High-throughput zebrafish intramuscular recording assay

Journal Pre-proof High-throughput Zebrafish Intramuscular Recording Assay Sung-Joon Cho, Yang Jun Kang, Sohee Kim

PII:

S0925-4005(19)31531-X

DOI:

https://doi.org/10.1016/j.snb.2019.127332

Reference:

SNB 127332

To appear in:

Sensors and Actuators: B. Chemical

Received Date:

29 May 2019

Revised Date:

21 October 2019

Accepted Date:

23 October 2019

Please cite this article as: Cho S-Joon, Kang YJ, Kim S, High-throughput Zebrafish Intramuscular Recording Assay, Sensors and Actuators: B. Chemical (2019), doi: https://doi.org/10.1016/j.snb.2019.127332

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High-throughput Zebrafish Intramuscular Recording Assay

Sung-Joon Cho†1,2, Yang Jun Kang3*, and Sohee Kim4*

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Division of Fundamental Neurobiology, Krembil Research Institute, University Health Network, Toronto,

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ON, Canada M5T 2S8. Collaborative Program in Neuroscience, University of Toronto, Toronto, Ontario, Canada, M5S 1A1

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Department of Mechanical Engineering, Chosun University, Gwangju 61452, Republic of Korea.

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Department of Robotics Engineering, Daegu Gyeongbuk Institute of Science and Technology (DGIST),

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Daegu 42988, Republic of Korea. †

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S.-J. Cho was with the School of Electrical Engineering and Computer Science at Gwangju Institute of

Corresponding authors. These authors contributed equally to this work.

Highlights

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*

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Science and Technology (GIST) in Gwangju, Republic of Korea.

Multiple zebrafish intramuscular recording was possible using a microfluidic chip

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Intramuscular activities of zebrafish larvae were obtained while they stayed in water Feasibility of the developed system was demonstrated using three different chemicals

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Abstract Despite tremendous efforts in utilizing zebrafish in neurological disease studies, owing to their easy handling, low cost, fast growth cycle, fecundity, high genetic similarity to humans, and transparency, the high-throughput electrophysiology methods for zebrafish are still absent. Although methods to detect intramuscular activities of adult and larval zebrafish have been previously introduced, the methods are

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complicated and time-consuming. Therefore, they have not been widely used in the zebrafish research

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community. We propose a high-throughput muscular activity measurement method using a simple and clever way of trapping zebrafish with microfluidic chip technology. Zebrafish larvae at 5 days post-fertilization

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were successfully retained in the designed microfluidic chip and were able to maintain their respiration for

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more than 24 h as long as the water was supplied through the inlet of the chip. The intramuscular activities of the larvae were obtained while they remained in the water. Significantly, this is the first reported method that

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can be used for measuring intramuscular activities while the larvae are inside the water. As a demonstration, we successfully modulated the locomotor activities of zebrafish using three different chemicals, proving that

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the developed method can be useful in monitoring intramuscular activities from multiple larvae and assessing the efficacy of pharmaceuticals.

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Introduction

The development of new electrophysiological tools provides brand new pathways for understanding neuromuscular diseases. Such new tools may allow us to improve the quality of lives of people who suffer from neuromuscular diseases that do not have effective treatments so far, such as myotonia and lumbar radiculopathy(Benditz et al., 2016; Matthews et al., 2010). Although the creation and development of new electrophysiological tools should be one of the most important ventures in neuroscience, there is widespread 2

apathy. Over the past decades, the use of zebrafish (Danio rerio) in human disease studies has received a great deal of attention. Zebrafish has become a rising champion in animal models despite having disadvantages such as being a poikilothermic non-mammalian animal. As a non-mammal, zebrafish are evolutionarily less similar to humans than rodents; they lack some mammalian organs such as lungs, mammary glands, prostate and skin(Thisse, 2002). Even with these limitations, zebrafish have gained substantial popularity in disease

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studies owing to their easy handling, low cost, fast growth cycle, fecundity, high genetic similarity to

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humans, and transparent bodies(Cho et al., 2017a; Santoriello and Zon, 2012). The complete genome sequence of zebrafish was determined, revealing that 70% of their genes are associated with human

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diseases(Santoriello and Zon, 2012). They are also known to be a perfect model for high-throughput genotyping and congenital diseases as they do not require cardiovascular functions and they absorb an

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adequate amount of oxygen through their skin by diffusion in their first ten days of life(Schwerte, 2006).

