We thank Fred Roberts&#39; technical support and revision of the manuscript.

All procedures involving zebrafish were approved by our Institutional Animal Care and Use Committee and are in compliance with the USDA Animal Welfare Act.
1. Experimental set-up
<ol>
	<li>Setting up the platform for image acquisition
	<ol>
		<li>Using small scissors or a scalpel make an incision on a sponge at the 12 o&#39;clock position to hold the fish during scanning. Place the sponge in a glass container (Figure 2A). 
		NOTE: The position of the incision should allow enough room to move transducer and also to keep the fish bellow the water line when the platform is tilted for scanning (Figure 2). The incision can vary depending on the size of the fish; however, for a standard size and weight, the incision should be approximately 2.5 cm x 0.7 cm x 0.5 cm (length, width, and depth, respectively). The glass container should be at least 6 cm deep to avoid water leakage while imaging the fish.</li>
		<li>Affix the glass box containing the sponge on the ultrasound platform, for instance using double-sided tape. Ensure the glass box is at the center of the platform and firmly attached (Figure 2B).</li>
		<li>Tilt the platform forward about 30&#176; using the knob on the left side of the platform holder (Figure 2B). Fill the glass square with 200-250 mL of fish system water containing 0.2 mg/mL tricaine methanesulfonate (MS222). 
		NOTE: Tricaine can be prepared as a 4 mg/mL stock solution in Tris 40 mM pH 7 and further diluted to the desired concentration in fish system water; 0.2 mg/mL was found to be the best concentration16. The 4 mg/mL tricaine stock solution can be stored for a long period of time at -20 &#176;C or at 4 &#176;C for one month.</li>
		<li>Insert the transducer within the micromanipulator holder on the working rail station, turning the notch of the transducer towards the operator. Keep the array parallel to the ground with the working side longitudinal with respect to the stage (see Figure 2B). Leave enough room (10 cm on both sides) for the now connected transducer-rail system to move along the x- and y-axes.</li>
		<li>Log in to the control software and choose Mouse (Small) Vascular. Create a new study as well as a new series for each animal included in the study. Find the new study button located on the bottom left side of the screen on the browser page (the view starts in B-mode).</li>
	</ol>
	</li>
</ol>
2. Handling the Fish
NOTE: Zebrafish used in this study were adult, 11-month-old males of the wild-type strain AB/Tuebingen (AB/TU). Zebrafish were maintained in a stand-alone flow-through aquarium system at 28 &#176;C in a constant light cycle set as 14 h light/10 h dark. Zebrafish were fed twice daily with brine shrimp (Artemia nauplii) and dry food flakes.
<ol>
	<li>Using a fish net, transfer the fish into a small tank containing system water with 0.2 mg/mL tricaine. Wait until the fish is fully anesthetized (no movement and no response to touch).</li>
	<li>Using a plastic teaspoon, gently and quickly transfer the fish into the glass box containing the sponge into the previously made incision with ventral side of the fish facing up. 
	NOTE: Make sure the head of the fish is positioned towards the operator (same direction as the notch of the transducer) and at a slightly higher level compared to the rest of the body to achieve better heart visualization.</li>
	<li>Gently lower the transducer (keeping its original position) using the handle on the rail system, placing it longitudinally and close to the ventral side of the fish with the notch of the transducer facing the operator. Leave 2-3 mm (no more than 1 cm) clearance from the fish. Adjust the platform in respect to the transducer using the micromanipulator in all 3 axes until the fish heart is visualized and then start image acquisition. The angle of the transducer should not be changed during the entire image acquisition (Figure 2C). 
