We would like to acknowledge Maurine Hobbs and the Centralized Zebrafish Animal Resource (CZAR) at the University of Utah for providing zebrafish husbandry, laboratory space, and equipment to carry out portions of this research. Expansion of the CZAR is supported in part by NIH grant # 1G20OD018369-01. We would also like to thank Rodney Stewart, Chloe Lim, Lance Graham, Cody James, Garrett Nickum, and the Huntsman Cancer Institute (HCI) Zebrafish Facility for zebrafish care. We would like to thank Kenneth Kompass for help with R programming. This work was funded in part by grants from the Huntsman Cancer Foundation (in conjunction with grant P30 CA042014 awarded to Huntsman Cancer Institute) (KJE) and NIH/NCI R01CA222570 (KJE).

Animal studies are carried out following procedures approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Utah.
1. Fixing Larvae
<ol>
	<li>At 3&#8211;7 days post fertilization (dpf), euthanize larvae with tricaine methanesulfonate (0.03%) and collect up to 15 larvae in a 2 mL tube using a glass pipette and pipette pump.</li>
	<li>Wash larvae twice with 1 mL of cold (4 &#176;C) 1x phosphate-buffered saline (PBS) on ice. For each wash, remove as much liquid as possible from the tube with a glass pipette and pipette pump, and then add 1 mL of cold PBS to tube.</li>
	<li>Remove as much PBS as possible using a glass pipette and pipette pump, and add 1 mL of cold (4 &#176;C) 4% paraformaldehyde (PFA) in PBS. 
	CAUTION: PFA is an irritant and suspected carcinogen. Gloves should be worn when handling PFA, and concentrated solutions should be handled in a chemical fume hood.</li>
	<li>Incubate at 4 &#176;C at least overnight (but up to several months) with gentle rocking.</li>
</ol>
2. Dissecting Tissues Surrounding Liver
<ol>
	<li>Remove larvae from PFA by rinsing 3x with 1 mL of cold (4 &#176;C) PBS and rocking for 5 min in between rinses. 
	NOTE: It is okay to keep the rinsed larvae in PBS for a day or two at 4 &#176;C.</li>
	<li>Pipette several larvae in PBS into one well of a 9-well round-bottom glass dish.</li>
	<li>Remove skin surrounding liver.
	<ol>
		<li>Use fine forceps to hold larva on its back (belly up), gripping on either side of the head as gently as possible. Then use very fine forceps in your other hand to grab the skin just overlying the heart.</li>
		<li>Pull skin down diagonally towards the tail of the fish and the bottom of the dish on the left or right side of the fish. Repeat for other side (right or left side).</li>
		<li>Continue grabbing flaps of skin and pulling down/back until all of the skin and melanophores overlying or near the liver have been removed.</li>
	</ol>
	</li>
	<li>Remove yolk, if present.
	<ol>
		<li>For 5&#8211;6 dpf larvae, lift yolk off in one piece by holding the fish with fine forceps on its back and using the very fine forceps to prod the yolk gently.</li>
		<li>For 3&#8211;4 dpf larvae, scrape the yolk off in pieces. Hold the fish with fine forceps on its back and use the very fine forceps to stroke the yolk, starting from the ventral side.</li>
	</ol>
	</li>
	<li>Place dissected larvae into fresh cold PBS using a glass pipette and pipette pump.</li>
</ol>
3. Imaging
<ol>
	<li>To mount larvae, pour a few mL of 3% methyl cellulose onto the lid of a clean plastic Petri dish.</li>
	<li>Use a glass pipette and pipette pump to add larvae to the methyl cellulose, adding as little PBS with the larvae as possible.</li>
	<li>Under a dissecting microscope at low magnification, use fine forceps to orient the fish so they are laying on their right side, facing left. 
	NOTE: Make sure the fish are oriented perfectly on their side or the liver measurements may not be accurate.</li>
	<li>Take a picture of each fish.
	<ol>
		<li>Confirm that the fish to be photographed is aligned perfectly, with one eye directly on top of the other eye. If necessary, use fine forceps to tap lightly on head or tail of fish to adjust orientation. If fish&#39;s tail is bent, remove the tail by pinching it forcefully with forceps to remove it so the fish lays flat.</li>
		<li>Zoom in to high magnification and focus on the liver, making sure that the liver&#39;s outline is clearly visible.</li>
		<li>Snap a picture and save the file.</li>
		<li>Repeat for all fish, making sure the magnification is the same for each picture.</li>
		<li>Take a picture of a micrometer using the same magnification (see Figure 2H).</li>
	</ol>
	</li>
</ol>
4. Image Analysis
<ol>
	<li>Measure the area of each fish&#39;s liver using image processing software.
	<ol>
		<li>Blind all liver pictures to avoid potential investigator bias and promote scientific rigor15. This step can be done manually by another lab member or using a computer program (Supplementary Material). Rename files randomly and create a &#34;randomization file&#34; containing a list of the original file names and corresponding blinded file names.</li>
		<li>Open randomized files in order, starting with file 1.</li>
		<li>Choose the freehand selections tool and outline each liver.</li>
		<li>Press Ctrl-M to measure the area of each liver.</li>
		<li>For any livers that cannot be accurately measured, insert a placeholder measurement (very small or very large, so it can be easily excluded later on).</li>
		<li>Save the measurements in a text file (&#34;measurements file&#34;).</li>
	</ol>
	</li>
	<li>Un-blind and analyze data
	<ol>
		<li>Open &#34;measurements file&#34; and &#34;randomization file&#34; in a spreadsheet program.</li>
		<li>Insert a new column in the &#34;measurements file&#34; and add the original file names for the blinded files, using the &#34;randomization file&#34;. Save this file as &#34;unblinded measurements file&#34;.</li>
		<li>Sort data by original file name.</li>
		<li>Be sure to exclude any liver measurements for which pictures were inadequate (see Figure 2A&#8211;G).</li>
		<li>If necessary, convert measurement values into desired scale (mm2, for example).
		<ol>
			<li>Open the scale bar in the image processing software.</li>
			<li>Use the straight line tool to measure 1 mm on the scale bar. The image processing software will measure in the same units as the livers (pixels), giving a conversion factor.</li>
			<li>Use the conversion factor to convert measurements in the &#34;unblinded measurements file&#34;.</li>
		</ol>
		</li>
		<li>Using the spreadsheet program or pasting data into a scientific graphing and statistics software, determine mean and standard deviation and calculate p value(s).</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Quantification of liver size is crucial in studies aimed at understanding liver development, regeneration, and oncogenesis. The protocol described here is a relatively quick, easy, and cheap technique for liver size quantification in larval zebrafish. Exercising appropriate caution while performing certain aspects of the protocol can aid in increased accuracy of results and decreased frustration.
Proper fixation of the larvae is crucial towards getting well-preserved biological samples and pre...

