We are grateful for the support from the NIH SBIR Phase II #R44OD024874 to H.C. and M.P.H.L. The authors would also like to acknowledge the financial support provided by the Consejo Nacional de Humanidades, Ciencias y Tecnologias (CONAHCYT) in Mexico through the fellowship titled Beca CONAHCYT Para Estudios de Doctorado en el Extranjero.

Adhering strictly to ethical guidelines ensures the well-being and appropriate handling of the zebrafish involved in our experiments. All experimental activities were conducted in accordance with the standards set forth by the Institutional Animal Care and Use Committees (IACUC). The experiments and findings detailed here were executed with the approval of IACUC under the authorized protocol AUP-21-066.
1. Preparation and storage of stock solutions
CAUTION: Prepare the tricaine stock solution in a fume hood, wearing appropriate personal protective equipment (PPE).
<ol>
	<li>Measure 400 mg of tricaine powder and dissolve it in 97.9 mL of deionized (DI) water. Place a magnetic stirrer into the container and gradually mix the solution.</li>
	<li>Adjust the pH by incrementally adding less than 17 mL of NaOH solution. Continue adding NaOH drops until the pH reaches a range of 6.8.-7.2.</li>
	<li>Determine the final concentration of tricaine solution, accounting for the added NaOH necessary to adjust the pH and store. 
	NOTE: Tricaine solution is light sensitive and should be stored in an amber-colored container at 4 &#176;C when not in use.</li>
	<li>Measure 258.13 mg of Amiodarone and dissolve it in 1 L of DI water. Insert a magnetic stirrer into the container and stir for a few minutes until the solution is completely dissolved and clear of any powder. Store the solution inside a refrigerator at 4&#176;C when not in use. 
	NOTE: Amiodarone must be prepared inside a fume hood and stored inside a refrigerator at 4 &#176;C. Always wear the appropriate PPE.</li>
</ol>
2. Fish husbandry
<ol>
	<li>Maintain the fish in individual tanks linked to a circulating water system, keeping the temperature between 28-29&#176;C.</li>
	<li>Ensure continuous monitoring of water quality and adhere to a 14 h/10 h day-night cycle.</li>
	<li>Divide the 40 fish into four groups, each consisting of 10 fish, and house them in separate tanks.</li>
</ol>
3. Setting up the device
NOTE: The procedure outlined in step 3.2 must be performed for each chamber of the device (Figure 1). While each chamber can be set up differently, for consistency in the experiment, it is advisable to maintain uniform settings across all chambers.
<ol>
	<li>Connect the device to a power source and initiate operation by pressing the Red Button located on the front panel.</li>
	<li>Navigate through the device&#39;s touchscreen to choose the chamber to be configured.
	<ol>
		<li>Adjust the low-pass filter setting to 0.1 Hz and the high-pass filter setting to 500 Hz. Adjust the gain setting to 10,000.</li>
	</ol>
	</li>
	<li>Lift the front lid to access the four chambers. Prior to starting the experiment, immerse each chamber&#39;s sponge (standard cellulose sponge) in Tricaine solution, then squeeze out any excess liquid.</li>
	<li>Rotate the four-electrode array to ensure the sponge is accessible before introducing the fish.</li>
</ol>
4. Amiodarone treatment
NOTE: Utilize the water from the fish tanks (fish water) to prepare diluted solutions of Amiodarone and Tricaine. Steps 4.3 and 4.4 must be repeated for each group separated in step 2.3 for each solution prepared in step 4.1.
<ol>
	<li>Follow the steps described below to dilute the Amiodarone stock solution.
	<ol>
		<li>Dilute 8.75 mL of stock solution into 41.25 mL of fish water to achieve a 70 &#181;M concentration.</li>
		<li>Dilute 12.5 mL of stock solution into 37.5 mL of fish water to achieve a 100 &#181;M concentration.</li>
		<li>Dilute 25 mL of stock solution into 25 mL of fish water to achieve a 200 &#181;M concentration.</li>
