Name: stab wound injury of the zebrafish adult telencephalon: a method to investigate vertebrate brain neurogenesis and regeneration
Uploaded: 2014-08-04T23:00:00.000+00:00
Duration: 9 min 16 s
Description: Watch this Scientific Journal Video about Stab Wound Injury of the Zebrafish Adult Telencephalon: A Method to Investigate Vertebrate Brain Neurogenesis and Regeneration at JoVE.com

We thank N. Borel and her fish house team, M. Rastegar for help with microscopy, and T. Dickmeis for comments on the manuscript. We are grateful for support by the EU IP ZF-Health, FP7-HEALTH-2007-B2, NeuroXsys, the Interreg Network for Synthetic Biology in the Upper Rhine valley (NSB-Upper Rhine) and the BMBF funded network EraSysBio.

Zebrafish were maintained in the fish facility of the Institute of Toxicology and Genetics (ITG) at Karlsruhe Institute of Technology (KIT). Experiments on animals were performed in accordance with the German animal protection standards and were approved by the Government of Baden-W&#252;rttemberg, Regierungspr&#228;sidium Karlsruhe, Germany (Aktenzeichen 35-9185.81/G-272/12 &#39;Adulte Neurogenese&#39;).
1. Generating a Stab Wound in the Telencephalon
<ol>
	<li>Anesthesia
	<ol>
		<li>To prepare a stock solution of tricaine (3-amino benzoic acid ethyl ester, also called ethyl 3-aminobenzoate) use 400 mg of tricaine powder and dissolve it in 97.9 ml water and 2.1 ml 1 M Tris/HCl buffer (pH 9). Adjust the pH to 7. Make 5 ml aliquots and store them at -20 &#176;C until usage.</li>
		<li>Transfer an appropriate number of 0.5-1 year old zebrafish from their tanks into plastic mouse cages. Note: The skull of an adult zebrafish at about 12 months of age is still relatively soft and can be perforated easily with a needle (see section 1.2.3 and Figure 1A).</li>
		<li>To anesthetize the adult zebrafish, combine 5 ml tricaine (stock solution) in a plastic crossing cage with ~100 ml of clean fish tank water (final concentration of tricaine 0.02% (w/v) tricaine).</li>
		<li>Incubate the fish in the anesthetics until they do not move anymore (45-60 sec).</li>
	</ol>
	</li>
	<li>Telencephalon Injury
	<ol>
		<li>Place individual anesthetized zebrafish into a slit in a block of tricaine-soaked foam.</li>
		<li>Under a dissecting microscope with light from the top gently hold the fish with one hand and orient it in a way that allows access to the head from the top.</li>
		<li>With the other hand push a 30 G syringe (insulin syringe with integrated needle) vertically through the skull into the medial region of one telencephalic hemisphere (Figures&#160;1A-1B). The hemispheres of the telencephalon are visible through the skull. Make sure that the introduced lesion is not deeper than 2 mm (Figure 1C). Note: It is highly recommended to introduce the lesion always into the same hemisphere to ease the identification of the site of lesion when dissecting the brain out of the skull.</li>
		<li>After introducing the telencephalic injury, place the fish into fresh fish water. Keep up to 10 fish in a 2 L mouse cage filled with fish water for recovery and connect the mouse cage to a water flow system. Note: The survival rate is usually ~97% if fish are healthy and no other experiments were performed with the fish before. Addition of antibiotics to the fish water is not necessary. After recovery, fish behave normally: they swim, feed, and mate.</li>
	</ol>
	</li>
</ol>
2. Analyzing the Effect of Telencephalic Injury
<ol>
	<li>Brain Dissection and Fixation
	<ol>
		<li>Re-anaesthetize the recovered fish at different time points after lesion (typically, 1, 3, 5, and 14 days post lesion (dpl)) with 0.02% (w/v) tricaine as above and euthanize them by putting them into ice water for 5 min.</li>
		<li>Place the fish on a paper tissue soaked in phosphate-buffered saline (PBS) and separate the head from the body by cutting behind the gills with a sharp scissor.</li>
		<li>Keep the heads in 1x PBS for 5 min in order to allow bleeding. Note: This drains blood from the tissue, which may perturb further steps in the protocol.</li>
		<li>Fix the heads overnight at 4 &#176;C in 4% paraformaldehyde (PFA) in PBS or 4 hr at room temperature (RT).</li>
		<li>Wash fixed heads twice with 1x PBS in a petri dish.</li>
		<li>Dissect brains carefully in 1x PBS under a dissecting microscope23. The lesion is usually visible under the dissecting microscope. Brains without a visible lesion should be discarded.</li>
		<li>Transfer brains into 2 ml reaction tubes filled with 1.5 ml 100% Methanol (MeOH) and invert the tubes 5x&#160;before incubating them at -20 &#176;C for at least 16 hr. Note: The brains can be kept for several months in MeOH at -20 &#176;C until needed for immunohistochemistry or in situ hybridization.</li>
	</ol>
	</li>
	<li>Tissue Preparation for Immunohistochemistry
	<ol>
		<li>Rehydrate the brains through a descending methanol series in 2 ml plastic tubes (sequentially 75% MeOH (1x PBS, 0.1% Tween-20 = PTW); 50% MeOH, 25% MeOH. Incubate the brains (15 brains/2 ml reaction tubes) for 5 min in each solution.</li>
		<li>Wash the brains in PTW for 5 min. Repeat 4x&#160;with fresh PTW.</li>
	</ol>
	</li>
	<li>Embedding of the Brains for Sectioning
	<ol>
		<li>For embedding, transfer the brains from the 2 ml reaction tube in PTW into the cavities of a polyethylene molding cup tray with a transfer pipet (5 mm diameter).</li>
		<li>Prepare 2% agarose (DNA grade) in 1x PBS in an Erlenmeyer flask. Heat it in the microwave at 600 W until the agarose is dissolved completely. Prepare 50 ml for 10 brains. Let the agarose cool down for 3 min at room temperature before usage. Keep the agarose on a pre-heated heating block (60 &#176;C) while embedding the brains in order to prevent solidification.</li>
		<li>Remove the 1x PBS from the mold containing one brain by using a Pasteur pipette (1 mm diameter). Be careful not to damage the brain.</li>
		<li>Pour agarose into the mold and fill it entirely. Then use a dissection needle to swirl the brain in the agarose to wash away the PTW making it easier to embed the brain.</li>
		<li>Orient the brain in the mold ventral side down, dorsal side up and place it straight.</li>
		<li>Let the agarose cool down. For faster cooling, place the mold on ice. Repeat steps.</li>
		<li>Repeat 2.3.3-2.3.6 for all brains.</li>
	</ol>
	</li>
	<li>Trimming of the Agarose Block
	<ol>
		<li>Using a dissection needle, pop out the agarose block.</li>
		<li>With a sharp razor blade, cut the agarose at the posterior end of the brain. Make sure this plane is parallel to the telencephalon.</li>
