Name: behavioral and physiological analysis in a zebrafish model of epilepsy
Uploaded: 2021-10-19T13:00:00.000+00:00
Duration: 8 s
Description: Watch this Scientific Journal Video about Behavioral And Physiological Analysis In A Zebrafish Model Of Epilepsy at JoVE.com

We would like to thank the staff of the ICM electrophysiology platform where the neurophysiology experiments were performed. We also thank Anca Marian for technical help. SC was supported by the Trampoline Grant #21488. EK was supported by the AFM Grant #18469 and ERC Consolidator Grant (ALS-Networks). HC was supported by PhD awards from the Fondation pour la Recherche M&#233;dicale (PLP20141031462) and ARSLA. For AD and RM, this work was supported by three grants from the Romanian National Authority for Scientific Research and Innovation, CNCS-UEFISCDI (project numbers PN-III-P4-ID-PCE-2016-0010, PN-III-P2-2.1-PED-2016-0007, and COFUND-NEURON-NMDAR-PSY), a grant by the European Union&#8217;s Horizon 2020 research and innovation programme &#8211; grant agreement no. 668863-SyBil-AA, and a National Science Foundation grant NSF-IOS-1656830 funded by the US Government.&#160;

Experimental procedures were approved by the National and Institutional Ethical Committees.
1. Transient Knock-down of DEPDC5 Gene in Zebrafish Embryo&#160;
<ol>
	<li>Preparation of tools:
	<ol>
		<li>Prepare silicon elastomer-coated injection Petri dishes: Mix the base and curing agent of the kit (see Table of Materials) in a 10:1 ratio. Fill a 35 mm Petri dish halfway with the mixture. Wait for the silicon to harden before using it (this can take several days).&#160;</li>
		<li>Prepare 1.2 mmol/L stock solutions of antisense morpholino oligonucleotides (AMO; see Table of Materials). Add 250 &#181;L of sterile water to 300 nmole lyophilized AMO to obtain a 1.2 mmol/L stock solution. For complete dissolution, heat the vials for 5 min at 65 &#176;C. Vortex briefly. Seal the tube cap with a plastic film (see Table of Materials).&#160;</li>
		<li>For the control rescue experiments, prepare an expression plasmid containing the human cDNA of DEPDC5 (see Table of Materials) cloned in the pCS2 backbone or a similar zebrafish-compatible expression plasmid. As a negative control, a mutation causing an early stop codon (p.Arg487*) was introduced in the cDNA.</li>
		<li>Prepare embryo water: 0.06 g/L aquarium salt (see Table of Materials) in reverse osmosis water + 0.5 mg/L methylene blue.</li>
		<li>The day of the injection, prepare microinjection borosilicate glass needles using a puller (see Table of Materials). Establish appropriate temperature settings on the needle puller. Use a 10 cm long, 1/0.5 OD/ID mm borosilicate glass capillary to generate two ~5 cm capillaries with thin tips with a length of approximately 1 cm.&#160;</li>
		<li>If the tip of the needles is very fine, preventing the ejection of solution, break the very end of the tapered tip using forceps under a microscope.</li>
		<li>Just prior to injection, prepare the working solutions of AMOs. Always prepare fresh solution to ensure the reproducibility of the results. Heat the AMO stock vials at 65 &#176;C for 5 min. Prepare a 5 &#181;L injection sample containing Fast Green dye (0.02% final concentration, see Table of Materials) and the AMO diluted at the working concentration in water.&#160;</li>
		<li>Determine the working concentration of the AMO empirically for each gene using a dose response curve. The working concentration represents a concentration where the AMO is effective in knocking down the gene without causing general toxicity, such as gross morphological defects. Typically, AMO working concentrations will be in a range of 0.2 mmol/L to 1 mmol/L (0.4 mmol/L was determined as the effective concentration for this study18). Inject the control Mismatch morpholino at the same concentration as the effective AMO.&#160;</li>
		<li>Vortex the tubes and centrifuge briefly to bring the droplets to the bottom of the tubes.</li>
		<li>For rescue experiments, prepare a 5 &#181;L injection sample with the AMO diluted at the working concentration and the cDNA expression plasmid diluted in water to a final concentration to be determined empirically. For the expression of DEPDC5 and the negative control plasmid, 100 ng/&#181;L was effective for phenotypic rescue.&#160;</li>
	</ol>
	</li>
	<li>Embryo preparation:
	<ol>
		<li>The day prior to microinjection, set up the zebrafish mating tanks. The morning of the injection, remove the dividers to enable spawning. Collect the eggs in 100 mm Petri dishes filled with embryo water using a fine sieve. Inject within 20-30 min from collection, while the eggs are at the one cell stage.&#160;</li>
		<li>Pick 60-80 eggs with a plastic Pasteur pipette and arrange them in the silicon-coated Petri dish for injection. The silicon surface will prevent the eggs from sliding during the injections. Remove most of the embryo water, leaving just enough to cover the eggs halfway.</li>
	</ol>
	</li>
	<li>Microinjections:
	<ol>
		<li>Fill a glass needle with injection solution. Place the needle vertically in one of the tubes containing the injection solution, ensuring that the bottom end of the needle is touching the solution. Wait several minutes until the colored injection solution rises by capillarity and is visible at the tip of the needle.</li>
		<li>Mount the filled needle on the injecting handle of the microinjector (see Table of Materials).</li>
		<li>Turn on the air compressor and adjust the pressure setting to generate an injection volume of ~2 nL.&#160;</li>
		<li>To calculate the volume of the injected solution, place a drop of mineral oil on a microtome slide. Inject the dye-containing solution using the set pressure and time parameters. Measure the diameter of the injected fluid sphere and calculate total volume using the formula Volume=4/3*&#960;*(d/2)3, where d=the measured diameter of the injected bolus.</li>
		<li>Using a dissecting binocular microscope with a 4X magnification, inject the eggs at the single cell stage by passing through the chorion and the yolk, and projecting the solution directly within the cell.</li>
		<li>Collect the injected embryos in a 100 mm Petri dish with embryo water, label the dish and incubate them at 28 &#176;C.&#160;</li>
		<li>Ensure that the incubator temperature is stable over time, as embryos development rate is thermosensitive. For example, the growth would be accelerated at higher temperatures and the developmental stage is critical for properly assessing the phenotype24.</li>
		<li>Check the quality of the eggs 6-8 h after injection and remove dead and unfertilized embryos using a plastic Pasteur pipette.</li>
		<li>The next morning, count and remove dead embryos in each dish with a plastic Pasteur pipette.</li>
	</ol>
	</li>
</ol>
2. Behavior Analysis&#160;
<ol>
	<li>Global activity analysis at 28 hpf:
	<ol>
		<li>Conduct the test the afternoon of the day after microinjection (28 hpf), ensuring that the time of day at which the test is conducted is consistent over experiments to perform valid statistical analysis as embryos development is very rapid.</li>
		<li>Fill a 35 mm dish (test dish) with embryo water and allow it to warm up in the incubator (28 &#176;C) for at least 15 min prior to starting the test.</li>
		<li>Place a plastic mesh grid (1.2x1.2 mm) cut to size, on the bottom of the test dish.</li>
		<li>Have another experimenter randomize the order of testing of the embryos and mask the names of the conditions to be tested.&#160;</li>
		<li>Make sure that the mortality rate does not change among conditions and compared to non-injected embryos to ensure the specificity of the phenotype. The percent of dead embryos in all conditions shall not exceed 10-13%18.</li>