	</ol>
	</li>
	<li>Prepare the anesthesia solution by diluting 4 mL of Tricaine stock solution into 100 mL of fish water, resulting in a Tricaine concentration of 0.16 mg/mL</li>
	<li>Submerge each fish individually in the prepared Amiodarone solution for 5 min. Use timers to accurately monitor the immersion duration.</li>
	<li>Transfer the fish into Tricaine solution for 2-3 min for anesthesia. Ensure the treatment does not cause the death of the fish. If a fish exhibits abnormal behavior, such as losing color or a significant reduction in gill movement, remove it from the solution immediately and place it back into clean fish water.</li>
</ol>
5. ECG data collection
NOTE: Immediately follow these instructions after completing step 4.4. Repeat the process for each chamber before sealing the device. The chambers are designed to accommodate fish ranging from 4-10 mm in width and 25-50 mm in length.
<ol>
	<li>Lift the fish out of the Tricaine solution and place it onto the sponge, ensuring its ventral side faces up.</li>
	<li>Position the electrode array in a rhombus shape over the fish&#39;s chest area to capture ECG signals from four different points simultaneously (Figure 2A,B). Make sure to adjust each electrode carefully so every pin makes contact with the fish&#39;s skin.
	<ol>
		<li>Rotate each electrode, lowering the pin until it makes contact with the fish&#39;s underside. If the pin is too high, turn the corresponding thumbscrew clockwise to lower it until it touches the fish.</li>
		<li>Perform the adjustment described above for the electrodes in all chambers.</li>
	</ol>
	</li>
	<li>Close the chambers&#39; lid once all electrode arrays are correctly positioned over the fish in the operational chambers.</li>
	<li>Mark each chamber for recording by selecting it on the touchscreen, then initiate the recording by clicking the Record button on-screen.</li>
	<li>Click the Stop button after 2 min and then click on Save button from the prompt windows that pop ups on-screen after stopping. Save the information either on a disk or to the cloud, depending on specific research needs.</li>
</ol>
6. Troubleshooting the four-electrode array positioning system
NOTE: Occasionally, one or several electrodes might fail to capture a clear ECG signal, necessitating repositioning. Use the following steps to troubleshoot the electrode array when needed.
<ol>
	<li>Press the Stop button on the device interface to halt the recording process and identify the channel or channels with inconsistent signals.</li>
	<li>Lift the front lid for access to the electrode chambers. Identify which chamber(s) and specific electrodes are causing issues.</li>
	<li>Turn the top thumbscrew clockwise to adjust the electrode&#39;s proximity to the fish. This action will lower the electrode closer to the fish.</li>
	<li>Repeat step 6.3 as needed. If merely moving closer does not solve the issue, rotate the electrode to a different spot on the fish until a satisfactory signal is achieved.</li>
</ol>
7. Extracting and analyzing data
NOTE: All the basic data analysis presented in this paper uses the Wavelet technique21 to reduce various types of noise and standard deviation of normal sinus beats (SDNN) to calculate the beat-to-beat variance of HR in one-minute segments.
<ol>
	<li>Once data has been saved to either the disk or the cloud, retrieve it for analysis or send it directly to a cloud server for further processing.</li>
	<li>Navigate to the saved data by clicking the Browse button on the screen and select the data sets to use.</li>
	<li>Select the Extract button on the options on-screen and attach a USB drive to the back part of the device to extract the data.</li>
	<li>Follow the device prompts to save the selected data sets onto the external drive for subsequent retrieval and analysis.</li>
</ol>

