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 border="0" cellpadding="0" cellspacing="0" style="width:751px;" width="750">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:197px;">Agarose</td>
			<td style="width:163px;">Sigma-Aldrich, France</td>
			<td style="width:84px;">A9539</td>
			<td style="width:307px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Aquarium salt&#160;</td>
			<td>Instant Ocean, Blacksburg, VA</td>
			<td>SS15-10</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Borosilicate glass with filament</td>
			<td>Sutter Instruments</td>
			<td>BF100-50-10</td>
			<td>OD: 1,5mm, ID: 0,5 mm</td>
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		<tr height="21">
			<td height="21" style="height:21px;">CaCl2</td>
			<td>Sigma-Aldrich, France</td>
			<td>C1016</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Depdc5-atg antisense morpholino</td>
			<td>GeneTools, OR, USA</td>
			<td>N/A</td>
			<td>sequence 5&#8217;- TGCCTTCATGGTGACCGTCATTTTA -3&#8217;&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Depdc5-mis antisense morpholino</td>
			<td>GeneTools, OR, USA</td>
			<td>N/A</td>
			<td>sequence 5&#8217;- TGCgTTgATcGTGACCcTgATTTTA -3&#8217;&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Depdc5-splice antisense morpholino</td>
			<td>GeneTools, OR, USA</td>
			<td>N/A</td>
			<td>sequence 5&#8217;- ACATTCCTGTTTCACCATAGATGAT -3&#8217;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Digitizer</td>
			<td>Molecular Devices, CA, USA</td>
			<td>Digidata 1550</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Fast Green Dye</td>
			<td>Sigma-Aldrich, France</td>
			<td>F7258</td>
			<td>Stock solution of 0.2%</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Glass-bottom petri dishes</td>
			<td>Ibidi, Germany</td>
			<td>81218</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">glucose</td>
			<td>Sigma-Aldrich, France</td>
			<td>68270</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Grasshopper 2 camera</td>
			<td style="width:163px;">FLIR, BC, Canada</td>
			<td>GRAS-03K2M-C</td>
			<td>formerly Point Grey Research</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">HEPES</td>
			<td>Sigma-Aldrich, France</td>
			<td>H3375</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">human wild-type DEPDC5 cDNA</td>
			<td>Dharmacon, France</td>
			<td>NM_001242897.1</td>
			<td>&#160;Accession: BC144291 Clone ID 905</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">ImageJ software</td>
			<td>NIH, USA</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">KCl</td>
			<td>Sigma-Aldrich, France</td>
			<td>P9333&#160;</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Matlab software</td>
			<td>MathWorks, MA, USA</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">MgCl2</td>
			<td>Sigma-Aldrich, France</td>
			<td>M2670</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">NaCl</td>
			<td>Sigma-Aldrich, France</td>
			<td>S7653&#160;</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">NaOH</td>
			<td>Sigma-Aldrich, France</td>
			<td>71687</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Pancuronium bromide&#160;</td>
			<td style="width:163px;">Alomone Labs</td>
			<td style="width:84px;">P-130</td>
			<td style="width:307px;">Stock solution of 60mM in water</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Parafilm</td>
			<td>Sigma-Aldrich, France</td>
			<td>P7793</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Patch clamp amplifier</td>
			<td>Molecular Devices, CA, USA</td>
			<td>MultiClamp 700B</td>
			<td>Computer-controled patch clamp amplifier</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">pClamp10 acquisition software</td>
			<td>Molecular Devices</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Pentylenetetrazol (PTZ)</td>
			<td>Sigma-Aldrich, France</td>
			<td>P6500</td>
			<td>Stock solution of 300mM (dissolved in recording solution)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Pipette puller</td>
			<td>Narishige, Japan</td>
			<td>PC-10</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Pneumatic PicoPump&#160;</td>
			<td>WPI, France</td>
			<td>PV 820</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sylgard 184 kit</td>
			<td>Sigma-Aldrich Intl.</td>
			<td>761036</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Transfer plastic pipettes</td>
			<td>Sigma-Aldrich, France</td>
			<td>Z350605</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">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

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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