	NOTE: As long as there is enough proximity (up to 1 cm), the water on top of the fish will provide a contact surface via liquid surface tension that allows transmission of the ultrasound waves between the probe and the fish. Therefore, there is no need to push the transducer against the fish. Try to complete this step and finish the scan in less than 3 minutes to prevent fish death or a decrease of the heart rate during image acquisition. If needed, use a timer. The heart can be found on the upper side of the screen towards the left side of the eye, which can be easily visualized if moving the x-axis all the way to the right. If there is continued difficulty in finding the heart while in B-Mode, switch to color Doppler mode, which will allow for tracking blood flow (red indicates blood flowing towards the operator) and locating the heart.</li>
</ol>
3. Image acquisition
NOTE: See Table of Materials for imaging system and image analysis software.
<ol>
	<li>Longitudinal View B-Mode
	<ol>
		<li>After localizing the heart, select or stay in B-Mode (found at the bottom left side of the touchscreen after having initiated a new series) and reduce the field in order to zoom in and have a closer look at the heart for easier tracing during analysis.</li>
		<li>In order to have a closer and clearer view of the heart in B-Mode image acquisition, reduce the field by zooming in. Use the touchscreen to manually narrow the field on both the x- and y-axes.</li>
		<li>If needed, enhance the quality/contrast of the image by setting the dynamic range to 45-50 dB. Go to the B-mode controls in the More Controls option and subsequently save the change to Mode Presets. Tap Mode Presets to select the optimized image acquisition setting every time before starting to image a new series.</li>
		<li>Take as many images as desired in the long axis plane by selecting Save Image. 
		NOTE: More detailed information and training resources on image acquisition can be found at <a href="https://www.visualsonics.com/product/software/vevo-lab" target="_blank">https://www.visualsonics.com/product/software/vevo-lab</a> and <a href="https://www.visualsonics.com/Learning-hub-online-video-training-our-users" target="_blank">https://www.visualsonics.com/Learning-hub-online-video-training-our-users</a></li>
	</ol>
	</li>
	<li>Longitudinal View Pulse Wave
	<ol>
		<li>Switch to Color Doppler for blood flow detection (select Color button) and acquisition (found at the bottom left side of the touchscreen after having initiated a new series).</li>
		<li>Using the touch screen position the quadrant on top of the atrioventricular valve and localize the inflow, which will be distinguished by the red color signal (Figure 3A). Reduce the quadrant area as much as possible to increase the frame rate. 
		NOTE: Lower the Color pulse-repetition-frequency (Color PRF) (velocity range) to ensure yellow color can be seen in the velocity profile of the Color Doppler image. This will increase the range of velocities that can be seen and will help to create a mosaic of color that will allow to visualize more clearly the peak velocities.</li>
		<li>Activate pulse wave (select PW) Doppler Mode to sample ventricular blood inflow velocity. Position the sample volume gate at the center of the atrioventricular valve (where the red color signal becomes more yellowish) to detect the maximum flow velocity. Adjust the PW angle on the screen using your fingers so it aligns with the direction of the blood inflow. Press start or update to start sampling the velocity of blood flowing into the ventricle. 
		NOTE: Make sure the angle correct line is parallel to the blood flow in order to provide consistent and reproducible results. Placing the angle correct line so it matches the direction of blood flow will ensure that velocities are accurately captured.</li>
