrafish staging series 40. 
	Note: The embryos should be transferred to E3/PTU at 24 hr post fertilization (hpf) to prevent pigment development if manipulations at older stages necessitate optical clarity.</li>
</ol>
8. Bead Implantation
<ol>
	<li>Incubate the microbeads in desired test molecule concentration or the corresponding vehicle solution for the required time in an appropriate volume. 
	Note: The researcher can use sterile Petri dishes ranging from 3-6 cm in diameter, or multi-well plates, both of which enable visualization of the microbeads within the solution. Time may vary depending on the type and amount of molecule of interest, and the researcher should refer to manufacturer or published recommendations for duration of incubation and other aspects such as sensitivity to light and temperature.</li>
	<li>Once the embryos have reached the desired developmental time point, remove the embryos from their chorions using fine forceps.</li>
	<li>Incubate the embryos in the filtered Ringer solution with antibiotics for 10 min to acclimate them to the solution.</li>
	<li>While the embryos are acclimating, transfer the microbeads into the microbead plate nestled within the implantation tray and rinse them in Ringer solution for 10 min. 
	Note: Implant the beads within 50 min after reaching this point. This will decrease the likelihood that one will implant a bead with a smaller concentration of the molecule of interest.</li>
	<li>Array the embryos into the wells of the microbead implantation tray, taking care to orient them so the area of interest where the microbead will be implanted is accessible.</li>
	<li>Make a small incision on the embryo using the tungsten needle.</li>
	<li>Transfer a microbead onto the embryo using the microcapillary transfer pipette: depressing the bulb of the pipette, gently pull the microbead into the capillary column, and then release the pressure to transfer the microbead at the desired destination.</li>
	<li>Use the whisker/lash tool to position the microbead inside the embryo. Note: Be sure to position the microbead into the tissue away from the site of incision so the microbead will not be expelled from the embryo as it heals.</li>
</ol>

The authors have nothing to disclose.

Over the past century, the understanding of body plan patterning and organogenesis has undergone monumental advances. Tissue manipulation techniques were critical in uncovering key information about these vital processes. Genetic modification is one of the most commonly used methods for ascertaining gene function, and methods of mosaic analysis, such as cell transplantation, provide useful approaches to understand autonomy of gene function. Microbead implantation provides another venue to interrogate how particular molecules alter developmental dynamics, as this method enables the researcher to alter a local tissue environment by introducing signaling molecules or inhibitors. A range of different microbeads are commercially available, that have varying sizes and other physical properties exist (e.g., charge) such that they can be employed to for the experimental conditions of interest. Thus, by implanting microbeads that are soaked in a protein or chemical of interest within an organism, researchers can investigate localized effects during development and find associations between the gene or molecule of interest and particular biological phenotype(s).
Studies such as the one performed by Wada and colleagues used microbead implantation in order to assess the effect of increased Hedgehog signaling in skeletal patterning in the anterior neurocranium (ANC) in zebrafish 23. Previous studies have shown that Shh is required for ANC formation 14. Using Shh-coated microbeads researchers identified that this signal promotes cartilage formation in the ANC. The microbead implantation process was used to demonstrate a clear link between Hedgehog signaling and cartilage formation in the ANC. Another key example of this tissue manipulation technique in zebrafish is observed in studies where scientists investigated the transcriptional control of the Erythroblastoma Twenty-Six (ETS) domain factors ETS-related molecule (Erm) and Polyoma enhancer activator 3 (Pea3) by Fgf signaling during early zebrafish brain development 19. Through microbead implantation experiments, they were able to show that Fgf8 and Fgf3 can ectopically activate expression of erm and pea3. These examples illustrate the utility of microbeads to provide insights into the developmental mechanisms that collaborate during tissue formation, which can be well-characterized by the use of methods to assess gene expression 41. Thus, microbead transplantation can be a viable method to explore other tissues, such as the intermediate mesoderm (IM) which gives rise to the kidney. Specifically, it would aid in renal developmental studies, to investigate how different molecules affect nephron segmentation 42 and tubulogenesis 43,44, processes that are only superficially understood at present. Furthermore, microbead implantation has begun to be used to study regeneration processes in zebrafish 29] and could be adapted for use with any number of organ regeneration models, such as following laser ablation of embryonic tissues like the nephron 45 or in conjunction with methods that have been formulated to perform research with the corresponding adult structures 46-49. Finally, microbead implantation has the potential to be used in models of human disease, such as cancer 50,51 or tissue degeneration 52,53.
In the present protocol, we demonstrate the method of microbead implantation in zebrafish embryos, which has also been similarly described by other researchers, but not shown through video protocols 1. With minimal practice we were able to implant microbeads at an approximate rate of 8-10 embryos/hr, which relates the feasibility of this procedure once the researcher has some experience. The results shown herein demonstrate that beads of various dimensions can be implanted at early stages, and that with care, this tissue manipulation technique can be implemented with minimal physical disruption to the embryo. One improvement that should be emphasized is the use of a whisker/lash tool to position the microbead into the embryo. This relatively inexpensive and simple piece of equipment is about the same diameter as the microbead, is easy to obtain and helps speed up the implantation process. The whisker/lash can be cut to a desired length to produce a firm yet delicate bead-handling tool, depending on researcher dexterity and preference. Finally, while we described here how to physically manipulate microbeads and zebrafish embryos to perform implantation, this protocol does not outline specific handling procedures for different drugs or peptides. In general, chemically treated microbeads should be implanted into the animal with expediency, to avoid undesired effects in other areas of the organism, and researchers should be well informed about the possible safety concerns associated with any such chemicals before initiating their studies.
In sum, we have demonstrated a relatively simple and efficient microbead implantation method with a wide range of applications using materials that are readily available in the lab. Ultimately, we hope that this manual will aid researchers with the delicate nature of zebrafish tissue manipulation.

