Name: automated high-throughput behavioral analyses in zebrafish larvae
Uploaded: 2013-07-04T14:00:00
Duration: 9 min 28 s
Description: Watch this Scientific Journal Video about Automated High-throughput Behavioral Analyses in Zebrafish Larvae at JoVE.com

We thank Sean Pelkowski for assistance in optimization of the behavioral assay. This work was supported by the National Institute of Child Health and Human Development, R01 HD060647 and the National Institute of Environmental Health Sciences, F32 ES021342 and R03 ES017755.

1. Collection of Zebrafish Embryos and Raising Larvae
<ol>
	<li>Glass Pyrex dishes with "fake" grass (made from green yarn) (Figure 1) should be inserted into the tanks at dawn and left in for two hours in order to collect zebrafish embryos. The glass dishes containing the embryos should be poured over a handheld strainer and rinsed with deionized water. The embryos should then be grown in egg water. The egg water contains 60 mg/L of Instant Ocean in deionized water and 0.25 mg/L of methylene blue, which is used as a mold inhibitor.</li>
	<li>Depending upon the hypothesis of the individual experiment, embryos can be treated immediately or during specific stages of development using toxicants or pharmaceuticals. The toxicants and pharmaceuticals are usually dissolved in DMSO (at a 1,000X concentration) and should be further diluted directly in the egg water medium.</li>
	<li>During embryonic and larval treatment with pharmaceuticals or toxicants, the larvae and embryos can be housed in deep Petri dishes at a density of about 50-60 larvae per 50 ml until the behavioral analyses at 7 dpf ( egg water solution should be changed at least every other day to avoid fungal/bacterial growth from dead embryos).</li>
</ol>
2. Preparing Molds for Behavioral Analyses
<ol>
	<li>Specially developed plastic molds which measure 11.7 cm x 7.6 cm x 5 mm were custom-built in-house. The molds are needed to create lanes using agarose which is poured into single well plastic plates from Thermo Scientific. The single well plates measure 12.4 cm x 8.1 cm x 1.2 cm.</li>
	<li>The molds contain five lanes in which the sides are angled at 60&deg;. The lanes in the molds are 3.5 mm high with a base of 18 mm at the top which is the widest whereas the bottom width is 14 mm. There is a 4 mm gap between lanes in the mold (Figure 2).</li>
	<li>To prepare the lanes pour 50 ml of melted agarose (0.8% agarose in deionized water with 60 mg/L Instant Ocean) in a single well plate. The mold should then be placed very slowly on top of the liquid agarose to eliminate any bubble formation and can be removed when the agarose has cooled (which takes roughly 45 min).</li>
	<li>The agarose lanes should be made no earlier than one day before the behavioral experiment is to be run (to avoid the agarose drying out) The lanes can be stored at room temperature with the lids on the dishes for up to roughly 36 hr. The agarose lanes should only be used for one experiment and should then be discarded.</li>
</ol>
3. Image Capture
<ol>
	<li>Up to 20 larvae can be placed in each lane of the plates. Typically 5 larvae per lane are used to facilitate the most accurate tracking of swim speed and to reduce the number of larvae that are needed per experiment. The lanes can be filled with egg water with or without pharmaceuticals or toxicants depending upon the experiment. However, lanes should not be filled up all the way until they are placed in the imaging cabinets; this will prevent overflow. For consistency, the larvae should have an acclimatization period of ten minutes after they are placed in the agarose molds and positioned on top of the laptop screen. Efficiently moving the larvae from the Petri dish to the agarose lane will help to reduce larvae stress. This is easiest when the larvae are housed in shallow tanks or Petri dishes.</li>
	<li>The imaging cabinets include a Canon digital camera used for time-lapse photography and a laptop. The camera should be placed at the top of the cabinet and aimed towards to the bottom of the cabinet where a 15.6 inch screen laptop should be placed with screen facing up (Figure 3). Four plates should be positioned by hand directly on top of the laptop screen. At this time the lanes can be topped off with egg water or chemical treatment so that it is level with the top of the lane (to eliminate shadows on the edges of the lanes in the images).</li>
	<li>A PowerPoint presentation is used as an aversive stimulus for the larvae. In the past moving red balls were shown to zebrafish larvae in 6 or 12 multi-well plates 15,17,18. The current PowerPoint starts with a blank white background for 15 min, followed by 15 min of a moving red bar on the top half of the plate (Figure 4). In order to eliminate overheating the larvae, it is best to purchase a laptop with a screen temperature that does not go above 28 &deg;C. To avoid evaporation of the liquid within the agarose lanes, the maximal imaging time should be kept to below one hour.</li>
	<li>The digital camera should be programmed for time-lapse photography, taking pictures every 6 sec for a total of 300 images per experiment. However, the frequency and length can be adjusted depending on the experiment and behavioral quantification. In the past extended imaging times were employed using longer intervals between each image. The camera can be set at a lower resolution for imaging at video speed (30 frames per second). While the lower resolution limits the recordings to a single multiwell plate, the video recordings are appropriate for imaging rapid swimming events 15.</li>
</ol>
4. Image Analysis
<ol>
	<li>The images should be opened with ImageJ and used with a macro that was specifically written in-house. The macro automatically splits the color channels so that the red color can be removed, subtracts the background, applies a threshold, and identifies the larvae by particle analysis. The most current macro for five lane larval analysis is Zebrafish_macro25k. Use the prompts in the macro to set number of images, color to be subtracted, image threshold, etc.</li>
	<li>After all of the images are run through the ImageJ macro, a results file will be displayed and will contain x,y coordinates of the individual larvae for each image along with the image number and the lane number.</li>
	<li>The results file should be saved in an Excel format and sorted based upon blank background vs. moving bar background and then the well number. An Excel template should be used that has equations built in that automatically determines placement of larvae in the wells, distance between larvae, speed of movement, and amount of rest. The most current Excel template is 25ib created in the Creton lab, which is available upon request. Graphs showing various treatment groups should be built into the Excel sheet along with t-tests for comparison between treatment groups and controls. Further statistical analysis can be performed using SPSS.</li>
</ol>