		<li>Repeat step 3.2.3 to determine the outflow velocity by placing the Color Doppler quadrant at the junction between the ventricle and the bulbus (bulbuventricular valve) and localize the flow, which will be distinguished by a blue color signal (Figure 3B). Position the sample volume gate right before the ventricle-bulbus junction and adjust the angle correction line to match the direction of the blood flow. 
		NOTE: As mentioned before, to achieve accurate velocity values, make sure the PW angle is aligned with the blood flow.</li>
		<li>Adjust the baseline (bar), lowering or raising it in the flow velocity panel, in order to detect and trace completely the signal peaks (Figure 3C,D). Identify the inflow peaks in the upper/positive quadrant (signal going towards the probe) and the outflow peaks in the lower/negative quadrant (signal going away from the probe).</li>
	</ol>
	</li>
</ol>
4. Fish recovery
<ol>
	<li>As soon as image acquisition is complete, using a teaspoon, transfer the fish into regular system aerated water free of tricaine and let the fish recover (usually takes 30 s to 2 min to resume gill movement and swimming).</li>
	<li>To help recovery, squirt water repeatedly over the gills using a transfer pipette to promote aeration of the water and oxygen transfer.</li>
</ol>
5. Image analysis
<ol>
	<li>Open the image analysis software.</li>
	<li>Select an image and click on the image processing icon (Figure 4). Using the available scale (Figure 4), adjust brightness and contrast of the image to allow clear visualization of ventricular walls or blood flow pattern.</li>
	<li>Using the B-mode image, open the drop-down list from the PSLAX (parasternal long axis) option on the cardiac package/measurements (Figure 4). Select LV trace and trace the ventricular inner wall at systole and diastole to obtain the ventricular area (VA) in systole (VAs) and diastole (VAd), end diastolic volume (EDV), and end systolic volume (ESV) (Figure 5A,B). 
	NOTE: Volume values are extrapolated from 2D image tracings and might deviate from the 3D entity. For all measurements, average at least 3 representative cardiac cycles per animal.</li>
	<li>Note the stroke volume and ejection fraction that will be automatically calculated and displayed by the software. 
	NOTE: Stroke volume, and ejection fraction can also be manually calculated using the formulas 
	SV = EDV-ESV 
	EF = (EDV-ESV)/EDV 
	where SV is stroke volume, EDV is end diastolic volume, ESV is end systolic volume, and EF is ejection fraction</li>
	<li>Calculate fractional area change using the formula 
	FAC = (VAd - VAs)/ VAd 
	where FAC is fractional area change, VAd is ventricular area in diastole, and VAs is ventricular area in systole.</li>
	<li>Calculate the cardiac output using the formula 
	CO = HR x SV 
	where CO is cardiac output, HR is heart rate, and SV is stroke volume</li>
	<li>Using the Pulsed Wave Doppler Mode image, measure the inflow blood velocity by selecting the MV Flow option under the cardiac package (Figure 4). Select E or A for early diastole and late diastole, respectively, and determine the peak-velocities on the graph (Figure 3C).</li>
	<li>Measure the outflow blood velocity by selecting AoV Flow and determine the peaks on the tracing (Figure 3D).</li>
	<li>Measure the heart rate using 2 different methodologies for a more reliable assessment:
	<ol>
		<li>When the heart is visualized on the screen during image acquisition, count the beats within 10 s and multiply it by 6.</li>
		<li>Using the Pulse Wave Doppler image on the Vevo LAB software, choose the heart rate button and trace intervals between 3 consecutive aortic &#64258;ow peaks (Figure 4 and Figure 6).</li>
		<li>To export data to a spreadsheet after having traced the LV and the peaks of the blood flow, click on report | export | save as | excel.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