		<li>Cut the agarose at the anterior end of the brain. Then flip the block so that it stands on the posterior plane.</li>
		<li>Remove the excess of agarose by cutting in parallel to the brain&#8217;s dorsal and ventral sides. Then flip the brain back so that it lies on its ventral side.</li>
		<li>Cut the block to a truncated triangle in which the telencephalon is located at the smaller plane (Figure 2).</li>
	</ol>
	</li>
	<li>Preparing the Vibratome for Sectioning
	<ol>
		<li>Prepare the vibratome according to the manufacturer&#8217;s instructions. Use a new razor blade and replace it after 10 brains.</li>
		<li>Fill the buffer bath with 1x PBS to a level that just reaches the bottom of the blade.</li>
		<li>Place a small dot of superglue on the top of the specimen disc of the vibratome.</li>
		<li>Carefully place the plane that is located posterior to the brain on the superglue. The glue will harden as soon as it is in contact with the 1x PBS in the vibratome buffer bath.</li>
		<li>Insert the specimen disc with the specimen into the buffer tray using the manipulator of the microtome.</li>
		<li>Use the manipulator to rotate the specimen disc into the desired position. Orient the block in the buffer bath in a way that locates the telencephalon just under the surface, with the dorsal part of the brain facing the blade. Then tighten the scrw with a 3 mm Allen key and remove the manipulator. Note: The clamping screw or one of the clamping devices must not be located over the gap in the specimen disc; in these positions clamping the specimen disc is not possible.</li>
	</ol>
	</li>
	<li>Sectioning the Brain
	<ol>
		<li>Prepare a 24-well plate to collect the sections. Per brain to be sectioned, fill one well with 1 ml of blocking buffer (0.1% (v/v) 10% Tween-20, 0.2% (w/v) bovine serum albumin (BSA), 1% (v/v) dimethyl sulfoxide (DMSO) in PBS).</li>
		<li>Start sectioning at 50 &#956;m thickness, 1 mm/sec speed, and 70 Hz frequency using a vibrating microtome.</li>
		<li>Use a synthetic brush (1 mm) to pick up the thin slices of agarose as they come off the blade and collect them in the 24-well plate.</li>
	</ol>
	</li>
	<li>Immunohistochemistry on Vibratome Sections
	<ol>
		<li>Block non-specific binding sites for 1 hr at RT in blocking buffer (0.1% (v/v) 10% Tween 20, 0.2% (w/v) BSA, 1% (v/v) DMSO in PBS).</li>
		<li>Remove the supernatant and add 250 &#956;l of antibody diluted in blocking buffer overnight at 4 &#176;C. Alternatively, incubate the primary antibody for 2 hr at RT. Then wash 3x for 10 min in PTW. Note: For PCNA staining use a 1:400 dilution (mouse anti-PCNA. For details see table of materials). Multiple primary antibodies can be incubated at the same time.</li>
		<li>Incubate the sections in secondary antibody for 2 hr at RT then wash 3x for 10 min in PTW. Note: 1:1,000 dilution in PTW, for detail see table of materials. Multiple secondary antibodies can be incubated at the same time.</li>
		<li>Optional: The best results are observed when sections are kept in PTW overnight at 4 &#176;C in order to allow washing out of non-specifically bound secondary antibodies. In this case, replace the PTW once again before mounting.</li>
	</ol>
	</li>
	<li>Adult Brain RNA in situ&#160;Hybridization (alternatively to immunohistochemistry)
	<ol>
		<li>Brain Dissection and Fixation (same procedure as for immunohistochemistry):
		<ol>
			<li>Hybridization of probe.</li>
			<li>Rehydrate brains in 2 ml reaction tubes as described for immunohistochemistry.</li>
			<li>Incubate brains in proteinase K (ProtK; 10 &#956;g/ml final conc.) in PTW for 30 min. After incubation time remove ProtK/PTW and fix the brains again for 20 min in BT-Fix (4% PFA, 4% sucrose, 0.12 mM CaCl2, 77 mM Na2HPO4, 23 mM NaH2PO4).</li>
			<li>Wash brains 5x for 5 min in PTW (0.1% (v/v) Tween-20 in PBS).</li>
			<li>Remove PTW and rinse the brains in 500 &#956;l HYB buffer (50% (v/v) deionized formamide, 5x SSC, 500 &#956;g/ml yeast tRNA, 50 &#956;g/ml heparin, 0.1% Tween-20, 9 mM citricacid). Wait until brains sink to the bottom of the tube then discard the supernatant and add 500 &#956;l of clean HYB buffer and incubate at 65-70 &#176;C for 3-4 hr.</li>
			<li>Dilute the RNA probe to 1:200 or 1:400 (depending on the probe) in 400 &#956;l final volume of HYB buffer. Heat the diluted RNA probe at 70 &#176;C for 5 min.</li>
			<li>Remove supernatant from the brains and add the RNA probe, incubate at 65-70 &#176;C (O/N). All wash buffer solutions should be prewarmed and kept at 70 &#176;C.&#160;</li>
		</ol>
		</li>
		<li>Labeling
		<ol>
			<li>Wash the brains at 65-70 &#176;C (during all 4 washing steps) with 500 &#956;l of wash buffer 1 (50% formamide, 1x SSC, 0.05% Tween 20) twice for 30 min. Replace buffer 1 by buffer 2 (2x SSC, 0.1% Tween 20), and incubate for 15 min. Remove buffer 2 and wash brains in buffer 3 (0.2x SSC, 0.1% Tween 20) for 30 min, repeat this step 2 twice. Replace buffer 3 by buffer 4 (0.1x SSC, 0.05% Tween 20 in PTW), incubate for 5 min.</li>
			<li>Remove supernatant and wash for 5 min in 500 &#956;l of PTW at RT.</li>
			<li>Embed brains in 2% agarose (in 1x PBS) and cut vibratome sections (50-100 &#956;m) as described for immunohistochemistry.</li>
			<li>Wash 1x for 5 min in 500 &#956;l blocking buffer (1x PBS, 0.1% Tween-20, 0.2% (w/v) BSA, 1% (v/v) DMSO).</li>
			<li>Remove supernatant and add 500 &#956;l blocking buffer for 2 hr at RT.</li>
			<li>Remove supernatant and add 300 &#956;l anti-Dig AP-coupled antibody diluted 1:4,000 in blocking buffer.</li>
			<li>Incubate o/n at 4 &#176;C.</li>
		</ol>
		</li>
		<li>NBT/BCIP Staining
		<ol>
			<li>Wash 5x for 15 min with PTW buffer at RT.</li>
			<li>Rinse 1x in staining buffer. Wash twice 5 min in staining buffer (100 mM Tris/HCl pH 9.5, 50 mM MgCl2, 100 mM NaCl, 0.1% Tween-20).</li>
			<li>Remove supernatant and stain with 500 &#956;l staining solution (3.5 &#956;l of 4-Nitroblue tetrazolium chloride (NBT) and 5-Bromo-4-chloro-3-indolyl-phosphate (BCIP) per 1 ml staining buffer).</li>
			<li>Stop staining by rinsing 3x to 5x with PTW.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Mounting of Sections
	<ol>
		<li>Mount the sections on slides using a water-soluble, non-fluorescent mounting medium (see the&#160;Materials table).</li>
		<li>Keep slides in the fridge at 4 &#176;C. Note: The staining is stable for several weeks or even months.</li>
		<li>Analyze the slides under a compound or confocal microscope.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

