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.

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			<td height="21" style="height:21px;width:333px;">Name of Material/ Equipment</td>
			<td style="width:163px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
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			<td height="21" style="height:21px;">Tricaine</td>
			<td>Sigma</td>
			<td align="right">335.2</td>
			<td></td>
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			<td height="21" style="height:21px;">10x DPBS (-/-)</td>
			<td>Life technologies</td>
			<td>14200-083</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">30 G syringe (Omnican 40)</td>
			<td>B.Braun Melsungen</td>
			<td align="right">916162</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Polyethylene molding cup tray&#160;</td>
			<td>Polysciences, Inc</td>
			<td>17177C-3</td>
			<td></td>
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			<td height="21" style="height:21px;">PeqGOLD Universal Agarose&#160;</td>
			<td>Peqlab</td>
			<td>35-1020</td>
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			<td height="21" style="height:21px;">Bovine serum albumin</td>
			<td>Biochrom</td>
			<td>W 2835919108</td>
			<td></td>
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			<td height="21" style="height:21px;">Aqua Poly/Mount</td>
			<td>Polysciences, Inc</td>
			<td>18606-20</td>
			<td></td>
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			<td height="21" style="height:21px;">Tween 20</td>
			<td>Carl Roth</td>
			<td align="right">9127.1</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">DMSO</td>
			<td>Carl Roth</td>
			<td>A994.2</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Dissection needle</td>
			<td>Carl Roth</td>
			<td>KX93.1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Paraformaldehyde</td>
			<td>Carl Roth</td>
			<td align="right">335.2</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Loctite 414</td>
			<td>Sch&#246;ffler und W&#246;rner</td>
			<td><a href="http://62.157.117.162/BP2NAV500/ItemDetail_NAV.aspx?ItemID=06%201362%200020">06 1362 0020</a></td>
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			<td height="21" style="height:21px;">Glass slides</td>
			<td>Labonord</td>
			<td align="right">5305103</td>
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			<td height="21" style="height:21px;">Cover slips 24 x 60 mm</td>
			<td style="width:163px;">Labonord</td>
			<td align="right">5305060</td>
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			<td height="21" style="height:21px;">Petri dishes 94 mm</td>
			<td>Greiner bio-one</td>
			<td align="right">632180</td>
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			<td height="21" style="height:21px;">Falcons 15 ml</td>
			<td style="width:163px;">Greiner bio-one</td>
			<td align="right">188261</td>
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			<td height="21" style="height:21px;">24-well plate</td>
			<td>Greiner bio-one</td>
			<td align="right">662160</td>
			<td></td>
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			<td height="21" style="height:21px;">Anti-S100 (1:500)</td>
			<td>Dako</td>
			<td>Z031101-2</td>
			<td></td>
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			<td height="21" style="height:21px;">Anti-PCNA (1:400)</td>
			<td>Dako</td>
			<td>M087901</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Anti-GFP (1&#160;:1000)</td>
			<td>Aves Labs</td>
			<td>GFP-1020</td>
			<td></td>
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			<td height="21" style="height:21px;">anti-Dig AP-coupled antibody&#160;</td>
			<td>Roche Diagnostics</td>
			<td align="right">11093274910</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">5-Bromo-4-chloro-3-indolyl-phosphate (BCIP)&#160;</td>
			<td>Roche Diagnostics</td>
			<td align="right">1383221</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">4-Nitroblue tetrazolium chloride (NBT)&#160;</td>
			<td>Roche Diagnostics</td>
			<td align="right">1383213</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Proteinase K</td>
			<td>Roche Diagnostics</td>
			<td align="right" style="width:119px;">1092766</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Polyethylene molding cup tray&#160;</td>
			<td style="width:163px;">Polysciences</td>
			<td style="width:119px;">17177C-3</td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Vibratome VT1000S</td>
			<td>Leica</td>
			<td style="width:119px;"></td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Confocal microscope Sp5 DM6000</td>
			<td>Leica</td>
			<td style="width:119px;"></td>
			<td></td>
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		<tr height="21">
			<td height="21" style="height:21px;">Dissecting microscope Nikon SMZ 645</td>
			<td>Nikon</td>
			<td style="width:119px;"></td>
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		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">24-well flat bottomed tissue culture plates&#160;</td>
			<td>Falcon catalog&#160;</td>
			<td align="right">353226</td>
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			<td height="21" style="height:21px;">Student Dumont #5 Forceps</td>
			<td>Fine Science Tools</td>
			<td>91150-20</td>
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			<td height="21" style="height:21px;">Surgical scissors</td>
			<td>Fine Science Tools</td>
			<td>91460-11</td>
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			<td height="25" style="height:25px;">Danio rerio Tg(olig2:EGFP)</td>
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			<td height="21" style="height:21px;">Danio rerio WT strains (e.g. AB2O2)</td>
			<td style="width:163px;"></td>
			<td style="width:119px;"></td>
			<td></td>
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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/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 ...

