This work was supported by JSPS KAKENHI Grant Number 18K14824 and 21K15195 and an internal grant of AIST, Japan.

All experimental protocols were approved by the Institutional Animal Care and Use Committee at the National Institute of Advanced Industrial Science and Technology. Zebrafish and medaka were maintained according to standard procedures28.
1. Stab wound injury in the adult optic tectum
<ol>
	<li>Prepare a 0.4% (w/v) tricaine stock solution for anesthesia. For a 100 mL stock solution, dissolve 400 mg tricaine methanesulfonate (see Table of Materials) in 90 mL of distilled water and adjust the pH to 7.0 using 1 M Tris HCl buffer (pH 9.0). Once the pH is adjusted, add water up to 100 mL, then make appropriate aliquots and store them at -20 &#176;C.</li>
	<li>To anesthetize the adult zebrafish or medaka, prepare a 0.02 % (w/v) tricaine solution by diluting the 0.4 % tricaine stock solution with fish facility water.</li>
	<li>Anesthetize the fish with 0.02% tricaine solution; they become anesthetized when they do not move or respond to touch (1-2 min).</li>
	<li>Place the anesthetized fish on a Styrofoam tray (see Table of Materials) and fix the fish upright between two 30 G needles inserted vertically into the Styrofoam.</li>
	<li>Hold the fish body to prevent the fish head from moving and vertically insert a 30 G needle through the skull into the medial region of one of the optic tectum hemispheres (Figure 1A,B). The insertion depth is ~0.75 mm, almost equal to half the length of the 30 G bevel. This insertion depth induces brain injury on zebrafish and medaka because of the similar body size of these fish. 
	NOTE: Medaka has scales on the skull. Scratch and remove the scales using the 30 G needle to efficiently and accurately induce the stab wound injury (Figure 1C).</li>
	<li>Transfer the injured fish into a fish tank with fresh fish facility water and move the tank to the fish breeding system once they have fully recovered from the anesthesia. 
	NOTE: Fish will recover and start freely moving within a few minutes.</li>
	<li>For lineage analysis after the injury, prepare fresh fish facility water containing 5 mM bromodeoxyuridine (BrdU) (see Table of Materials) and then incubate the fish with this 5 mM BrdU solution for a predetermined amount of time13,19 (as determined by an experimental design). Then dissect these fish as per step 2 and evaluate BrdU incorporation and cell lineage identity as appropriate. 
	&#8203;NOTE: This step is an optional step for cell lineage analysis. To detect BrdU signals, perform antigen retrieval as shown in step 4.3.</li>
</ol>
2. Brain dissection
<ol>
	<li>Prepare 0.02% tricaine solution, a Styrofoam tray for dissection, and a 10 mL syringe containing phosphate-buffered saline (1x PBS) with an extension tube and 30 G needles (see Table of Materials) for intracardial perfusion.</li>
	<li>Anesthetize the injured fish at selected time points.</li>
	<li>Place paper towels on the Styrofoam tray and place the anesthetized fish on the paper towels.</li>
	<li>Hold the fish body in place by vertically inserting two 30 G needles on either side of the anal fin (Figure 2A).</li>
	<li>Make a ventral incision from the anus to the heart (Figure 2B) and keep the heart visible by inserting another two 30 G needles on each side of this organ (Figure 2C). 
	NOTE: The heart is covered in the silvery epithelial layer of the hypodermis in both zebrafish and medaka (Figure 2C).</li>
	<li>Gently remove the silver epithelial layer with the tip of the forceps (Figure 2D).</li>
	<li>Insert the 30 G needle into the ventricle, keeping the bevel up, and make an incision into the atrium using the forceps (Figure 2E). Carefully press down on the syringe to push the 1x PBS and confirm that the atrium flushed the blood. 
	NOTE: To prevent the needle from penetrating the ventricle, carefully insert the needle to the depth of the half bevel and then adjust the insertion depth. If the perfusion is well done, gills turn white (Figure 2F,G).</li>
	<li>Stop the perfusion after the blood is drained and the gills turn white.</li>
	<li>Remove the needles fixing the fish body in place and fix the fish ventral side down (Figure 2H). Cut the spinal cord and remove the skull from the optic tectum and telencephalon (Figure 2I). Then, cut the optic nerves and dissect the brain carefully. 
	NOTE: Watch out for the optic nerves during the dissection because the nerves are connected to the tectum. When the optic nerve is pulled, the optic tectum can be detached from the brain. If the blood removal is not complete, the brain looks light pink (the right brain in Figure 2J).</li>
	<li>Transfer the brains into 1.5 mL tubes with 1 mL of 4 % paraformaldehyde in 1x PBS and fix them overnight at 4 &#176;C.</li>
</ol>
3. Preparation of frozen sections
<ol>
	<li>Wash the fixed brains three times in 1x PBS for 5 min each.</li>
	<li>To cryoprotect the brains, transfer them into 1.5 mL tubes with 1 mL of 30 % (w/v) sucrose in 1x PBS and incubate them overnight at 4 &#176;C. Prepare the embedding compound (2:1 mixture of OCT compound and 30% sucrose, see Table of Materials) by combining 30 mL of OCT and 15 mL of 30 % sucrose. Store this at 4 &#176;C. 
	NOTE: To remove the bubbles from the mixed compound, centrifuge the 50 mL tubes at 10,000 x g for 2 min at room temperature or place the tubes upright overnight.</li>