We describe here a method to induce a telencephalic injury by pushing a needle through the skull of the adult zebrafish, and to analyze the cellular and molecular reaction to this injury by immunohistochemistry and in situ hybridization. The main advantage of this technique over other existing approaches is its simplicity, speed and cost efficiency. Therefore, this technique, which allows production of many injured brains in a short time, can be easily combined with a large scale in situ hybridization screen to identify genes with a role in regeneration and neurogenesis.
The age of the adult zebrafish on which the injury is performed is crucial. If the fish are older than 1.5 years, the skull is too hard and it is possible that the skull is being pushed down or fractured rather than cleanly perforated. In contrast, in younger fish (younger than 4 months), there is still a lot of neurogenesis going on, which might lead to false positive results regarding reactive neurogenesis.
Another critical step in this approach is to verify the efficiency of the stab wound and to exclude that both hemispheres are wounded. This can be monitored by staining the sections with the anti-PCNA antibody. In a correctly introduced lesion, PCNA should be significantly up-regulated in only one of the hemispheres.
It is important to bear in mind that the induced mechanical injury will have several side effects such as non-specific cell ablation, increased cell death because of secondary degeneration, inflammation and destruction of the blood-brain barrier, and damage of ventricular zone and as a possible consequence flow of the cerebrospinal fluid into the brain parenchyma. However, the injured fish appear to suffer very little from the injury. Within 2 min&#160;they are free swimming again and normal feeding behavior is restored within about 4 hr. After 3 weeks the injury is morphologically not visible anymore. In our hands, from several hundred surgeries we did not lose a single fish despite the fact that we had a 100% injury rate as assessed by histology at 5 days post-lesion.
In the last years other techniques such as transgenic approaches have been developed in zebrafish to produce tissue or cell type specific ablation2. Although these techniques are very elegant and minimally invasive they are based on the generation of complicated DNA constructs and the establishment of several stable zebrafish transgenic lines which can be very tedious and time consuming. Therefore, simple mechanical approaches such as the assay described in this protocol remain an important tool to study adult brain neurogenesis and regeneration.