stab-wound-injury-zebrafish-adult-telencephalon-method-to-investigate

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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

Stereotaxic Surgery for Excitotoxic Lesion of Specific Brain Areas in the Adult Rat

Many behavioral functions in mammals, including rodents and humans, are mediated principally by discrete brain regions. A common method for discerning the function of various brain regions for behavior or other experimental outcomes is to implement a localized ablation of function. In humans, patient populations with localized brain lesions are often studied for deficits, in hopes of revealing the underlying function of the damaged area. In rodents, one can experimentally induce lesions of specific brain regions.
Lesion can be accomplished in several ways. Electrolytic lesions can cause localized damage but will damage a variety of cell types as well as traversing fibers from other brain regions that happen to be near the lesion site. Inducible genetic techniques using cell-type specific promoters may also enable site-specific targeting. These techniques are complex and not always practical depending on the target brain area. Excitotoxic lesion using stereotaxic surgery, by contrast, is one of the most reliable and practical methods of lesioning excitatory neurons without damaging local glial cells or traversing fibers.
Here, we present a protocol for stereotaxic infusion of the excitotoxin, N-methyl-D-aspartate (NMDA), into the basolateral amygdala complex. Using anatomical indications, we apply stereotaxic coordinates to determine the location of our target brain region and lower an injection needle in place just above the target. We then infuse our excitotoxin into the brain, resulting in excitotoxic death of nearby neurons. While our experimental subject of choice is a rat, the same methods can be applied to other mammals, with the appropriate adjustments in equipment and coordinates.
This method can be used on a variety of brain regions, including the basolateral amygdala1-6, other amygdala nuclei6, 7, hippocampus8, entorhinal cortex9 and prefrontal cortex10. It can also be used to infuse biological compounds such as viral vectors1, 11. The basic stereotaxic technique could also be adapted for implantation of more permanent osmotic pumps, allowing more prolonged exposure to a compound of interest.

Targeted ablation of specific brain region(s) by infusion of an excitotoxin using stereotaxic coordinates is described. This technique could also be adapted for infusion of other chemicals into the rat brain.

Many behavioral functions in mammals, including rodents and humans, are mediated principally by discrete brain regions. A common method for discerning the ...

Micromanipulation of Gene Expression in the Adult Zebrafish Brain Using Cerebroventricular Microinjection of Morpholino Oligonucleotides