Hepatocellular carcinoma (HCC) is the most common primary malignancy of the liver1 and the third leading cause of cancer-related mortality2. To better understand mechanisms of hepatocarcinogenesis and identify potential HCC therapeutics, we and others have developed transgenic zebrafish in which hepatocyte-specific expression of oncogenes such as &#946;-catenin3,4, Kras(V12)5,6, Myc7, or Yap18 leads to HCC in adult animals. In these zebrafish, liver enlargement is noted as early as 6 days post fertilization (dpf), providing a facile platform for testing the effects of drugs and genetic alterations on oncogene-driven liver overgrowth. Accurate and precise measurement of larval liver size is essential for determining the effects of these manipulations.
Liver size and shape can be assessed semi-quantitatively in fixed zebrafish larvae by CY3-SA labeling9 or in live zebrafish larvae using hepatocyte-specific fluorescent reporters and fluorescence dissecting microscopy5,6. The latter method is relatively quick, and its lack of precision can be addressed by photographing and measuring the area of each liver using image processing software7,10. However, it can be technically challenging to uniformly position all live larvae in an experiment such that two-dimensional liver area is an accurate representation of liver size. A similar technique for quantifying liver size involves using light sheet fluorescence microscopy to quantify larval liver volume8, which may be more accurate for detecting size differences when the liver is expanded non-uniformly in different dimensions. Fluorescence-activated cell sorting (FACS) can be used to count the number of fluorescently labeled hepatocytes and other liver cell types in larval livers8,11. In this method, larval livers are pooled and dissociated, so information about individual liver size and shape is lost. In combination with another liver size determination method, FACS enables differentiation between increased cell number (hyperplasia) and increased cell size (hypertrophy). All of these methods employ expensive fluorescence technology (microscope or cell sorter) and, except for CY3-SA labeling, require labeling of hepatocytes with a fluorescent reporter.
Here we describe in detail a method for quantifying zebrafish larval liver area using bright-field microscopy and image processing software3,12,13,14. This protocol enables precise quantification of the area of individual livers in situ without the use of fluorescence microscopy. While analyzing liver size, we blind the image identity to reduce investigator bias and improve scientific rigor15.