	<li>Incubate an aluminum block overnight at -80 &#176;C in order to cool the cryomold (see Table of Materials) immediately after the dissected brain is embedded in the cryomold with the embedding compound. 
	NOTE: Liquid nitrogen can also be used to cool the aluminum block. In that case, the aluminum block should be placed in a Styrofoam box filled with enough liquid nitrogen to cover the aluminum block just before embedding. After confirming the liquid nitrogen sublimation, place the cryomold on the precooled aluminum block. It is better to confirm how many cryomolds can be placed on the aluminum block at once. It is advisable to use liquid nitrogen or an additional precooled block to ensure enough space to cool all samples quickly.</li>
	<li>Put the precooled aluminum block in a Styrofoam box. Use a Styrofoam box with a lid to prevent the aluminum block from warming.</li>
	<li>For coronal sections, embed the brain in a cryomold and adjust the orientation using forceps under the microscope (Figure 3A).</li>
	<li>Place the cryomold on the precooled aluminum block in the Styrofoam box and allow the OCT compound to freeze (Figure 3B). When the OCT compound starts to freeze, it becomes white.</li>
	<li>Apply a small circle of OCT compound on a specimen disc, and mount the cryoblock to attach the cryoblock to the specimen disc.</li>
	<li>Place the specimen disc in the cryostat and freeze the OCT compound completely.</li>
	<li>Place the specimen disc on the specimen head in the cryostat.</li>
	<li>Set the orientation of the cryoblock and trim the OCT to remove any extra regions. 
	NOTE: Adjust the orientation of the cutting plane while cutting the telencephalon. The blade is equipped with a cryostat. It is possible to adjust the orientation of the cutting plane by changing the angle of the specimen head.</li>
	<li>Cut 14 &#181;m thick serial sections through the whole optic tectum using a cryostat. Store the slides at -25 &#176;C. 
	&#8203;NOTE: Long-term storage should be at -80 &#176;C.</li>
</ol>
4. Fluorescent immunostaining
<ol>
	<li>Dry glass slides in a slide rack for 30 min at room temperature (RT). Wash slides in 1x PBS for 30 min at RT. If immunostaining with anti-PCNA or BrdU antibodies is to be performed, proceed to step 4.2 or step 4.3.</li>
	<li>Prepare 500 mL of 10 mM sodium citrate in a 500 mL beaker and warm the citrate buffer to 85 &#176;C on a hot plate magnetic stirrer, then place the slides in a slide staining rack and incubate the rack in the citrate buffer at 85 &#176;C for 30 min. To avoid boiling, keep the temperature around 85 &#176;C. After antigen retrieval, wash the slides three times in 0.1 % Triton in 1x PBS (PBSTr), with each wash taking 5 min at RT. 
	NOTE: This step is an optional step for proliferating cell nuclear antigen (PCNA) retrieval.</li>
	<li>Prepare a 2 N HCl solution by diluting 12 N HCl in distilled water. Using a liquid blocker (see Table of Materials), draw a line as a hydrophobic barrier to keep the 2 N HCl solution around the sections.
	<ol>
		<li>Place the slides on immunostaining trays and apply 200-300 &#181;L of the 2N HCl solution. Incubate the trays at 37 &#176;C for 30 min. After antigen retrieval, wash three times in 0.1 % PBSTr with each wash taking 5 min at RT. 
		NOTE: This step is an optional step for BrdU antigen retrieval.</li>
	</ol>
	</li>
	<li>Absorb and remove the remaining solution using paper towels. Then, use a liquid blocker to draw a hydrophobic barrier to keep the antibody solution around the sections as necessary.</li>
	<li>Place the slides on trays and apply 200-300 &#181;L of blocking solution, 3% (v/v) horse serum in PBSTr) to every slide, and incubate the slides for 1 h at RT.</li>
	<li>Wash the sections with PBSTr for 5 min at RT and repeat three times.</li>
	<li>Prepare the primary antibodies (see Table of Materials) using the blocking solution (200-300 &#181;L of antibody solution for each slide). Remove the remaining PBSTr with paper towels and place the slides on the immunostaining trays. Apply the primary antibody solution to every slide and incubate trays overnight at 4 &#176;C.</li>
	<li>Wash the sections with PBSTr for 5 min at RT and repeat three times.</li>
	<li>Prepare fluorescent secondary antibody solution (see Table of Materials) using the blocking buffer (1:500). Remove the remaining PBSTr with paper towels and place the slides on the immunostaining trays. Apply the secondary antibody solution to every slide and shade trays with aluminum foil. Incubate for 1-2 h at RT.</li>
	<li>Wash the sections with PBSTr for 5 min at RT three times without exposing them to light.</li>
	<li>Prepare Hoechst solution (1:500 dilution in 1x PBS, see Table of Materials) for nuclear staining. Remove the remaining PBSTr with paper towels and place the slides on the immunostaining trays. Apply the Hoechst solution to every slide and cover the trays with aluminum foil. Incubate for 30 min at RT.</li>
	<li>Wash the sections with 1x PBS for 10 min at RT without exposing them to light.</li>
	<li>Absorb and remove the remaining 1x PBS using paper towels and mount the sections on slides using a water-soluble mounting medium (see Table of Materials) for fluorescent immunostaining. Slowly set a 24 x 45 mm coverslip onto the sections. Store at 4 &#176;C, without exposing the slides to light.</li>
	<li>Observe and image the sections under fluorescent microscopy or confocal microscopy.</li>
</ol>