We describe a systematic method for echocardiographic imaging and assessment of cardiac function in adult zebrafish. Echocardiography is the only available non-invasive and most robust method for live adult fish cardiac imaging and functional analysis, and it is becoming increasingly popular in zebrafish cardiovascular research. The amount of time needed is short and allows for high-throughput and longitudinal studies. However, there is considerable variation in the methodology employed and data analysis. Standardization...

The zebrafish (Danio rerio) is a well-established vertebrate model for studies of developmental processes and human diseases1. Zebrafish have high genetic similarity to humans (70%), genetic tractability, high fecundity, and optical transparency during embryonic development, which allows direct visual analysis of organs and tissues, including the heart. Despite having just one atrium and one ventricle, the zebrafish heart (Figure 1) is physiologically similar to mammalian four-chambered hearts. Importantly, the zebrafish heart rate, electrocardiogram morphology, and action potential shape resemble those of humans more than murine species2. These features have made zebrafish an excellent model for cardiovascular research and have provided major insights into cardiac development3,4, regeneration5, and pathologic conditions1,3,4, including arteriosclerosis, cardiomyopathies, arrhythmias, congenital heart diseases, and amyloid light chain cardiotoxicity1,4,6. Assessment of cardiac function has been possible during the embryonic stage (1-days post fertilization) through direct video analysis using high-speed video microscopy7,8. However, zebrafish lose their transparency beyond the embryonic stage, limiting functional evaluations of normal mature hearts and late-onset heart conditions. To overcome this limitation, echocardiography has been successfully employed as a high-resolution, real-time, noninvasive imaging alternative to evaluate adult zebrafish heart function9,10,11,12,13,14,15.
In zebrafish, the heart is located ventrally in the thoracic cavity immediately posterior to the gills with the atrium located dorsal to the ventricle. The atrium collects venous blood from the sinus venosus and transfers it to the ventricle where it is further pumped to the bulbus arteriosus (Figure 1). Here, we describe a physiological, underwater, protocol to assess cardiac function in adult zebrafish by non-invasive echocardiography using a linear array ultrasound probe with a center frequency of 50 MHz for B-mode imaging at a resolution of 30 &#181;m. Since ultrasound waves can easily travel through water, keeping close proximity between the fish and the scanning probe underwater provides enough contact surface for heart detection with no need for ultrasound gel and is overall less stressful for the fish. Although alternative zebrafish echocardiography systems were reported by several authors9,12,13, here we present the general and most commonly used setup that applies to high-frequency ultrasound in animals.
The method allows high resolution imaging of the adult zebrafish heart, tracing of cardiac structures, and quantification of peak-velocities from Doppler blood flow measurements. We show reliable in vivo quantification of important systolic and diastolic parameters, such as ejection fraction (EF), fractional area change (FAC), ventricular blood inflow and outflow velocities, heart rate (HR), and cardiac output (CO). We contribute to establishing a reliable range of normal healthy adult zebrafish cardiac functional and dimensional parameters to allow a more precise evaluation of pathologic states. Overall, we provide a robust method to assess cardiac function in zebrafish, which has proven extremely useful in establishing and validating zebrafish heart disease models6,16, heart injury and recovery10,13, and regeneration11,12, and can be further used to evaluate potential drugs.

<table><tbody><tr><td>Double sided tape</td><td></td><td></td><td></td></tr><tr><td>Fish net</td><td></td><td></td><td></td></tr><tr><td>Glass container - 100 inch high</td><td></td><td></td><td></td></tr><tr><td>High frequency transducer</td><td>Fujifilm/VisualSonics</td><td>MX700</td><td>Band width 29-71 MHz, Centre transmit 50 MHz, Axial resolution 30 &amp;micro;m</td></tr><tr><td>Plastic teaspoon</td><td></td><td></td><td></td></tr><tr><td>Scalpel or scissors</td><td></td><td></td><td></td></tr><tr><td>Small fish tanks</td><td></td><td></td><td></td></tr><tr><td>Sponge (kitchen sponge)</td><td></td><td></td><td></td></tr><tr><td>Transfer pipets (graduated 3 mL)</td><td>Samco Scientific</td><td>212</td><td></td></tr><tr><td>Tricaine (MS-222)</td><td>Sigma-Aldrich</td><td>A5040</td><td></td></tr><tr><td>Vevo 3100 Imaging system and imaging station</td><td>Fujifilm/VisualSonics</td><td></td><td></td></tr><tr><td>Vevo LAB sofware v 1.7.1</td><td>Fujifilm/VisualSonics</td><td></td><td></td></tr></tbody></table>

high-frequency ultrasound echocardiography to assess zebrafish cardiac function

The zebrafish (Danio rerio) has become a very popular model organism in cardiovascular research, including human cardiac diseases, largely due to its embryonic transparency, genetic tractability, and amenity to rapid, high-throughput studies. However, the loss of transparency limits heart function analysis at the adult stage, which complicates modeling of age-related heart conditions. To overcome such limitations, high-frequency ultrasound echocardiography in zebrafish is emerging as a viable option. Here, we present a detailed protocol to assess cardiac function in adult zebrafish by non-invasive echocardiography using high-frequency ultrasound. The method allows visualization and analysis of zebrafish heart dimension and quantification of important functional parameters, including heart rate, stroke volume, cardiac output, and ejection fraction. In this method, the fish are anesthetized and kept underwater and can be recovered after the procedure. Although high-frequency ultrasound is an expensive technology, the same imaging platform can be used for different species (e.g., murine and zebrafish) by adapting different transducers. Zebrafish echocardiography is a robust method for cardiac phenotyping, useful in the validation and characterization of disease models, particularly late-onset diseases; drug screens; and studies of heart injury, recovery, and regenerative capacity.