Developmental biology researchers utilize a vast array of cellular, molecular, and genetic methods to uncover the mechanisms that control how an organism is formed. Among these approaches, tissue manipulation is a key tool in deciphering complex questions about cell fate, cellular movement, and the organization of tissues. One way to alter local tissue environments is through the surgical application of microbeads that are used to deliver a focal source of proteins or other signaling molecules 1. This type of experimental manipulation has been widely implemented in classical vertebrate embryology models, such as the frog and chick 2.
The zebrafish has become an important vertebrate model organism for the study of organogenesis and also provides many unique advantages for disease modeling 3-5 as they share high genetic conservation with humans 6. In particular, the optical transparency and external development of the zebrafish embryo offers an unmatched viewpoint for the observation of tissue ontogeny 3-5. The implementation of large-scale forward genetic screens has generated a powerful repository of zebrafish mutant strains for further study 7,8, and the identification of alternative screening techniques that can be efficiently conducted at reduced scale in single laboratories 9,10. Further experimental work with zebrafish has been facilitated through advances in transgenic methodologies and reverse genetic approaches 11,12, as well as chemical genetics 13-15.
Tissue manipulation techniques, such as the implementation of microbeads,&#160;have not been as widely employed in the zebrafish, but nevertheless provide a useful tool to further understand cell signaling during development. Microbead implantation has been used to interrogate the processes of organ formation in the zebrafish retina 16,17, heart 18, brain 19-22, neural crest 23, and fin 24,25. In these and other studies, beads have been applied during development to understand the diffusion of signaling molecules 26, how gradients affect cell migration 27 and axial patterning 28. More recently, microbeads have been utilized to evaluate regeneration mechanisms in zebrafish adults 29. In developmental studies, for example, zebrafish microbead work has provided insights into the mechanisms of limb formation through studies of the pectoral fin 25. The zebrafish pectoral fin bud is homologous to the forelimb bud in the mouse 30 and chick 31. The vertebrate limb bud has two essential signaling nodes: the zone of polarizing activity (ZPA) that establishes the anterior-to-posterior axis through the expression of Sonic hedgehog (Shh) and downstream Hox gene targets, and the apical ectodermal ridge (AER) present at the tip of the limb bud, which acts to establish proximal to distal identity of the limb through expression of Fibroblast growth factors (Fgfs). By implanting Fgf soaked microbeads into zebrafish Shh genetic mutants, investigators identified Fgf as essential to progression of the cell cycle and growth of the vertebrate limb 25. In addition to the Fgf and Shh signaling cascades that establish positional identity, pioneering studies using the chick limb bud identified retinoic acid (RA) as a molecule that could mimic the action of the polarizing region &#160;to establish anterior to posterior identity 32. These experiments involved placing small strips of RA-soaked Whatman paper into the chick limb to assess digit patterning 32. Further, researchers have performed other elegant studies employing the use of microbeads, cell transplantation, and exogenous RA treatments in zebrafish to determine that RA acts to provide long-range positional cues within the zebrafish hindbrain and mesoderm 28. However, at present many questions remain about the roles of signaling factors like Fgf and RA during numerous aspects of vertebrate development. The signaling effects of RA, acting as a morphogen, impact many organs 33, such as the developing heart 34 and the renal progenitors, where RA specifies proximal kidney cells type fates 35-39. Further understanding of such topics&#160;could benefit significantly from experimental studies using tissue manipulation and microbead implantation techniques.
While fewer studies have been performed with microbead implantation in the zebrafish, when compared to models like the chick, those that have been carried out have been highly informative. One reason for the paucity of microbead implantation based-research in the zebrafish embryo is likely the notion that there are difficult technical challenges, based on the size of the embryo, which constitute an impediment to successfully performing such manipulations. However, microbead implantation in zebrafish embryos can be learned with practice and assisted through visual observation of the technique, and thus can be pursued as a means to interrogate the mechanisms of development. Here, we demonstrate the precise application of a microbead into the zebrafish embryo, which can be utilized for conducting a wide variety of assays on tissue formation and cellular morphogenesis.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Sodium Chloride</td>