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Owing to these advantages, zebrafish are being utilized in a wide range of medicine research including muscle studies(Chen et al., 2016; Gilbert et al., 2017; Savchenko et al., 2018). With regards to muscles,

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zebrafish larvae fully develop their muscles by 2 days post-fertilization (dpf)(Sztal et al., 2016). Zebrafish mutational screens on muscle structure have been studied since 1990(Felsenfeld et al., 1990), and many muscular mutations have been introduced since then. For example, Lin et al. introduced a severe muscular dystrophy model that was impaired in 48 h post-fertilization (hpf), and Granato et al. introduced the sapje

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mutations, which exhibit muscle degenerations in 72 hpf(Granato et al., 1996; Lin et al., 2011). Electromyography (EMG) examines the functional and electrical activity of muscles and is an effective tool in diagnosing and assessing the efficacy of therapeutics on neuromuscular disorders. However, EMG signals recorded from the skin (surface EMG) not only measure superficial muscles, but they are also easily influenced by motion artifacts, myoelectric crosstalk, and other environmental conditions. Therefore, 3

measuring intramuscular activity is preferred for more accurate diagnosis(Cho et al., 2015). Intramuscular measurements on freely swimming adult zebrafish were first reported in 1988(Liu and Westerfield, 1988). A set of hooked needle electrodes were inserted into adult zebrafish muscles, and the intramuscular signals were measured while the fish was freely swimming in a tank. In 2015, Cho et al. demonstrated microfabricated silicon needle electrodes that were fine and durable enough to take measurements from zebrafish larvae at 5 dpf. During the recording, the larvae were immobilized inside the

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coil-shaped electrodes, which also served as stable reference and ground electrodes(Cho et al., 2015).

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However, this method requires sophisticated engineering techniques to fabricate microneedle electrodes and manual handling to place the larva into the coil-shaped electrode. Most importantly, measurements are

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performed on fish in contact with a minimal amount of water, which is a stressful environment for zebrafish. Although these previous methods are significant in enabling intramuscular measurements of single zebrafish,

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because of the lower efficiency in zebrafish electrophysiology, behavioral testing is still preferred to

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measuring intramuscular activities from zebrafish. Although exploring an electrophysiological biomarker is essential, tools for high-throughput muscle activity detection in zebrafish are desired to overcome this

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limitation of single fish electrophysiology.

In the last decade, microfluidics technology has become an important tool in biomedical applications from drug delivery to organ-on-a-chip. Well-fabricated microfluidic devices can accelerate and facilitate a wide range of medical applications. Following such a trend, microfluidics technology has also been applied in

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zebrafish research and demonstrated distinct advantages. Microfluidics technology can be used to perform all the processes in parallel, which expedites the efficiency of using zebrafish in biomedical studies(Yang et al., 2016). The biggest limitation of using zebrafish is that it is an aquatic organism. Because zebrafish need to be in the water all the time during a research procedure, handling, surgery, and electrophysiological experiments with zebrafish are more difficult than with rodents(Cho et al., 2017b). By adapting microfluidic technology, 4

the manipulation and immobilization of zebrafish becomes painless; handling of zebrafish such as using pipettes or embedding them in agarose gel for immobilization are no longer necessary. Zebrafish can be controlled by water flowing through the microfluidic chip(Bischel et al., 2013; Lin et al., 2015). Owning to these advantages, microfluidics has been widely used in zebrafish studies such as perfusion, culturing, and imaging(Swain et al., 2013; Yang et al., 2016). Previous approaches have focused on trapping and observing behavioral aspects of zebrafish using microfluidic chips. Microfluidic technology allowed high-throughput

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neuronal activity recording in zebrafish(Hong et al., 2016; Turrini et al., 2017), but other types of in vivo

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electrophysiological monitoring using microfluidic chips have not been adopted.

Thus, an efficient method, ideally without using anesthetic agents and without taking the fish out of water, is

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still necessary. The need to record signals from multiple zebrafish at once arises to maximize the efficiency of using zebrafish. Furthermore, anesthetic agents and placing the fish out of water may affect the activity of

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zebrafish. In this paper, we implemented microfluidic devices to enable long-term high-throughput muscle

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activity monitoring. Using the developed monitoring system, swimming zebrafish larvae were easily transported and trapped in each channel from an inlet pool, which allowed them to be restrained for more

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than 24 hours as long as the water was provided to the microfluidic channels to maintain a healthy environment. Then, needle electrodes that locked up with a customized micromanipulator were delicately inserted into the skeletal muscles of larvae. Such a setup allowed continuous monitoring of muscle activities from multiple zebrafish with high sensitivity and low electrical noise. The feasibility of the monitoring

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system was demonstrated by comparing four different groups—Control, Tricaine (MS-222), N-methyl-Daspartate (NMDA), and dizocilpine (MK-801)—using three different drugs that manipulate muscle activities. We believe that the developed system can facilitate a rigid and reliable analysis of neuromuscular zebrafish studies, and provide a more economical and time-efficient method for assessing the efficacy of therapeutics.