		<li>Place 10-12 embryos still within their chorion on the plastic mesh using a plastic Pasteur pipette. Fill the test dish with enough embryo water to keep the embryos submerged but not floating. If needed, move the embryos with care using a plastic tip to position them on the grid.</li>
		<li>Using a video camera (see Table of Materials) attached to a dissection microscope, record the spontaneous coiling activity for a defined length of time (10-20 min long videos are usually sufficient to obtain representative samples of activity bursts for the quantification)&#160;</li>
		<li>Return the embryos to their respective dish and put them back in the incubator. Repeat the experiment with as many embryos as needed for each condition (as determined by a 90% power analysis).</li>
		<li>To analyze total spontaneous movement, use a ZebraLab system (see Table of Materials). Using the activity quantification module, upload the recorded video and design the tracking arenas around each embryo as appropriate. Set the freeze and burst threshold to 10 and 50, respectively.&#160;</li>
		<li>Run the automated video analysis, which quantifies total activity inside each of the defined arenas, then recover the data set as a spreadsheet and perform the analysis using a data analysis software.</li>
	</ol>
	</li>
	<li>Touch-Evoked-Escape-Response (TEER) at 48 hpf:
	<ol>
		<li>Conduct the test in the morning two days after the injection (48 h after the fertilization).</li>
		<li>At least 2 h prior to testing, dechorionate the embryos using fine forceps. Ensure that the time of day for the dechorionation and the behavior test is consistent over experiments.&#160;</li>
		<li>Fill a 130 mm dish (test dish) with embryo water and allow it to warm up in the incubator (28 &#176;C) for at least 15 min prior to starting the test.</li>
		<li>Count and remove dead and morphologically deformed larvae. Record the numbers for each condition.</li>
		<li>Have another experimenter randomize the order and codify the names of the conditions to be tested.</li>
		<li>Mount the camera (see Table of Materials) over the test dish making sure that the entire test dish is within the field of view. Placing a ruler within the field of view provides an internal calibration for distance.</li>
		<li>With a plastic Pasteur pipette, put an embryo in the center of the test dish and begin the recording using an acquisition rate of 30 fps.</li>
		<li>With a fine plastic tip, touch slightly the tail of the embryo with a flicking motion.</li>
		<li>Stop the recording when the larva has terminated its movement.</li>
		<li>Remove the embryo from the test dish and place it in a new dish filled with embryo water. Repeat the test with as many embryos as needed for each condition (as determined by a 90% power analysis).</li>
		<li>Return the embryos to their original dish and put them back in the incubator.</li>
		<li>To analyze the parameters of the swimming behavior, load the recorded video to ImageJ analysis software. Download and install the Manual Tracking plugin of ImageJ (see Table of Materials). Launch the plugin by choosing Tools | Plugin | Manual Tracking in the menu.&#160;</li>
		<li>In the dialog window, introduce the calibrated scale of the image. Including a ruler in the camera field facilitates the conversion of cm to pixels.&#160;</li>
		<li>Select Add track and start the trajectory tracing by clicking on the image of the zebrafish larva in the first frame. The frames advance automatically with each point that is added to the trace.&#160;</li>
		<li>Continue tracing the movement until the end of the swimming episode.&#160;</li>
		<li>Select End track on tracking window, retrieve the X-Y coordinates and calculate total distance, velocity and turning angle.</li>
	</ol>
	</li>
</ol>
3. Electrophysiological Analysis
<ol>
	<li>Reagent and tool preparation:
	<ol>
		<li>Prepare 1% agarose in embryo water (see section 1.1.4). Aliquot the liquid agarose in microcentrifuge tubes and keep these on a heating block at 42 &#176;C to prevent the agarose from hardening.</li>
		<li>Prepare the recording solution (in mmol/L): NaCl 134, KCl 2.9, CaCl2 2.1, MgCl2 1.2, glucose 10, HEPES 10, pH 7.8.</li>
		<li>Pull borosilicate glass micropipettes with a tip opening of 1.5-2 &#181;m (5-6 Mm2&#183;kg&#183;s&#8722;3&#183;A&#8722;2&#160;resistance) unpolished.</li>
	</ol>
	</li>
	<li>Preparation of zebrafish larvae for electrophysiology:&#160;
	<ol>
		<li>Place the fish in a glass-bottom Petri dish (see Table of Materials) and remove excess extracellular media in order to ensure the fish is brought as close to the cover slip as possible.&#160;</li>
		<li>Using a plastic Pasteur pipette, add warm liquid agarose on and around the larva. Use just enough agarose to cover the fish. While the agarose hardens, use fine forceps to orient the fish in a straight position, ventral side down, in the center of the dish.&#160;</li>
		<li>Add 2 mL of the recording solution containing 10 &#181;M Pancuronium bromide (see Table of Materials) to block neuromuscular transmission. The addition of the paralyzer is necessary to eliminate artefacts due to small movements during the recordings.&#160;</li>
	</ol>
	</li>
	<li>Electrophysiological recording
	<ol>
		<li>Fill the micropipette with recording solution.</li>
		<li>With the patch clamp amplifier (see Table of Materials) in voltage clamp configuration, measure electrode resistance in bath to confirm its correct value.</li>
		<li>Using a 20x objective, position the head of the larva in the central field of view and lower the micropipette to reach the recording position in the brain, within the optic tectum.</li>
		<li>Switch the patch clamp amplifier to current clamp and fix the holding current to 0 mA.</li>
		<li>Using a low-pass filter of 1 kHz, an acquisition rate of 1 kHz and a digital gain of 10, record spontaneous activity for 60 min for determining baseline activity levels.</li>
		<li>After 1 h of baseline recording, add 200 &#181;L pentylenetetrazol (PTZ, see Table of Materials) solution 300 mmol/L to the bath for a final concentration of 20 mmol/L PTZ.</li>
		<li>Record the neuronal activity in PTZ for another 120 min.</li>
	</ol>
	</li>
	<li>Depolarization event determination
	<ol>
		<li>Field recording events have very slow dynamics (frequencies of interest are in the range of 0.005-0.2 s-1). Therefore, filter the signal with a low pass (Butterworth 5th order LPF at 100 s-1) in order to avoid aliasing. Subsample the recorded voltage data from the acquisition frame rate (in this case, 1 ks-1) down to 250 s-1 (RAW SIGNAL).</li>
		<li>To identify the timestamps for each depolarization event, use a DETECTION SIGNAL, which is a high-pass filtered version of the recorded signal (Butterworth 1st order HPF at 0.01 s-1).&#160;</li>
		<li>By eliminating the low frequency components, the detection of depolarization events can be performed using a simple thresholding method. Use a fixed threshold for noise elimination and event detection (0.3 mV was used for this study).</li>
		<li>Characterize the depolarization event by a series of thresholds crossings that occur at time intervals smaller than 4 s. Compute the start and the end of depolarization event as determined from consecutive sequences of threshold crossings. Events that are shorter than 40 ms can be discarded as noise.</li>
		<li>Compute the amplitude of the events in the unfiltered (RAW SIGNAL) to eliminate errors due to the effect of the low pass filtering on the peak of the event. Select the depolarization wavelet from the raw signal using the timestamps determined in the filtered signal. Measure the amplitude as the difference between the maximum and minimum values of the wavelet selected from the raw signal. 
		NOTE: The script files to perform step 3.4 &#8212; Depolarization event determination &#8212; and to obtain Figure 1 are provided as Supplementary File attached to this article.</li>
	</ol>
	</li>
</ol>