The described protocol allows for measurement of important cardiac dimensional and functional parameters, analogous to the technique used in human and animal echocardiography. The B-Mode images allow for tracing of ventricular inner wall in systole and diastole (Figure 5) and obtaining of dimensional data, such as chamber and wall dimensions, and functional data, such as heart rate, stroke volume, and cardiac output as well as parameters of ventricular systol...

Watch this Scientific Journal Video about High-Frequency Ultrasound Echocardiography to Assess Zebrafish Cardiac Function at JoVE.com

High-Frequency Ultrasound Echocardiography to Assess Zebrafish Cardiac Function

We describe a protocol to assess heart morphology and function in adult zebrafish using high-frequency echocardiography. The method allows visualization of the heart and subsequent quantification of functional parameters, such as heart rate (HR), cardiac output (CO), fractional area change (FAC), ejection fraction (EF), and blood inflow and outflow velocities.

The zebrafish (Danio rerio) has become a very popular model organism in cardiovascular research, including human cardiac diseases, largely due to its ...

high-frequency-ultrasound-echocardiography-to-assess-zebrafish

Research

JoVE Journal

Developmental Biology

8.7K Views.  Stanford University. Het protocol maakt niet-invasieve en hoge resolutie representatie en analyse van volwassen zebravis hartfunctie. Deze methode kan worden gebruikt voor de validatie van volwassen hart-en vaatziekten modellen en potentiële drug schermen. Deze techniek maakt een snelle positionering van de vis mogelijk voor een correcte beeldvorming die nauwkeurige en reproduceerbare resultaten oplevert.Het is niet invasief en uitgevoerd onder water die het mogelijk maakt de snelle terugwinning van de vi...

Video: High-Frequency Ultrasound Echocardiography to Assess Zebrafish Cardiac Function

Large-scale Zebrafish Embryonic Heart Dissection for Transcriptional Analysis

The zebrafish embryonic heart is composed of only a few hundred cells, representing only a small fraction of the entire embryo. Therefore, to prevent the cardiac transcriptome from being masked by the global embryonic transcriptome, it is necessary to collect sufficient numbers of hearts for further analyses. Furthermore, as zebrafish cardiac development proceeds rapidly, heart collection and RNA extraction methods need to be quick in order to ensure homogeneity of the samples. Here, we present a rapid manual dissection protocol for collecting functional/beating hearts from zebrafish embryos. This is an essential prerequisite for subsequent cardiac-specific RNA extraction to determine cardiac-specific gene expression levels by transcriptome analyses, such as quantitative real-time polymerase chain reaction (RT-qPCR). The method is based on differential adhesive properties of the zebrafish embryonic heart compared with other tissues; this allows for the rapid physical separation of cardiac from extracardiac tissue by a combination of fluidic shear force disruption, stepwise filtration and manual collection of transgenic fluorescently labeled hearts.

To analyse cardiac gene expression profiles during zebrafish heart development, total RNA has to be extracted from isolated hearts. Here, we present a protocol for collecting functional/beating hearts by rapid manual dissection from zebrafish embryos to obtain cardiac-specific mRNA.

The zebrafish embryonic heart is composed of only a few hundred cells, representing only a small fraction of the entire embryo. Therefore, to prevent the ...