Mammals have a very limited capacity to generate new neurons during adult life, and adult neurogenesis in the brain is mostly restricted to two telencephalic regions situated at the subventricular zone (SVZ) of the lateral ventricles, and the subgranular zone (SGZ) of the dentate gyrus in the hippocampus1. Mammals also cannot repair damage of the brain caused by neurodegenerative diseases, stroke, or injuries, efficiently. Lost neurons are not replaced and instead proliferation of different types of glial cells, including astrocytes, microglia, and oligodendrocytes, is observed. As a consequence of this proliferative process called reactive gliosis, the injured brain tissue is sealed off by the formation of a glial scar, which inhibits neuronal and axonal regeneration2-4.
In contrast to mammals, teleost fish like the zebrafish form new neurons abundantly in adult stages and have a high regenerative and proliferative potential to repair lesions of the central nervous system2,5-7. The zebrafish adult brain contains 16 distinct neural stem cell niches that generate a large number of new neurons during the entire life8-11. Neurons born in these specific proliferation zones of the adult zebrafish brain migrate to their target areas, where they will mature and integrate into the existing neuronal networks8,10,12-15.
Among the progenitor zones identified in adult zebrafish, the telencephalic ventricular zone is one of the most studied neuronal stem cell niches. Cells in this progenitor niche can be classified into three distinct types regarding their morphology, division rate and expression of different glial or neuronal markers. Type I and type II cells are radial glial like cells which have long processes. They express stem cells markers including GFAP, nestin, and S100&#946;. Type I cells do not proliferate while type II cells proliferate slowly, as evidenced by their expression of proliferation markers such as proliferating cell nuclear antigen (PCNA) and the incorporation of desoxythymidine analogues. The type III cells are fast dividing cells that are committed to become neural progenitors and express neural marker genes, such as PSA-NCAM5,16.
Although the regenerative capacities of teleost fish are well documented, only few publications have described and investigated in detail the cellular and molecular mechanisms involved in neuronal regeneration in the adult telencephalon in response to injury15,17-22. To decipher the mechanisms implicated in zebrafish adult central nervous system regeneration and repair and to understand the similarities and the differences between constitutive and regenerative neurogenesis, we developed a zebrafish model of adult telencephalic injury. Here, we will describe visually how to generate an injury in one of the telencephalic hemispheres with the help of a syringe needle, while keeping the contralateral unlesioned side as an internal control. We will also show how to monitor changes upon injury in cell proliferation and gene expression by immunohistochemistry and in situ hybridization assays.

<table><tbody><tr><td>Tricaine</td><td>Sigma</td><td>335.2</td><td></td></tr><tr><td>10x DPBS (-/-)</td><td>Life technologies</td><td>14200-083</td><td></td></tr><tr><td>30 G syringe (Omnican 40)</td><td>B.Braun Melsungen</td><td>916162</td><td></td></tr><tr><td>Polyethylene molding cup tray&amp;nbsp;</td><td>Polysciences, Inc</td><td>17177C-3</td><td></td></tr><tr><td>PeqGOLD Universal Agarose&amp;nbsp;</td><td>Peqlab</td><td>35-1020</td><td></td></tr><tr><td>Bovine serum albumin</td><td>Biochrom</td><td>W 2835919108</td><td></td></tr><tr><td>Aqua Poly/Mount</td><td>Polysciences, Inc</td><td>18606-20</td><td></td></tr><tr><td>Tween 20</td><td>Carl Roth</td><td>9127.1</td><td></td></tr><tr><td>DMSO</td><td>Carl Roth</td><td>A994.2</td><td></td></tr><tr><td>Dissection needle</td><td>Carl Roth</td><td>KX93.1</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Carl Roth</td><td>335.2</td><td></td></tr><tr><td>Loctite 414</td><td>Sch&amp;ouml;ffler und W&amp;ouml;rner</td><td>&lt;a href=&quot;http://62.157.117.162/BP2NAV500/ItemDetail_NAV.aspx?ItemID=06%201362%200020&quot;&gt;06 1362 0020&lt;/a&gt;</td><td></td></tr><tr><td>Glass slides</td><td>Labonord</td><td>5305103</td><td></td></tr><tr><td>Cover slips 24 x 60 mm</td><td>Labonord</td><td>5305060</td><td></td></tr><tr><td>Petri dishes, 94 mm</td><td>Greiner bio-one</td><td>632180</td><td></td></tr><tr><td>Falcon tubes, 15 ml</td><td>Greiner bio-one</td><td>188261</td><td></td></tr><tr><td>24-well plate</td><td>Greiner bio-one</td><td>662160</td><td></td></tr><tr><td>Anti-S100 (1:500)</td><td>Dako</td><td>Z031101-2</td><td></td></tr><tr><td>Anti-PCNA (1:400)</td><td>Dako</td><td>M087901</td><td></td></tr><tr><td>Anti-GFP (1:1,000)</td><td>Aves Labs</td><td>GFP-1020</td><td></td></tr><tr><td>anti-Dig AP-coupled antibody&amp;nbsp;</td><td>Roche Diagnostics</td><td>11093274910</td><td></td></tr><tr><td>5-Bromo-4-chloro-3-indolyl-phosphate (BCIP)&amp;nbsp;</td><td>Roche Diagnostics</td><td>1383221</td><td></td></tr><tr><td>4-Nitroblue tetrazolium chloride (NBT)&amp;nbsp;</td><td>Roche Diagnostics</td><td>1383213</td><td></td></tr><tr><td>Proteinase K</td><td>Roche Diagnostics</td><td>1092766</td><td></td></tr><tr><td>Polyethylene molding cup tray&amp;nbsp;</td><td>Polysciences</td><td>17177C-3</td><td></td></tr><tr><td>Vibratome VT1000S</td><td>Leica</td><td></td><td></td></tr><tr><td>Confocal microscope Sp5 DM6000</td><td>Leica</td><td></td><td></td></tr><tr><td>Dissecting microscope Nikon SMZ 645</td><td>Nikon</td><td></td><td></td></tr><tr><td>24-well flat bottomed tissue culture plates&amp;nbsp;</td><td>Falcon catalog&amp;nbsp;</td><td>353226</td><td></td></tr><tr><td>Student Dumont #5 Forceps</td><td>Fine Science Tools</td><td>91150-20</td><td></td></tr><tr><td>Surgical scissors</td><td>Fine Science Tools</td><td>91460-11</td><td></td></tr><tr><td>&lt;em&gt;Danio rerio Tg(olig2:EGFP)&lt;/em&gt;</td><td>Shin &lt;em&gt;et al.&lt;/em&gt;&lt;sup&gt;24&lt;/sup&gt;</td><td></td><td></td></tr><tr><td>&lt;em&gt;Danio rerio&lt;/em&gt; WT strains (&lt;em&gt;e.g.&lt;/em&gt;, AB2O2)</td><td></td><td></td><td></td></tr></tbody></table>

stab wound injury of the zebrafish adult telencephalon: a method to investigate vertebrate brain neurogenesis and regeneration