Zebra II: Advanced ECG System for Multichannel Zebrafish Cardiovascular Studies

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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Atrial+fibrillation+and+non-cardiovascular+diseases:+A+systematic+review.">Atrial fibrillation and non-cardiovascular diseases: A systematic review.</a> Arq Bras Cardiol. 105 (5), 519-526 (2015).</li><li>Hu, Z., Ding, L., Yao, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Atrial+fibrillation:+mechanism+and+clinical+management.">Atrial fibrillation: mechanism and clinical management.</a> Chin Med J. 136 (22), 2668-2676 (2023).</li><li>Babapoor-Farrokhran, S., Rasekhi, R. T., Gill, D., Babapoor, S., Amanullah, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Arrhythmia+in+COVID-19.">Arrhythmia in COVID-19.</a> SN Comp Clin Med. 2 (9), 1430-1435 (2020).</li><li>Arnaout, R., 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=Zebrafish+model+for+human+long+QT+syndrome.">Zebrafish model for human long QT syndrome.</a> Proc Natl Acad Sci. 104 (27), 11316-11321 (2007).</li><li>Koopman, C. D., 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=The+zebrafish+grime+mutant+uncovers+an+evolutionarily+conserved+role+for+Tmem161b+in+the+control+of+cardiac+rhythm.">The zebrafish grime mutant uncovers an evolutionarily conserved role for Tmem161b in the control of cardiac rhythm.</a> Proc Natl Acad Sci. 118 (9), e2018220118 (2021).</li><li>Genge, C. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Reviews of Physiology, Biochemistry and Pharmacology. , (2016).</li><li>Nemtsas, P., Wettwer, E., Christ, T., Weidinger, G., Ravens, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adult+zebrafish+heart+as+a+model+for+human+heart?+An+electrophysiological+study.">Adult zebrafish heart as a model for human heart? An electrophysiological study.</a> J Mol Cell Cardiol. 48 (1), 161-171 (2010).</li><li>Sun, P., 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=Micro-electrocardiograms+to+study+post-ventricular+amputation+of+Zebrafish+heart.">Micro-electrocardiograms to study post-ventricular amputation of Zebrafish heart.</a> Ann Biomed Engg. 37 (5), 890-901 (2009).</li><li>Lin, M. H., 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=Development+of+a+rapid+and+economic+in+vivo+electrocardiogram+platform+for+cardiovascular+drug+assay+and+electrophysiology+research+in+adult+zebrafish.">Development of a rapid and economic in vivo electrocardiogram platform for cardiovascular drug assay and electrophysiology research in adult zebrafish.</a> Sci Rep. 8 (1), 15986 (2018).</li><li>Chaudhari, G. H., Chennubhotla, K. S., Chatti, K., Kulkarni, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optimization+of+the+adult+zebrafish+ECG+method+for+assessment+of+drug-induced+QTc+prolongation.">Optimization of the adult zebrafish ECG method for assessment of drug-induced QTc prolongation.</a> J Pharmacol Toxicol Methods. 67 (2), 115-120 (2013).</li><li>Liu, C. C., Li, L., Lam, Y. W., Siu, C. W., Cheng, S. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improvement+of+surface+ECG+recording+in+adult+zebrafish+reveals+that+the+value+of+this+model+exceeds+our+expectation.">Improvement of surface ECG recording in adult zebrafish reveals that the value of this model exceeds our expectation.</a> Sci Rep. 6, 25073 (2016).</li><li>Stoyek, M. R., Rog-Zielinska, E. A., Quinn, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Age-associated+changes+in+electrical+function+of+the+zebrafish+heart.">Age-associated changes in electrical function of the zebrafish heart.</a> Prog Biophys Mol Biol. 138, 91-104 (2018).</li><li>Cao, H., 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=Wearable+multi-channel+microelectrode+membranes+for+elucidating+electrophysiological+phenotypes+of+injured+myocardium.">Wearable multi-channel microelectrode membranes for elucidating electrophysiological phenotypes of injured myocardium.</a> Integ Biol. 6 (8), 789-795 (2014).</li><li>Lenning, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Real-time+monitoring+and+analysis+of+zebrafish+electrocardiogram+with+anomaly+detection.">Real-time monitoring and analysis of zebrafish electrocardiogram with anomaly detection.</a> Sensors. 18 (1), 61 (2018).</li><li>Yu, F., 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=Flexible+microelectrode+arrays+to+interface+epicardial+electrical+signals+with+intracardial+calcium+transients+in+zebrafish+hearts.">Flexible microelectrode arrays to interface epicardial electrical signals with intracardial calcium transients in zebrafish hearts.</a> Biomed Microdev. 14 (2), 357-366 (2012).</li><li>Cho, S. J., 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=Zebrafish+as+an+animal+model+in+epilepsy+studies+with+multichannel+EEG+recordings.">Zebrafish as an animal model in epilepsy studies with multichannel EEG recordings.</a> Sci Rep. 7 (1), 3099 (2017).</li><li>Le, 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=Continuous+electrocardiogram+monitoring+in+Zebrafish+with+prolonged+mild+anesthesia.">Continuous electrocardiogram monitoring in Zebrafish with prolonged mild anesthesia.</a> , 2610-2613 (2020).</li><li>Le, 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=A+novel+wireless+ECG+system+for+prolonged+monitoring+of+multiple+zebrafish+for+heart+disease+and+drug+screening+studies.">A novel wireless ECG system for prolonged monitoring of multiple zebrafish for heart disease and drug screening studies.</a> Biosensors Bioelectron. 197, 113808 (2022).</li><li>Kouzehgar, K., Eslamian, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+2+-+Application+of+experimental+data+and+soft+computing+techniques+in+determining+the+outflow+and+breach+characteristics+in+embankments+and+landslide+dams.">Chapter 2 - Application of experimental data and soft computing techniques in determining the outflow and breach characteristics in embankments and landslide dams.</a> Handbook Hydroinfo. 3, 11-31 (2023).</li><li>Milan, D. 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Hung Cao and Michael Lau are the founders of Sensoriis, Inc. Sensoriis's system Zebra II was developed under NIH support, SBIR Phase II grant #R44OD024874. The University of California, Irvine, has reviewed and approved Hung Cao's relationship with Sensoriis, Inc. in accordance with its conflict-of-interest policies. The other authors declare no conflict of interest.

