Name: microbead implantation in the zebrafish embryo
Uploaded: 2015-07-30T14:00:00.000+00:00
Duration: 5 min 54 s
Description: Watch this Scientific Journal Video about Microbead Implantation in the Zebrafish Embryo at JoVE.com

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><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 (Tricaine)</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>250 ml 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>60 mm x 15 mm</td><td>VWR</td><td>25373-085</td><td></td></tr><tr><td>100 mm x 15 mm</td><td>VWR</td><td>25373-100</td><td></td></tr><tr><td>150 mm x 15 mm</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 &amp;micro;m)</td><td>Biorad</td><td>140-1454 AG1-X8</td><td></td></tr><tr><td>Microbeads (45 &amp;micro;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.&amp;nbsp;</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

Research

JoVE Journal

Developmental Biology

11.2K Views.  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.

Video: Microbead Implantation in the Zebrafish Embryo

Infection of Zebrafish Embryos with Intracellular Bacterial Pathogens

Zebrafish (Danio rerio) embryos are increasingly used as a model for studying the function of the vertebrate innate immune system in host-pathogen interactions 1. The major cell types of the innate immune system, macrophages and neutrophils, develop during the first days of embryogenesis prior to the maturation of lymphocytes that are required for adaptive immune responses. The ease of obtaining large numbers of embryos, their accessibility due to external development, the optical transparency of embryonic and larval stages, a wide range of genetic tools, extensive mutant resources and collections of transgenic reporter lines, all add to the versatility of the zebrafish model. Salmonella enterica serovar Typhimurium (S. typhimurium) and Mycobacterium marinum can reside intracellularly in macrophages and are frequently used to study host-pathogen interactions in zebrafish embryos. The infection processes of these two bacterial pathogens are interesting to compare because S. typhimurium infection is acute and lethal within one day, whereas M. marinum infection is chronic and can be imaged up to the larval stage 2, 3. The site of micro-injection of bacteria into the embryo (Figure 1) determines whether the infection will rapidly become systemic or will initially remain localized. A rapid systemic infection can be established by micro-injecting bacteria directly into the blood circulation via the caudal vein at the posterior blood island or via the Duct of Cuvier, a wide circulation channel on the yolk sac connecting the heart to the trunk vasculature. At 1 dpf, when embryos at this stage have phagocytically active macrophages but neutrophils have not yet matured, injecting into the blood island is preferred. For injections at 2-3 dpf, when embryos also have developed functional (myeloperoxidase-producing) neutrophils, the Duct of Cuvier is preferred as the injection site. To study directed migration of myeloid cells towards local infections, bacteria can be injected into the tail muscle, otic vesicle, or hindbrain ventricle 4-6. In addition, the notochord, a structure that appears to be normally inaccessible to myeloid cells, is highly susceptible to local infection 7. A useful alternative for high-throughput applications is the injection of bacteria into the yolk of embryos within the first hours after fertilization 8. Combining fluorescent bacteria and transgenic zebrafish lines with fluorescent macrophages or neutrophils creates ideal circumstances for multi-color imaging of host-pathogen interactions. This video article will describe detailed protocols for intravenous and local infection of zebrafish embryos with S. typhimurium or M. marinum bacteria and for subsequent fluorescence imaging of the interaction with cells of the innate immune system.

Transparent zebrafish embryos have proved useful model hosts to visualize and functionally study interactions between innate immune cells and intracellular bacterial pathogens, such as Salmonella typhimurium and Mycobacterium marinum. Micro-injection of bacteria and multi-color fluorescence imaging are essential techniques involved in the application of zebrafish embryo infection models.

Zebrafish (Danio rerio) embryos are increasingly used as a model for studying the function of the vertebrate innate immune system in host-pathogen ...

Immunology and Infection

Microinjection of mRNA and Morpholino Antisense Oligonucleotides in Zebrafish Embryos.