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The authors have nothing to disclose.

Here a set of methods is described which can be used to induce stab wound injuries in the optic tectum utilizing a needle to facilitate the evaluation of RGC proliferation and differentiation after brain injury. Needle-mediated stab wounds are a simple, efficiently implemented method that can be applied to many experimental samples using a standard set of tools. Stab wound injury models for several regions of the zebrafish brain have been developed3,19,</sup...

Zebrafish (Danio rerio) have an increased ability to regenerate their central nervous system (CNS) compared to other mammals1,2,3. Recently, to better understand the molecular mechanisms underlying this increased regenerative capacity, comparative analyses of tissue regeneration using next-generation sequencing technology have been performed4,5,6. The brain structures in zebrafish and tetrapods are quite different7,8,9. This means that several brain injury models using small fish with similar brain structures and biological features have been developed to facilitate the investigation of the underlying molecular mechanisms contributing to this increased regenerative capacity.
In addition, medaka (Oryzias latipes) is a popular laboratory animal with a low capacity for heart and neuronal regeneration10,11,12,13 compared with zebrafish. Zebrafish and medaka have similar brain structures and niches for adult neural stem cells (NSCs)14,15,16,17. In zebrafish and medaka, the optic tectum includes two types of NSCs, neuroepithelial-like stem cells and radial glial cells (RGCs)15,18. A stab wound injury for the optic tectum of adult zebrafish was previously developed, and this model was used to investigate the molecular mechanisms regulating brain regeneration in these animals19,20,21,22,23. This young adult zebrafish stab wound injury model induced regenerative neurogenesis from RGCs19,24,25. This stab wound injury in the optic tectum is a robust and reproducible method13,19,20,21,22,23,24,25. When the same injury model was applied to adult medaka, the low neurogenic capacity of RGCs in medaka optic tectum was revealed via the comparative analysis of RGC proliferation and differentiation following injury13.
Stab wound injury models in the optic tectum have also been developed in mummichog models26, but details of the tectum injury have been not well documented when compared with telencephalic injury27. The stab wound injury in the optic tectum using zebrafish and medaka allows the investigation of the differential cellular responses and gene expression between species with differential regenerative capacity. This protocol describes how to perform a stab wound injury in the optic tectum using an injection needle. This method can be applied to small fish like zebrafish and medaka. The processes for sample preparation for histological analysis and cellular proliferation and differentiation analysis using fluorescent immunohistochemistry and cryosections are explained here.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>10 mL syringe</td>
			<td>TERUMO</td>
			<td>SS-10ESZ</td>
			<td></td>
		</tr>
		<tr>
			<td>1M Tris-HCl (pH 9.0)</td>
			<td>NIPPON GENE</td>
			<td>314-90381</td>
			<td></td>
		</tr>
		<tr>
			<td>30 G needle</td>
			<td>Dentronics</td>
			<td>HS-2739A</td>
			<td></td>
		</tr>
		<tr>
			<td>4% Paraformaldehyde Phosphate Buffer Solution</td>
			<td>Wako</td>
			<td>163-20145</td>
			<td></td>
		</tr>
		<tr>
			<td>Aluminum block</td>