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Teratol. 33, 742-751 (2011).</li><li>Stewart, A., Wu, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pharmacological+modulation+of+anxiety-like+phenotypes+in+adult+zebrafish+behavioral+models.">Pharmacological modulation of anxiety-like phenotypes in adult zebrafish behavioral models.</a> Prog. Neuropsychopharmacol. Biol. Psychiatry. 35, 1421-1431 (2011).</li><li>Eddins, D., Cerutti, D., Williams, P., Linney, E., Levin, E. 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(20), e923 (2008).</li><li>Pelkowski, S., Kapoor, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+high-throughput+imaging+system+for+automated+analyses+of+avoidance+behavior+in+zebrafish+larvae.">A novel high-throughput imaging system for automated analyses of avoidance behavior in zebrafish larvae.</a> Behav. Brain Res. 223, 135-144 (2011).</li><li>Richendrfer, H., Pelkowski, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Assessment+of+developmental+toxicity+by+automated+analyses+of+behavior+in+zebrafish+larvae.">Assessment of developmental toxicity by automated analyses of behavior in zebrafish larvae.</a> Unpublished observations. , (2012).</li><li>Richendrfer, H., Pelkowski, S., Colwill, R. 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The authors declare that they have no competing financial interests.

While we are continually improving our novel behavioral assay, it has always been useful for the detection of avoidance and thigmotaxis behavior in zebrafish larvae 15. Many trials have been performed to optimize the results of the assay, such as color of stimulus used, ideal number of larvae per lane, and length of behavioral assay. Previously, we used multi-well plates (with 6 or 12 wells) 15,17,18. However, recently we have created the novel lane mold to create a larger swimming space for the larvae allowing us to gather a larger number of behavioral measures in a single assay 16 (Figure 5). Other modifications include variations of the PowerPoint shown (altered movement or length of assay) and the size of the lanes used (we also have molds for more narrow lanes).
Currently, this high-throughput automated system is unique in its ability to measure a large range of behaviors in zebrafish larvae at the same time such as speed, avoidance, proximity to other larvae, and thigmotaxis in multi-lane plates. Results can be obtained quickly and a large number of larvae can be analyzed at the time of imaging. The system is both inexpensive to build and quick and easy to set up. A limitation of this system is that 3-D movements cannot be assessed in the zebrafish larvae. Automated systems that track adult zebrafish have the 3-D capability and can identify a wider range of behaviors such as movement up or down within the water column 10,19. Another limitation is that our imaging system is currently not optimized for high-throughput analyses at video speed. Video speed imaging is possible when setting the camera to a lower resolution 15, but this restricts the analysis to a single plate.
In using the newly created "lane" method, several parts of the assay needed to be executed in a precise manner. When placing the larvae in the lanes, it is critical to make sure the level of liquid is very shallow until the plates are positioned on top of the laptop screen. If the lanes are too full of liquid, the larvae will escape into the periphery of the plate. In addition, when inserting the mold into the agarose, care must be taken to lower the mold very slowly. If the mold is inserted too quickly, bubbles will form in the agarose and will be identified by the Image J macro as additional larvae. It is advised that if the agarose lanes have even a few bubbles, it is best to make new ones.
In the future, we would like to optimize our behavioral assay to analyze other complex behaviors such as learning in zebrafish larvae and examine how learning may be affected by exposure to toxicants and pharmaceuticals in early development. We are currently working on assays that may be useful for analyzing learning behavior in which the behavioral results may facilitate determining which brain areas are affected by certain toxicants or pharmaceuticals during development. Automated assays have been developed for measuring learning behaviors in zebrafish larvae 20 and these assays may be amendable for high-throughput screening by using the robust avoidance response in multi-lane plates.
We propose that this behavioral assay could be used in future studies for testing the developmental effects of a large number of pharmaceuticals and toxicants. Such studies would provide a wealth of information on specific risk factors and contribute to setting better health and safety guidelines for pregnant women and children.