The current steps were modified and improved according to this new device version from a previous study made by our team20. Here, a labeled computer-aided design (CAD) of the device is also included.
As highlighted in the protocol's troubleshooting guide, the device necessitates the user to execute certain critical steps with diligence and precision, as these steps crucially affect the ECG signal's quality. Consistent with findings from our prior study...

Cardiovascular diseases (CVDs) are the leading cause of death worldwide. According to the 2024 AHA Update Report, around 2552 people die every day of CVDs in the U.S., and the average direct and indirect cost of CVD was estimated at $378.0 billion in 2017-20181. Atrial fibrillation (AF) is one of the most common cardiac arrhythmias in the U.S.2 and is associated with an unfavorable prognosis, increasing the risk of stroke and death in both CVDs and non-cardiovascular disease scenarios, leading to an increased risk of adverse outcomes3. This type of arrhythmia is distinguished by its rapid and irregular electrical activity in the atria, which usually results in a fast and irregular ventricular rhythm4. Further, given the recent events of COVID-19, there has been an increased number of studies and evidence that show an increased appearance of arrhythmia in relation to COVID-19, mostly to sinus bradycardia5. On the opposite side of AF, sinus bradycardia presents a decreased and slower heart rhythm, with a heart rate lower than 60 beats per minute (bpm). This diagnosis requires an electrocardiogram (ECG) showing a normal sinus rhythm at a rate lower than 60 bpm.
The zebrafish, Danio rerio, has proven to be an ideal model for cardiovascular and drug development studies because of its homology to humans in both morphology, physiology, and genetics6,7. Despite only having two discernible chambers compared to the four-chambered heart in humans, the zebrafish possesses a similar contractile structure with an analogous conduction system8,9. Hence, the zebrafish model is suitable for researching cardiac arrhythmias and exploring the association between related genetic pathways and electrophysiological characteristics through ECG analysis. Regarding the sensor design for ECG recording in zebrafish, conventional needle electrodes are commonly used10. Other studies presented alternatives for conventional needles by designing a different type of needle made of different materials11, these incorporated materials such as tungsten filament, stainless steel, and silver wire to assess the quality of the recorded data. Together with the portable ECG kit, they sought to establish a universal platform for both research and educational labs.
The needle system was also deployed in other studies11,12,13,14 for carrying out research on biological and/or drug-induced effects, in which it showed encouraging outcomes. However, to obtain optimal signals, it was necessary to carefully insert the needles through the zebrafish&#39;s dermis. This invasive procedure can cause injury to the fish&#39;s heart due to the sharpness of the needles, thus possibly changing signal morphology13. Furthermore, accurately placing the electrode on the small heart without causing harm to ensure clear ECG capture presents a significant learning curve. Consequently, various substitute probe systems have been introduced, such as microelectrode arrays (MEA) and 3D-printed sensors. Both our team and other researchers have successfully utilized MEA for data acquisition, achieving data that offer a good signal-to-noise ratio (SNR) along with high spatial and temporal resolution15,16,17. For example, we introduced an MEA that envelops the heart of the fish, allowing the capture of site-specific ECG signals15,16. In contrast, another team developed a different kind of MEA based on a flexible printed circuit board (FPCB) crafted from polyimide film aimed at recording multiple electroencephalograms (EEGs) for epilepsy research18. While MEAs enable the recording of numerous signals, they are limited by the capacity to assess only one fish at a time due to the restricted number of electrode channels. In response to this limitation, we have recently showcased an advanced system capable of simultaneous ECG recordings from multiple fish19. This system employs two electrodes, each 125 &#181;m in thickness and made of polyimide, with gold-sputtered electrodes positioned at the bottom of their housing to facilitate data acquisition. However, the inclusion of a water circulation pump introduced noise, affecting the signal-to-noise ratio (SNR) quality of the ECG signals. Additionally, the