			<td>American Bioanalytical</td>
			<td>AB01915</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Chloride</td>
			<td>American Bioanalytical</td>
			<td>AB01652</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium Chloride</td>
			<td>American Bioanalytical</td>
			<td>AB00366</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma Life Science</td>
			<td>H4034</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin solution</td>
			<td>Sigma-Aldrich</td>
			<td>P4333-20ML</td>
			<td></td>
		</tr>
		<tr>
			<td>N-Phenylthiourea (PTU)</td>
			<td>Aldrich Chemistry</td>
			<td>P7629</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethyl 3-aminobenzoate (Tricane)</td>
			<td>Fluka Analytical</td>
			<td>A5040</td>
			<td></td>
		</tr>
		<tr>
			<td>Borosilicate glass</td>
			<td>Sutter Instruments Co.</td>
			<td>BF100-50-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Flaming/Brown Micropipette puller</td>
			<td>Sutter Instruments Co.</td>
			<td>Mo. P097</td>
			<td></td>
		</tr>
		<tr>
			<td>UltraPure Agarose</td>
			<td>Invitrogen</td>
			<td>15510-027</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium Sulfate</td>
			<td>Sigma-Aldrich</td>
			<td>M7506</td>
			<td></td>
		</tr>
		<tr>
			<td>Methylene Blue</td>
			<td>Sigma-Aldrich</td>
			<td>M9140</td>
			<td></td>
		</tr>
		<tr>
			<td>250mL Erlenmeyer Flask</td>
			<td>Fischer Scientific</td>
			<td>FB-500-250</td>
			<td></td>
		</tr>
		<tr>
			<td>Falcon Diposable Petri Dishes, Sterile</td>
			<td>Corning</td>
			<td>430167</td>
			<td></td>
		</tr>
		<tr>
			<td>60mm x 15mm</td>
			<td>VWR</td>
			<td>25373-085</td>
			<td></td>
		</tr>
		<tr>
			<td>100mm x 15mm</td>
			<td>VWR</td>
			<td>25373-100</td>
			<td></td>
		</tr>
		<tr>
			<td>150mm x 15mm</td>
			<td>VWR</td>
			<td>25373-187</td>
			<td></td>
		</tr>
		<tr>
			<td>Saint-Gobain Chemware Microspatula</td>
			<td>Fischer Scientific</td>
			<td>21-401-50B</td>
			<td></td>
		</tr>
		<tr>
			<td>P-1000 Micropipette tips</td>
			<td>Fischer Scientific</td>
			<td>2707402</td>
			<td></td>
		</tr>
		<tr>
			<td>Low Temperature Incubator</td>
			<td>Fischer Scientific</td>
			<td>11 690 516DQ</td>
			<td></td>
		</tr>
		<tr>
			<td>Dimethly Sulfoxide</td>
			<td>American Bioanalytical</td>
			<td>AB00435</td>
			<td></td>
		</tr>
		<tr>
			<td>Microbeads (45-106 &#181;m)</td>
			<td>Biorad</td>
			<td>140-1454 AG1-X8</td>
			<td></td>
		</tr>
		<tr>
			<td>Microbeads (45 &#181;m)</td>
			<td>Polysciences</td>
			<td>7314</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro Dissecting Tweezer</td>
			<td>Roboz Surgical Instruments Co.</td>
			<td>RS-5010</td>
			<td></td>
		</tr>
		<tr>
			<td>Tungsten Needle</td>
			<td>Roboz Surgical Instruments Co.</td>
			<td>RS-6065</td>
			<td></td>
		</tr>
		<tr>
			<td>Capillary tube holder</td>
			<td>Globe Scientific Inc.&#160;</td>
			<td>51674</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Several types of manipulation tools are useful for handling microbeads during research applications (Figure 1). In addition, a simple agarose mold with small wells can be utilized to hold the zebrafish embryo, which is vital to perform these experiments in a timely and efficient manner (Figure 1). The implantation of a microbead can be performed at the spatial position and temporal stage of interest, which allows researchers to narrow a specific window of action for the molecule of interest.
Here, single microbeads were implanted into zebrafish embryos at the tail bud stage or at later time points during somitogenesis stages (Figure 3). Two different sized microbeads, rinsed with Ringer&#8217;s solution alone, were used in this protocol demonstration (Figure 3A, 3B and 3C). This microbead implantation in the absence of any conjugated small molecules demonstrates that bead implantation can be achieved without gross morphological effects during the proper development of the zebrafish embryo (Figure 4).
The experimentalist may encounter difficulties with maneuvering the microbead into the puncture hole made by a tungsten needle atop the embryo. To achieve better handling of the microbead once placed on the embryo by the micro-needle, a whisker/lash tool can be used. This can be crafted with naturally shed feline or canine whiskers, although other natural or synthetic lash filaments can be utilized (Figure 1). A gentle push of the microbead with either the tungsten needle or the whisker tool should see the microbead sink into the Ringer&#8217;s solution of the mold. Once the microbead has sunk into the mold it should be moved large distances if necessary using the tungsten&#160;needle and once atop the embryo the needle or whisker tool can be used to position the microbead into the embryo. The success of the microbead implantation can be immediately gauged through visualization at a&#160;stereomicroscope, as the bead will remain stably positioned in the tissue. To evaluate bead position over time, each embryo sample can be imaged through live time-course photography or checked at periodic intervals (as depicted in Figure 4) to ensure that the bead location has not shifted over the duration of the experiment due to endogenous tissue morphogenesis. Understanding the position of the bead over time is critical for the most informed interpretation of results, such as evaluating the effect of a small molecule on a target tissue or developmental process.