An essential tool for investigating the role of a gene during development is the ability to perform gene knockdown, overexpression, and misexpression studies. In zebrafish (Danio rerio), microinjection of RNA, DNA, proteins, antisense oligonucleotides and other small molecules into the developing embryo provides researchers a quick and robust assay for exploring gene function in vivo. In this video-article, we will demonstrate how to prepare and microinject in vitro synthesized EGFP mRNA and a translational-blocking morpholino oligo against pkd2, a gene associated with autosomal dominant polycystic kidney disease (ADPKD), into 1-cell stage zebrafish embryos. We will then analyze the success of the mRNA and morpholino microinjections by verifying GFP expression and phenotype analysis. Broad applications of this technique include generating transgenic animals and germ-line chimeras, cell-fate mapping and gene screening. Herein we describe a protocol for overexpression of EGFP and knockdown of pkd2 by mRNA and morpholino oligonucleotide injection.

Microinjection is a well-established and effective method for introducing foreign substances into fertilized zebrafish embryos. Here, we demonstrate a robust microinjection technique for performing mRNA overexpression, and morpholino oligonucleotide gene knockdown studies in zebrafish.

An essential tool for investigating the role of a gene during development is the ability to perform gene knockdown, overexpression, and misexpression ...

Biology

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

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 ...

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue morphogenesis goes in hand with molecular and structural changes at the single cell level that result in variations of subcellular mechanical properties. As a consequence, even within the same cell, different organelles and compartments resist differently to mechanical stresses; and mechanotransduction pathways can actively regulate their mechanical properties. The ability of a cell to adapt to the microenvironment of the tissue niche thus is in part due to the ability to sense and respond to mechanical stresses. We recently proposed a new mechanosensation paradigm in which nuclear deformation and positioning enables a cell to gauge the physical 3D environment and endows the cell with a sense of proprioception to decode changes in cell shape. In this article, we describe a new method to measure the forces and material properties that shape the cell nucleus inside living cells, exemplified on adherent cells and mechanically confined cells. The measurements can be performed non-invasively with optical traps inside cells, and the forces are directly accessible through calibration-free detection of light momentum. This allows measuring the mechanics of the nucleus independently from cell surface deformations and allowing dissection of exteroceptive and interoceptive mechanotransduction pathways. Importantly, the trapping experiment can be combined with optical microscopy to investigate the cellular response and subcellular dynamics using fluorescence imaging of the cytoskeleton, calcium ions, or nuclear morphology. The presented method is straightforward to apply, compatible with commercial solutions for force measurements, and can easily be extended to investigate the mechanics of other subcellular compartments, e.g., mitochondria, stress-fibers, and endosomes.

Here, we present a protocol to investigate the intracellular mechanical properties of isolated embryonic zebrafish cells in three-dimensional confinement with direct force measurement by an optical trap.

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue ...

A Zebrafish Embryo Model for In Vivo Visualization and Intravital Analysis of Biomaterial-associated Staphylococcus aureus Infection

Biomaterial-associated infection (BAI) is a major cause of the failure of biomaterials/medical devices. Staphylococcus aureus is one of the major pathogens in BAI. Current experimental BAI mammalian animal models such as mouse models are costly and time-consuming, and therefore not suitable for high throughput analysis. Thus, novel animal models as complementary systems for investigating BAI in vivo are desired. In the present study, we aimed to develop a zebrafish embryo model for in vivo visualization and intravital analysis of bacterial infection in the presence of biomaterials based on fluorescence microscopy. In addition, the provoked macrophage response was studied. To this end, we used fluorescent protein-expressing S. aureus and transgenic zebrafish embryos expressing fluorescent proteins in their macrophages and developed a procedure to inject bacteria alone or together with microspheres into the muscle tissue of embryos. To monitor bacterial infection progression in live embryos over time, we devised a simple but reliable method of microscopic scoring of fluorescent bacteria. The results from microscopic scoring showed that all embryos with more than 20 colony-forming units (CFU) of bacteria yielded a positive fluorescent signal of bacteria. To study the potential effects of biomaterials on infection, we determined the CFU numbers of S. aureus with and without 10 &#181;m polystyrene microspheres (PS10) as model biomaterials in the embryos. Moreover, we used the ObjectJ project file &#34;Zebrafish-Immunotest&#34; operating in ImageJ to quantify the fluorescence intensity of S. aureus infection with and without PS10 over time. Results from both methods showed higher numbers of S. aureus in infected embryos with microspheres than in embryos without microspheres, indicating an increased infection susceptibility in the presence of the biomaterial. Thus, the present study shows the potential of the zebrafish embryo model to study BAI with the methods developed here.