<table><tbody><tr><td>Camera for dissecting microscope</td><td>Leica, for example</td><td></td><td></td></tr><tr><td>Dissecting microscope</td><td>Leica, for example</td><td></td><td></td></tr><tr><td>Fine (Dumont #5) forceps</td><td>Fine Science Tools</td><td>11254-20</td><td></td></tr><tr><td>Glass pipets</td><td>VWR</td><td>14672-608</td><td></td></tr><tr><td>Image analysis software</td><td>Image J/FIJI</td><td></td><td>ImageJ/FIJI can be dowloaded for free: &lt;a href=&quot;https://imagej.net/Welcome&quot; target=&quot;_blank&quot;&gt;https://imagej.net/Welcome&lt;/a&gt;</td></tr><tr><td>Methyl cellulose</td><td>Sigma</td><td>M0387</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Sigma Aldrich</td><td>P6148</td><td></td></tr><tr><td>Phosphate-buffered saline</td><td>Various suppliers</td><td></td><td></td></tr><tr><td>Pipette pump</td><td>VWR</td><td>53502-233</td><td></td></tr><tr><td>Plastic Petri dishes</td><td>USA Scientific Inc</td><td>2906</td><td></td></tr><tr><td>Pyrex 9-well round-bottom glass dish</td><td>VWR</td><td>89090-482</td><td></td></tr><tr><td>Software for blinding files</td><td>R project</td><td></td><td>R can be downloaded for free: &lt;a href=&quot;https://www.r-project.org/&quot; target=&quot;_blank&quot;&gt;https://www.r-project.org/&lt;/a&gt;</td></tr><tr><td>Scientific graphing and statistics software</td><td>GraphPad Prism</td><td></td><td></td></tr><tr><td>Spreadsheet program</td><td>Microsoft Excel</td><td></td><td></td></tr><tr><td>Tricaine methanesulfonate (Tricaine-S)</td><td>Western Chemical</td><td>200-226</td><td></td></tr><tr><td>Very fine (Dumont #55) forceps</td><td>Fine Science Tools</td><td>11255-20</td><td></td></tr></tbody></table>

quantifying liver size in larval zebrafish using brightfield microscopy

In several transgenic zebrafish models of hepatocellular carcinoma (HCC), hepatomegaly can be observed during early larval stages. Quantifying larval liver size in zebrafish HCC models provides a means to rapidly assess the effects of drugs and other manipulations on an oncogene-related phenotype. Here we show how to fix zebrafish larvae, dissect the tissues surrounding the liver, photograph livers using bright-field microscopy, measure liver area, and analyze results. This protocol enables rapid, precise quantification of liver size. As this method involves measuring liver area, it may underestimate differences in liver volume, and complementary methodologies are required to differentiate between changes in cell size and changes in cell number. The dissection technique described herein is an excellent tool to visualize the liver, gut, and pancreas in their natural positions for myriad downstream applications including immunofluorescence staining and in situ hybridization. The described strategy for quantifying larval liver size is applicable to many aspects of liver development and regeneration.