Adult zebrafish have an amazing capacity to regenerate their central nervous system after injury. To investigate the cellular response and the molecular mechanisms involved in zebrafish adult central nervous system (CNS) regeneration and repair, we developed a zebrafish model of adult telencephalic injury.
In this approach, we manually generate an injury by pushing an insulin syringe needle into the zebrafish adult telencephalon. At different post injury days, fish are sacrificed, their brains are dissected out and stained by immunohistochemistry and/or in situ hybridization (ISH) with appropriate markers to observe cell proliferation, gliogenesis, and neurogenesis. The contralateral unlesioned hemisphere serves as an internal control. This method combined for example with RNA deep sequencing can help to screen for new genes with a role in zebrafish adult telencephalon neurogenesis, regeneration, and repair.

Using the workflow described in this article, a lesion was introduced into the right hemisphere of an adult zebrafish telencephalon (Figures&#160;1A-1B). A correct wounding leads to a lesion canal that extends from dorsal to ventral through the whole pallium of the telencephalon (Figure 3C). The lesion should be restricted to one of the hemispheres and should not cross the medial ventricle. The lesion canal is visible with bright field microscopy (Figure 3C) and will be healed after approximately 35 days15.
It was previously shown that after wounding an up-regulation of proliferating cell nuclear antigen (PCNA; Figure 3A) and S100&#946; (Figure 3B) at 3 to 7 dpl is detectable by immunohistochemistry15. The staining of PCNA and S100&#946; is overlapping, indicating proliferation of radial glial cells.
Furthermore, in order to visualize oligodendrocyte precursor cells (OPCs), a stab wound assay was performed on the Tg(olig2:EGFP) fish24. Upon staining with an anti-GFP antibody, accumulation of OPCs in close proximity to the lesion canal is detectable (Figure 3D). This accumulation is transient, and OPC clusters are not observed anymore at 35 dpl15.
If the lesion was not introduced properly, the lesion canal will not be visible, and up-regulation of PCNA and S100&#946; or accumulation of OPCs will not be detectable. PCNA is the most sensitive marker for brain injury. In order to verify the efficiency of the stab wound and to exclude that both hemispheres are wounded, it is strongly recommended to always stain the sections with an anti-PCNA antibody. In a correctly introduced lesion, PCNA should be significantly up-regulated (up to 4-fold15) in only one of the hemispheres. Although in most cases it is sufficient to only use the contralateral unlesioned side as an internal control to monitor changes in gene expression, it is highly recommended to compare the gene expression in the control hemisphere with the expression in telencephalic hemispheres of wild type brains. In this way, any systemic effects of the lesion on gene expression that would affect both hemispheres can be detected.
The stab wound assay in combination with a systematic expression screen of transcription regulators (TRs) (25 and Schmidt et al., unpublished) can also be used to identify TR genes with a possible role in neurogenesis and regeneration (Figures 4A-4C).
This approach has identified a number of factors that are expressed in the telencephalon and whose expression is specifically up-regulated in response to injury (Figure 4A-C).
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51753/51753fig1highres.jpg" width="500" /> 
Figure 1. Schematic representation of adult zebrafish telencephalic stab wounding. A) An insulin syringe needle is pushed through the skull of an adult zebrafish to generate a telencephalic injury. The white dashed line represents the border between the two telencephalic hemispheres and the green cross indicates the position of the stab wound. B) Dorsal view of an adult zebrafish brain. The major brain areas are indicated. The green cross shows the position of the induced injury. C) The tip of the needle, which serves to induce the telencephalic injury, measures 2 mm. Olfactory bulb (ob), telencephalon (tel) and optic tectum (ot).
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51753/51753fig2highres.jpg" width="500" /> 
Figure 2. Schematic representation of an adult zebrafish brain in a trimmed agarose block just before vibratome sectioning. Anterior is up. Olfactory bulb (ob), telencephalon (tel), optic tectum (ot), cerebellum (cb) and medulla (m). Scale bar: 500 &#956;m.
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/51753/51753fig3highres.jpg" width="500" /> 
Figure 3. Regenerative responses to stab wound injury. A-D) Adult zebrafish brain at 5 days post lesion (dpl). A-B) The proliferative marker PCNA&#160;(A) and the radial glial marker S100&#946; (B)&#160;are up-regulated at the ventricular zone (arrows) upon stab wounding. (C)&#160;The lesion channel is visible under bright field illumination (arrows). The ventricular zone is indicated by the green line. (D)&#160;Oligodendrocyte precursor cells (olig2:EGFP+) accumulate at the site of lesion (arrows). Scale bar: 200 &#956;m. In all panels, the right hemisphere is the lesioned hemisphere. Dorsal is up. The dashed line in A to D indicates the border between the pallium and the subpallium.
<img alt="Figure 4" fo:content-width="6in" src="/files/ftp_upload/51753/51753fig4highres.jpg" width="600" /> 
Figure 4. An example of genes up-regulated in response to stab injury. Note the strong up-regulation (arrows) of eomesa&#160;(A, B)&#160;and sox4a&#160;(C)&#160;mRNA expression in the right telencephalic hemisphere (stabbed hemisphere) as monitored by in situ hybridization at 5 dpl. The left hemisphere is uninjured and serves as control. (B) Shows a higher magnification of the dorsal telencephalic region of A. Dorsal is up. Scale bar: 200 &#956;m for A and C and 55 &#181;m for B. The dashed line in A and C indicates the separation between the pallium and the subpallium. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51753/51753fig4highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table>
	<tbody>
		<tr>
			<td colspan="2">ABBREVIATIONS</td>
		</tr>
		<tr>
			<td>AP</td>
			<td>Alkaline Phosphatase</td>
		</tr>
		<tr>
			<td>BCIP</td>
			<td>5-bromo-4-chloro-3&#39;-indolyphosphate</td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Bovine serum albumin</td>
		</tr>
		<tr>
			<td>CNS</td>
			<td>Central nervous system</td>
		</tr>
		<tr>
			<td>Dig</td>
			<td>Digoxygenin</td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Dimethyl sulfoxide</td>
		</tr>
		<tr>
			<td>dpl</td>
			<td>Days post lesion</td>
		</tr>
		<tr>
			<td>eomesa</td>
			<td>Eomesodermin homolog a</td>
		</tr>
		<tr>
			<td>GFAP</td>
			<td>Glial fibrillary acidic protein</td>
		</tr>
		<tr>
			<td>hr</td>
			<td>Hour</td>
		</tr>
		<tr>
			<td>HYB buffer</td>
			<td>Hybridization buffer</td>
		</tr>
		<tr>
			<td>ISH</td>
			<td>in situ hybridization</td>
		</tr>
		<tr>
			<td>MeOH</td>
			<td>Methanol</td>
		</tr>
		<tr>
			<td>NBT</td>
			<td>Nitro-blue tetrazolium</td>
		</tr>
		<tr>
			<td>OPCs</td>
			<td>Oligodendrocyte precursor cells</td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Phosphate buffered saline</td>
		</tr>
		<tr>
			<td>PCNA</td>
			<td>Proliferating cell nuclear antigen</td>
		</tr>
		<tr>
			<td>PFA</td>
			<td>Paraformaldehyde</td>
		</tr>
		<tr>
			<td>ProtK</td>
			<td>Proteinase K</td>
		</tr>
		<tr>
			<td>PSA-NCAM</td>
			<td>Polysialylated neuronal cell adhesion molecule</td>
		</tr>
		<tr>
			<td>PTW</td>
			<td>PBS, 0.1%&#160;Tween-20</td>
		</tr>
		<tr>
			<td>RNA</td>
			<td>Ribonucleic acid</td>
		</tr>
		<tr>
			<td>RT</td>
			<td>Room temperature</td>
		</tr>
		<tr>
			<td>SGZ</td>
			<td>Subgranular zone</td>
		</tr>
		<tr>
			<td>sox4a</td>
			<td>SRY-box containing gene 4a</td>
		</tr>
		<tr>
			<td>SSC</td>
			<td>Saline-sodium citrate</td>
		</tr>
		<tr>
			<td>SVZ</td>
			<td>Subventricular zone</td>
		</tr>
		<tr>
			<td>TRs</td>
			<td>Transcription regulators</td>
		</tr>
	</tbody>
</table>