The use of these above steps will provide an ideal environment in which to imbed microbeads into the zebrafish embryo at a desired time and position. In our experience, novice users of this microbead implantation protocol typically gained the skill necessary to perform consistent surgical implantation of microbeads into the zebrafish embryo after approximately&#160;a single week of practice.
<img alt="Figure 1" src="/files/ftp_upload/52943/52943fig1.jpg" /> 
Figure 1.&#160;Tools used to perform microbead implantation. (A) The bead implantation tray (left, 15 cm diameter) with embryo molds (on left) and a microbead incubation tray (on right, 6 cm diameter). This working tray enables the researcher to place the embryos into the desired orientation for microbead implantation, and to have the prepared microbeads for implantation in close proximity at the microscope station. Microbeads should be incubated in small molecule(s) and/or vehicle and then rinsed in a separate microbead plate (right) before placing them into the bead implantation tray that is positioned under the microscope station. (B) Two pairs of fine forceps will be used to remove the chorion surrounding each zebrafish embryo and also to break the end of the microcapillary needle. (C) A tungsten needle will be employed to make a very fine puncture into the zebrafish embryo in which to position the microbead. (D) The microcapillary transfer pipette will be used to pick up and position a microbead on top of the zebrafish embryo near the site of the puncture. (E) The whisker/lash tool will allow the gentle positioning of the microbead into the incision site within the embryo that was previously made by the tungsten needle. <a href="https://www.jove.com/files/ftp_upload/52943/52943fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52943/52943fig2.jpg" /> 
Figure 2.&#160;Schematic of the microbead implantation procedure. (A) A microbead implantation tray with microbeads soaking in the desired wash solution is set up under a stereomicroscope station, and the embryos positioned with tissue facing upwards. (B) Using a tungsten needle, a small incision into the embryo can be made with a slight yet forceful movement. (C) Using a microcapillary transfer pipette, the desired microbead can be picked up from the microbead incubation chamber in the tray, located on the right, and brought to the left compartment where the embryo working area is situated. The microbead should be deposited near the site of the incision, then gently positioned into the incision site made by the tungsten needle using the whisker/lash tool. Once a microbead is maneuvered&#160;into the incision site, it should be tucked into the tissue (away from the incision) to stably position the microbead and prevent subsequent movement out of the embryo as development continues. <a href="https://www.jove.com/files/ftp_upload/52943/52943fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52943/52943fig3.jpg" /> 
Figure 3.&#160;Microbeads can be implanted into embryos at different developmental stages. (A) Zebrafish embryos were implanted with a microbead (diameter of approximately 50 &#181;m) into the trunk at the tail bud stage. When examined at the 24 hr post bead implant (hpbi), the embryos had developed normally and the microbead showed minimal movement during the passage of developmental time. (B) Zebrafish embryos implanted with microbeads (with a diameter of approximately 50 &#181;m) into the yolk sac at the 3 somite stage (ss) displayed normal development with relatively minor bead movement over time. (C) Larger microbeads (diameter of approximately 70 &#181;m) were implanted into the 7 ss embryo similarly showed normal subsequent development, as well as minimal if any movement from the site of implantation. <a href="https://www.jove.com/files/ftp_upload/52943/52943fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/52943/52943fig4.jpg" /> 
Figure 4.&#160;Gross development of the zebrafish embryo is unperturbed by the presence of a microbead through 72 hpbi. Zebrafish embryos were implanted with microbeads soaked in Ringer&#8217;s solution (with a diameter of approximately 50 &#181;m) at the 7 somite stage (ss). Each microbead was surgically inserted into the trunk, between somites 3 and 4. The presence of the bead was not associated with overt developmental defects at 24 hr post bead implant (hpbi), 48 hpbi, or 72 hpbi. <a href="https://www.jove.com/files/ftp_upload/52943/52943fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Microbead Implantation in the Zebrafish Embryo at JoVE.com