A General Method for Evaluating Incubation of Sucrose Craving in Rats

For someone on a food-restricted diet, food craving in response to food-paired cues may serve as a key behavioral transition point between abstinence and relapse to food taking 1. Food craving conceptualized in this way is akin to drug craving in response to drug-paired cues. A rich literature has been developed around understanding the behavioral and neurobiological determinants of drug craving; we and others have been focusing recently on translating techniques from basic addiction research to better understand addiction-like behaviors related to food 2-4.
As done in previous studies of drug craving, we examine sucrose craving behavior by utilizing a rat model of relapse. In this model, rats self-administer either drug or food in sessions over several days. In a session, lever responding delivers the reward along with a tone+light stimulus. Craving behavior is then operationally defined as responding in a subsequent session where the reward is not available. Rats will reliably respond for the tone+light stimulus, likely due to its acquired conditioned reinforcing properties 5. This behavior is sometimes referred to as sucrose seeking or cue reactivity. In the present discussion we will use the term "sucrose craving" to subsume both of these constructs.
In the past decade, we have focused on how the length of time following reward self-administration influences reward craving. Interestingly, rats increase responding for the reward-paired cue over the course of several weeks of a period of forced-abstinence. This "incubation of craving" is observed in rats that have self-administered either food or drugs of abuse 4,6. This time-dependent increase in craving we have identified in the animal model may have great potential relevance to human drug and food addiction behaviors.
Here we present a protocol for assessing incubation of sucrose craving in rats. Variants of the procedure will be indicated where craving is assessed as responding for a discrete sucrose-paired cue following extinction of lever pressing within the sucrose self-administration context (Extinction without cues) or as responding for sucrose-paired cues in a general extinction context (Extinction with cues).

Responding for food or drug-paired cues increases over the course of a period of abstinence and this may relate to an increased susceptibility to relapse behaviors. Here we detail a procedure for evaluating this "incubation of craving" in rats that have self-administered sucrose.

For someone on a food-restricted diet, food craving in response to food-paired cues may serve as a key behavioral transition point between abstinence and ...

A Lightweight, Headphones-based System for Manipulating Auditory Feedback in Songbirds

Experimental manipulations of sensory feedback during complex behavior have provided valuable insights into the computations underlying motor control and sensorimotor plasticity1. Consistent sensory perturbations result in compensatory changes in motor output, reflecting changes in feedforward motor control that reduce the experienced feedback error. By quantifying how different sensory feedback errors affect human behavior, prior studies have explored how visual signals are used to recalibrate arm movements2,3 and auditory feedback is used to modify speech production4-7. The strength of this approach rests on the ability to mimic naturalistic errors in behavior, allowing the experimenter to observe how experienced errors in production are used to recalibrate motor output.
Songbirds provide an excellent animal model for investigating the neural basis of sensorimotor control and plasticity8,9. The songbird brain provides a well-defined circuit in which the areas necessary for song learning are spatially separated from those required for song production, and neural recording and lesion studies have made significant advances in understanding how different brain areas contribute to vocal behavior9-12. However, the lack of a naturalistic error-correction paradigm - in which a known acoustic parameter is perturbed by the experimenter and then corrected by the songbird - has made it difficult to understand the computations underlying vocal learning or how different elements of the neural circuit contribute to the correction of vocal errors13.
The technique described here gives the experimenter precise control over auditory feedback errors in singing birds, allowing the introduction of arbitrary sensory errors that can be used to drive vocal learning. Online sound-processing equipment is used to introduce a known perturbation to the acoustics of song, and a miniaturized headphones apparatus is used to replace a songbird's natural auditory feedback with the perturbed signal in real time. We have used this paradigm to perturb the fundamental frequency (pitch) of auditory feedback in adult songbirds, providing the first demonstration that adult birds maintain vocal performance using error correction14. The present protocol can be used to implement a wide range of sensory feedback perturbations (including but not limited to pitch shifts) to investigate the computational and neurophysiological basis of vocal learning.

We describe the design and assembly of miniaturized headphones suitable for replacing a songbird&#x2019;s natural auditory feedback with a manipulated acoustic signal. Online sound processing hardware is used to manipulate song output, introduce real-time errors in auditory feedback via the headphones, and drive vocal motor learning.