Materials and Methods 5

Fabrication of a microfluidic chip for trapping zebrafish As shown in Fig. 1A, a microfluidic chip for trapping zebrafish was composed of a single inlet, eight outlets, and eight trapping channels. The detailed dimensions are shown in Fig. S1 in the Supplementary Information. We conducted several preliminary experiments to find out the optimal design of the microfluidic device. First, we measured the head size of 5 dpf zebrafish, which resulted in an average of 685 μm with a standard

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deviation of 28 μm (n=10, Fig. S2). To effectively trap individual zebrafish, each trapping channel was designed to have a wavy-shaped channel section (with a maximum width of 1000 µm and a minimum width

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of 600 µm) and a narrowed channel section (with a width of 200 µm) connected in series, as shown in Fig.

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S1. To secure a channel height of 600 µm, six adhesive sheets, where each sheet had a thickness of 100 µm (Colorad, South Korea), were stacked and bonded vertically. A cutter blade (CE6000-40, Graphtec, USA) for

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xurography technique was used to cut the adhesive sheets(Bartholomeusz et al., 2006; Islam et al., 2015). The cover was peeled off from the liner. The master mold was prepared by attaching the liner on a glass slide.

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Polydimethylsiloxane (PDMS) was mixed with a curing agent at a weight ratio of 10:1, and the mixture was poured onto the master mold positioned in a Petri dish. Air bubbles dissolved in PDMS were removed by a

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vacuum pump. After curing in a convection oven at a temperature of 80 C for 1 h, the PDMS block was peeled off from the master mold. Using two biopsy punches, the inlet port and outlet ports were created with diameters of 1.5 mm and 0.5 mm, respectively. In addition, two more 0.5 mm holes were punctured for insertion of the ground and needle electrodes per channel. The PDMS block and a glass substrate were

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treated with oxygen plasma (CUTE-MPR, Femto Science Co., South Korea), and the microfluidic chip was prepared by bonding the PDMS block to the glass substrate. A pipette tip (~1 mL) was tightly fitted into the hole of the inlet port. In addition, the end of eight polyethylene tubes (inner diameter=250 µm and length=300 mm) were connected to the individual outlet ports. The other ends of the corresponding tubes were connected with disposable syringes (~1 mL). By manipulating a pipette, 0.5 mL of water containing 6

zebrafish was dropped into the inlet port. The water and zebrafish were then sucked into the microfluidic channel by a syringe one by one. Each zebrafish was completely trapped into the corresponding trapping channel. After all the fish were trapped in the channels, there was no water flow to maintain their positions and to prevent any external noises during electrophysiological recordings.

Electrodes and the needle insertion system

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M1 stainless steel screws with 1 mm diameter and 5 mm height (NasaKorea, South Korea) were prepared as

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ground electrodes. The ground electrodes were put into the punctured holes to be contacted with zebrafish heads (Fig. 1A). The tungsten needle electrodes (AM Systems, USA) were 75 mm long, had a base diameter

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of 127 m, and an impedance of 1 MΩ at 1 kHz. A customized insertion system was developed for micromanipulation of the needle electrodes with respect to the fabricated microfluidic chip containing

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zebrafish, as shown in Fig. 1A. The system consisted of two bodies: a chip manipulator and an electrode

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manipulator. The microfluidic chip containing zebrafish was placed on the chip holder, attached to an X-, Ydirectional stage with a resolution of motion of 0.1 μm. The needle electrodes were fastened by the electrode

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holder, which was attached to a Z-axis stage with a motion resolution of 0.1 μm. The needle electrodes were manually aligned so the needle electrodes can go through the 0.5 mm holes. This alignment process was required for each microfluidic chip change.