			<td></td>
			<td></td>
			<td>115 x 80 x 37 mm (W x D x H) is enough size to freeze 6 cryomolds</td>
		</tr>
		<tr>
			<td>Anti-BLBP</td>
			<td>Millipore</td>
			<td>ABN14</td>
			<td>1:500</td>
		</tr>
		<tr>
			<td>Anti-BrdU</td>
			<td>Abcam</td>
			<td>ab1893</td>
			<td>1:500</td>
		</tr>
		<tr>
			<td>Anti-HuC</td>
			<td>Invitrogen</td>
			<td>A21271</td>
			<td>1:100</td>
		</tr>
		<tr>
			<td>Anti-PCNA</td>
			<td>Santa Cruz Biotechnology</td>
			<td>sc-56</td>
			<td>1:200</td>
		</tr>
		<tr>
			<td>Brmodeoxyuridine</td>
			<td>Wako</td>
			<td>023-15563</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal microscope C1 plus</td>
			<td>Nikon</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Cryomold</td>
			<td>Sakura Finetek Japan</td>
			<td>4565</td>
			<td>10 x 10 x 5 mm (W x D x H)</td>
		</tr>
		<tr>
			<td>Cryostat</td>
			<td>Leica</td>
			<td>CM1960</td>
			<td></td>
		</tr>
		<tr>
			<td>Danio rerio WT strains RW</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Extension tube</td>
			<td>TERUMO</td>
			<td>SF-ET3520</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluoromount (TM) Aqueous Mounting Medium, for use with fluorescent dye-stained tissues</td>
			<td>SIGMA-ALDRICH</td>
			<td>F4680-25ML</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>DUMONT</td>
			<td>11252-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat anti-Mouse IgG (H+L) Highly Cross-Adsorbed Secondary Antibody, Alexa Fluor Plus 488</td>
			<td>Invitrogen</td>
			<td>A32723</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat anti-Rabbit IgG (H+L) Highly Cross-Adsorbed Secondary Antibody, Alexa Fluor 546</td>
			<td>Invitrogen</td>
			<td>A11035</td>
			<td></td>
		</tr>
		<tr>
			<td>Hoechst 33342 solution</td>
			<td>Dojindo</td>
			<td>23491-52-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrochloric Acid</td>
			<td>Wako</td>
			<td>080-01066</td>
			<td></td>
		</tr>
		<tr>
			<td>Incubation Chamber for 10 slides Dark Orange</td>
			<td>COSMO BIO CO., LTD.</td>
			<td>10DO</td>
			<td></td>
		</tr>
		<tr>
			<td>MAS coat sliding glass</td>
			<td>Matsunami glass</td>
			<td>MAS-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro cover glass</td>
			<td>Matsunami glass</td>
			<td>C024451</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscopy</td>
			<td>Nikon</td>
			<td>SMZ745T</td>
			<td></td>
		</tr>
		<tr>
			<td>Normal horse serum blocking solution</td>
			<td>VECTOR LABRATORIES</td>
			<td>S-2000-20</td>
			<td></td>
		</tr>
		<tr>
			<td>O.C.T Compound</td>
			<td>Sakura Finetek Japan</td>
			<td>83-1824</td>
			<td></td>
		</tr>
		<tr>
			<td>Oryzias latipes WT strains Cab</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PAP Pen Super-Liquid Blocker</td>
			<td>DAIDO SANGYO</td>
			<td>PAP-S</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate Buffered Saline (PBS) Tablets, pH 7.4</td>
			<td>TaKaRa</td>
			<td>T9181</td>
			<td></td>
		</tr>
		<tr>
			<td>Styrofoam tray</td>
			<td></td>
			<td></td>
			<td>100 x 100 x 10 mm (W x D x H) styrofoam sheet is available as tray</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Wako</td>
			<td>196-00015</td>
			<td>30 % (w/v) Sucrose in PBS</td>
		</tr>
		<tr>
			<td>Tricaine (MS-222)</td>
			<td>nacarai tesque</td>
			<td>14805-24</td>
			<td></td>
		</tr>
		<tr>
			<td>Trisodium Citrate Dihydrate</td>
			<td>Wako</td>
			<td>191-01785</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Wako</td>
			<td>04605-250</td>
			<td></td>
		</tr>
	</tbody>
</table>