Watch this Scientific Journal Video about Stab Wound Injury of the Zebrafish Adult Telencephalon: A Method to Investigate Vertebrate Brain Neurogenesis and Regeneration at JoVE.com

Stab Wound Injury of the Zebrafish Adult Telencephalon: A Method to Investigate Vertebrate Brain Neurogenesis and Regeneration

To shed light on the cellular and molecular mechanisms of zebrafish adult neurogenesis and regeneration, we developed a protocol for invasive surgery causing mechanical injuries in the zebrafish adult telencephalon and subsequent monitoring of changes in the stabbed hemisphere by immunohistochemistry or in situ hybridization.

Adult zebrafish have an amazing capacity to regenerate their central nervous system after injury. To investigate the cellular response and the molecular ...

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Unit 1

The Nervous System and Nervous Tissue

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15.3K Views.  Karlsruhe Institute of Technology. To shed light on the cellular and molecular mechanisms of zebrafish adult neurogenesis and regeneration, we developed a protocol for invasive surgery causing mechanical injuries in the zebrafish adult telencephalon and subsequent monitoring of changes in the stabbed hemisphere by immunohistochemistry or in situ hybridization.

Video: Stab Wound Injury of the Zebrafish Adult Telencephalon: A Method to Investigate Vertebrate Brain Neurogenesis and Regeneration

Mechanical Vessel Injury in Zebrafish Embryos

Zebrafish (Danio rerio) embryos have proven to be a powerful model for studying a variety of developmental and disease processes. External development and optical transparency make these embryos especially amenable to microscopy, and numerous transgenic lines that label specific cell types with fluorescent proteins are available, making the zebrafish embryo an ideal system for visualizing the interaction of vascular, hematopoietic, and other cell types during injury and repair in vivo. Forward and reverse genetics in zebrafish are well developed, and pharmacological manipulation is possible. We describe a mechanical vascular injury model using micromanipulation techniques that exploits several of these features to study responses to vascular injury including hemostasis and blood vessel repair. Using a combination of video and timelapse microscopy, we demonstrate that this method of vascular injury results in measurable and reproducible responses during hemostasis and wound repair. This method provides a system for studying vascular injury and repair in detail in a whole animal model.

This article describes a method for creating a mechanical vessel injury in zebrafish embryos. This injury model provides a platform for studying hemostasis, injury-related inflammation, and wound healing in an organism ideally suited for real-time microscopy.

Zebrafish (Danio rerio) embryos have proven to be a powerful model for studying a variety of developmental and disease processes.  External development ...