Zebrafish are becoming a popular model for genetic, developmental and behavioral sciences 1-4. They hatch from their chorions by 2-3 days post fertilization (dpf), develop fully functioning organs by 4-5 dpf, and exhibit a large number of behaviors by 7 dpf 5,6. Zebrafish larvae are ideally suited for high-throughput analyses because of their small size 7,8. Software is commercially available for automated analyses of behavior in larval and adult zebrafish 9-14. However, this software can be expensive and has limited options for measuring complex behaviors of zebrafish larvae in multi-well plates.
We created a novel high-throughput imaging system that is inexpensive to set up and can quantify a number of different behaviors in 7 dpf zebrafish larvae 15,16. The system allows us to quickly and efficiently test subtle behavioral abnormalities after embryonic exposure to a number of pharmaceuticals and environmental toxicants 16-18.
The system was built using wooden cabinets, which house a digital camera at the top of the cabinet. The camera faces downward to the bottom of the cabinet where a laptop is placed with the screen facing up 15. Time lapse imaging is used to capture the placement of the larvae in the lanes. Larvae can be housed in up to four multi-well or multi-lane plates which are positioned on top of the laptop screen. We use a PowerPoint presentation as an aversive stimulus to which the larvae respond by moving away (avoidance) and swimming towards the edge (thigmotaxis) 15,17. The images are imported into ImageJ in which an automated macro is used to split the color channels, subtract the background, and apply a threshold to identify individual larvae. Coordinates are listed for each larva in each image and can be inserted into an Excel file which we use to quantify avoidance and thigmotaxis behavior, fish to fish distance, swim speed, and amount of rest 16.

<table border="1">
		<tbody>
		 <tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Reagent</td>
		 </tr>
		 <tr>
			<td>Instant Ocean</td>
			<td>That Pet Place</td>
			<td>198262</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Agarose</td>
			<td>Fisher</td>
			<td>BP1356-100</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Methylene Blue</td>
			<td>That Pet Place</td>
			<td>214325</td>
			<td></td>
		 </tr>
		 <tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Equipment</td>
		 </tr>
		 <tr>
			<td>One well plates</td>
			<td>Fisher</td>
			<td>12-565-493</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Digital camera</td>
			<td>Canon </td>
			<td>EOS Rebel T1i</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Imaging Cabinets</td>
			<td>WoodCraft Towers</td>
			<td></td>
			<td></td>
		 </tr>
		 <tr>
			<td>Laptops</td>
			<td>Acer Aspire</td>
			<td>Any is good as long as it has a 15.6 in. LCD screen with 1366 x 768 pixel resolution and a brightness of 220 cd/m2.</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Camera Lens</td>
			<td>Canon</td>
			<td>EF-S 55 - 250 mm f/4.0 - 5.6 IS zoom lens</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Plastic diffuser</td>
			<td>Pendaflex </td>
			<td>52345</td>
			<td></td>
		 </tr>
		 <tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Software</td>
		 </tr>
		 <tr>
			<td>PowerPoint 2010</td>
			<td>Microsoft</td>
			<td></td>
			<td></td>
		 </tr>
		 <tr>
			<td>ImageJ</td>
			<td>NIH</td>
			<td></td>
			<td><a href="http://rsb.info.nih.gov/ij/" target="_blank">http://rsb.info.nih.gov/ij/</a></td>
		 </tr>
		 <tr>
			<td>Excel 2010</td>
			<td>Microsoft</td>
			<td></td>
			<td></td>
		 </tr>
		 <tr>
			<td>Statistical software</td>
			<td>SPSS 20</td>
			<td></td>
			<td></td>
		 </tr>
		</tbody>
 </table>