A Zebrafish Embryo Model for In Vivo Visualization and Intravital Analysis of Biomaterial-associated Staphylococcus aureus Infection

The present study describes a zebrafish embryo model for in vivo visualization and intravital analysis of biomaterial-associated infection over time based on fluorescence microscopy. This model is a promising system complementing mammalian animal models such as mouse models for studying biomaterial-associated infections in vivo.

Biomaterial-associated infection (BAI) is a major cause of the failure of biomaterials/medical devices. Staphylococcus aureus is one of the major ...

Bioengineering

Elucidating Microglia Behavior in Zebrafish Optic Tectum Development

Live Imaging and Characterization of Microglia Dynamics in the Zebrafish Embryo

Microglia are highly dynamic cells and their migration and colonization of the brain parenchyma is a crucial step for proper brain development and function. Externally developing zebrafish embryos possess optical transparency, which along with well-characterized transgenic reporter lines that fluorescently label microglia, make zebrafish an ideal vertebrate model for such studies. In this paper, we take advantage of the unique features of the zebrafish model to visualize the dynamics of microglia cells in vivo and under physiological conditions. We use confocal microscopy to record a timelapse of microglia cells in the optic tectum of the zebrafish embryo and then, extract tracking data using the IMARIS 10.0 software to obtain the cells' migration path, mean speed, and distribution in the optic tectum at different developmental stages. This protocol can be a useful tool to elucidate the physiological significance of microglia behavior in various contexts, contributing to a deeper characterization of these highly motile cells.

Author Spotlight: Using Zebrafish to Explore Microglia Migration During Brain Development

We demonstrate a method that takes advantage of fast scanning confocal microscopy to perform live imaging of microglia cells in the developing zebrafish optic tectum, allowing for the analysis of the dynamics of these cells in vivo.

Microglia are highly dynamic cells and their migration and colonization of the brain parenchyma is a crucial step for proper brain development and ...

Brain Ventricular Microinjections of Lipopolysaccharide into Larval Zebrafish to Assess Neuroinflammation and Neurotoxicity

Neuroinflammation is a key player in various neurological disorders, including neurodegenerative diseases. Therefore, it is of great interest to research and develop alternative in vivo neuroinflammation models to understand the role of neuroinflammation in neurodegeneration. In this study, a larval zebrafish model of neuroinflammation mediated by ventricular microinjection of lipopolysaccharide (LPS) to induce an immune response and neurotoxicity was developed and validated. The transgenic zebrafish lines elavl3:mCherry, ETvmat2:GFP, and mpo:EGFP were used for real-time quantification of brain neuron viability by fluorescence live imaging integrated with fluorescence intensity analysis. The locomotor behavior of zebrafish larvae was recorded automatically using a video-tracking recorder. The content of nitric oxide (NO), and the mRNA expression levels of inflammatory cytokines including interleukin-6 (IL-6), interleukin-1&#946; (IL-1&#946;), and human tumor necrosis factor &#945; (TNF-&#945;) were investigated to assess the LPS-induced immune response in the larval zebrafish head. At 24 h after the brain ventricular injection of LPS, loss of neurons and locomotion deficiency were observed in zebrafish larvae. In addition, LPS-induced neuroinflammation increased NO release and the mRNA expression of IL-6, IL-1&#946;, and TNF-&#945; in the head of 6 days post fertilization (dpf) zebrafish larvae, and resulted in the recruitment of neutrophils in the zebrafish brain. In this study, injection of zebrafish with LPS at a concentration of 2.5-5 mg/mL at 5 dpf was determined as the optimum condition for this pharmacological neuroinflammation assay. This protocol presents a new, quick, and efficient methodology for brain ventricle microinjection of LPS to induce LPS-mediated neuroinflammation and neurotoxicity in a zebrafish larva, which is useful for studying neuroinflammation and could also be used as a high-throughput in vivo drug screening assay.