Experimental manipulations of sensory feedback during complex behavior have provided valuable insights into the computations underlying motor control and ...

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 ...

Recording and Analysis of Circadian Rhythms in Running-wheel Activity in Rodents

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, including rats, hamsters, and mice, are active during the night and relatively inactive during the day. Many other behavioral and physiological measures also exhibit daily rhythms, but in rodents, running-wheel activity serves as a particularly reliable and convenient measure of the output of the master circadian clock, the suprachiasmatic nucleus (SCN) of the hypothalamus. In general, through a process called entrainment, the daily pattern of running-wheel activity will naturally align with the environmental light-dark cycle (LD cycle; e.g. 12 hr-light:12 hr-dark). However circadian rhythms are endogenously generated patterns in behavior that exhibit a ~24 hr period, and persist in constant darkness. Thus, in the absence of an LD cycle, the recording and analysis of running-wheel activity can be used to determine the subjective time-of-day. Because these rhythms are directed by the circadian clock the subjective time-of-day is referred to as the circadian time (CT). In contrast, when an LD cycle is present, the time-of-day that is determined by the environmental LD cycle is called the zeitgeber time (ZT).
Although circadian rhythms in running-wheel activity are typically linked to the SCN clock6-8, circadian oscillators in many other regions of the brain and body9-14 could also be involved in the regulation of daily activity rhythms. For instance, daily rhythms in food-anticipatory activity do not require the SCN15,16 and instead, are correlated with changes in the activity of extra-SCN oscillators17-20. Thus, running-wheel activity recordings can provide important behavioral information not only about the output of the master SCN clock, but also on the activity of extra-SCN oscillators. Below we describe the equipment and methods used to record, analyze and display circadian locomotor activity rhythms in laboratory rodents.

Circadian rhythms in voluntary wheel-running activity in mammals are tightly coupled to the molecular oscillations of a master clock in the brain. As such, these daily rhythms in behavior can be used to study the influence of genetic, pharmacological, and environmental factors on the functioning of this circadian clock.

When rodents have free access to a running wheel in their home cage, voluntary use of this wheel will depend on the time of day1-5. Nocturnal rodents, ...

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 ...

Functional Analysis of the Larval Feeding Circuit in Drosophila

The serotonergic feeding circuit in Drosophila melanogaster larvae can be used to investigate neuronal substrates of critical importance during the development of the circuit. Using the functional output of the circuit, feeding, changes in the neuronal architecture of the stomatogastric system can be visualized. Feeding behavior can be recorded by observing the rate of retraction of the mouth hooks, which receive innervation from the brain. Locomotor behavior is used as a physiological control for feeding, since larvae use their mouth hooks to traverse across an agar substrate. Changes in feeding behavior can be correlated with the axonal architecture of the neurites innervating the gut. Using immunohistochemistry it is possible to visualize and quantitate these changes. Improper handling of the larvae during behavior paradigms can alter data as they are very sensitive to manipulations. Proper imaging of the neurite architecture innervating the gut is critical for precise quantitation of number and size of varicosities as well as the extent of branch nodes. Analysis of most circuits allow only for visualization of neurite architecture or behavioral effects; however, this model allows one to correlate the functional output of the circuit with the impairments in neuronal architecture.

Functional Analysis of the Larval Feeding Circuit in Drosophila

The feeding circuit in Drosophila melanogaster larvae serves a simple yet powerful model that allows changes in feeding rate to be correlated with alterations in the stomatogastric neural circuitry. This circuit is composed of central serotonergic neurons that send projections to the mouth hooks as well as the foregut.

The serotonergic  feeding circuit in Drosophila  melanogaster larvae can be used to investigate neuronal substrates of  critical importance during the ...