automated high-throughput behavioral analyses in zebrafish larvae

We have created a novel high-throughput imaging system for the analysis of behavior in 7-day-old zebrafish larvae in multi-lane plates. This system measures spontaneous behaviors and the response to an aversive stimulus, which is shown to the larvae via a PowerPoint presentation. The recorded images are analyzed with an ImageJ macro, which automatically splits the color channels, subtracts the background, and applies a threshold to identify individual larvae placement in the lanes. We can then import the coordinates into an Excel sheet to quantify swim speed, preference for edge or side of the lane, resting behavior, thigmotaxis, distance between larvae, and avoidance behavior. Subtle changes in behavior are easily detected using our system, making it useful for behavioral analyses after exposure to environmental toxicants or pharmaceuticals.

In our earlier assays, using the bouncing ball aversive stimulus, wild type larvae that are untreated respond to the moving ball by swimming down in the well (avoidance behavior) and towards the edges of the well (thigmotaxis behavior) 15. We later confirmed that thigmotaxis behavior in this assay is a measure of anxiety-related behavior in zebrafish larvae 17. There were significant differences in the larvae movement away from the ball and preference for the edge when compared to the blank white background. These behaviors have also been confirmed in our new assay using the moving red bar and are even more robust 16. Moreover, we can now sample a larger number of behaviors in a single assay including swim speed, rest, preference for the end or side of the well, and distance between fish (Figure 5). Control larvae grown in egg water show an increased preference to be down in the dish and on the edge of the lane after they are presented with an aversive stimulus (moving red bar). Similar results are obtained when larvae are grown in egg water containing 1 &mu;g/ml DMSO, a solvent that is commonly used to dissolve various pharmaceuticals and toxicants as 1,000X stock solutions.
Representative results are shown in Figure 5 in larvae treated with egg water and DMSO (as controls) and varying concentrations of an organophosphate pesticide commonly found in non-organic foods. The results shown are a sampling from one experiment. However, when repeated, the results indicate that swim speed and thigmotaxis behavior is altered by low concentrations of organophosphate pesticides, which mimic levels in human food consumption 18.
<img alt="Figure 1" src="/files/ftp_upload/50622/50622fig1.jpg" /> 
 Figure 1. Collection Trays. Glass Pyrex dishes are used to collect embryos from the adult fish tanks. Lids from the Pyrex dishes were cut and inserted with plastic grids and green yarn was sewn onto the grids in the plastic. This creates a breeding atmosphere for the adult zebrafish by mimicking the natural environment.
<img alt="Figure 2" fo:content-width="/files/ftp_upload/50622/50622fig2highres.jpg" fo:width="6in" src="/files/ftp_upload/50622/50622fig2.jpg" /> 
 Figure 2. Plastic mold and agarose lanes. A) The mold is shown on the left. 0.8% agarose is poured into a one-well plate; the mold is slowly inserted and then removed when the agarose has cooled. B) The plate on the right shows the lanes created in agarose by the plastic mold.
<img alt="Figure 3" src="/files/ftp_upload/50622/50622fig3.jpg" /> 
 Figure 3. Imaging Cabinets. Imaging cabinets were specially built in our laboratory and used for high-throughput behavioral analyses. A 15 megapixel digital camera was attached to the top of the cabinet facing downwards in order to gather time lapse images of the larvae in multilane plates placed on top of the screen of a laptop. Between the plates and the screen there is a plastic diffuser that is used to prevent moir&eacute; patterns in the images collected.
<img alt="Figure 4" fo:content-width="/files/ftp_upload/50622/50622fig4highres.jpg" fo:width="4.5in" src="/files/ftp_upload/50622/50622fig4.jpg" /> 
 Figure 4. Blank background and PowerPoint aversive stimulus. This is the current PowerPoint that is used to evoke behavioral changes in zebrafish larvae. It provides robust behavioral differences between A) the blank background and B) the moving red bar.
<img alt="Figure 5" fo:content-width="/files/ftp_upload/50622/50622fig5highres.jpg" fo:width="5.5in" src="/files/ftp_upload/50622/50622fig5.jpg" /> 
 Figure 5. Behaviors Quantified in the high-throughput assay. Example of the behaviors that are quantified from our behavioral assay within the Excel sheet that we use for x,y coordinates of the larvae. The white bars show data from larvae exposed to a blank background and the red bars show data from the larvae exposed to the red moving bar in the PowerPoint. The graphs indicate the measurements that can be obtained from behavioral analysis A) Percentage of larvae down in the lane, B) Percentage of larvae on the end of the lane, C) Percentage of larvae on the edge of the lane, D) Distance between fish (mm), E) Swim speed of the larvae (mm/min), F) Percentage rest of the larvae. In the graphs shown, data is from treatment of larvae with DMSO control and several concentrations of a pesticide ranging from 0.001 to 0.1 &mu;M (levels commonly found in the human diet). <a href="https://www.jove.com/files/ftp_upload/50622/50622fig5large.jpg" target="_blank"> Click here to view larger figure</a>.