This protocol demonstrates the microinjection of lipopolysaccharide into the brain ventricular region in a zebrafish larval model to study the resulting neuroinflammatory response and neurotoxicity.

Neuroinflammation is a key player in various neurological disorders, including neurodegenerative diseases. Therefore, it is of great interest to research ...

Neuroscience

Zebrafish Breeding and Embryo Handling

Zebrafish (Danio rerio) are an important model organism that is particularly valuable for research in developmental biology. Zebrafish are extremely fertile and can produce hundreds of progeny per week, so it is relatively easy to collect a large number of embryos for high sample numbers. Furthermore, zebrafish undergo rapid development and embryos are transparent, allowing for easy visualization of developmental processes.
This video covers the steps required for the collection of newly fertilized zebrafish embryos. A brief overview of zebrafish mating behavior is presented, followed by instructions for setting up crosses in specialized laboratory breeding tanks that allow for controlled mating. Also covered are the conditions required to initiate the release of eggs (known as spawning) the morning after tanks are set. Next, essential techniques for working with embryos are presented, including the inhibition of pigment development with the chemical PTU, and dechorionation:&#160;a procedure in which the shell-like membrane surrounding the embryo (the chorion) is removed. Finally, the video concludes with some practical applications of these techniques in developmental research.

Zebrafish (Danio rerio) are an important model organism that is particularly valuable for research in developmental biology. Zebrafish are extremely ...

Education

JoVE Science Education

Basic Biology

Biology II: Mouse, Zebrafish, and Chick

Zebrafish Microinjection Techniques

One of the major advantages to working with zebrafish (Danio rerio) is that their genetics can be easily manipulated by microinjection of early stage embryos. Using this technique, solutions containing genetic material or knockdown constructs are delivered into the blastomeres: the embryonic cells sitting atop the yolk of the newly fertilized egg. Delivery into the cytoplasm is achieved either through direct injection into the blastomere, or via natural cytoplasmic movements that occur after a solution is injected into the yolk. Successful genetic manipulations are usually followed by quantification of embryonic phenotypes in order to elucidate the genetic mechanisms of development.
This video will provide an introduction to carrying out microinjections in zebrafish embryos. The discussion begins with a review of the essential tools for the technique, including the injection apparatus and the microinjector, which controls fluid movement with pressure pulses of air. Next, important preparatory steps are demonstrated, such as the pouring of agar plates to stabilize embryos during injection and calibration of the microinjection apparatus. The injection procedure is then presented along with tips on when and where injections should be performed. Finally, applications of the microinjection technique are discussed, including gene overexpression via mRNA injection, gene silencing by delivery of antisense morpholino oligonucleotides, and the generation of transgenic zebrafish using specially engineered plasmid DNA.

Introduction to Microinjection of Early Zebrafish Embryos

One of the major advantages to working with zebrafish (Danio rerio) is that their genetics can be easily manipulated by microinjection of early stage ...

Microbead Implantation in the Zebrafish Embryo

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Infection of Zebrafish Embryos with Intracellular Bacterial Pathogens

Microinjection of mRNA and Morpholino Antisense Oligonucleotides in Zebrafish Embryos.

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

A Simple and Effective Transplantation Device for Zebrafish Embryos

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

A Zebrafish Embryo Model for In Vivo Visualization and Intravital Analysis of Biomaterial-associated Staphylococcus aureus Infection

Live Imaging and Characterization of Microglia Dynamics in the Zebrafish Embryo

Brain Ventricular Microinjections of Lipopolysaccharide into Larval Zebrafish to Assess Neuroinflammation and Neurotoxicity

Zebrafish Breeding and Embryo Handling

Zebrafish Microinjection Techniques