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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+in+epilepsy+research:+Legacies+and+new+directions.">Animal models in epilepsy research: Legacies and new directions.</a> Nature Neuroscience. , (2015).</li><li>Zhang, Y., 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=Pharmacological+characterization+of+an+antisense+knockdown+zebrafish+model+of+Dravet+syndrome:+Inhibition+of+epileptic+seizures+by+the+serotonin+agonist+fenfluramine.">Pharmacological characterization of an antisense knockdown zebrafish model of Dravet syndrome: Inhibition of epileptic seizures by the serotonin agonist fenfluramine.</a> PLoS ONE. , (2015).</li><li>Swaminathan, A., 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=Non-canonical+mTOR-Independent+Role+of+DEPDC5+in+Regulating+GABAergic+Network+Development.">Non-canonical mTOR-Independent Role of DEPDC5 in Regulating GABAergic Network Development.</a> Current Biology. , (2018).</li><li>Samarut, &. #. 2. 0. 1. ;., 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=&#947;-Aminobutyric+acid+receptor+alpha+1+subunit+loss+of+function+causes+genetic+generalized+epilepsy+by+impairing+inhibitory+network+neurodevelopment.">&#947;-Aminobutyric acid receptor alpha 1 subunit loss of function causes genetic generalized epilepsy by impairing inhibitory network neurodevelopment.</a> Epilepsia. , 2061-2074 (2018).</li><li>Turrini, L., 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=Optical+mapping+of+neuronal+activity+during+seizures+in+zebrafish.">Optical mapping of neuronal activity during seizures in zebrafish.</a> Scientific Reports. , (2017).</li><li>Rosch, R. E., Hunter, P. R., Baldeweg, T., Friston, K. J., Meyer, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+imaging+and+dynamic+causal+modelling+reveal+brain-wide+changes+in+effective+connectivity+and+synaptic+dynamics+during+epileptic+seizures.">Calcium imaging and dynamic causal modelling reveal brain-wide changes in effective connectivity and synaptic dynamics during epileptic seizures.</a> PLoS Computational Biology. , (2018).</li><li>Baraban, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Forebrain+Electrophysiological+Recording+in+Larval+Zebrafish.">Forebrain Electrophysiological Recording in Larval Zebrafish.</a> Journal of Visualized Experiments. , (2013).</li><li>Afrikanova, T., 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=Validation+of+the+Zebrafish+Pentylenetetrazol+Seizure+Model:+Locomotor+versus+Electrographic+Responses+to+Antiepileptic+Drugs.">Validation of the Zebrafish Pentylenetetrazol Seizure Model: Locomotor versus Electrographic Responses to Antiepileptic Drugs.</a> PLoS ONE. , (2013).</li></ol>

The authors have nothing to disclose.