use of bulky and costly data collection tools, which required a wired connection to a computer, was necessitated. On the market, systems like iWORX offer improved portability through a compact amplifier, yet they face unresolved issues: (i) their recording time is too short (3-5 min) for experiments needing longer durations, such as those involving acute drug effects; (ii) the requirement for anesthetizing the animals, which can stress them and skew innate cardiac electrophysiological data; (iii) manual, sequential measurements hinder research with large numbers of fish; and (iv) the need for offline, labor-intensive ECG data processing. Importantly, no high-throughput systems incorporating microelectronics for analyzing mutant phenotypes have yet emerged. Therefore, the creation of high-throughput systems that enable extended ECG recording is crucial for uncovering links between arrhythmic phenotypes and mutant genotypes, identifying multiple arrhythmic phenotypes associated with single mutant genotypes, and testing the effectiveness of cardiac drugs using the zebrafish model. Consequently, we introduced, in 2022, the groundbreaking prototype of the Zebra II system. This novel system is equipped to perform extended ECG recordings from several fish at once. To achieve this, an in-house electronic system was crafted, utilizing the Internet of Things (IoT) technology for wireless data transmission and enabling data analysis through a mobile application. The inclusion of IoT technology not only bolstered the system&#39;s flexibility and portability but also facilitated remote collaborations for research on zebrafish models. In a prior study, we validated the system&#39;s efficacy through extensive testing, showcasing its ability to 1) acquire ECG data simultaneously from four fish; 2) perform continuous ECG recordings for up to 1 h, which significantly exceeds the capacity of existing systems that only capture a few minutes; 3) minimize the confounding effects of anesthesia by employing a 50% reduced concentration of Tricaine. Furthermore, the system demonstrated a strong capability for lengthy ECG recordings across various experiments, such as those involving sodium-induced sinoatrial (SA) block, temperature-induced heart rate changes, and drug-induced arrhythmias in both Tg(SCN5A-D1275N) mutant and wildtype fish under controlled experimental conditions. The system&#39;s adoption of multiple electrode channels for extended ECG recording further enables the execution of randomized controlled trials, allowing for two fish per experimental group to be studied under identical conditions20. In this paper, we introduce the upgraded version of the system, featuring a 4-point electrode. This enhancement enables users to capture ECG readings from various locations on the fish, significantly reducing the chance of obtaining low-quality ECG signals. Moreover, this iteration builds upon and refines the majority of the functionalities outlined in previous research19. This evolved ECG system shows great promise in addressing existing limitations, thereby substantially facilitating the analysis of arrhythmic phenotypes and the screening of drugs in zebrafish. The innovative addition of the 4-point electrode was rigorously tested through a brief study on drug-induced arrhythmia in wild-type zebrafish, further validating its effectiveness.

<table><tbody><tr><td>Amiodarone Hydrochloride</td><td>Sigma-Aldrich</td><td>PHR1164-1G</td><td>Amiodarone powder</td></tr><tr><td>Benchtop pH/mV Meter</td><td>Sper Scientific</td><td>10500308</td><td></td></tr><tr><td>Ethyle3-Aminobenzoate, Methanesulfonic Acid Salt, 98%, ACROS Organics</td><td>Fisher Scientific</td><td>AC118002500</td><td>Tricaine powder</td></tr><tr><td>Isotemp Hot Plate Stirrer</td><td>Fisher Scientific</td><td>SP88854200</td><td></td></tr><tr><td>Zebra II System</td><td>Sensoriis</td><td>N/A</td><td>Commercial Zebrafish ECG Recording System</td></tr><tr><td>Zebrafish ECG System</td><td>iWorx</td><td>ZS-200</td><td>Commercial Zebrafish ECG Recording System</td></tr></tbody></table>

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 ability to simultaneously record ECG signals from multiple fish significantly distinguishes this device, offering a considerable advantage in reducing the time required for drug or cardiac research on zebrafish or other similar species (Figure 1). Recording accurate ECG signals can be challenging due to the anatomical variations among fish, especially when it comes to electrode placement. Traditionally, obtaining a clear ECG signal involves positioning the active electrode over the heart...