stab wound injury model of the adult optic tectum using zebrafish and medaka for the comparative analysis of regenerative capacity

While zebrafish have a superior capacity to regenerate their central nervous system (CNS), medaka has a lower CNS regenerative capacity. A brain injury model was developed in the adult optic tectum of zebrafish and medaka and comparative histological and molecular analyses were performed to elucidate the molecular mechanisms regulating the high regenerative capacity of this tissue across these fish species. Here a stab wound injury model is presented for the adult optic tectum using a needle and histological analyses for proliferation and differentiation of the neural stem cells (NSCs). A needle was manually inserted into the central region of the optic tectum, and then the fish were intracardially perfused, and their brains were dissected. These tissues were then cryosectioned and evaluated using immunostaining against the appropriate NSC proliferation and differentiation markers. This tectum injury model provides robust and reproducible results in both zebrafish and medaka, allowing for comparing NSC responses after injury. This method is available for small teleosts, including zebrafish, medaka, and African killifish, and enables us to compare their regenerative capacity and investigate unique molecular mechanisms.

Stab wound injury in the optic tectum using needle insertion into the right hemisphere (Figure 1, Figure 4A, and Figure 5A) induces various cellular responses, including radial glial cell (RGC) proliferation and the generation of newborn neurons. Similarly, aged populations of zebrafish and medaka were used to counteract any aging effects in the regenerative response. Then fluorescent immunostaining was performed on the frozen secti...

Watch this Scientific Journal Video about Stab Wound Injury Model of the Adult Optic Tectum Using Zebrafish and Medaka for the Comparative Analysis of Regenerative Capacity at JoVE.com

Stab Wound Injury Model of the Adult Optic Tectum Using Zebrafish and Medaka for the Comparative Analysis of Regenerative Capacity

A mechanical brain injury model in the adult zebrafish is described to investigate the molecular mechanisms regulating their high regenerative capacity. The method explains to create a stab wound injury in the optic tectum of multiple species of small fish to evaluate the regenerative responses using fluorescent immunostaining.

While zebrafish have a superior capacity to regenerate their central nervous system (CNS), medaka has a lower CNS regenerative capacity. A brain injury ...

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Research

JoVE Journal

Developmental Biology

1.7K Views.  National Institute of Advanced Industrial Science and Technology. A mechanical brain injury model in the adult zebrafish is described to investigate the molecular mechanisms regulating their high regenerative capacity. The method explains to create a stab wound injury in the optic tectum of multiple species of small fish to evaluate the regenerative responses using fluorescent immunostaining.