Displacement Analysis of Myocardial Mechanical Deformation (DIAMOND) Reveals Segmental Heterogeneity of Cardiac Function in Embryonic Zebrafish

Zebrafish are increasingly utilized as a model organism for cardiomyopathies and regeneration. Current methods evaluating cardiac function fail to reliably detect segmental mechanics and are not readily feasible in zebrafish. Here we present a semiautomated, open-source method for the quantitative assessment of four-dimensional (4D) segmental cardiac function: displacement analysis of myocardial mechanical deformation (DIAMOND). Transgenic embryonic zebrafish were imaged in vivo using a light-sheet fluorescence microscopy system with 4D cardiac motion synchronization. Acquired 3D digital hearts were reconstructed at end-systole and end-diastole, and the ventricle was manually segmented into binary datasets. Then, the heart was reoriented and isotropically resampled along the true short axis, and the ventricle was evenly divided into eight portions (I&#8211;VIII) along the short axis. Due to the different resampling planes and matrices at end-systole and end-diastole, a transformation matrix was applied for image registration to restore the original spatial relationship between the resampled systolic and diastolic image matrices. After image registration, the displacement vector of each segment from end-systole to end-diastole was calculated based on the displacement of mass centroids in three dimensions (3D). DIAMOND shows that basal myocardial segments adjacent to the atrioventricular canal undergo the highest mechanical deformation and are the most susceptible to doxorubicin-induced cardiac injury. Overall, DIAMOND provides novel insights into segmental cardiac mechanics in zebrafish embryos beyond traditional ejection fraction (EF) under both physiological and pathological conditions.

Displacement Analysis of Myocardial Mechanical Deformation DIAMOND Reveals Segmental Heterogeneity of Cardiac Function in Embryonic Zebrafish

The goal of this protocol is to detail a novel method for the assessment of segmental cardiac function in embryonic zebrafish under both physiological and pathological conditions.

Zebrafish are increasingly utilized as a model organism for cardiomyopathies and regeneration. Current methods evaluating cardiac function fail to ...

Design and Microinjection of Morpholino Antisense Oligonucleotides and mRNA into Zebrafish Embryos to Elucidate Specific Gene Function in Heart Development

The morpholino oligomer-based knockdown system has been used to identify the function of various gene products through loss or reduced expression. Morpholinos (MOs) have the advantage in biological stability over DNA oligos because they are not susceptible to enzymatic degradation. For optimal effectiveness, MOs are injected into 1-4 cell stage embryos. The temporal efficacy of knockdown is variable, but MOs are believed to lose their effects due to dilution eventually. Morpholino dilution and injection amount should be closely controlled to minimize the occurrence of off-target effects while maintaining on-target efficacy. Additional complementary tools, such as CRISPR/Cas9 should be performed against the target gene of interest to generate mutant lines and to confirm the morphant phenotype with these lines. This article will demonstrate how to design, prepare, and microinject a translation-blocking morpholino against hand2 into the yolk of 1-4 cell stage zebrafish embryos to knockdown hand2 function and rescue these "morphants" by co-injection of mRNA encoding the corresponding cDNA. Subsequently, the efficacy of the morpholino microinjections is assessed by first verifying the presence of morpholino in the yolk (co-injected with phenol red) and then by phenotypic analysis. Moreover, cardiac functional analysis to test for knockdown efficacy will be discussed. Finally, assessing the effect of morpholino-induced blockage of gene translation via western blotting will be explained.

The present protocol describes designing, preparing, and microinjecting a translational-blocking morpholino against a representative cardiac gene; Heart And Neural Crest Derivatives Expressed2 (hand2) into the yolk of newly fertilized zebrafish embryos to knock down gene function. It also shows a transient rescue of these "morphants" by co-injection of mRNA encoding this gene product.

The morpholino oligomer-based knockdown system has been used to identify the function of various gene products through loss or reduced expression.  ...