Microbead Implantation in the Zebrafish Embryo

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 ...

microbead-implantation-in-the-zebrafish-embryo

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Developmental Biology

11.1K ビュー.  University of Notre Dame. 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.

ビデオ: Microbead Implantation in the Zebrafish Embryo

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 ...

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin condensation, microtubule-kinetochore attachment, chromosome segregation, and cytokinesis in a small time frame. Errors in the delicate process can result in human disease, including birth defects and cancer. Traditional approaches investigating human mitotic disease states often rely on cell culture systems, which lack the natural physiology and developmental/tissue-specific context advantageous when studying human disease. This protocol overcomes many obstacles by providing a way to visualize, with high resolution, chromosome dynamics in a vertebrate system, the zebrafish. This protocol will detail an approach that can be used to obtain dynamic images of dividing cells, which include: in vitro transcription, zebrafish breeding/collecting, embryo embedding, and time-lapse imaging. Optimization and modifications of this protocol are also explored. Using H2A.F/Z-EGFP (labels chromatin) and mCherry-CAAX (labels cell membrane) mRNA-injected embryos, mitosis in AB wild-type, auroraBhi1045, and esco2hi2865 mutant zebrafish is visualized. High resolution live imaging in zebrafish allows one to observe multiple mitoses to statistically quantify mitotic defects and timing of mitotic progression. In addition, observation of qualitative aspects that define improper mitotic processes (i.e., congression defects, missegregation of chromosomes, etc.) and improper chromosomal outcomes (i.e., aneuploidy, polyploidy, micronuclei, etc.) are observed. This assay can be applied to the observation of tissue differentiation/development and is amenable to the use of mutant zebrafish and pharmacological agents. Visualization of how defects in mitosis lead to cancer and developmental disorders will greatly enhance understanding of the pathogenesis of disease.