Watch this Scientific Journal Video about Automated High-throughput Behavioral Analyses in Zebrafish Larvae at JoVE.com

Automated High-throughput Behavioral Analyses in Zebrafish Larvae

Our laboratory developed a novel high-throughput automated imaging system that is useful for the detection of several different behaviors in 7-day-old zebrafish larvae. The system can be used for detecting subtle changes in behavior after the larvae have been exposed to environmental toxicants or pharmaceuticals.

We have created a novel high-throughput imaging system for the analysis of behavior in 7-day-old zebrafish larvae in multi-lane plates. This system ...

automated-high-throughput-behavioral-analyses-in-zebrafish-larvae

Research

JoVE Journal

Neuroscience

In vivo Electroporation of Developing Mouse Retina

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge. Gene targeting to generate constitutive or conditional loss of function knockouts remains cost and labor intensive, as well as time consuming. Adding to these challenges, retina expressed genes may have essential roles outside the retina leading to unintended confounds when using a knockout approach. Furthermore, the ability to ectopically express a gene in a gain of function experiment can be extremely valuable when attempting to identify a role in cell fate specification and/or terminal differentiation.
We present a method for the rapid and efficient incorporation of DNA plasmids into the neonatal mouse retina by electroporation. The application of short electrical impulses above a certain field strength results in a transient increase in plasma membrane permeability, facilitating the transfer of material across the membrane 1,2,3,4. Groundbreaking work demonstrated that electroporation could be utilized as a method of gene transfer into mammalian cells by inducing the formation of hydrophilic plasma membrane pores allowing the passage of highly charged DNA through the lipid bilayer 5. Continuous technical development has resulted in the viability of electroporation as a method for in vivo gene transfer in multiple mouse tissues including the retina, the method for which is described herein 6, 7, 8, 9, 10. 
DNA solution is injected into the subretinal space so that DNA is placed between the retinal pigmented epithelium and retina of the neonatal (P0) mouse and electrical pulses are applied using a tweezer electrode. The lateral placement of the eyes in the mouse allows for the easy orientation of the tweezer electrode to the necessary negative pole-DNA-retina-positive pole alignment. Extensive incorporation and expression of transferred genes can be identified by postnatal day 2 (P2). Due to the lack of significant lateral migration of cells in the retina, electroporated and non-electroporated regions are generated. Non-electroporated regions may serve as internal histological controls where appropriate. 
Retinal electroporation can be used to express a gene under a ubiquitous promoter, such as CAG, or to disrupt gene function using shRNA constructs or Cre-recombinase. More targeted expression can be achieved by designing constructs with cell specific gene promoters. Visualization of electroporated cells is achieved using bicistronic constructs expressing GFP or by co-electroporating a GFP expression construct. Furthermore, multiple constructs may be electroporated for the study of combinatorial gene effects or simultaneous gain and loss of function of different genes. Retinal electroporation may also be utilized for the analysis of genomic cis-regulatory elements by generating appropriate expression constructs and deletion mutants. Such experiments can be used to identify cis-regulatory regions sufficient or required for cell specific gene expression 11. Potential experiments are limited only by construct availability.

In vivo Electroporation of Developing Mouse Retina

A method for the incorporation of plasmid DNA into murine retinal cells for the purpose of performing either gain- or loss of function studies in vivo is presented. This method capitalizes on the transient increase in permeability of cell plasma membranes induced by the application of an external electrical field.

The functional characterization of genes expressed during mammalian retinal development remains a significant challenge.  Gene targeting to generate ...

FIM Imaging and FIMtrack: Two New Tools Allowing High-throughput and Cost Effective Locomotion Analysis

The analysis of neuronal network function requires a reliable measurement of behavioral traits. Since the behavior of freely moving animals is variable to a certain degree, many animals have to be analyzed, to obtain statistically significant data. This in turn requires a computer assisted automated quantification of locomotion patterns. To obtain high contrast images of almost translucent and small moving objects, a novel imaging technique based on frustrated total internal reflection called FIM was developed. In this setup, animals are only illuminated with infrared light at the very specific position of contact with the underlying crawling surface. This methodology results in very high contrast images. Subsequently, these high contrast images are processed using established contour tracking algorithms. Based on this, we developed the FIMTrack software, which serves to extract a number of features needed to quantitatively describe a large variety of locomotion characteristics. During the development of this software package, we focused our efforts on an open source architecture allowing the easy addition of further modules. The program operates platform independent and is accompanied by an intuitive GUI guiding the user through data analysis. All locomotion parameter values are given in form of csv files allowing further data analyses. In addition, a Results Viewer integrated into the tracking software provides the opportunity to interactively review and adjust the output, as might be needed during stimulus integration. The power of FIM and FIMTrack is demonstrated by studying the locomotion of Drosophila larvae.