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Animal Preparation

Wild-type (AB strain) zebrafish larvae were a kind gift from Dr. Seok-Yong Choi, Chonnam National University Medical School. Zebrafish were maintained under a 14-hr:10-hr light:dark cycle and staged in dpf as per standard criteria in accordance with the standard guidelines at the zebrafish facility of the Chonnam National University Medical School. All animal care and experiments were approved by the Institutional Animal Care and Use Committee of Gwangju Institute of Science and Technology and conducted in 7

accordance with relevant guidelines and regulations in Republic of Korea.

Pharmacology As a proof-of-concept of the proposed method, spontaneous muscle movements were measured from four groups of larvae (n = 12 each) – Control, Tricaine, NMDA, and MK-801. The control group was not exposed to any drugs. Larvae were preincubated in Tricaine for 5 min, and in NMDA and MK-801 for 20 min before

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and throughout the recording sessions. The drugs were absorbed through the skin and via swallowing in the

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stagnant solutions. Note that zebrafish were submerged in Tricaine for only 5 min because prolonged immersion in Tricaine can cause hypoxemia and hypoventilation, which may result in death(Neiffer and

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Stamper, 2009). Tricaine (A-5040, Sigma-Aldrich, USA) was prepared in a final concentration of 0.61 mM while N-methyl-D-aspartate (M3262, Sigma-Aldrich) and MK-801 (M107, Sigma-Aldrich) were prepared in

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a final concentration of 80 μM. The concentrations were determined by referring to other zebrafish

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research(Matthews and Varga, 2011; Mussulini et al., 2013; Savchenko et al., 2018).

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Intramuscular recording and analysis

Under direct visual guidance aided by magnifying lenses, the needle electrodes were inserted in the axial myomeres of trapped larvae in the microfluidic chip while the needle electrodes and the ground electrodes were connected to the signal acquisition system for intramuscular recording (MP36, Biopac Systems, USA).

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The accurate placement of the electrodes was ascertained by observing the changes in signal morphology during the electrode insertion process and the presence of the insertional activity as shown in Fig. S3. The presence of the insertional activity is critical in clinical EMG to confirm that the needle was inserted into the muscle (Maronian et al., 2004). The signals were band-pass filtered with a frequency range from 0.15 to 5 kHz. Then, the signals were displayed and stored on a laptop using Biopac Student Lab 4.1 software. All recordings were performed inside a Faraday cage in a dark room and terminated after 5 min. For quantitative 8

analysis, interburst interval and average number of bursts per group were analyzed using Biopac Student Lab 4.1 and MATLAB (MathWorks, Inc., USA).

Statistical Analysis All statistical analysis were performed using OriginPro 2016 (OriginLab Corp., USA). The relationship between the interburst intervals and the number of bursts were represented as mean ± S.E.M and analyzed by

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one-way ANOVA followed by Bonferroni’s test post-hoc. All tests resulting in p<0.05 were considered

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statistically significant.

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Microfluidic chip for trapping zebrafish

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Results and discussion

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Fig. 1A represents the design of the whole recording system. It consisted of three main components of the microfluidic chip, the needle electrodes and the micromanipulator. The microfluidic chip was optimized to

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trap eight larvae (5 dpf) without any artificial manipulations such as using anesthetic agents or agarose block to minimize stress. Because the minimum channel width and the channel height were smaller than the size of the fish head, as shown in Figs. S1 and S2, the fish were not able to voluntarily move into the trapping channel. For this reason, the suctional force at each outlet by a syringe was applied to forcibly move and trap

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individual fish at a specific position in the trapping channel. After the head of the fish was positioned in a wider region with the width of 1000 μm, it did not move to a narrower region with the width of 600 μm due to the geometrical constriction of the trapping channel. The cross-sectional views of the wide guide channel with a width of 1500 μm and the wavy-shaped trapping channel are shown in Fig. S4. Although the vertical wall did not show a fine straight surface when compared with conventional microfabrication results, the bottom showed a straight and fine surface. As all the fish were unexceptionally trapped in the trapping 9

channels, we were able to confirm that the wavy-shaped trapping channels demonstrated the ability to trap the fish effectively. Zebrafish larvae at 5 dpf in water were first collected using a pipette. Then, a droplet of water containing larvae was added into the conical pipette tip. Larvae were successfully transported and trapped in each channel (Fig. 1B and Fig. S2) by applying suction from a syringe that was connected to the outlet port while the fish were freely swimming in the inlet. Note that each outlet was connected with an individual syringe, so

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the larvae were delivered to each channel one by one. This allowed us to prevent unwanted tail-forwarded