Manipulation of gene expression in tissues is required to perform functional studies. In this paper, we demonstrate the cerebroventricular microinjection (CVMI) technique as a means to modulate gene expression in the adult zebrafish brain. By using CVMI, substances can be administered into the cerebroventricular fluid and be thoroughly distributed along the rostrocaudal axis of the brain. We particularly focus on the use of antisense morpholino oligonucleotides, which are potent tools for knocking down gene expression in vivo. In our method, when applied, morpholino molecules are taken up by the cells lining the ventricular surface. These cells include the radial glial cells, which act as neurogenic progenitors. Therefore, knocking down gene expression in the radial glial cells is of utmost importance to analyze the widespread neurogenesis response in zebrafish, and also would provide insight into how vertebrates could sustain adult neurogenesis response. Such an understanding would also help the efforts for clinical applications in human neurodegenerative disorders and central nervous system regeneration. Thus, we present the cerebroventricular microinjection method as a quick and efficient way to alter gene expression and neurogenesis response in the adult zebrafish forebrain. We also provide troubleshooting tips and other useful information on how to carry out the CVMI procedure.

In this article, we demonstrate a method for manipulation of gene expression in the ventricular cells of the adult zebrafish telencephalon using antisense morpholino oligonucleotides. We present this method as an efficient and quick protocol that can be used for functional studies in the adult vertebrate brain.

Manipulation of gene expression in tissues is  required to perform functional studies. In this paper, we demonstrate the  cerebroventricular ...

Functional Interrogation of Adult Hypothalamic Neurogenesis with Focal Radiological Inhibition

The functional characterization of adult-born neurons remains a significant challenge. Approaches to inhibit adult neurogenesis via invasive viral delivery or transgenic animals have potential confounds that make interpretation of results from these studies difficult. New radiological tools are emerging, however, that allow one to noninvasively investigate the function of select groups of adult-born neurons through accurate and precise anatomical targeting in small animals. Focal ionizing radiation inhibits the birth and differentiation of new neurons, and allows targeting of specific neural progenitor regions. In order to illuminate the potential functional role that adult hypothalamic neurogenesis plays in the regulation of physiological processes, we developed a noninvasive focal irradiation technique to selectively inhibit the birth of adult-born neurons in the hypothalamic median eminence. We describe a method for Computer tomography-guided focal irradiation (CFIR) delivery to enable precise and accurate anatomical targeting in small animals. CFIR uses three-dimensional volumetric image guidance for localization and targeting of the radiation dose, minimizes radiation exposure to nontargeted brain regions, and allows for conformal dose distribution with sharp beam boundaries. This protocol allows one to ask questions regarding the function of adult-born neurons, but also opens areas to questions in areas of radiobiology, tumor biology, and immunology. These radiological tools will facilitate the translation of discoveries at the bench to the bedside.

The function of adult-born mammalian neurons remains an active area of investigation. Ionizing radiation inhibits the birth of new neurons. Using computer tomography-guided focal irradiation (CFIR), three-dimensional anatomical targeting of specific neural progenitor populations can now be used to assess the functional role of adult neurogenesis.

The functional characterization of adult-born neurons remains a significant challenge. Approaches to inhibit adult neurogenesis via invasive viral ...

Heat-Induced Antigen Retrieval: An Effective Method to Detect and Identify Progenitor Cell Types during Adult Hippocampal Neurogenesis

Traditional methods of immunohistochemistry (IHC) following tissue fixation allow visualization of various cell types. These typically proceed with the application of antibodies to bind antigens and identify cells with characteristics that are a function of the inherent biology and development. Adult hippocampal neurogenesis is a sequential process wherein a quiescent neural stem cell can become activated and proceed through stages of proliferation, differentiation, maturation and functional integration. Each phase is distinct with a characteristic morphology and upregulation of genes. Identification of these phases is important to understand the regulatory mechanisms at play and any alterations in this process that underlie the pathophysiology of debilitating disorders. Our heat-induced antigen retrieval approach improves the intensity of the signal that is detected and allows correct identification of the progenitor cell type. As discussed in this paper, it especially allows us to circumvent current problems in detection of certain progenitor cell types.

During adult hippocampal neurogenesis, a distinct set of genetic markers are expressed when a quiescent neural stem cell sequentially progresses and develops into a functionally integrated neuron in the circuit. Using heat-induced antigen retrieval, progenitor cell types that are otherwise difficult to detect are identified with improved effectiveness.