Transgenic zebrafish expressing hepatocyte-specific activated &#946;-catenin (Tg(fabp10a:pt-&#946;-cat) zebrafish)3 and non-transgenic control siblings were euthanized at 6 dpf and liver area was quantified using brightfield microscopy and image processing software. Transgenic zebrafish have significantly increased liver size (0.0006 cm2) as compared to their non-transgenic siblings (0.0004 cm2, p &#60; 0.0001; Figure 1...

Watch this Scientific Journal Video about Quantifying Liver Size in Larval Zebrafish Using Brightfield Microscopy at JoVE.com

Quantifying Liver Size in Larval Zebrafish Using Brightfield Microscopy

Here we demonstrate a method for quantifying liver size in larval zebrafish, providing a way to assess the effects of genetic and pharmacologic manipulations on liver growth and development.

In several transgenic zebrafish models of hepatocellular carcinoma (HCC), hepatomegaly can be observed during early larval stages. Quantifying larval ...

quantifying-liver-size-larval-zebrafish-using-brightfield

Research

JoVE Journal

Developmental Biology

6.5K Views.  University of Utah School of Medicine. Here we demonstrate a method for quantifying liver size in larval zebrafish, providing a way to assess the effects of genetic and pharmacologic manipulations on liver growth and development.

Video: Quantifying Liver Size in Larval Zebrafish Using Brightfield Microscopy

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two layers of skin that enclose undifferentiated mesenchyme, and because the larval tail fin regenerates rapidly within 2-3 days. Using this system, we demonstrate a method for capturing the repair dynamics of the amputated tail fin with time-lapse video brightfield stereomicroscopy. We demonstrate that fin amputation triggers a contraction of the amputation wound and extrusion of cells around the wound margin, leading to their subsequent clearance. Fin regeneration proceeds from proximal to distal direction after a short delay. In addition, developmental growth of the larva can be observed during all stages. The presented method provides an opportunity for observing and analyzing whole tissue-scale behaviors such as fin development and growth in a simple microscope setting, which is easily adaptable to any stereomicroscope with time-lapse capabilities.

We present a protocol for capturing the dynamics of zebrafish larval tail fin regeneration on a whole-tissue scale using brightfield-based stereomicroscopy. This technique enables capturing the regeneration dynamics with single cell resolution. This methodology can be adapted to any stereomicroscope equipped with a CCD camera and time-lapse software.

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two ...

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM compared to confocal systems are its low phototoxicity, gentle mounting strategies, fast acquisition with high signal to noise ratio and the possibility of imaging samples from various angles (views) for long periods of time. Imaging from multiple views unleashes the full potential of LSFM, but at the same time it can create terabyte-sized datasets. Processing such datasets is the biggest challenge of using LSFM. In this protocol we outline some solutions to this problem. Until recently, LSFM was mostly performed in laboratories that had the expertise to build and operate their own light sheet microscopes. However, in the last three years several commercial implementations of LSFM became available, which are multipurpose and easy to use for any developmental biologist. This article is primarily directed to those researchers, who are not LSFM technology developers, but want to employ LSFM as a tool to answer specific developmental biology questions. 
Here, we use imaging of zebrafish eye development as an example to introduce the reader to LSFM technology and we demonstrate applications of LSFM across multiple spatial and temporal scales. This article describes a complete experimental protocol starting with the mounting of zebrafish embryos for LSFM. We then outline the options for imaging using the commercially available light sheet microscope. Importantly, we also explain a pipeline for subsequent registration and fusion of multiview datasets using an open source solution implemented as a Fiji plugin. While this protocol focuses on imaging the developing zebrafish eye and processing data from a particular imaging setup, most of the insights and troubleshooting suggestions presented here are of general use and the protocol can be adapted to a variety of light sheet microscopy experiments.

Light sheet fluorescence microscopy is an excellent tool for imaging embryonic development. It allows recording of long time-lapse movies of live embryos in near physiological conditions. We demonstrate its application for imaging zebrafish eye development across wide spatio-temporal scales and present a pipeline for fusion and deconvolution of multiview datasets.

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM ...