Video: Stab Wound Injury Model of the Adult Optic Tectum Using Zebrafish and Medaka for the Comparative Analysis of Regenerative Capacity

Kidney Regeneration in Adult Zebrafish by Gentamicin Induced Injury

The kidney is essential for fluid homeostasis, blood pressure regulation and filtration of waste from the body. The fundamental unit of kidney function is the nephron. Mammals are able to repair existing nephrons after injury, but lose the ability to form new nephrons soon after birth. In contrast to mammals, adult fish produce new nephrons (neonephrogenesis) throughout their lives in response to growth requirements or injury. Recently, lhx1a has been shown to mark nephron progenitor cells in the adult zebrafish kidney, however mechanisms controlling the formation of new nephrons after injury remain unknown. Here we show our method for robust and reproducible injury in the adult zebrafish kidney by intraperitoneal (i.p.) injection of gentamicin, which uses a noninvasive visual screening process to select for fish with strong but nonlethal injury. Using this method, we can determine optimal gentamicin dosages for injury and go on to demonstrate the effect of higher temperatures on kidney regeneration in zebrafish.

Here we present a reliable method to study adult kidney regeneration by inducing acute kidney injury by gentamicin injection. We show that injury is dependent on gentamicin dosage and environmental temperature using in situ hybridization to label lhx1a+ developing new nephrons.

The kidney is essential for fluid homeostasis, blood pressure regulation and filtration of waste from the body. The fundamental unit of kidney function is ...

Visualization of Chondrocyte Intercalation and Directional Proliferation via Zebrabow Clonal Cell Analysis in the Embryonic Meckel&#8217;s Cartilage

Development of the vertebrate craniofacial structures requires precise coordination of cell migration, proliferation, adhesion and differentiation. Patterning of the Meckel&#39;s cartilage, a first pharyngeal arch derivative, involves the migration of cranial neural crest (CNC) cells and the progressive partitioning, proliferation and organization of differentiated chondrocytes. Several studies have described CNC migration during lower jaw&#160;morphogenesis, but the details of how the chondrocytes achieve organization in the growth and extension of Meckel&#8217;s cartilage remains unclear. The sox10 restricted and chemically induced Cre recombinase-mediated recombination generates permutations of distinct fluorescent proteins (RFP, YFP and CFP), thereby creating a multi-spectral labeling of progenitor cells and their progeny, reflecting distinct clonal populations. Using confocal time-lapse photography, it is possible to observe the chondrocytes behavior during the development of the zebrafish Meckel&#8217;s cartilage.
Multispectral cell labeling enables scientists to demonstrate extension of the Meckel&#8217;s chondrocytes.&#160;During extension phase of the Meckel&#8217;s cartilage, which prefigures the mandible, chondrocytes intercalate to effect extension as they stack in an organized single-cell layered row. Failure of this organized intercalating process to mediate cell extension provides the cellular mechanistic explanation for hypoplastic mandible that we observe in mandibular malformations.

Visualization of Chondrocyte Intercalation and Directional Proliferation via Zebrabow Clonal Cell Analysis in the Embryonic Meckel&amp;#8217;s Cartilage

Cell organization of craniofacial bones has long been hypothesized but never directly visualized. Multi-spectral cell labeling and in vivo live imaging allows visualization of dynamic cell behavior in zebrafish lower jaw. Here, we detail the protocol to manipulate Zebrabow transgenic fish and directly observe cell intercalation and morphological changes of chondrocytes in the Meckel&#8217;s cartilage.

Development of the vertebrate craniofacial structures requires precise coordination of cell migration, proliferation, adhesion and differentiation. ...

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 ...