Light-sheet Fluorescence Microscopy to Capture 4-Dimensional Images of the Effects of Modulating Shear Stress on the Developing Zebrafish Heart

The hemodynamic forces experienced by the heart influence cardiac development, especially trabeculation, which forms a network of branching outgrowths from the myocardium. Genetic program defects in the Notch signaling cascade are involved in ventricular defects such as Left Ventricular Non-Compaction Cardiomyopathy or Hypoplastic Left Heart Syndrome. Using this protocol, it can be determined that shear stress driven trabeculation and Notch signaling are related to one another. Using Light-sheet Fluorescence Microscopy, visualization of the developing zebrafish heart was possible. In this manuscript, it was assessed whether hemodynamic forces modulate the initiation of trabeculation via Notch signaling and thus, influence contractile function occurs. For qualitative and quantitative shear stress analysis, 4-D (3-D+time) images were acquired during zebrafish cardiac morphogenesis, and integrated light-sheet fluorescence microscopy with 4-D synchronization captured the ventricular motion. Blood viscosity was reduced via gata1a-morpholino oligonucleotides (MO) micro-injection to decrease shear stress, thereby, down-regulating Notch signaling and attenuating trabeculation. Co-injection of Nrg1 mRNA with gata1a MO rescued Notch-related genes to restore trabeculation. To confirm shear stress driven Notch signaling influences trabeculation, cardiomyocyte contraction was further arrested via tnnt2a-MO to reduce hemodynamic forces, thereby, down-regulating Notch target genes to develop a non-trabeculated myocardium. Finally, corroboration of the expression patterns of shear stress-responsive Notch genes was conducted by subjecting endothelial cells to pulsatile flow. Thus, the 4-D light-sheet microscopy uncovered hemodynamic forces underlying Notch signaling and trabeculation with clinical relevance to non-compaction cardiomyopathy.

Here, we present a protocol to visualize developing hearts in zebrafish in 4-Dimensions (4-D). 4-D imaging, via light-sheet fluorescence microscopy (LSFM), takes 3-Dimensional (3-D) images over time, to reconstruct developing hearts. We show qualitatively and quantitatively that shear stress activates endocardial Notch signaling during chamber development, which promotes cardiac trabeculation.

The hemodynamic forces experienced by the heart influence cardiac development, especially trabeculation, which forms a network of branching outgrowths ...

Bioengineering

Analysis of Zebrafish Cardiac Contraction with Virtual Reality Interaction

4D Light-sheet Imaging of Zebrafish Cardiac Contraction

Zebrafish is an intriguing model organism known for its remarkable cardiac regeneration capacity. Studying the contracting heart in vivo is essential for gaining insights into structural and functional changes in response to injuries. However, obtaining high-resolution and high-speed 4-dimensional (4D, 3D spatial + 1D temporal) images of the zebrafish heart to assess cardiac architecture and contractility remains challenging. In this context, an in-house light-sheet microscope (LSM) and customized computational analysis are&#160;used to overcome these technical limitations. This strategy, involving LSM system construction, retrospective synchronization, single cell tracking, and user-directed analysis, enables one to investigate the micro-structure and contractile function across the entire heart at the single-cell resolution in the transgenic Tg(myl7:nucGFP) zebrafish larvae. Additionally, we are able to further incorporate microinjection of small molecule compounds to induce cardiac injury in a precise and controlled manner. Overall, this framework allows one to track physiological and pathophysiological changes, as well as the regional mechanics at the single-cell level during cardiac morphogenesis and regeneration.

Author Spotlight: High-Resolution 4D Light-Sheet Imaging and Virtual Reality in Zebrafish for Single-Cell Analysis of Heart Function

This protocol utilizes light-sheet imaging to investigate cardiac contractile function in zebrafish larvae and gain insights into cardiac mechanics through cell tracking and interactive analysis.

Zebrafish is an intriguing model organism known for its remarkable cardiac regeneration capacity. Studying the contracting heart in vivo is essential for ...