Mitosis is critical to every living organism and defects often lead to cancer and developmental disorders. Using this imaging protocol and zebrafish as a model system, researchers can visualize mitosis in a live vertebrate organism and the multitude of defects that arise when mitotic processes are defective.

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin ...

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance to man of conventional invertebrate model systems and the complexity of genomes in vertebrates are critical challenges to modeling human normal and pathological conditions. The chordate Ciona intestinalis is an invertebrate chordate that emerged from a common ancestor with the vertebrates and may represent features at the interface between invertebrates and vertebrates. A common body plan with much simpler cellular and genomic composition should unveil gene regulatory network (GRN) links and functional genomics readouts explaining phenomena in the vertebrate condition. The compact genome of Ciona, a fixed embryonic lineage with few divisions and large cells, combined with versatile community tools foster efficient gene functional analyses in this organism. Here, we present several crucial methods for this promising model organism, which belongs to the closest sister group to vertebrates. We present protocols for transient transgenesis by electroporation, along with microinjection-mediated gene knockdown, which together provide the means to study gene function and genomic regulatory elements. We extend our protocols to provide information on how community databases are utilized for in silico design of gene regulatory or gene functional experiments. An example study demonstrates how novel information can be gained on the interplay, and its quantification, of selected neural factors conserved between Ciona and man. Furthermore, we show examples of differential subcellular localization in embryonic cells, following DNA electroporation in Ciona zygotes. Finally, we discuss the potential of these protocols to be adapted for tissue specific gene interference with emerging gene editing methods. The in vivo approaches in Ciona overcome major shortcomings of classical model organisms in the quest of unraveling conserved mechanisms in the chordate developmental program, relevant to stem cell research, drug discovery, and subsequent clinical application.

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

We present transient transgenesis and gene knockdown in Ciona intestinalis, a chordate sister group to vertebrates, using microinjection and electroporation techniques. Such methods facilitate functional genomics in this simple invertebrate that features rudimentary characteristics of vertebrates, including notochord and head sensory epithelia, and many orthologs of human disease associated genes.

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance ...

Use of Immunolabeling to Analyze Stable, Dynamic, and Nascent Microtubules in the Zebrafish Embryo

Microtubules (MTs) are dynamic and fragile structures that are challenging to image in vivo, particularly in vertebrate embryos. Immunolabeling methods are described here to analyze distinct populations of MTs in the developing neural tube of the zebrafish embryo. While the focus is on neural tissue, this methodology is broadly applicable to other tissues. The procedures are optimized for early to mid-somitogenesis-stage embryos (1 somite to 12 somites), however they can be adapted to a range of other stages with relatively minor adjustments. The first protocol provides a method to assess the spatial distribution of stable and dynamic MTs and perform a quantitative analysis of these populations with image-processing software. This approach complements existing tools to image microtubule dynamics and distribution in real-time, using transgenic lines or transient expression of tagged constructs. Indeed, such tools are very useful, however they do not readily distinguish between dynamic and stable MTs. The ability to image and analyze these distinct microtubule populations has important implications for understanding mechanisms underlying cell polarization and morphogenesis. The second protocol outlines a technique to analyze nascent MTs specifically. This is accomplished by capturing the de novo growth properties of MTs over time, following microtubule depolymerization with the drug nocodazole and a recovery period after drug washout. This technique has not yet been applied to the study of MTs in zebrafish embryos, but is a valuable assay for investigating the in vivo function of proteins implicated in microtubule assembly.

Immunolabeling methods to analyze distinct populations of microtubules in the developing zebrafish brain are described here, which are broadly applicable to other tissues. The first protocol outlines an optimized method for immunolabeling stable and dynamic microtubules. The second protocol provides a method to image and quantify nascent microtubules specifically.

Microtubules (MTs) are dynamic and fragile structures that are challenging to image in vivo, particularly in vertebrate embryos. Immunolabeling methods ...