Salinity-dependent Toxicity Assay of Silver Nanocolloids Using Medaka Eggs

Salinity is an important characteristic of the aquatic environment. For aquatic organisms it defines the habitats of freshwater, brackish water, and seawater. Tests of the toxicity of chemicals and assessments of their ecological risks to aquatic organisms are frequently performed in freshwater, but the toxicity of chemicals to aquatic organisms depends on pH, temperature, and salinity. There is no method, however, for testing the salinity dependence of toxicity to aquatic organisms. Here, we used medaka (Oryzias latipes) because they can adapt to freshwater, brackish water, and seawater. Different concentrations of embryo-rearing medium (ERM) (1x, 5x, 10x, 15x, 20x, and 30x) were employed to test the toxicity of silver nanocolloidal particles (SNCs) to medaka eggs (1x ERM and 30x ERM have osmotic pressures equivalent to freshwater and seawater, respectively). In six-well plastic plates, 15 medaka eggs in triplicate were exposed to SNCs at 10 mg/L&#8722;1 in different concentrations of ERM at pH 7 and 25 &#176;C in the dark.
We used a dissecting microscope and a micrometer to measure heart rate per 15 sec and eye diameter on day 6 and full body length of the larvae on hatching day (section 4). The embryos were observed until hatching or day 14; we then counted the hatching rate every day for 14 days (section 4). To see silver accumulation in embryos, we used inductively coupled plasma mass spectrometry to measure the silver concentration of test solutions (section 5) and dechorionated embryos (section 6).The toxicity of the SNCs to medaka embryos obviously increased with increasing salinity. This new method allows us to test the toxicity of chemicals in different salinities.

Embryonic stages are the most susceptible to xenobiotics. Although chemical toxicity depends on salinity, no method exists to test the salinity dependence of toxicity to aquatic organisms. Here, we describe a new and high-throughput method for determining the salinity dependence of toxicity to aquatic embryos.

Salinity is an important characteristic of the aquatic environment. For aquatic organisms it defines the habitats of freshwater, brackish water, and ...

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

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

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

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

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

A Seminiferous Tubule Squash Technique for the Cytological Analysis of Spermatogenesis Using the Mouse Model

Meiotic progression in males is a process that requires the concerted action of a number of highly regulated cellular events. Errors occurring during meiosis can lead to infertility, pregnancy loss or genetic defects. Commencing at the onset of puberty and continuing throughout adulthood, continuous semi-synchronous waves of spermatocytes undergo spermatogenesis and ultimately form haploid sperm. The first wave of mouse spermatocytes undergoing meiotic initiation appear at day 10 post-partum (10 dpp) and are released into the lumen of seminiferous tubules as mature sperm at 35 dpp. Therefore, it is advantageous to utilize mice within this developmental time-window in order to obtain highly enriched populations of interest. Analysis of rare cell stages is more difficult in older mice due to the contribution of successive spermatogenic waves, which increase the diversity of the cellular populations within the tubules. The method described here is an easily implemented technique for the cytological evaluation of the cells found within the seminiferous tubules of mice, including spermatogonia, spermatocytes, and spermatids. The tubule squash technique maintains the integrity of isolated male germ cells and allows examination of cellular structures that are not easily visualized with other techniques. To demonstrate the possible applications of this tubule squash technique, spindle assembly was monitored in spermatocytes progressing through the prophase to metaphase I transition (G2/MI transition). In addition, centrosome duplication, meiotic sex chromosome inactivation (MSCI), and chromosome bouquet formation were assessed as examples of the cytological structures that can be observed using this tubule squash method. This technique can be used to pinpoint specific defects during spermatogenesis that are caused by mutation or exogenous perturbation, and thus, contributes to our molecular understanding of spermatogenesis.

The goal of this tubule squash technique is to rapidly assess cytological features of developing mouse spermatocytes while preserving cellular integrity. This method allows for the study of all stages of spermatogenesis, and can be easily implemented alongside other biochemical and molecular biological approaches for the study of mouse meiosis.

Meiotic progression in males is a process that requires the concerted action of a number of highly regulated cellular events. Errors occurring during ...

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling processes of excitable neuronal and muscular cells and many other biological processes, such as embryonic developmental patterning. However, there is a need for in vivo electrical activity monitoring in vertebrate embryogenesis. The advances of genetically encoded fluorescent voltage indicators (GEVIs) have made it possible to provide a solution for this challenge. Here, we describe how to create a transgenic voltage indicator zebrafish using the established voltage indicator, ASAP1 (Accelerated Sensor of Action Potentials 1), as an example. The Tol2 kit and a ubiquitous zebrafish promoter, ubi, were chosen in this study. We also explain&#160;the processes of Gateway site-specific cloning, Tol2 transposon-based zebrafish transgenesis, and the imaging process for early-stage fish embryos and fish tumors using regular epifluorescent microscopes. Using this fish line, we found that there are cellular electric voltage changes during zebrafish embryogenesis, and fish larval movement. Furthermore, it was observed that in a few zebrafish malignant peripheral nerve sheath tumors, the tumor cells were generally polarized compared to the surrounding normal tissues.