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position. We confirmed that the trapped zebrafish maintained their respiration for 24 h. The heads of the trapped larvae were contacted to the pre-installed ground electrodes (Fig. 1C). After that, the needle

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electrodes were successfully inserted in the axial myomeres of each larva. The four tungsten needle

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electrodes hold together by the electrode holder (Fig. 1A) were aligned with each hole created in the microfluidic chip before trapping the animals. This alignment process was only required once for each

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microfluidic chip. The whole process from trapping four animals to inserting needle electrodes took around two minutes, which is about 7 times faster than our previous method(Cho et al., 2015). A detailed illustration

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of the inside of the microfluidic chip is shown in Fig. 1C. With the X-, Y-, and Z- directional stages (Fig. 1A), precise manipulation of the fish and the needle electrodes was possible with micrometre-range precision. The tungsten needle electrodes were sharp and rigid enough to penetrate the larvae’s skin. Additionally, the electrical characteristics of the electrodes were appropriate to acquire zebrafish

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intramuscular signals.

Intramuscular activity recording from multiple zebrafish Fig. 1C shows the schematic view of the zebrafish and electrodes inside a microfluidic channel. Even though the zebrafish and electrodes were inside the water, we could obtain stable and independent intramuscular

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signals from each zebrafish, as shown in Fig. 2. Although the microfluidic chip was designed to trap eight zebrafish simultaneously, the intramuscular activities were measured from four zebrafish due to the limitation of the recording capability of the used signal acquisition system. However, the measurement of more larvae is possible by simply increasing the number of recording channels of the signal acquisition system. To demonstrate the feasibility of monitoring the activities of multiple locomotors of trapped larvae in the microfluidic chip, we first recorded the signals from a drug-free control group, followed by groups of fish

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pre-incubated in Tricaine, NMDA, and MK-801 solutions as a proof of principle. Like other mammalian animal models, zebrafish locomotor activity can be pharmacologically manipulated. Tricaine is a sodium

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blocker that is known to eliminate spontaneous contractions of muscles(Matthews and Varga, 2011), which is

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the only licensed anesthetic agent approved for fish by the US Food and Drug Administration. NMDA is an excitatory amino acids, which is known to induce rhythmic motor output with an increased synaptic drive to

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motoneurons (Dale and Roberts, 1984; Gabriel et al., 2007). MK-801, a noncompetitive NMDA receptor antagonist, has been widely used in learning and memory research; it has been reported that the

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administration of MK-801 decreases the amplitude and frequency of locomotor activities in both mammals

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and zebrafish (F. Fenaux, M. Corio, R. Palisses, 1991; Kyriakatos et al., 2011; Sison and Gerlai, 2011). After trapping the zebrafish larvae, the electrical activities from multiple larvae muscles were measured using the needle electrodes. The recording was performed in a quiet and dark condition for 5 min, as we found that the larvae were not active and had a tendency to maintain its position straight in such conditions. Although a

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more extended recording was possible using the current setup, it is rare to record more than five minutes of needle EMG. It is recommended to obtain around 20 activities for EMG analysis, which only takes a minute to obtain them from an object(Daube and Rubin, 2009). The recording was attempted on 64 larvae in total, and we were able to successfully obtain the signals from 48 larvae, which yielded 75% of success rate in the recording. The low yield was primarily speculated to originate from the two factors. First, the distance between the ground electrode and the needle electrode was not adjustable depending on individual animals. 11

Second, the channel size was larger than the body size of larvae, as the chip was designed to trap their heads. Because of this, the larvae were able to wag its tails during the needle insertion process. Fig. 2 shows representative examples of the recorded muscle activities of 5 dpf zebrafish from each group (total n = 48, see Fig. S5 for signals in other time scales. Note that the presented signals are from all different fish.). The recorded signals were acceptably stable. Because the trapped zebrafish heads were well contacted with the ground electrodes, the low electrical noise was achieved with high signal-to-noise-ratio of 23dB. Fig. 3

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represents the average number of muscle activities that were detected and the interburst intervals of three groups. Significant differences between groups were observed under the 40 ms of interburst interval. The

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burst pattern of the spontaneous activities from the control group corresponded with those reported in

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previous studies of zebrafish(Cho et al., 2015). The control group showed electrical signals of less than 200 µV in peak-to-peak amplitude, which corresponded well with the intramuscular signals of adult zebrafish that