Developmental Biology

Zebrafish Keratocyte Explants to Study Collective Cell Migration and Reepithelialization in Cutaneous Wound Healing

Due to their unique motile properties, fish keratocytes dissociated from explant cultures have long been used to study the mechanisms of single cell migration. However, when explants are established, these cells also move collectively, maintaining many of the features which make individual keratocytes an attractive model to study migration: rapid rates of motility, extensive actin-rich lamellae with a perpendicular actin cable, and relatively constant speed and direction of migration. In early explants, the rapid interconversion of cells migrating individually with those migrating collectively allows the study of the role of cell-cell adhesions in determining the mode of migration, and emphasizes the molecular links between the two modes of migration. Cells in later explants lose their ability to migrate rapidly and collectively as an epithelial to mesenchymal transition occurs and genes associated with wound healing and inflammation are differentially expressed. Thus, keratocyte explants can serve as an in vitro model for the reepithelialization that occurs during cutaneous wound healing and can represent a unique system to study mechanisms of collective cell migration in the context of a defined program of gene expression changes. A variety of mutant and transgenic zebrafish lines are available, which allows explants to be established from fish with different genetic backgrounds. This allows the role of different proteins within these processes to be uniquely addressed. The protocols outlined here describe an easy and effective method for establishing these explant cultures for use in a variety of assays related to collective cell migration.

Zebrafish keratocytes migrate in cell sheets from explants and provide an in vitro model for the study of the mechanisms of collective cell migration in the context of epithelial wound healing. These protocols detail an effective way to establish primary explant cultures for use in collective cell migration assays.

Due to their unique motile properties, fish keratocytes dissociated from explant cultures have long been used to study the mechanisms of single cell ...

Motor Nerve Transection and Time-lapse Imaging of Glial Cell Behaviors in Live Zebrafish

The nervous system is often described as a hard-wired component of the body even though it is a considerably fluid organ system that reacts to external stimuli in a consistent, stereotyped manner, while maintaining incredible flexibility and plasticity. Unlike the central nervous system (CNS), the peripheral nervous system (PNS) is capable of significant repair, but we have only just begun to understand the cellular and molecular mechanisms that govern this phenomenon. Using zebrafish as a model system, we have the unprecedented opportunity to couple regenerative studies with in vivo imaging and genetic manipulation. Peripheral nerves are composed of axons surrounded by layers of glia and connective tissue. Axons are ensheathed by myelinating or non-myelinating Schwann cells, which are in turn wrapped into a fascicle by a cellular sheath called the perineurium. Following an injury, adult peripheral nerves have the remarkable capacity to remove damaged axonal debris and re-innervate targets. To investigate the roles of all peripheral glia in PNS regeneration, we describe here an axon transection assay that uses a commercially available nitrogen-pumped dye laser to axotomize motor nerves in live transgenic zebrafish. We further describe the methods to couple these experiments to time-lapse imaging of injured and control nerves. This experimental paradigm can be used to not only assess the role that glia play in nerve regeneration, but can also be the platform for elucidating the molecular mechanisms that govern nervous system repair.

Although the peripheral nervous system (PNS) is capable of significant repair after injury, little is known about the cellular and molecular mechanisms that govern this phenomenon. Using live, transgenic zebrafish and a reproducible nerve transection assay, we can study dynamic glial cell behaviors during nerve degeneration and regeneration.

The  nervous system is often described as a hard-wired component of the body even  though it is a considerably fluid organ system that reacts to external ...

Electroporation Method for In Vivo Delivery of Plasmid DNA in the Adult Zebrafish Telencephalon

Electroporation is a transfection method in which an electrical field is applied to cells to create temporary pores in a cell membrane and increase its permeability, thereby allowing different molecules to be introduced to the cell. In this paper, electroporation is used to introduce plasmids to ependymoglial cells, which line the ventricular zone of the adult zebrafish telencephalon. A fraction of these cells shows stem cell properties and generates new neurons in the zebrafish brain; therefore, studying their behavior is essential to determine their roles in neurogenesis and regeneration. The introduction of plasmids via electroporation enables long-term labeling and tracking of a single ependymoglial cell. Furthermore, plasmids such as Cre recombinase or Cas9 can be delivered to single ependymoglial cells, which enables gene recombination or gene editing and provides a unique opportunity to assess the cell&#8217;s autonomous gene function in a controlled, natural environment. Finally, this detailed, step-by-step electroporation protocol is used to obtain successful introduction of plasmids into a large number of single ependymoglial cells.

Presented here is an electroporation method for plasmid DNA delivery and ependymoglial cell labeling in the adult zebrafish telencephalon. This protocol is a quick and efficient method to visualize and trace individual ependymoglial cells and opens up new possibilities to apply electroporation to a broad range of genetic manipulations.

Electroporation is a transfection method in which an electrical field is applied to cells to create temporary pores in a cell membrane and increase its ...

Spinal Cord Transection in the Larval Zebrafish

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

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

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

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.

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 ...

Modeling Amyloid-&#946;42 Toxicity and Neurodegeneration in Adult Zebrafish Brain

Alzheimer's disease (AD) is a debilitating neurodegenerative disease in which accumulation of toxic amyloid-&#946;42 (A&#946;42) peptides leads to synaptic degeneration, inflammation, neuronal death, and learning deficits. Humans cannot regenerate lost neurons in the case of AD in part due to impaired proliferative capacity of the neural stem/progenitor cells (NSPCs) and reduced neurogenesis. Therefore, efficient regenerative therapies should also enhance the proliferation and neurogenic capacity of NSPCs. Zebrafish (Danio rerio) is a regenerative organism, and we can learn the basic molecular programs with which we could design therapeutic approaches to tackle AD. For this reason, the generation of an AD-like model in zebrafish was necessary. Using our methodology, we can introduce synthetic derivatives of A&#946;42 peptide with tissue penetrating capability into the adult zebrafish brain, and analyze the disease pathology and the regenerative response. The advantage over the existing methods or animal models is that zebrafish can teach us how a vertebrate brain can naturally regenerate, and thus help us to treat human neurodegenerative diseases better by targeting endogenous NSPCs. Therefore, the amyloid-toxicity model established in the adult zebrafish brain may open new avenues for research in the field of neuroscience and clinical medicine. Additionally, the simple execution of this method allows for cost-effective and efficient experimental assessment. This manuscript describes the synthesis and injection of A&#946;42 peptides into zebrafish brain.