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

zebra-ii-as-novel-system-to-record-electrophysiological-signals

Research

JoVE Journal

Bioengineering

335 Views.  UC Irvine. 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. 

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

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 ...

Neuroscience

Electroretinogram Recording in Larval Zebrafish using A Novel Cone-Shaped Sponge-tip Electrode

The zebrafish (Danio rerio) is commonly used as a vertebrate model in developmental studies and is particularly suitable for visual neuroscience. For functional measurements of visual performance, electroretinography (ERG) is an ideal non-invasive method, which has been well established in higher vertebrate species. This approach is increasingly being used for examining the visual function in zebrafish, including during the early developmental larval stages. However, the most commonly used recording electrode for larval zebrafish ERG to date is the glass micropipette electrode, which requires specialized equipment for its manufacture, presenting a challenge for laboratories with limited resources. Here, we present a larval zebrafish ERG protocol using a cone-shaped sponge-tip electrode. The novel electrode is easier to manufacture and handle, more economical, and less likely to damage the larval eye than the glass micropipette. Like previously published ERG methods, the current protocol can assess outer retinal function through photoreceptor and bipolar cell responses, the a- and b-wave, respectively. The protocol can clearly illustrate the refinement of visual function throughout the early development of zebrafish larvae, supporting the utility, sensitivity, and reliability of the novel electrode. The simplified electrode is particularly useful when establishing a new ERG system or modifying existing small-animal ERG apparatus for zebrafish measurement, aiding researchers in the visual neurosciences to use the zebrafish model organism.

Here, we present a protocol that simplifies the measurement of light evoked electroretinogram responses from larval zebrafish. A novel cone-shaped sponge-tip electrode can help to make the study of visual development in larval zebrafish using the electroretinogram ERG easier to achieve with reliable outcomes and lower cost.

The zebrafish (Danio rerio) is commonly used as a vertebrate model in developmental studies and is particularly suitable for visual neuroscience. For ...

Developmental Biology

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 ...

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing organisms, combined with the embryonic optical clarity, served as initial compelling attributes of this model. Over the past two decades, the success of this model has been further propelled by its amenability to large-scale mutagenesis screens and by the ease of transgenesis. More recently, gene-editing approaches have extended the power of the model.
For neurodevelopmental studies, the zebrafish embryo and larva provide a model to which multiple methods can be applied. Here, we focus on methods that allow the study of an essential property of neurons, electrical excitability. Our preparation for the electrophysiological study of zebrafish spinal neurons involves the use of veterinarian suture glue to secure the preparation to a recording chamber. Alternative methods for recording from zebrafish embryos and larvae involve the attachment of the preparation to the chamber using a fine tungsten pin1,2,3,4,5. A tungsten pin is most often used to mount the preparation in a lateral orientation, although it has been used to mount larvae dorsal-side up4. The suture glue has been used to mount embryos and larvae in both orientations. Using the glue, a minimal dissection can be performed, allowing access to spinal neurons without the use of an enzymatic treatment, thereby avoiding any resultant damage. However, for larvae, it is necessary to apply a brief enzyme treatment to remove the muscle tissue surrounding the spinal cord. The methods described here have been used to study the intrinsic electrical properties of motor neurons, interneurons, and sensory neurons at several developmental stages6,7,8,9.

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

This manuscript describes methods for electrophysiological recordings from spinal neurons of zebrafish embryos and larvae. The preparation maintains neurons in situ and often involves minimum dissection. These methods allow for the electrophysiological study of a variety of spinal neurons, from the initial electrical excitability acquisition through the early larval stages.

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing ...