Epilepsy is a complex neurological disease, featuring a wide range of etiologies that are starting to be elucidated with the advent of genetic sequencing technologies25,26,27. Versatile animal models are essential for an efficient translational strategy that will yield both insights into the pathological mechanisms of genetically linked epilepsies, as well as targeted therapies for the distinct forms of this condition. Zebrafish models have been very effective at reproducing major features of epilepsy and providing reliable readouts for anti-epileptic drug screening5,28. Spontaneous seizures can be detected in genetically modified zebrafish15,29,30,31 and neurophysiological analysis in these models28 has confirmed the neuronal basis of the epileptic-like behavior32,33. Small-sized zebrafish larvae are amenable to chemical screens in 96-well format using automated detection of simple behavior, such as spontaneous swimming, which allows for rapid detection of potential therapeutics.
The DEPDC5 knock-down model presented here is obtained by injection of AMO into the zebrafish embryo to block gene expression during development. This model presents several keystone phenotypic features during different time points of larval development, which can be used as indicators of therapy efficiency during a chemical or genetic screening protocol. The AMO-mediated gene knock-down is a powerful technique, displaying advantages over chemically-induced seizure models, as it specifically targets the expression of a gene of interest, thus allowing the identification of the underlying pathogenic mechanisms triggered by a genetic mutation. Chemical inducers, which are nevertheless potent tools for drug screenings, can act through multiple cellular pathways that might not be always relevant to the genetic mutation under study. While AMO injection is in itself a simple technique when mastered by the experimenter, it also presents a number of limitations. The injections have to be performed at the one cell stage embryo; in our hands, injections at later stages greatly increased the variability of the phenotype. This limits the time available for injection; therefore, a strategy of generating eggs for injection in a time sequence is useful. We routinely use 4-5 crosses that we open at 15-20 min intervals, allowing the injection of one clutch before obtaining the next one. Further, care must be taken to assess the phenotype at the same time points between different experiments, as stereotyped behaviors evolve rapidly during the first days of development. The volume and concentration of AMOs must also be carefully controlled, as general toxicity due to injecting excessive amounts will mask the specific phenotype. The different controls presented in the introduction are essential for determining the right injection dose and the corresponding phenotype.
Field recordings of the larval zebrafish brain are a useful tool for investigating the deleterious effects of genetic mutations involved in different brain disorders on the global neuronal activity34. Depolarization events seen under these experimental conditions are an established method for assessing electrophysiological effects of drugs in different epileptic conditions15,35. However, the assessment of these effects has mostly been done qualitatively rather than quantitatively, and having a subjective observer as an actor in the analysis. Here, we develop an automatic detection strategy that can objectively quantify the rate of depolarizations, their amplitude and duration, and can evaluate the progress of these parameters across time, or with different genetic or pharmacologic interventions.&#160;
The representative results presented here show the expected field activity of the DEPDC5 knock-down genetic model in comparison to a Mismatch control in 4-6 dpf zebrafish, before and after the application of PTZ to introduce epileptiform-like electrographic activity. Previously, we have shown a significant increase in the basal activity of the DEPDC5 knockdown condition18. Here, we show that the response of these two conditions to PTZ, a chemical epileptiform activity inducer, has a similar trajectory in time, starting with a period of relatively low frequency, high amplitude depolarization events and continuing with a period of higher frequency, lower amplitude depolarization events. Field recording events have slow dynamics (frequencies of interest are in the range of 0.005-0.2 s-1), therefore both low-pass and high-pass filters are used in this protocol to isolate the events of interest. After eliminating the low frequency noise, the detection of depolarization events is performed using a simple threshold. Since the statistics of the signal are greatly affected by the presence of depolarization events, we could not use the standard deviation of the total signal to determine this threshold. The variability of the value of the standard deviation across datasets was greater than the observed recording noise levels. Therefore, after visual inspection of the traces, we used a fixed value of the threshold of 0.3 mV, in order to avoid the biasing induced by different levels of depolarization activity.
The described protocol provides a standardized and simple method for evaluating the motor behavior and the neuronal field activity, via extracellular current clamp voltage recording coupled with automatic detection of depolarization events in the optic tectum, to characterize epileptiform-like phenotypes in zebrafish models.