Identification of Dopamine D1-Alpha Receptor Within Rodent Nucleus Accumbens by an Innovative RNA In Situ Detection Technology

In the central nervous system, the D1-alpha subtype receptor (Drd1&#945;) is the most abundant dopamine (DA) receptor, which plays a vital role in regulating neuronal growth and development. However, the mechanisms underlying Drd1&#945; receptor abnormalities mediating behavioral responses and modulating working memory function are still unclear. Using a novel RNA in situ hybridization assay, the current study identified dopamine Drd1&#945; receptor and tyrosine hydroxylase (TH) RNA expression from DA-related circuitry in the nucleus accumbens (NAc) area and substantia nigra region (SNR), respectively. Drd1&#945; expression in the NAc shows a &#34;discrete dot&#34; staining pattern. Clear sex differences in Drd1&#945; expression were observed. In contrast, TH shows a &#34;clustered&#34; staining pattern. Regarding TH expression, female rats displayed a higher signal expression per cell relative to male animals. The methods presented here provide a novel in situ hybridization technique for investigating changes in dopamine system dysfunction during the progression of central nervous system diseases.

Identification of Dopamine D1-Alpha Receptor Within Rodent Nucleus Accumbens by an Innovative RNA In Situ Detection Technology

Identification of dopamine D1-alpha receptor in the nucleus accumbens is critical for clarifying D1 receptor dysfunction during a central nervous system disease. We performed a novel RNA in situ hybridization assay to visualize single RNA molecules in a specific brain area.

In the central nervous system, the D1-alpha subtype receptor (Drd1&#945;) is the most abundant dopamine (DA) receptor, which plays a vital role in ...

The Power of Interstimulus Interval for the Assessment of Temporal Processing in Rodents

Temporal processing deficits have been implicated as a potential elemental dimension of higher-level cognitive processes, commonly observed in neurocognitive disorders. Despite the popularization of prepulse inhibition (PPI) in recent years, many current protocols promote using a percent of control measure, thereby precluding the assessment of temporal processing. The present study used cross-modal PPI and gap prepulse inhibition (gap-PPI) to demonstrate the benefits of employing a range of interstimulus intervals (ISIs) to delineate effects of sensory modality, psychostimulant exposure, and age. Assessment of sensory modality, psychostimulant exposure, and age reveals the utility of an approach varying the interstimulus interval (ISI) to establish the shape of the ISI function, including increases (sharper curve inflections) or decreases (flattening of the response amplitude curve) in startle amplitude. Additionally, shifts in peak response inhibition, suggestive of a differential sensitivity to the manipulation of ISI, are often revealed. Thus, the systematic manipulation of ISI affords a critical opportunity to evaluate temporal processing, which may reveal the underlying neural mechanisms involved in neurocognitive disorders.

Temporal processing, a preattentive process, may underlie deficits in higher-level cognitive processes, including attention, commonly observed in neurocognitive disorders. Using prepulse inhibition as an exemplar paradigm, we present a protocol for manipulating interstimulus interval (ISI) to establish the shape of the ISI function to provide an assessment of temporal processing.

Temporal processing deficits have been implicated as a potential elemental dimension of higher-level cognitive processes, commonly observed in ...

Ballistic Labeling of Pyramidal Neurons in Brain Slices and in Primary Cell Culture

It has been reported that the size and shape of dendritic spines is related to their structural plasticity. To identify the morphological structure of pyramidal neurons and dendritic spines, a ballistic labeling technique can be utilized. In the present protocol, pyramidal neurons are labeled with DilC18(3) dye and analyzed using neuronal reconstruction software to assess neuronal morphology and dendritic spines. To investigate neuronal structure, dendritic branching analysis and Sholl analysis are performed, allowing researchers to draw inferences about dendritic branching complexity and neuronal arbor complexity, respectively. The evaluation of dendritic spines is conducted using an automatic assisted classification algorithm integral to the reconstruction software, which classifies spines into four categories (i.e., thin, mushroom, stubby, filopodia). Furthermore, an additional three parameters (i.e., length, head diameter, and volume) are also chosen to assess alterations in dendritic spine morphology. To validate the potential of wide application of the ballistic labeling technique, pyramidal neurons from in vitro cell culture were successfully labeled. Overall, the ballistic labeling method is unique and useful for visualizing neurons in different brain regions in rats, which in combination with sophisticated reconstruction software, allows researchers to elucidate the possible mechanisms underlying neurocognitive dysfunction.

We present a protocol to label and analyze pyramidal neurons, which is critical for evaluating potential morphological alterations in neurons and dendritic spines that may underlie neurochemical and behavioral abnormalities.

It has been reported that the size and shape of dendritic spines is related to their structural plasticity. To identify the morphological structure of ...

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 ...