Zebra II: Advanced ECG System for Multichannel Zebrafish Cardiovascular Studies

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

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

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

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

Anesthesia-free Heartbeat Measurements in Freely Moving Zebrafish

Zebrafish (Danio rerio) is a widely used model organism in physiological, pharmacological, and toxicological research due to its genetic similarity to humans and transparent embryonic stage, which facilitates non-invasive cardiovascular studies. However, current methods for heart rate assessment in zebrafish often rely on anesthesia to immobilize the subject, introducing physiological alterations that compromise data accuracy and reproducibility. This study presents a novel, anesthesia-free technique for measuring heartbeat in freely moving zebrafish larvae, addressing a critical limitation in cardiovascular research. The proposed approach integrates shortwave-infrared imaging with machine-learning-based heart tracking, allowing for precise and continuous cardiac activity monitoring in non-immobilized specimens. A convolutional neural network was trained to detect the heart region, and a photoplethysmographic signal was extracted from image sequences to determine heart rate. Experimental validation demonstrated the method's reliability and consistency across multiple test conditions. A key benefit of the methodology is its ability to preserve the natural physiological state of zebrafish, minimizing stress-induced artifacts. This non-invasive, label-free technique offers significant advantages for studying cardiovascular physiology, drug cardiotoxicity, and environmental toxicology, expanding the potential applications of zebrafish as a model for biomedical research.

The study aims to develop technology for anesthesia-free heartbeat measurements in moving zebrafish. Our approach combines shortwave-infrared imaging and machine-learning-based tracking of the heart. It is a non-invasive, label-free, and user-friendly technique that suits a wide range of studies on the zebrafish model.

Zebrafish (Danio rerio) is a widely used model organism in physiological, pharmacological, and toxicological research due to its genetic similarity to ...

Recovery of Adult Zebrafish Hearts for High-throughput Applications

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and purification of RNA, DNA, and proteins. All of these applications demand the rapid recovery of significant numbers of zebrafish hearts to avoid gene regulatory, metabolic, and other changes that begin after death. Adult zebrafish hearts are also required for studying heart structure for a variety of mutants and for studying heart regeneration. However, the traditional zebrafish heart dissection is slow and difficult and requires specialized tools, making large-scale dissection of adult zebrafish hearts tedious. Traditional methods also harbor the risk of damaging the heart during the dissection. Here, we describe a method for dissection of adult zebrafish hearts that is fast, reproducible, and preserves heart architecture. Furthermore, this method does not require specialized tools, is painless for the zebrafish, can be performed on fresh or fixed specimens, and can be performed on zebrafish as young as one month old. The approach described expands the use of adult zebrafish for cardiovascular research.

Use of zebrafish for cardiovascular research is expanding towards research on adult hearts. For these applications, quick and simple isolation of cardiac tissues is key to avoid post-mortem changes and to obtain an adequate number of samples. Here, we describe a fast and reproducible method for dissecting adult zebrafish hearts.

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and ...

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 ...

Light Sheet Microscopy of Fast Cardiac Dynamics in Zebrafish Embryos

Embryonic cardiac research has greatly benefited from advances in fast in vivo light sheet fluorescence microscopy (LSFM). Combined with the rapid external development, tractable genetics, and translucency of the zebrafish, Danio rerio, LSFM has delivered insights into cardiac form and function at high spatial and temporal resolution without significant phototoxicity or photobleaching. Imaging of beating hearts challenges existing sample preparation and microscopy techniques. One needs to maintain a healthy sample in a constricted field of view and acquire ultrafast images to resolve the heartbeat. Here we describe optimized tools and solutions to study the zebrafish heart in vivo. We demonstrate the applications of bright transgenic lines for labeling the cardiac constituents and present novel gentle embedding and immobilization techniques that avoid developmental defects and changes in heart rate. We also propose a data acquisition and analysis pipeline adapted to cardiac imaging. The entire workflow presented here focuses on zebrafish embryonic heart imaging but can also be applied to various other samples and experiments.

We describe optimized tools to study the zebrafish heart in vivo with light sheet fluorescence microscopy. Specifically, we suggest bright cardiac transgenic lines and present new gentle embedding and immobilization techniques that avoid developmental and heart defects. A possible data acquisition and analysis pipeline adapted to cardiac imaging is also provided.

High-Frequency Ultrasound Echocardiography to Assess Zebrafish Cardiac Function

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