Traditional  methods of immunohistochemistry (IHC) following tissue fixation allow visualization  of various cell types. These typically proceed with the ...

Methods for the Modulation and Analysis of NF-&#954;B-dependent Adult Neurogenesis

The hippocampus plays a pivotal role in the formation and consolidation of episodic memories, and in spatial orientation. Historically, the adult hippocampus has been viewed as a very static anatomical region of the mammalian brain. However, recent findings have demonstrated that the dentate gyrus of the hippocampus is an area of tremendous plasticity in adults, involving not only modifications of existing neuronal circuits, but also adult neurogenesis. This plasticity is regulated by complex transcriptional networks, in which the transcription factor NF-&#954;B plays a prominent role. To study and manipulate adult neurogenesis, a transgenic mouse model for forebrain-specific neuronal inhibition of NF-&#954;B activity can be used.
In this study, methods are described for the analysis of NF-&#954;B-dependent neurogenesis, including its structural aspects, neuronal apoptosis and progenitor proliferation, and cognitive significance, which was specifically assessed via a dentate gyrus (DG)-dependent behavioral test, the spatial pattern separation-Barnes maze (SPS-BM). The SPS-BM protocol could be simply adapted for use with other transgenic animal models designed to assess the influence of particular genes on adult hippocampal neurogenesis. Furthermore, SPS-BM could be used in other experimental settings aimed at investigating and manipulating DG-dependent learning, for example, using pharmacological agents.

Methods for the manipulation and analysis of NF-&#954;B-dependent adult hippocampal neurogenesis are described. A detailed protocol is presented for a dentate gyrus-dependent behavioral test (termed the spatial pattern separation-Barnes maze) for the investigation of cognitive outcome in mice. This technique should also help enable investigations in other experimental settings.

The hippocampus plays a pivotal role in the formation and consolidation of episodic memories, and in spatial orientation. Historically, the adult ...

A Novel Light Damage Paradigm for Use in Retinal Regeneration Studies in Adult Zebrafish

Light-induced retinal degeneration (LIRD) is commonly used in both rodents and zebrafish to damage rod and cone photoreceptors. In adult zebrafish, photoreceptor degeneration triggers M&#252;ller glial cells to re-enter the cell cycle and produce transient-amplifying progenitors. These progenitors continue to proliferate as they migrate to the damaged area, where they ultimately give rise to new photoreceptors. Currently, there are two widely-used LIRD paradigms, each of which results in varying degrees of photoreceptor loss and corresponding differences in the regeneration response. As more genetic and pharmacological tools are available to test the role of individual genes of interest during regeneration, there is a need to develop a robust LIRD paradigm. Here we describe a LIRD protocol that results in widespread and consistent loss of both rod and cone photoreceptors in which we have combined the use of two previously established LIRD techniques. Furthermore, this protocol can be extended for use in pigmented animals, which eliminates the need to maintain transgenic lines of interest on the albino background for LIRD studies.

Multiple light damage protocols have been described to damage photoreceptors and consequently induce a retinal regeneration response in adult zebrafish. This protocol describes an improved method that can be used in pigmented animals and that damages the vast majority of rod and cone photoreceptors across the entire retina.

Light-induced retinal degeneration (LIRD) is commonly used in both rodents and zebrafish to damage rod and cone photoreceptors. In adult zebrafish, ...

Electrophysiological Recording in the Brain of Intact Adult Zebrafish

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram recordings. This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments, allowing recording of neural activity. Immobilization of the adult requires a mechanism to deliver dissolved oxygen to the gills in lieu of buccal and opercular movement. With our technique, animals are immobilized and perfused with habitat water to fulfill this requirement. A craniotomy is performed under tricaine methanesulfonate (MS-222; tricaine) anesthesia to provide access to the brain. The primary electrode is then positioned within the craniotomy window to record extracellular brain activity. Through the use of a multitube perfusion system, a variety of pharmacological compounds can be administered to the adult fish and any alterations in the neural activity can be observed. The methodology not only allows for observations to be made regarding changes in neurological activity, but it also allows for comparisons to be made between larval and adult zebrafish. This gives researchers the ability to identify the alterations in neurological activity due to the introduction of various compounds at different life stages.