Here, we show the process of creating a cellular electric voltage reporter zebrafish line to visualize embryonic development, movement, and fish tumor cells in vivo.

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling ...

Adult Zebrafish Injury Models to Study the Effects of Prednisolone in Regenerating Bone Tissue

Zebrafish are able to regenerate various organs, including appendages (fins) after amputation. This involves the regeneration of bone, which regrows within roughly two weeks after injury. Furthermore, zebrafish are able to heal bone rapidly after trepanation of the skull, and repair fractures that can be easily introduced into zebrafish bony fin rays. These injury assays represent feasible experimental paradigms to test the effect of administered drugs on rapidly forming bone. Here, we describe the use of these 3 injury models and their combined use with systemic glucocorticoid treatment, which exerts bone inhibitory and immunosuppressive effects. We provide a workflow on how to prepare for immunosuppressive treatment in adult zebrafish, illustrate how to perform fin amputation, trepanation of calvarial bones, and fin fractures, and describe how the use of glucocorticoids affects both bone forming osteoblasts and cells of the monocyte/macrophage lineage as part of innate immunity in bone tissue.

Here, we describe 3 adult zebrafish injury models and their combined use with immunosuppressive drug treatment. We provide guidance on imaging of regenerating tissues and on detecting bone mineralization therein.

Zebrafish are able to regenerate various organs, including appendages (fins) after amputation. This involves the regeneration of bone, which regrows ...

Visualization of Tangential Cell Migration in the Developing Chick Optic Tectum

Time-lapse imaging is a powerful method to analyze migrating cell behavior. After fluorescent cell labeling, the movement of the labeled cells in culture can be recorded under video microscopy. For analyzing cell migration in the developing brain, slice culture is commonly used to observe cell migration parallel to the slice section, such as radial cell migration. However, limited information can be obtained from the slice culture method to analyze cell migration perpendicular to the slice section, such as tangential cell migration. Here, we present the protocols for time-lapse imaging to visualize tangential cell migration in the developing chick optic tectum. A combination of cell labeling by electroporation in ovo and a subsequent flat-mount culture on the cell culture insert enables detection of migrating cell movement in the horizontal plane. Moreover, our method facilitates detection of both individual cell behavior and the collective action of a group of cells in the long term. This method can potentially be applied to detect the sequential change of the fluorescent-labeled micro-structure, including the axonal elongation in the neural tissue or cell displacement in the non-neural tissue.

We describe the methods for fluorescent labeling of tangentially migrating cells by electroporation, and for time-lapse imaging of the labeled cell movement in a flat-mount culture in order to visualize migrating cell behavior in the developing chick optic tectum.

Time-lapse imaging is a powerful method to analyze migrating cell behavior. After fluorescent cell labeling, the movement of the labeled cells in culture ...

Mapping the Emergent Spatial Organization of Mammalian Cells using Micropatterns and Quantitative Imaging

A fundamental goal in biology is to understand how patterns emerge during development. Several groups have shown that patterning can be achieved in vitro when stem cells are spatially confined onto micropatterns, thus setting up experimental models which offer unique opportunities to identify, in vitro, the fundamental principles of biological organisation.
Here we describe our own implementation of the methodology. We adapted a photo-patterning technique to reduce the need for specialized equipment to make it easier to establish the method in a standard cell biology laboratory. We also developed a free, open-source and easy to install image analysis framework in order to precisely measure the preferential positioning of sub-populations of cells within colonies of standard shapes and sizes. This method makes it possible to reveal the existence of patterning events even in seemingly disorganized populations of cells. The technique provides quantitative insights and can be used to decouple influences of the environment (e.g., physical cues or endogenous signaling), on a given patterning process.

The method presented here uses micropatterning together with quantitative imaging to reveal spatial organization within mammalian cultures. The technique is easy to establish in a standard cell biology laboratory and offers a tractable system to study patterning in vitro.

A fundamental goal in biology is to understand how patterns emerge during development. Several groups have shown that patterning can be achieved in vitro ...