AFM and Microrheology in the Zebrafish Embryo Yolk Cell

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. Various propositions claim to have been important contributions to the understanding of the mechanical properties of cells and tissues in their spatiotemporal organization in different developmental and morphogenetic processes. However, due to the lack of reliable and accessible tools to measure material properties and tensional parameters in vivo, validating these hypotheses has been difficult. Here we present methods employing atomic force microscopy (AFM) and particle tracking with the aim of quantifying the mechanical properties of the intact zebrafish embryo yolk cell during epiboly. Epiboly is an early conserved developmental process whose study is facilitated by the transparency of the embryo. These methods are simple to implement, reliable, and widely applicable since they overcome intrusive interventions that could affect tissue mechanics. A simple strategy was applied for the mounting of specimens, AFM recording, and nanoparticle injections and tracking. This approach makes these methods easily adaptable to other developmental times or organisms.

The lack of tools to measure material properties and tensional parameters in vivo prevents validating their roles during development. We employed atomic force microscopy (AFM) and nanoparticle-tracking to quantify mechanical features on the intact zebrafish embryo yolk cell during epiboly. These methods are reliable and widely applicable avoiding intrusive interventions.

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. ...

Effectiveness of the Air Stripping in Two Salmonid Fish, Rainbow Trout (Oncorhynchus Mykiss) and Brown Trout (Salmo Trutta Morpha fario)

Egg collection is one of the most crucial procedures during fish reproduction in salmonid hatcheries. Classic methods involve the use of hand massage on fish abdomens to expel the eggs. An alternative method uses the pressure of gas injected into the body cavity, which causes the subsequent release of the eggs. This method is believed to have less negative effects on both the welfare and egg quality of the broodstocks. Herein, we compare the results of air and hand stripping methods with respect to one-year survival and egg quantity and quality in two salmonid fish, rainbow trout (Oncorhynchus mykiss) and brown trout (Salmo trutta morpha fario). Our results indicate that air stripping yielded a better quality of eggs and higher one-year survival rate in rainbow trout. In addition, air stripping resulted in lower mortality rate than the group subjected to hand stripping (25% vs. 35%). The pH and hatching rate of the hand stripped group was lower than those of the air stripped group. In the case of brown trout, the quality of eggs obtained by both hand and air-stripping methods was similar; however, the one-year losses in fish were higher in air stripped group (15% compared to 0% in hand stripped fish). Although the advantages of air stripping method over hand stripping in terms of egg quality might not be observed in all salmonid species, the air-stripping procedure might be a promising option to be adopted in hatcheries as it ensures a high level of reproducibility and efficiency.

Effectiveness of the Air Stripping in Two Salmonid Fish, Rainbow Trout Oncorhynchus Mykiss and Brown Trout Salmo Trutta Morpha fario

The principal aim of this study is to standardize and test the pneumatic method (air stripping) of collecting eggs in rainbow trout and brown trout. This method allows effective and simple collection of the eggs without the necessity of fish abdomen massage.

Egg collection is one of the most crucial procedures during fish reproduction in salmonid hatcheries. Classic methods involve the use of hand massage on ...

Following Endocardial Tissue Movements via Cell Photoconversion in the Zebrafish Embryo

During embryogenesis, cells undergo dynamic changes in cell behavior, and deciphering the cellular logic behind these changes is a fundamental goal in the field of developmental biology. The discovery and development of photoconvertible proteins have greatly aided our understanding of these dynamic changes by providing a method to optically highlight cells and tissues. However, while photoconversion, time-lapse microscopy, and subsequent image analysis have proven to be very successful in uncovering cellular dynamics in organs such as the brain or the eye, this approach is generally not used in the developing heart due to challenges posed by the rapid movement of the heart during the cardiac cycle. This protocol consists of two parts. The first part describes a method for photoconverting and subsequently tracking endocardial cells (EdCs) during zebrafish atrioventricular canal (AVC) and atrioventricular heart valve development. The method involves temporally stopping the heart with a drug in order for accurate photoconversion to take place. Hearts are allowed to resume beating upon removal of the drug and embryonic development continues normally until the heart is stopped again for high-resolution imaging of photoconverted EdCs at a later developmental time point. The second part of the protocol describes an image analysis method to quantify the length of a photoconverted or non-photoconverted region in the AVC in young embryos by mapping the fluorescent signal from the three-dimensional structure onto a two-dimensional map. Together, the two parts of the protocol allows one to examine the origin and behavior of cells that make up the zebrafish AVC and atrioventricular heart valve, and can potentially be applied for studying mutants, morphants, or embryos that have been treated with reagents that disrupt AVC and/or valve development.