This paper describes how an adult zebrafish can be immobilized, intubated, and used for in vivo electrophysiological experiments to allow recordings and manipulation of neural activity in an intact animal.

Previously, electrophysiological studies in adult zebrafish have been limited to slice preparations or to eye cup preparations and electrorentinogram ...

A Method to Inflict Closed Head Traumatic Brain Injury in Drosophila

Traumatic brain injury (TBI) affects millions of people each year, causing impairment of physical, cognitive, and behavioral functions and death. Studies using Drosophila have contributed important breakthroughs in understanding neurological processes. Thus, with the goal of understanding the cellular and molecular basis of TBI pathologies in humans, we developed the High Impact Trauma (HIT) device to inflict closed head TBI in flies. Flies subjected to the HIT device display phenotypes consistent with human TBI such as temporary incapacitation and progressive neurodegeneration. The HIT device uses a spring-based mechanism to propel flies against the wall of a vial, causing mechanical damage to the fly brain. The device is inexpensive and easy to construct, its operation is simple and rapid, and it produces reproducible results. Consequently, the HIT device can be combined with existing experimental tools and techniques for flies to address fundamental questions about TBI that can lead to the development of diagnostics and treatments for TBI. In particular, the HIT device can be used to perform large-scale genetic screens to understand the genetic basis of TBI pathologies.

A Method to Inflict Closed Head Traumatic Brain Injury in Drosophila

Here we describe a method to inflict closed head traumatic brain injury (TBI) in Drosophila. This method provides a gateway to investigate the cellular and molecular mechanisms that underlie TBI pathologies using the vast array of experimental tools and techniques available for flies.

Traumatic brain injury (TBI) affects millions of people each year, causing impairment of physical, cognitive, and behavioral functions and death. Studies ...

Dorsal Root Ganglion Injection and Dorsal Root Crush Injury as a Model for Sensory Axon Regeneration

Achieving axon regeneration after nervous system injury is a challenging task. As different parts of the central nervous system (CNS) differ from each other anatomically, it is important to identify an appropriate model to use for the study of axon regeneration. By using a suitable model, we can formulate a specific treatment based on the severity of injury, the neuronal cell type of interest, and the desired spinal tract for assessing regeneration. Within the sensory pathway, DRG neurons are responsible for relaying sensory information from the periphery to the CNS. We present here a protocol that uses a DRG injection with a viral vector and a concurrent dorsal root crush injury in the lower cervical spinal cord of an adult rat as a model to study sensory axon regeneration. As demonstrated using a control virus, AAV5-GFP, we show the effectiveness of a direct DRG injection in transducing DRG neurons and tracing sensory axons into the spinal cord. We also show the effectiveness of the dorsal root crush injury in denervating the forepaw as an injury model for evaluating axon regeneration. Despite the requirement for specialized training to perform this invasive surgical procedure, the protocol is flexible, and potential users can modify many parts to accommodate their experimental requirements. Importantly, it can serve as a foundation for those in search of a suitable animal model for their studies. We believe that this article will help new users to learn the procedure in a very efficient and effective manner.

This protocol presents the use of a dorsal root ganglion (DRG) injection with a viral vector and a concurrent dorsal root crush injury in an adult rat as a model to study sensory axon regeneration. This model is suitable for investigating the use of gene therapy to promote sensory axon regeneration.

Achieving axon regeneration after nervous system injury is a challenging task. As different parts of the central nervous system (CNS) differ from each ...

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.