Generation of Zebrafish Larval Xenografts and Tumor Behavior Analysis

Zebrafish larval xenografts are being widely used for cancer research to perform in vivo and real-time studies of human cancer. The possibility of rapidly visualizing the response to anti-cancer therapies (chemo, radiotherapy, and biologicals), angiogenesis and metastasis with single cell resolution, places the zebrafish xenograft model as a top choice to develop preclinical studies.

The zebrafish larval xenograft assay presents several experimental advantages compared to other models, but probably the most striking is the reduction of size scale and consequently time. This reduction of scale allows single cell imaging, the use of a relatively low number of human cells (compatible with biopsies), medium-high-throughput drug screenings, but most importantly enables a significant reduction of the time of the assay. All these advantages make the zebrafish xenograft assay extremely attractive for future personalized medicine applications.

Many zebrafish xenograft protocols have been developed with a wide diversity of human tumors; however, a general and standardized protocol to efficiently generate zebrafish larval xenografts is still lacking. Here we provide a step-by-step protocol, with tips to generate xenografts and guidelines for tumor behavior analysis, whole-mount immunofluorescence, and confocal imaging quantification.

Here, we provide a step-by-step protocol, with tips to generate xenografts and guidelines for tumor behavior analysis, whole-mount immunofluorescence, and confocal imaging quantification.

Zebrafish larval xenografts are being widely used for cancer research to perform in vivo and real-time studies of human cancer. The possibility of rapidly ...

Cancer Research

Analysis of Liver Microenvironment During Early Progression of Non-Alcoholic Fatty Liver Disease-Associated Hepatocellular Carcinoma in Zebrafish

Liver cancer is currently the third&#160;leading cause of cancer related death worldwide, and Hepatocellular Carcinoma (HCC) accounts for 75-90% of all liver cancer cases. With the introduction of effective treatments to prevent and treat hepatitis B/C, non-alcoholic fatty liver disease (NAFLD), and the more aggressive form know as non-alcoholic steatohepatitis (NASH), are quickly becoming the number one risk factors to develop HCC in modern societies. To better understand the role NASH has on the development of HCC we designed a NASH-associated HCC zebrafish. The optical clarity and genetic tractability of the zebrafish larvae make them an appealing and powerful model to study the liver microenvironment and immune cell composition using non-invasive fluorescent live imaging. This protocol describes how to use a NASH-associated HCC zebrafish model to investigate the effect of cholesterol surplus in the liver microenvironment and its impact on immune cell composition at early stages of the disease. First, we feed HCC larvae (s704Tg), which express hepatocyte-specific activated beta-catenin, with a 10% high cholesterol diet for 8 days to develop a NASH-associated HCC model. Here we describe how to make use of different transgenic lines to evaluate several early malignancy features in the liver by non-invasive confocal microscopy, such as liver area, cell, and nuclear morphology (hepatocytes area, nuclear area, nuclear:cytoplasmic ratio (N:C ratio), nuclear circularity, micronuclei/nuclear herniation scoring) and angiogenesis. Then, using transgenic lines with tagged immune cells (neutrophils, macrophages, and T cells) we show how to analyze liver immune cell composition in NASH-associated HCC larvae. The described techniques are useful to evaluate liver microenvironment and immune cell composition at early hepatocarcinogenesis stages, but they can also be modified to study such features in other liver disease models.

Here, we present how to generate a non-alcoholic fatty liver disease (NAFLD)-associated Hepatocellular Carcinoma (HCC) zebrafish model to study the impact of cholesterol surplus on liver microenvironment and immune cell landscape.

Liver cancer is currently the third&#160;leading cause of cancer related death worldwide, and Hepatocellular Carcinoma (HCC) accounts for 75-90% of all ...