FIM is a novel, cost effective imaging system designed to track small moving objects such as C. elegans, planaria or Drosophila larvae. The accompanying FIMTrack program is designed to deliver fast and efficient data analysis. Together, these tools allow high-throughput analysis of behavioral traits.

The analysis of neuronal network function requires a reliable measurement of behavioral traits. Since the behavior of freely moving animals is variable to ...

Electroretinogram Analysis of the Visual Response in Zebrafish Larvae

The electroretinogram (ERG) is a noninvasive electrophysiological method for determining retinal function. Through the placement of an electrode on the surface of the cornea, electrical activity generated in response to light can be measured and used to assess the activity of retinal cells in vivo. This manuscript describes the use of the ERG to measure visual function in zebrafish. Zebrafish have long been utilized as a model for vertebrate development due to the ease of gene suppression by morpholino oligonucleotides and pharmacological manipulation. At 5-10 dpf, only cones are functional in the larval retina. Therefore, the zebrafish, unlike other animals, is a powerful model system for the study of cone visual function in vivo. This protocol uses standard anesthesia, micromanipulation and stereomicroscopy protocols that are common in laboratories that perform zebrafish research. The outlined methods make use of standard electrophysiology equipment and a low light camera to guide the placement of the recording microelectrode onto the larval cornea. Finally, we demonstrate how a commercially available ERG stimulator/recorder originally designed for use with mice can easily be adapted for use with zebrafish. ERG of larval zebrafish provides an excellent method of assaying cone visual function in animals that have been modified by morpholino oligonucleotide injection as well as newer genome engineering techniques such as Zinc Finger Nucleases (ZFNs), Transcription Activator-Like Effector Nucleases (TALENs), and Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas9, all of which have greatly increased the efficiency and efficacy of gene targeting in zebrafish. In addition, we take advantage of the ability of pharmacological agents to penetrate zebrafish larvae to evaluate the molecular components that contribute to the photoresponse. This protocol outlines a setup that can be modified and used by researchers with various experimental goals.

We present a method for the electroretinographic (ERG) analysis of zebrafish larvae utilizing micromanipulation and electroretinography techniques. This is a simple and straightforward method for assaying visual function of zebrafish larvae in vivo.

The electroretinogram (ERG) is a noninvasive electrophysiological method for determining retinal function. Through the placement of an electrode on the ...

Ablation of a Neuronal Population Using a Two-photon Laser and Its Assessment Using Calcium Imaging and Behavioral Recording in Zebrafish Larvae

To identify the role of a subpopulation of neurons in behavior, it is essential to test the consequences of blocking its activity in living animals. Laser ablation of neurons is an effective method for this purpose when neurons are selectively labeled with fluorescent probes. In the present study, protocols for laser ablating a subpopulation of neurons using a two-photon microscope and testing of its functional and behavioral consequences are described. In this study, prey capture behavior in zebrafish larvae is used as a study model. The pretecto-hypothalamic circuit is known to underlie this visually-driven prey catching behavior. Zebrafish pretectum were laser-ablated, and neuronal activity in the inferior lobe of the hypothalamus (ILH; the target of the pretectal projection) was examined. Prey capture behavior after pretectal ablation was also tested.

Here, we present a protocol to ablate a genetically labeled subpopulation of neurons by a two-photon laser from Zebrafish larvae.

To identify the role of a subpopulation of neurons in behavior, it is essential to test the consequences of blocking its activity in living animals. Laser ...