This protocol describes the synthesis, characterization, and injection of monomeric amyloid-&#946;42 peptides for generating amyloid toxicity in adult zebrafish to establish an Alzheimer&#39;s disease model, followed by histological analyses and detection of aggregations.

Alzheimer's disease (AD) is a debilitating neurodegenerative disease in which accumulation of toxic amyloid-&#946;42 (A&#946;42) peptides leads to ...

A Scalable Model to Study the Effects of Blunt-Force Injury in Adult Zebrafish

Blunt-force traumatic brain injuries (TBI) are the most common form of head trauma, which spans a range of severities and results in complex and heterogenous secondary effects. While there is no mechanism to replace or regenerate the lost neurons following a TBI in humans, zebrafish possess the ability to regenerate neurons throughout their body, including the brain. To examine the breadth of pathologies exhibited in zebrafish following a blunt-force TBI and to study the mechanisms underlying the subsequent neuronal regenerative response, we modified the commonly used rodent Marmarou weight drop for the use in adult zebrafish. Our simple blunt-force TBI model is scalable, inducing a mild, moderate, or severe TBI, and recapitulates many of the phenotypes observed following human TBI, such as contact- and post-traumatic seizures, edema, subdural and intracerebral hematomas, and cognitive impairments, each displayed in an injury severity-dependent manner. TBI sequelae, which begin to appear within minutes of the injury, subside and return to near undamaged control levels within 7 days post-injury. The regenerative process begins as early as 48 hours post-injury (hpi), with the peak cell proliferation observed by 60 hpi. Thus, our zebrafish blunt-force TBI model produces characteristic primary and secondary injury TBI pathologies similar to human TBI, which allows for investigating disease onset and progression, along with the mechanisms of neuronal regeneration that is unique to zebrafish.

We modified the Marmarou weight drop model for adult zebrafish to examine a breadth of pathologies following blunt-force traumatic brain injury (TBI) and the mechanisms underlying subsequent neuronal regeneration. This blunt-force TBI model is scalable, induces a mild, moderate, or severe TBI, and recapitulates injury heterogeneity observed in human TBI.

Blunt-force traumatic brain injuries (TBI) are the most common form of head trauma, which spans a range of severities and results in complex and ...

Assessment of Swim Endurance and Swim Behavior in Adult Zebrafish

Due to their renowned regenerative capacity, adult zebrafish are a premier vertebrate model to interrogate mechanisms of innate spinal cord regeneration. Following complete transection of their spinal cord, zebrafish extend glial and axonal bridges across severed tissue, regenerate neurons proximal to the lesion, and regain their swim capacities within 8 weeks of injury. Recovery of swim function is thus a central readout for functional spinal cord repair. Here, we describe a set of behavioral assays to quantify zebrafish motor capacity inside an enclosed swim tunnel. The goal of these methods is to provide quantifiable measurements of swim endurance and swim behavior in adult zebrafish. For swim endurance, zebrafish are subjected to a constantly increasing water current velocity until exhaustion, and time at exhaustion is reported. For swim behavior assessment, zebrafish are subjected to low current velocities and swim videos are captured with a dorsal view of the fish. Percent activity, burst frequency, and time spent against the water current provide quantifiable readouts of swim behavior. We quantified swim endurance and swim behavior in wild-type zebrafish before injury and after spinal cord transection. We found that zebrafish lose swim function after spinal cord transection and gradually regain that capacity between 2 and 6 weeks post-injury. The methods described in this study could be applied to neurobehavioral, musculoskeletal, skeletal muscle regeneration, and neural regeneration studies in adult zebrafish.

Capable of functional recovery after spinal cord injury, adult zebrafish is a premier model system to elucidate innate mechanisms of neural regeneration. Here, we describe swim endurance and swim behavior assays as functional readouts of spinal cord regeneration.

Due to their renowned regenerative capacity, adult zebrafish are a premier vertebrate model to interrogate mechanisms of innate spinal cord regeneration. ...

Controlled Semi-Automated Laser-Induced Injuries for Studying Spinal Cord Regeneration in Zebrafish Larvae

Zebrafish larvae possess a fully functional central nervous system (CNS) with a high regenerative capacity only a few days after fertilization. This makes this animal model very useful for studying spinal cord injury and regeneration. The standard protocol for inducing such lesions is to transect the dorsal part of the trunk manually. However, this technique requires extensive training and damages additional tissues. A protocol was developed for laser-induced lesions to circumvent these limitations, allowing for high reproducibility and completeness of spinal cord transection over many animals and between different sessions, even for an untrained operator. Furthermore, tissue damage is mainly limited to the spinal cord itself, reducing confounding effects from injuring different tissues, e.g., skin, muscle, and CNS. Moreover, hemi-lesions of the spinal cord are possible. Improved preservation of tissue integrity after laser injury facilitates further dissections needed for additional analyses, such as electrophysiology. Hence, this method offers precise control of the injury extent that is unachievable manually. This allows for new experimental paradigms in this powerful model in the future.

The present protocol describes a method to induce tissue-specific and highly reproducible injuries in zebrafish larvae using a laser lesion system combined with an automated microfluidic platform for larvae handling.

Zebrafish larvae possess a fully functional central nervous system (CNS) with a high regenerative capacity only a few days after fertilization. This makes ...

Stab Wound Injury of the Zebrafish Adult Telencephalon: A Method to Investigate Vertebrate Brain Neurogenesis and Regeneration

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