Automated High-throughput Behavioral Analyses in Zebrafish Larvae

We have created a novel high-throughput imaging system for the analysis of behavior in 7-day-old zebrafish larvae in multi-lane plates. This system measures spontaneous behaviors and the response to an aversive stimulus, which is shown to the larvae via a PowerPoint presentation. The recorded images are analyzed with an ImageJ macro, which automatically splits the color channels, subtracts the background, and applies a threshold to identify individual larvae placement in the lanes. We can then import the coordinates into an Excel sheet to quantify swim speed, preference for edge or side of the lane, resting behavior, thigmotaxis, distance between larvae, and avoidance behavior. Subtle changes in behavior are easily detected using our system, making it useful for behavioral analyses after exposure to environmental toxicants or pharmaceuticals.

Our laboratory developed a novel high-throughput automated imaging system that is useful for the detection of several different behaviors in 7-day-old zebrafish larvae. The system can be used for detecting subtle changes in behavior after the larvae have been exposed to environmental toxicants or pharmaceuticals.

We have created a novel high-throughput imaging system for the analysis of behavior in 7-day-old zebrafish larvae in multi-lane plates. This system ...

Neuroscience

Real Time and Repeated Measurement of Skeletal Muscle Growth in Individual Live Zebrafish Subjected to Altered Electrical Activity

A number of methods can be used to visualize individual cells throughout the body of live embryonic, larval or juvenile zebrafish. We show that live fish with fluorescently-marked plasma membranes can be scanned in a confocal laser scanning microscope in order to determine the volume of muscle tissue and the number of muscle fibers present. Efficient approaches for the measurement of cell number and size in live animals over time are described and validated against more arduous segmentation methods. Methods are described that permit the control of muscle electrical, and thus contractile, activity. Loss of skeletal muscle contractile activity greatly reduced muscle growth. In larvae, a protocol is described that allows reintroduction of patterned electrical-evoked contractile activity. The described methods minimize the effect of inter-individual variability and will permit analysis of the effect of electrical, genetic, drug, or environmental stimuli on a variety of cellular and physiological growth parameters in the context of the living organism. Long-term follow-up of the measured effects of a defined early-life intervention on individuals can subsequently be performed.

Optical clarity is a major advantage for cell biological and physiological work in zebrafish. Robust methods for measurement of cell growth in individual animals are described that permit novel insights into how growth of skeletal muscle and neighboring tissues are integrated with whole body growth.

A number of methods can be used to visualize individual cells throughout the body of live embryonic, larval or juvenile zebrafish. We show that live fish ...

Microgavage of Zebrafish Larvae

The zebrafish has emerged as a powerful model organism for studying intestinal development1-5, physiology6-11, disease12-16, and host-microbe interactions17-25. Experimental approaches for studying intestinal biology often require the in vivo introduction of selected materials into the lumen of the intestine. In the larval zebrafish model, this is typically accomplished by immersing fish in a solution of the selected material, or by injection through the abdominal wall. Using the immersion method, it is difficult to accurately monitor or control the route or timing of material delivery to the intestine. For this reason, immersion exposure can cause unintended toxicity and other effects on extraintestinal tissues, limiting the potential range of material amounts that can be delivered into the intestine. Also, the amount of material ingested during immersion exposure can vary significantly between individual larvae26. Although these problems are not encountered during direct injection through the abdominal wall, proper injection is difficult and causes tissue damage which could influence experimental results. We introduce a method for microgavage of zebrafish larvae. The goal of this method is to provide a safe, effective, and consistent way to deliver material directly to the lumen of the anterior intestine in larval zebrafish with controlled timing. Microgavage utilizes standard embryo microinjection and stereomicroscopy equipment common to most laboratories that perform zebrafish research. Once fish are properly positioned in methylcellulose, gavage can be performed quickly at a rate of approximately 7-10 fish/ min, and post-gavage survival approaches 100% depending on the gavaged material. We also show that microgavage can permit loading of the intestinal lumen with high concentrations of materials that are lethal to fish when exposed by immersion. To demonstrate the utility of this method, we present a fluorescent dextran microgavage assay that can be used to quantify transit from the intestinal lumen to extraintestinal spaces. This test can be used to verify proper execution of the microgavage procedure, and also provides a novel zebrafish assay to examine intestinal epithelial barrier integrity under different experimental conditions (e.g. genetic manipulation, drug treatment, or exposure to environmental factors). Furthermore, we show how gavage can be used to evaluate intestinal motility by gavaging fluorescent microspheres and monitoring their subsequent transit. Microgavage can be applied to deliver diverse materials such as live microorganisms, secreted microbial factors/toxins, pharmacological agents, and physiological probes. With these capabilities, the larval zebrafish microgavage method has the potential to enhance a broad range of research fields using the zebrafish model system.