Behavioral And Physiological Analysis In A Zebrafish Model Of Epilepsy

Epilepsy represents one of the most common neurological disorders, affecting an estimated 50 million people worldwide. Recent advances in genetic research have uncovered a large spectrum of genes implicated in various forms of epilepsy, highlighting the heterogeneous nature of this disorder. Appropriate animal models are essential for investigating the pathological mechanisms triggered by genetic mutations implicated in epilepsy and for developing specialized, targeted therapies. In recent years, zebrafish has emerged as a valuable vertebrate organism for modeling epilepsies, with the use of both genetic manipulation and exposure to known epileptogenic drugs, such as pentylenetetrazole (PTZ), to identify novel anti-epileptic therapeutics. Deleterious mutations in the mTOR regulator DEPDC5 have been associated with various forms of focal epilepsies and knock-down of the zebrafish orthologue causes hyperactivity associated with spontaneous seizure-like episodes, as well as enhanced electrographic activity and characteristic turn wheel swimming. Here, we described the method involved in generating the DEPDC5 loss-of-function model and illustrate the protocol for assessing motor activity at 28 and 48 h post fertilization (hpf), as well as a method for recording field activity in the zebrafish optic tectum. An illustration of the effect of the epileptogenic drug PTZ on neuronal activity over time is also provided.

Here, we present a protocol for the development and the characterization of a zebrafish model of epilepsy resulting from the transient inhibition of the DEPDC5 gene.

Epilepsy represents one of the most common neurological disorders, affecting an estimated 50 million people worldwide. Recent advances in genetic research ...

Using Zebrafish Larvae to Study the Pathological Consequences of Hemorrhagic Stroke

Despite being the most severe subtype of stroke with high global mortality, there is no specific treatment for patients with intracerebral hemorrhage (ICH). Modelling ICH pre-clinically has proven difficult, and current rodent models poorly recapitulate the spontaneous nature of human ICH. Therefore, there is an urgent requirement for alternative pre-clinical methodologies for study of disease mechanisms in ICH and for potential drug discovery.
The use of zebrafish represents an increasingly popular approach for translational research, primarily due to a number of advantages they possess over mammalian models of disease, including prolific reproduction rates and larval transparency allowing for live imaging. Other groups have established that zebrafish larvae can exhibit spontaneous ICH following genetic or chemical disruption of cerebrovascular development. The aim of this methodology is to utilize such models to study the pathological consequences of brain hemorrhage, in the context of pre-clinical ICH research. By using live imaging and motility assays, brain damage, neuroinflammation and locomotor function following ICH can be assessed and quantified.
This study shows that key pathological consequences of brain hemorrhage in humans are conserved in zebrafish larvae highlighting the model organism as a valuable in vivo&#160;system for pre-clinical investigation of ICH. The aim of this methodology is to enable the pre-clinical stroke community to employ the zebrafish larval model as an alternative complementary model system to rodents.

Here we present a protocol to quantify brain injury, locomotor deficits and neuroinflammation following bleeding in the brain in zebrafish larvae, in the context of human intracerebral hemorrhage (ICH).

Despite being the most severe subtype of stroke with high global mortality, there is no specific treatment for patients with intracerebral hemorrhage ...

Alternate Immersion in Glucose to Produce Prolonged Hyperglycemia in Zebrafish

Zebrafish (Danio rerio) are an excellent model to investigate the effects of chronic hyperglycemia, a hallmark of Type II Diabetes Mellitus (T2DM). This alternate immersion protocol is a noninvasive, step-wise method of inducing hyperglycemia for up to eight weeks. Adult zebrafish are alternately exposed to sugar (glucose) and water for 24 hours each. The zebrafish begin treatment in a 1% glucose solution for 2 weeks, then a 2% solution for 2 weeks, and finally a 3% solution for the remaining 4 weeks. Compared to water-treated (stress) and mannitol-treated (osmotic) controls, glucose-treated zebrafish have significantly higher blood sugar levels. The glucose-treated zebrafish show blood sugar levels of 3-times that of controls, suggesting that after both four and eight weeks hyperglycemia can be achieved. Sustained hyperglycemia was associated with increased Glial Fibrillary Acidic Protein (GFAP) and increased nuclear factor Kappa B (NF-kB)&#160;levels in retina and decreased physiological responses, as well as cognitive deficits suggesting this protocol can be used to model disease complications.

This protocol noninvasively induces hyperglycemia in zebrafish for up to 8 weeks. Using this protocol, an in-depth study of the adverse effects of hyperglycemia can be made.

Zebrafish (Danio rerio) are an excellent model to investigate the effects of chronic hyperglycemia, a hallmark of Type II Diabetes Mellitus (T2DM). This ...