Due to its small size, oviparous development and transparency at early stages of development, zebrafish has emerged as a valuable vertebrate organism for modeling human diseases as diverse as cardiovascular, cancer or neurological disorders1,2. Zebrafish combines the advantages of a vertebrate, including the high conservation of organ architecture and genetic code, with the small size and ease of genetic manipulation of simpler model organisms, therefore facilitating both fundamental studies and translational applications. In particular, its amenability to high-throughput automated screening of behavior and fluorescent markers of cellular processes has made zebrafish a particularly attractive model for epilepsy research. This has been demonstrated by a high increase in the last decade of the number of publications featuring chemically-induced and/or genetic models of epilepsy3,4,5 and, more recently, reports of promising therapeutics obtained from chemical screens in these models6,7,8.&#160;
DEPDC5 is a member of the GATOR1 complex, a negative regulator of mTOR signaling9. Mutations in the DEPDC5 gene have been first discovered in 2013 in probands suffering from autosomal dominant focal epilepsies10,11, and have since been reported in a number of clinical conditions associated with focal epileptic manifestations and focal cortical dysplasia12. The large majority of reported mutations are predicted to cause the loss-of-function of the gene12, and this was formally demonstrated for a number of DEPDC5 mutated transcripts which are targeted by nonsense mediated mRNA decay12,13. In agreement, knock-down of the gene orthologue in zebrafish using antisense morpholino oligonucleotides (AMOs) results in a number of features that are common to epileptic models in this organism, including hyperactivity, turn wheel-like swimming, spontaneous seizures and enhanced neuronal activity14,15,16,17,18. Interestingly, treatment with rapamycin, an inhibitor of mTOR signaling, reversed the behavioral features of this model18, supporting the hypothesis that DEPDC5 loss-of-function can trigger epilepsy due to a misregulation of the mTOR pathway9,19.&#160;
Transient knock-down of gene expression in vivo using antisense oligonucleotides carrying the morpholino modification has been an invaluable tool for studying the role of specific genes, on par with si/shRNA-based techniques. Recently, AMO-based strategies have also found clinical applications, with a first AMO therapy receiving the FDA approval for the treatment of Duchenne muscular atrophy in 201620. While it was reported that in zebrafish the phenotype of acute AMO-based gene knock-down does not always correlate with the constitutive knock-out models21, this can be due at least in some instances to compensatory mechanisms engendered by constitutive genetic modifications22. However, the issue of specificity of the AMO-induced phenotype is an undisputable concern that has to be diligently addressed in studies using this technology23. In order to ensure the specificity of the AMO-based knock-down phenotype, several key controls are necessary. These include a dose-response curve that allows the selection of the lowest dose of AMO effective for gene knock-down, avoiding overall toxicity due to the introduction of an excess of genetic material. The use of a Mismatch AMO that does not target any particular region in the genome is also required for establishing an appropriate dose and in identifying a specific phenotype. A second AMO which targets a different region of the same gene, such as a splice-blocking AMO, is necessary to confirm that the phenotype is due to the knock-down of the target gene. Rescue of the knock-down phenotype with the cDNA of the gene, either the human orthologue or a codon-modified version of the zebrafish gene that cannot be targeted by the AMO, provides a strong argument in favor of the phenotype specificity. Lack of rescue with the same cDNA containing loss-of-function mutations (such as the introduction of an early stop codons) is a further proof in this direction.&#160;
Here, we present a method for generating a zebrafish DEPDC5 loss-of-function model and the protocol for behavioral phenotyping at 28 and 48 h post fertilization (hpf). At 28 hpf, DEPDC5 loss-of-function causes overall hyperactivity, as evidenced by enhanced coiling and twitching movements of the embryos within the chorion. An automated motion detection system can be used at this stage to quantify the overall activity per embryo. At 48 hpf, zebrafish exhibit stereotyped escape swimming in response to touch. In zebrafish with downregulated expression of DEPDC5, the swimming trajectory is significantly more tortuous than in controls, the fish exhibiting a &#8220;cork-screw&#8221; or &#8220;turn-wheel&#8221; like pattern, similar to other reported epilepsy models in this organism3,4. Electrophysiological recordings were obtained in the optic tectum in zebrafish larvae between 4-6 days post fertilization (dpf) and show a baseline increase in neuronal activity in the DEPDC5 knock-down animals. The advantage of this model is that it presents several phenotypic features at different time points, which can be useful in monitoring and assessing the efficacy of drug therapies during development.&#160;

<table><tbody><tr><td>Agarose</td><td>Sigma-Aldrich, France</td><td>A9539</td><td></td></tr><tr><td>Aquarium salt&amp;nbsp;</td><td>Instant Ocean, Blacksburg, VA</td><td>SS15-10</td><td></td></tr><tr><td>Borosilicate glass with filament</td><td>Sutter Instruments</td><td>BF100-50-10</td><td>OD: 1.5mm, ID: 0.5 mm</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich, France</td><td>C1016</td><td></td></tr><tr><td>Depdc5-atg antisense morpholino</td><td>GeneTools, OR, USA</td><td>N/A</td><td>sequence 5&amp;rsquo;- TGCCTTCATGGTGACCGTCATTTTA -3&amp;rsquo;&amp;nbsp;</td></tr><tr><td>Depdc5-mis antisense morpholino</td><td>GeneTools, OR, USA</td><td>N/A</td><td>sequence 5&amp;rsquo;- TGCgTTgATcGTGACCcTgATTTTA -3&amp;rsquo;&amp;nbsp;</td></tr><tr><td>Depdc5-splice antisense morpholino</td><td>GeneTools, OR, USA</td><td>N/A</td><td>sequence 5&amp;rsquo;- ACATTCCTGTTTCACCATAGATGAT -3&amp;rsquo;</td></tr><tr><td>Digitizer</td><td>Molecular Devices, CA, USA</td><td>Digidata 1550</td><td></td></tr><tr><td>Fast Green Dye</td><td>Sigma-Aldrich, France</td><td>F7258</td><td>Stock solution of 0.2%</td></tr><tr><td>Glass-bottom petri dishes</td><td>Ibidi, Germany</td><td>81218</td><td></td></tr><tr><td>Glucose</td><td>Sigma-Aldrich, France</td><td>68270</td><td></td></tr><tr><td>Grasshopper 2 camera</td><td>FLIR, BC, Canada</td><td>GRAS-03K2M-C</td><td>formerly Point Grey Research</td></tr><tr><td>HEPES</td><td>Sigma-Aldrich, France</td><td>H3375</td><td></td></tr><tr><td>Human wild-type DEPDC5 cDNA</td><td>Dharmacon, France</td><td>NM_001242897.1</td><td>&amp;nbsp;Accession: BC144291 Clone ID 905</td></tr><tr><td>ImageJ software</td><td>NIH, USA</td><td>N/A</td><td></td></tr><tr><td>KCl</td><td>Sigma-Aldrich, France</td><td>P9333&amp;nbsp;</td><td></td></tr><tr><td>Matlab software</td><td>MathWorks, MA, USA</td><td>N/A</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich, France</td><td>M2670</td><td></td></tr><tr><td>NaCl</td><td>Sigma-Aldrich, France</td><td>S7653&amp;nbsp;</td><td></td></tr><tr><td>NaOH</td><td>Sigma-Aldrich, France</td><td>71687</td><td></td></tr><tr><td>Pancuronium bromide&amp;nbsp;</td><td>Alomone Labs</td><td>P-130</td><td>Stock solution of 60 mM in water</td></tr><tr><td>Parafilm</td><td>Sigma-Aldrich, France</td><td>P7793</td><td></td></tr><tr><td>Patch clamp amplifier</td><td>Molecular Devices, CA, USA</td><td>MultiClamp 700B</td><td>Computer-controled patch clamp amplifier</td></tr><tr><td>pClamp10 acquisition software</td><td>Molecular Devices</td><td>N/A</td><td></td></tr><tr><td>Pentylenetetrazol (PTZ)</td><td>Sigma-Aldrich, France</td><td>P6500</td><td>Stock solution of 300 mM (dissolved in recording solution)</td></tr><tr><td>Pipette puller</td><td>Narishige, Japan</td><td>PC-10</td><td></td></tr><tr><td>Pneumatic PicoPump&amp;nbsp;</td><td>WPI, France</td><td>PV 820</td><td></td></tr><tr><td>Sylgard 184 kit</td><td>Sigma-Aldrich Intl.</td><td>761036</td><td></td></tr><tr><td>Transfer plastic pipettes</td><td>Sigma-Aldrich, France</td><td>Z350605</td><td></td></tr><tr><td>Zebralab</td><td>Viewpoint, France</td><td>N/A</td><td></td></tr></tbody></table>