This protocol describes a method for the photoconversion of Kaede fluorescent protein in endocardial cells of the living zebrafish embryo that enables the tracking of endocardial cells during atrioventricular canal and atrioventricular heart valve development.

During embryogenesis, cells undergo dynamic changes in cell behavior, and deciphering the cellular logic behind these changes is a fundamental goal in the ...

Minimally Invasive Embryo Transfer and Embryo Vitrification at the Optimal Embryo Stage in Rabbit Model

Assisted reproductive techniques (ARTs), such as in vitro embryo culture or embryo cryopreservation, affect natural development patterns with perinatal and postnatal consequences. To ensure the innocuousness of ART applications, studies in animal models are necessary. In addition, as a last step, embryo development studies require evaluation of their capacity to develop full-term healthy offspring. Here, embryo transfer to the uterus is indispensable to perform any ARTs-related experiment.
The rabbit has been used as a model organism to study mammalian reproduction for over a century. In addition to its phylogenetic proximity to the human species and its small size and low maintenance cost, it has important reproductive characteristics such as induced ovulation, a chronology of early embryonic development similar to humans and a short gestation that allow us to study the consequences of ART application easily. Moreover, ARTs (such as intracytoplasmic sperm injection, embryo culture, or cryopreservation) are applied with suitable efficiency in this species.
Using the laparoscopic embryo transfer technique and the cryopreservation protocol presented in this article, we describe 1) how to transfer embryos through an easy, minimally invasive technique and 2) an effective protocol for long-term storage of rabbit embryos to provide time-flexible logistical capacities and the ability to transport the sample. The outcomes obtained after transferring rabbit embryos at different developmental stages indicate that morula is the ideal stage for rabbit embryo recovery and transfer. Thus, an oviductal embryo transfer is required, justifying the surgical procedure. Furthermore, rabbit morulae are successfully vitrified and laparoscopically transferred, proving the effectiveness of the described techniques.

Assisted reproductive techniques (ARTs) are in continuous evaluation to improve outcomes and reduce the associated risks. This manuscript describes a minimally invasive embryo transfer procedure with an efficient cryopreservation protocol that allows the use of rabbits as an ideal animal model of human reproduction.

Assisted reproductive techniques (ARTs), such as in vitro embryo culture or embryo cryopreservation, affect natural development patterns with perinatal ...

Microinjection-based System for In Vivo Implantation of Embryonic Cardiomyocytes in the Avian Embryo

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a general challenge in developmental biology research. In the embryonic heart, this can be particularly difficult as regional differences in the expression of transcriptional regulators, paracrine/juxtacrine signaling cues, and hemodynamic force are all known to influence cardiomyocyte maturation. A simplified method to alter a developing cardiomyocyte's molecular and biomechanical microenvironment would, therefore, serve as a powerful technique to examine how local conditions influence cell fate and function. To address this, we have optimized a method to physically transplant juvenile cardiomyocytes into ectopic locations in the heart or the surrounding embryonic tissue. This allows us to examine how microenvironmental conditions influence cardiomyocyte fate transitions at single cell resolution within the intact embryo. Here, we describe a protocol in which embryonic myocytes can be isolated from a variety of cardiac sub-domains, dissociated, fluorescently labeled, and microinjected into host embryos with high precision. Cells can then be directly analyzed in situ using a variety of imaging and histological techniques. This protocol is a powerful alternative to traditional grafting experiments that can be prohibitively difficult in a moving tissue such as the heart. The general outline of this method can also be adapted to a variety of donor tissues and host environments, and its ease of use, low cost, and speed make it a potentially useful application for a variety of developmental studies.

In this method, embryonic cardiac tissues are surgically microdissected, dissociated, fluorescently labeled, and implanted into host embryonic tissues. This provides a platform for studying the individual or tissue level developmental organization under ectopic hemodynamic conditions, and/or altered paracrine/juxtacrine environments.

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a ...