We present a novel method for microgavage of larval zebrafish utilizing standard embryo microinjection and stereomicroscopy equipment. We demonstrate that microgavage is a safe and efficient technique useful for delivering controlled amounts of diverse materials specifically into the larval zebrafish intestinal lumen.

The zebrafish has emerged as a powerful model organism for studying intestinal development1-5, physiology6-11, disease12-16, and host-microbe ...

Biology

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 ...

Bioengineering

In Vivo Whole-Brain Imaging of Zebrafish Larvae

In Vivo Whole-Brain Imaging of Zebrafish Larvae Using Three-Dimensional Fluorescence Microscopy

As a vertebrate model animal, larval zebrafish are widely used in neuroscience and provide a unique opportunity to monitor whole-brain activity at the cellular resolution. Here, we provide an optimized protocol for performing whole-brain imaging of larval zebrafish using three-dimensional fluorescence microscopy, including sample preparation and immobilization, sample embedding, image acquisition, and visualization after imaging. The current protocol enables in vivo imaging of the structure and neuronal activity of a larval zebrafish brain at a cellular resolution for over 1 h using confocal microscopy and custom-designed fluorescence microscopy. The critical steps in the protocol are also discussed, including sample mounting and positioning, preventing bubble formation and dust in the agarose gel, and avoiding motion in images caused by incomplete solidification of the agarose gel and paralyzation of the fish. The protocol has been validated and confirmed in multiple settings. This protocol can be easily adapted for imaging other organs of a larval zebrafish.

Author Spotlight: In Vivo Whole-Brain Imaging of Zebrafish Larvae Using Three-Dimensional Fluorescence Microscopy  

Presented here is a protocol for in vivo whole-brain imaging of larval zebrafish using three-dimensional fluorescence microscopy. The experimental procedure includes sample preparation, image acquisition, and visualization.

As a vertebrate model animal, larval zebrafish are widely used in neuroscience and provide a unique opportunity to monitor whole-brain activity at the ...

High-fat Feeding Paradigm for Larval Zebrafish: Feeding, Live Imaging, and Quantification of Food Intake

Zebrafish are emerging as a model of dietary lipid processing and metabolic disease. This protocol describes how to feed larval zebrafish a lipid-rich meal, which consists of an emulsion of chicken egg yolk liposomes created by sonicating egg yolk in embryo media. Detailed instructions are provided to screen larvae for egg yolk consumption so that larvae that fail to feed will not confound experimental results. The chicken egg yolk liposomes can be spiked with fluorescent lipid analogs, including fatty acids and cholesterol, enabling both systemic and subcellular visualization of dietary lipid processing. Several methods are described to mount larvae that are conducive to short- and long-term live imaging with both upright and inverted objectives at high and low magnification. Additionally presented is an assay to quantify larval food intake by extracting the lipids of larvae fed fluorescent lipid analogs, spotting the lipids on a thin layer chromatography plate, and quantifying the fluorescence. Finally, critical aspects of the procedures, important controls, options for modifying the protocols to address specific experimental questions, and potential limitations are discussed. These techniques can be applied not only to focused, hypothesis driven inquiries, but also to a variety of screens and live imaging techniques to study dietary lipid metabolism and the control of food intake.

Zebrafish are emerging as a valuable model of dietary lipid processing and metabolic disease. Described are protocols of lipid-rich larval feeds, live imaging of dietary fluorescent lipid analogs, and quantification of food intake. These techniques can be applied to a variety of screening, imaging, and hypothesis driven inquiry techniques.

Zebrafish are emerging as a model of dietary lipid processing and metabolic disease. This protocol describes how to feed larval zebrafish a lipid-rich ...