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.

Figure 1 shows representative voltage traces of 4-6 dpf zebrafish larva extracellular field recordings in the case of two genetic conditions: Mismatch control and DEPDC5 knock-down. In the baseline period of the recording, DEPDC5 knock-down shows a higher occurrence of spontaneous events, while the Mismatch control displays very few fluctuations. These activity patterns are representative of the significant increase in neuronal activity due to loss-of-function of DEPDC5, as we have previously reported18. After PTZ application, both Mismatch control and DEPDC5 knock-down show an increased number of depolarization events. During the first period after PTZ application (10 &#8211; 60 min), a rate of 0.8 events per min is observed in both Mismatch control and DEPDC5 knock-down, where the majority of events are of high amplitude (&#62;1 mV). During the latter response period (60 &#8211; 120 min after PTZ application), the rate of depolarization events increases to around 1 event per min, and the majority of the events are of low amplitude (&#8804;1 mV).&#160;
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58837/58837fig1.jpg" /> 
Figure 1: Example traces of field recordings in the zebrafish larvae brain. (A) Overview of 180 min recording for a Mismatch control larva and a DEPDC5 Knock-down. First, spontaneous baseline activity was recorded, then PTZ was applied in bath (red bar). (B) Peri-stimulus time histograms of the depolarization events for Mismatch control and DEPDC5 knock-down. The events were classified as high amplitude (&#62;1 mV - blue) and low amplitude (&#8804;1 mV - black). (C-E) Example traces of the different periods of the recording: (C) spontaneous activity, (D) High amplitude events during the first period after PTZ application, (E) Low amplitude events during the latter period after PTZ application. Note that the script files to obtain these figures are provided as Supplementary File. <a href="https://www.jove.com/files/ftp_upload/58837/58837fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary File: Script files for step 3.4. <a href="https://www.jove.com/files/ftp_upload/58837/Supplementary_File.zip" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Behavioral And Physiological Analysis In A Zebrafish Model Of Epilepsy at JoVE.com

Behavioral And Physiological Analysis In A Zebrafish Model Of Epilepsy

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 ...

behavioral-and-physiological-analysis-in-a-zebrafish-model-of-epilepsy

Research

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Neuroscience

5.5K Views.  Institut Imagine. 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.

Video: Behavioral And Physiological Analysis In A Zebrafish Model Of Epilepsy

Simultaneous Video-EEG-ECG Monitoring to Identify Neurocardiac Dysfunction in Mouse Models of Epilepsy

In epilepsy, seizures can evoke cardiac rhythm disturbances such as heart rate changes, conduction blocks, asystoles, and arrhythmias, which can potentially increase risk of sudden unexpected death in epilepsy (SUDEP). Electroencephalography (EEG) and electrocardiography (ECG) are widely used clinical diagnostic tools to monitor for abnormal brain and cardiac rhythms in patients. Here, a technique to simultaneously record video, EEG, and ECG in mice to measure behavior, brain, and cardiac activities, respectively, is described. The technique described herein utilizes a tethered (i.e., wired) recording configuration in which the implanted electrode on the head of the mouse is hard-wired to the recording equipment. Compared to wireless telemetry recording systems, the tethered arrangement possesses several technical advantages such as a greater possible number of channels for recording EEG or other biopotentials; lower electrode costs; and greater frequency bandwidth (i.e., sampling rate) of recordings. The basics of this technique can also be easily modified to accommodate recording other biosignals, such as electromyography (EMG) or plethysmography for assessment of muscle and respiratory activity, respectively. In addition to describing how to perform the EEG-ECG recordings, we also detail methods to quantify the resulting data for seizures, EEG spectral power, cardiac function, and heart rate variability, which we demonstrate in an example experiment using a mouse with epilepsy due to Kcna1 gene deletion. Video-EEG-ECG monitoring in mouse models of epilepsy or other neurological disease provides a powerful tool to identify dysfunction at the level of the brain, heart, or brain-heart interactions.