Behavioral And Physiological Analysis In A Zebrafish Model Of Epilepsy

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

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

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

Using an Automated 3D-tracking System to Record Individual and Shoals of Adult Zebrafish

Like many aquatic animals, zebrafish (Danio rerio) moves in a 3D space. It is thus preferable to use a 3D recording system to study its behavior. The presented automatic video tracking system accomplishes this by using a mirror system and a calibration procedure that corrects for the considerable error introduced by the transition of light from water to air. With this system it is possible to record both single and groups of adult zebrafish. Before use, the system has to be calibrated. The system consists of three modules: Recording, Path Reconstruction, and Data Processing. The step-by-step protocols for calibration and using the three modules are presented. Depending on the experimental setup, the system can be used for testing neophobia, white aversion, social cohesion, motor impairments, novel object exploration etc. It is especially promising as a first-step tool to study the effects of drugs or mutations on basic behavioral patterns. The system provides information about vertical and horizontal distribution of the zebrafish, about the xyz-components of kinematic parameters (such as locomotion, velocity, acceleration, and turning angle) and it provides the data necessary to calculate parameters for social cohesions when testing shoals.

The use of a 3D automatic video system that can track individual and groups of zebrafish is described. As application example we explore the effects of the NMDA-receptor antagonist MK-801 on shoals of zebrafish.

Like many aquatic animals, zebrafish (Danio rerio) moves in a 3D space. It is thus preferable to use a 3D recording system to study its behavior. The ...

Biology

In Vivo Surface Electrocardiography for Adult Zebrafish

The electrocardiogram waveforms of adult zebrafish and those of humans are remarkably similar. These electrocardiogram similarities enhance the value of zebrafish not only as a research model for human cardiac electrophysiology and myopathies but also as a surrogate model in high throughput pharmaceutical screening for potential cardiotoxicities to humans, such as QT prolongation. As such, in vivo electrocardiography for adult zebrafish is an electrical phenotyping tool that is necessary, if not indispensable, for cross-sectional or longitudinal in vivo electrophysiological characterizations. However, too often, the lack of a reliable, practical, and cost-effective recording method remains a major challenge preventing this in vivo diagnostic tool from becoming more readily accessible. Here, we describe a practical, straightforward approach to in vivo electrocardiography for adult zebrafish using a low-maintenance, cost-effective, and comprehensive system that yields consistent, reliable recordings. We illustrate our protocol using healthy adult male zebrafish of 12-18 months of age. We also introduce a rapid real-time interpretation strategy for quality validation to ensure data accuracy and robustness early in the electrocardiogram recording process.

Here, we present a reliable, minimally invasive, and cost-effective method to record and interpret electrocardiograms in live anesthetized adult zebrafish.

The electrocardiogram waveforms of adult zebrafish and those of humans are remarkably similar. These electrocardiogram similarities enhance the value of ...

Zebrafish Electrocardiography (ECG): A Minimally Invasive Assay to Evaluate Cardiac Function

Source: Zhao, Y., et al. <a href="https://www.jove.com/video/60011/" target="_blank">In Vivo Surface Electrocardiography for Adult Zebrafish</a>. J. Vis. Exp. (2019).
This video shows how&#160;to perform electrocardiography (ECG or EKG) measurements, a test which measures the electrical activity of the heart, on a zebrafish.

Source: Zhao, Y., et al. In Vivo Surface Electrocardiography for Adult Zebrafish. J. Vis. Exp. (2019).
This video shows how&#160;to perform ...

JoVE Encyclopedia of Experiments

Danio rerio (zebrafish)

Adult

Adult Zebrafish Electrophysiology: A Method to Measure the Extracellular Electrical Activity of Neurons In Vivo

Source: Johnston, L., et al. <a href="https://www.jove.com/v/51065/" target="_blank">Electrophysiological Recording in the Brain of Intact Adult Zebrafish</a>. &#160;J. Vis. Exp. (2013)
The protocol describes the procedure to perform electrophysiology in adult zebrafish. This method can be used for measuring neural activity in vivo.

Source: Johnston, L., et al. Electrophysiological Recording in the Brain of Intact Adult Zebrafish. &#160;J. Vis. Exp. (2013)
The protocol describes the ...

Monitoring Electrical Signals from the Afferent Neurons in an Immobilized Zebrafish Larva

Source: Lunsford, E. T., et al. <a href="https://app.jove.com/v/62233">Activity of Posterior Lateral Line Afferent Neurons during Swimming in Zebrafish.</a> J. Vis. Exp. (2021).
This video demonstrates the recording of electrical signals from the afferent neurons in a paralyzed zebrafish larva by positioning a micropipette near the lateral line ganglion and forming a loose seal for signal capture. The extracellular solution maintains neuronal viability during this process.

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
1. ...

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

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