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have been previously reported(Liu and Westaerfield, 1988), as expected. Predictably, no muscle activities were detected in the Tricaine group (Fig. 2B). In the NMDA group, rhythmic locomotor activity was

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initiated, as shown in Fig. 2C. The NMDA group showed increased signal amplitudes by approximately 200% and increased locomotors frequency by 150%, which was consistent with the findings in juvenile and

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adult zebrafish and other animal models(Dale and Roberts, 1984; Giroux, 2003; Muller et al., 2000; Müller and van Leeuwen, 2004). The MK-801 group showed decreased amplitudes by 12% and decreased the frequency of the bursts by 95% compared to the control group, similar to the motor pattern observed in adult

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zebrafish through electrophysiological and behavioral analysis(Kyriakatos et al., 2011; Sison and Gerlai, 2011). Interestingly, the most common interburst interval in the MK-801 group was 20 ms, compared to 10 ms for the control and NMDA groups.

Conclusions Previously, only the EMG of single zebrafish was recorded due to technical limitations. In this study, high12

throughput intramuscular recordings from multiple zebrafish larvae were demonstrated for the first time. The larvae were successfully restrained in water inside the microfluidic chip, without using anesthetic agents, which is a healthier environment for zebrafish than taking them out of water in previous studies. Longer-term electrophysiological recordings would be only possible by establishing such a fish-friendly environment. In addition, the design of the microfluidic chip is easily expandable to accommodate more larvae by increasing the number of trapping channels in the chip. As a proof-of-concept, we performed EMG recordings in four

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different conditions, with larvae preincubated in three different chemicals affecting the muscle activity and one with no treatment. We could clearly recognize the effects of the chemicals by observing distinctive EMG

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signals per chemical. Although the signals were recorded only from four larvae at a time due to the limitation

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of the used signal acquisition system, the results prove that this new technique provides a way to highthroughput electrophysiological recording and a healthier environment for zebrafish, suggesting the potential

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for long-term intramuscular recording. This integrated electrophysiological microfluidic system can be used

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Declaration of interests

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to validate neuromuscular disease models and investigate the efficacy of pharmaceuticals.

The authors declare that they have no known competing financial interests or personal relationships that could have

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appeared to influence the work reported in this paper.

The authors declare the following financial interests/personal relationships which may be considered as potential competing interests:

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Acknowledgments We thank Dr. Seok-Yong Choi (Chonnam National University Medical School) for providing zebrafish. This research was supported by the Brain Research Program (NRF-2018M3C7A1022309) and the Basic Science Research Program (NRF-2018R1A1A1A05020389) through the National Research Foundation funded by the Ministry of Science and ICT of Korea. This research was also supported in part by Canadian Institutes of

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Health Research (MFE-164732).

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FIGURES

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Figure 1 A. Schematic illustration of the developed multiple zebrafish EMG system. The microfluidic chip contains a small well to hold the zebrafish suspension, outlets, trapping channels, and electrodes. The microfluidic chip is placed on the micromanipulator including X-, Y-, and Z- stages. The manipulator sturdily holds the needle electrodes and the microfluidic chip. The insertion of the needle electrodes is controlled by the Z-axis stage, and the microfluidic chip is controlled by the X- and Y- stages, which allows for precise needle insertion. B. Picture of trapped zebrafish in the fabricated microfluidic chip. C. Schematic illustration of zebrafish and electrodes inside a microfluidic channel. The head of the trapped zebrafish makes contact with the ground electrode, and the needle electrode is vertically inserted under the direct visual guidance.

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Figure 2 Typical intramuscular activities detected in zebrafish larvae preincubated in solutions containing different chemicals. Spontaneous muscle activity recordings from 5 dpf zebrafish that were captured in the microfluidic chip. The control group was not exposed to any drugs (A). Larvae were exposed to 0.61 mM of Tricaine for 5 min (B), and 80 μM of NMDA and MK-801 for 20 min prior to recordings and throughout the recording sessions (C, D). The numbers in black circles represent that the signals were obtained from four different zebrafish in one microfluidic chip.

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Figure 3 Interburst interval and number of bursts. Average number of detected muscle activities (Yaxis), and interburst intervals (X-axis) in the control (A), NMDA (B), and MK-801 (C) groups (n = 12 each). Note that the Tricaine group was excluded because no muscle activity was detected. Data are presented as mean ± SEM and analyzed by one-way ANOVA followed by Bonferroni’s test as post-hoc. * indicates significant difference between groups with p<0.05.

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