Here, we present a protocol to record brain and heart bio signals in mice using simultaneous video, electroencephalography (EEG), and electrocardiography (ECG). We also describe methods to analyze the resulting EEG-ECG recordings for seizures, EEG spectral power, cardiac function, and heart rate variability.

In epilepsy, seizures can evoke cardiac rhythm disturbances such as heart rate changes, conduction blocks, asystoles, and arrhythmias, which can ...

Genetics

High-throughput DNA Extraction and Genotyping of 3dpf Zebrafish Larvae by Fin Clipping

Zebrafish (Danio rerio) possess orthologues for 84% of the genes known to be associated with human diseases. In addition, these animals have a short generation time, are easy to handle, display a high reproductive rate, low cost, and are easily amenable to genetic manipulations by microinjection of DNA in embryos. Recent advances in gene editing tools are enabling precise introduction of mutations and transgenes in zebrafish. Disease modeling in zebrafish often leads to larval phenotypes and early death which can be challenging to interpret if genotypes are unknown. This early identification of genotypes is also needed in experiments requiring sample pooling, such as in gene expression or mass spectrometry studies. However, extensive genotypic screening is limited by traditional methods, which in most labs are performed only on adult zebrafish or in postmortem larvae. We addressed this problem by adapting a method for the isolation of PCR-ready genomic DNA from live zebrafish larvae that can be achieved as early as 72 h post-fertilization (hpf). This time and cost-effective technique, improved from a previously published genotyping protocol, allows the identification of genotypes from microscopic fin biopsies. The fins quickly regenerate as the larvae develop. Researchers are then able to select and raise the desired genotypes to adulthood by utilizing this high-throughput PCR-based genotyping procedure.

Zebrafish have been used as reliable genetic model organisms in biomedical research, especially with the advent of gene-editing technologies. When larval phenotypes are expected, DNA extraction and genotype identification can be challenging. Here, we describe an efficient genotyping procedure for zebrafish larvae, by tail clipping, as early as 72-h post-fertilization.

Zebrafish (Danio rerio) possess orthologues for 84% of the genes known to be associated with human diseases. In addition, these animals have a short ...

Behavioral Approaches to Studying Innate Stress in Zebrafish

Responding appropriately to stressful stimuli is essential for survival of an organism. Extensive research has been done on a wide spectrum of stress-related diseases and psychiatric disorders, yet further studies into the genetic and neuronal regulation of stress are still required to develop better therapeutics. The zebrafish provides a powerful genetic model to investigate the neural underpinnings of stress, as there exists a large collection of mutant and transgenic lines. Moreover, pharmacology can easily be applied to zebrafish, as most drugs can be added directly to water. We describe here the use of the 'novel tank test' as a method to study innate stress responses in zebrafish, and demonstrate how potential anxiolytic drugs can be validated using the assay. The method can easily be coupled with zebrafish lines harboring genetic mutations, or those in which transgenic approaches for manipulating precise neural circuits are used. The assay can also be used in other fish models. Together, the described protocol should facilitate the adoption of this simple assay to other laboratories.

This manuscript describes a simple method to measure stress behaviorally in adult zebrafish. The approach takes advantage of the innate tendency that zebrafish prefer the bottom half of a tank when in a stressful state. We also describe methods for coupling the assay with pharmacology.

Responding appropriately to stressful stimuli is essential for survival of an organism. Extensive research has been done on a wide spectrum of ...

Behavior

Forebrain Electrophysiological Recording in Larval Zebrafish

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked seizures, epilepsy can be acquired as a result of an insult to the brain or a genetic mutation. Efforts to model seizures in animals have primarily utilized acquired insults (convulsant drugs, stimulation or brain injury) and genetic manipulations (antisense knockdown, homologous recombination or transgenesis) in rodents. Zebrafish are a vertebrate model system1-3 that could provide a valuable alternative to rodent-based epilepsy research. Zebrafish are used extensively in the study of vertebrate genetics or development, exhibit a high degree of genetic similarity to mammals and express homologs for ~85% of known human single-gene epilepsy mutations. Because of their small size (4-6 mm in length), zebrafish larvae can be maintained in fluid volumes as low as 100 &mu;l during early development and arrayed in multi-well plates. Reagents can be added directly to the solution in which embryos develop, simplifying drug administration and enabling rapid in vivo screening of test compounds4. Synthetic oligonucleotides (morpholinos), mutagenesis, zinc finger nuclease and transgenic approaches can be used to rapidly generate gene knockdown or mutation in zebrafish5-7. These properties afford zebrafish studies an unprecedented statistical power analysis advantage over rodents in the study of neurological disorders such as epilepsy. Because the "gold standard" for epilepsy research is to monitor and analyze the abnormal electrical discharges that originate in a central brain structure (i.e., seizures), a method to efficiently record brain activity in larval zebrafish is described here. This method is an adaptation of conventional extracellular recording techniques and allows for stable long-term monitoring of brain activity in intact zebrafish larvae. Sample recordings are shown for acute seizures induced by bath application of convulsant drugs and spontaneous seizures recorded in a genetically modified fish.

A simple method to record extracellular field potentials in the larval zebrafish forebrain is described. The method provides a robust in vivo read-out of seizure-like activity. This technique can be used with genetically modified zebrafish larvae carrying epilepsy-related genes or seizures evoked by administration of convulsant drugs.

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked ...

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.

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 ...

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 ...