A Rat Model of EcoHIV Brain Infection

It has been well studied that the EcoHIV infected mouse model is of significant utility in investigating HIV associated neurological complications. Establishment of the EcoHIV infected rat model for studies of drug abuse and neurocognitive disorders, would be beneficial in the study of neuroHIV and HIV-1 associated neurocognitive disorders (HAND). In the present study, we demonstrate the successful creation of a rat model of active HIV infection using chimeric HIV (EcoHIV). First, the lentiviral construct of EcoHIV was packaged in cultured 293 FT cells for 48 hours. Then, the conditional medium was concentrated and titered. Next, we performed bilateral stereotaxic injections of the EcoHIV-EGFP into F344/N rat brain tissue. One week after infection, EGFP fluorescence signals were detected in the infected brain tissue, indicating that EcoHIV successfully induces an active HIV infection in rats. In addition, immunostaining for the microglial cell marker, Iba1, was performed. The results indicated that microglia were the predominant cell type harboring EcoHIV. Furthermore, EcoHIV rats exhibited alterations in temporal processing, a potential underlying neurobehavioral mechanism of HAND as well as synaptic dysfunction eight weeks after infection. Collectively, the present study extends the EcoHIV model of HIV-1 infection to the rat offering a valuable biological system to study HIV-1 viral reservoirs in the brain as well as HAND and associated comorbidities such as drug abuse.

Here, we present a protocol to establish a new rat model of active HIV infection using chimeric HIV (EcoHIV), which is critical for enhancing our understanding of HIV-1 viral reservoirs in the brain and offering a system to study HIV-associated neurocognitive disorders and associated comorbidities (i.e., drug abuse).

It has been well studied that the EcoHIV infected mouse model is of significant utility in investigating HIV associated neurological complications. ...

Optogenetic Phase Transition of TDP-43 in Spinal Motor Neurons of Zebrafish Larvae

Abnormal protein aggregation and selective neuronal vulnerability are two major hallmarks of neurodegenerative diseases. Causal relationships between these features may be interrogated by controlling the phase transition of a disease-associated protein in a vulnerable cell type, although this experimental approach has been limited so far. Here, we describe a protocol to induce phase transition of the RNA/DNA-binding protein TDP-43 in spinal motor neurons of zebrafish larvae for modeling cytoplasmic aggregation of TDP-43 occurring in degenerating motor neurons in amyotrophic lateral sclerosis (ALS). We describe a bacterial artificial chromosome (BAC)-based genetic method to deliver an optogenetic TDP-43 variant selectively to spinal motor neurons of zebrafish. The high translucency of zebrafish larvae allows for the phase transition of the optogenetic TDP-43 in the spinal motor neurons by a simple external illumination using a light-emitting diode (LED) against unrestrained fish. We also present a basic workflow of live imaging of the zebrafish spinal motor neurons and image analysis with freely available Fiji/ImageJ software to characterize responses of the optogenetic TDP-43 to the light illumination. This protocol enables the characterization of TDP-43 phase transition and aggregate formation in an ALS-vulnerable cellular environment, which should facilitate an investigation of its cellular and behavioral consequences.

We describe a protocol to induce phase transition of TAR DNA-binding protein 43 (TDP-43) by light in the spinal motor neurons using zebrafish as a model.

Abnormal protein aggregation and selective neuronal vulnerability are two major hallmarks of neurodegenerative diseases. Causal relationships between ...

&lt;em&gt;In Vivo&lt;/em&gt; Whole-Brain Imaging of Zebrafish Larvae

In Vivo Whole-Brain Imaging of Zebrafish Larvae Using Three-Dimensional Fluorescence Microscopy

As a vertebrate model animal, larval zebrafish are widely used in neuroscience and provide a unique opportunity to monitor whole-brain activity at the cellular resolution. Here, we provide an optimized protocol for performing whole-brain imaging of larval zebrafish using three-dimensional fluorescence microscopy, including sample preparation and immobilization, sample embedding, image acquisition, and visualization after imaging. The current protocol enables in vivo imaging of the structure and neuronal activity of a larval zebrafish brain at a cellular resolution for over 1 h using confocal microscopy and custom-designed fluorescence microscopy. The critical steps in the protocol are also discussed, including sample mounting and positioning, preventing bubble formation and dust in the agarose gel, and avoiding motion in images caused by incomplete solidification of the agarose gel and paralyzation of the fish. The protocol has been validated and confirmed in multiple settings. This protocol can be easily adapted for imaging other organs of a larval zebrafish.

Author Spotlight: In Vivo Whole-Brain Imaging of Zebrafish Larvae Using Three-Dimensional Fluorescence Microscopy  

Presented here is a protocol for in vivo whole-brain imaging of larval zebrafish using three-dimensional fluorescence microscopy. The experimental procedure includes sample preparation, image acquisition, and visualization.

As a vertebrate model animal, larval zebrafish are widely used in neuroscience and provide a unique opportunity to monitor whole-brain activity at the ...