Pharmacological Validation of the Prepulse Inhibition of Startle Response in Larval Zebrafish using a Commercial Automated System and Software

While there is an abundance of commercial and standardized automated systems and software for performing the prepulse inhibition (PPI) assay in rodents, to the best of our knowledge, all PPI assays performed in the zebrafish have, until now, been done using custom made systems which were only available to individual groups. This has thereby presented challenges, particularly with regard to issues of data reproducibility and standardization. In the present work, we generated a protocol that utilizes commercially available automated systems to pharmacologically validate the PPI&#160;assay in larval zebrafish. Consistent with published findings, we were able to replicate the results of apomorphine, haloperidol and ketamine on the PPI response of 6 days post-fertilization zebrafish larvae.

Here we describe a protocol that utilizes commercially available automated systems to pharmacologically validate the prepulse inhibition (PPI) assay in larval zebrafish.

While there is an abundance of commercial and standardized automated systems and software for performing the prepulse inhibition (PPI) assay in rodents, ...

Zebra II: Advanced ECG System for Multichannel Zebrafish Cardiovascular Studies

Zebra II as A Novel System to Record Electrophysiological Signals in Zebrafish

Zebrafish and their mutant lines have been extensively used in biomedical investigations, cardiovascular studies, and drug screening. In the current study, the commercial version of the novel system, Zebra II, is presented. The protocol demonstrates electrocardiogram (ECG) acquisition and analysis from multiple zebrafish within controllable working environments. The device is composed of an external and independent perfusion system, a 4-point electrode, temperature sensors, and an embedded electronic system. In previous studies, the device prototype underwent validation against the established iWORX system through several tests, demonstrating similar data quality and ECG response to drug interventions. Following this, the study delved into examining the impact of anesthetic drugs and temperature fluctuations on zebrafish ECG, necessitating instant data evaluation. Thanks to the apparatus&#39;s capacity for consistent delivery of anesthetics and drugs, it was possible to extend ECG data collection up to 1 h, markedly longer than the 5 min duration supported by current systems. This paper introduces a pioneering, cloud-based, automated analysis utilizing data from four zebrafish, offering an efficient method for conducting combination experiments and significantly reducing time and effort. The system proved effective in capturing and analyzing ECG, especially in detecting drug-induced arrhythmias in wild-type zebrafish. Additionally, the capability to gather data across multiple channels facilitated the execution of randomized controlled trials with zebrafish models. The developed ECG system overcomes existing limitations, showing the potential to greatly expedite drug discovery and cardiovascular research involving zebrafish.

The study introduces Zebra II, an advanced system for prolonged ECG acquisition and analysis in zebrafish. This system features an independent perfusion system and multiple-point electrodes for handling multiple fish in controlled environments. Acute effects of Amiodarone were assessed and analyzed with wild-type zebrafish.

Zebrafish and their mutant lines have been extensively used in biomedical investigations, cardiovascular studies, and drug screening. In the current ...

Bioengineering

Behavior Tracking and Neural Activity Pattern in the Zebrafish Forebrain

Two-Photon Calcium Imaging of Forebrain Activity in Behaving Adult Zebrafish

Adult zebrafish (Danio rerio) exhibit a rich repertoire of behaviors for studying cognitive functions. They also have a miniature brain that can be used for measuring activities across brain regions through optical imaging methods. However, reports on the recording of brain activity in behaving adult zebrafish have been scarce. The present study describes procedures to perform two-photon calcium imaging in the dorsal forebrain of adult zebrafish. We focus on steps to restrain adult zebrafish from moving their heads, which provides stability that enables laser scanning imaging of the brain activity. The head-restrained animals can freely move their body parts and breathe without aids. The procedure aims to shorten the time of head restraint surgery, minimize brain motion, and maximize the number of neurons recorded. A setup for presenting an immersive visual environment during calcium imaging is also described here, which can be used to study neural correlates underlying visually triggered behaviors.

Author Spotlight: Advancements in Adult Zebrafish Brain Research 

Here, we present a protocol to perform two-photon calcium imaging in the dorsal forebrain of adult zebrafish.

Adult zebrafish (Danio rerio) exhibit a rich repertoire of behaviors for studying cognitive functions. They also have a miniature brain that can be used ...

Startle Assay for Toxicity Evaluation in Zebrafish Embryo

Evaluating Toxicity of Chemicals using a Zebrafish Vibration Startle Response Screening System

We developed a simple screening system for the evaluation of neuromuscular and general toxicity in zebrafish embryos. The modular system consists of electrodynamic transducers above which tissue culture dishes with embryos can be placed. Multiple such loudspeaker-tissue culture dish pairs can be combined. Vibrational stimuli generated by the electrodynamic transducers induce a characteristic startle and escape response in the embryos. A belt-driven linear drive sequentially positions a camera above each loudspeaker to record the movement of the embryos. In this way, alterations to the startle response due to lethality or neuromuscular toxicity of chemical compounds can be visualized and quantified. We present an example of the workflow for chemical compound screening using this system, including the preparation of embryos and treatment solutions, operation of the recording system, and data analysis to calculate benchmark concentration values of compounds active in the assay. The modular assembly based on commercially available simple components makes this system both economical and flexibly adaptable to the needs of particular laboratory setups and screening purposes.

Author Spotlight: High-Throughput Toxicity Screening Using Zebrafish Embryo Startle Response Assay

We describe a screening system's workflow and data analysis for evaluating chemical compound toxicity based on the zebrafish embryo vibration startle response. The system records the movements of zebrafish embryos upon exposure to a vibration stimulus and allows for an integrated evaluation of general toxicity/lethality and neuromuscular toxicity.

We developed a simple screening system for the evaluation of neuromuscular and general toxicity in zebrafish embryos. The modular system consists of ...