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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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: <a href="https://imagej.net/Welcome" target="_blank">https://imagej.net/Welcome</a></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: <a href="https://www.r-project.org/" target="_blank">https://www.r-project.org/</a></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

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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 ...

Hepatocyte-specific Ablation in Zebrafish to Study Biliary-driven Liver Regeneration

The liver has a great capacity to regenerate. Hepatocytes, the parenchymal cells of the liver, can regenerate in one of two ways: hepatocyte- or biliary-driven liver regeneration. In hepatocyte-driven liver regeneration, regenerating hepatocytes are derived from preexisting hepatocytes, whereas, in biliary-driven regeneration, regenerating hepatocytes are derived from biliary epithelial cells (BECs). For hepatocyte-driven liver regeneration, there are excellent rodent models that have significantly contributed to the current understanding of liver regeneration. However, no such rodent model exists for biliary-driven liver regeneration. We recently reported on a zebrafish liver injury model in which BECs extensively give rise to hepatocytes upon severe hepatocyte loss. In this model, hepatocytes are specifically ablated by a pharmacogenetic means. Here we present in detail the methods to ablate hepatocytes and to analyze the BEC-driven liver regeneration process. This hepatocyte-specific ablation model can be further used to discover the underlying molecular and cellular mechanisms of biliary-driven liver regeneration. Moreover, these methods can be applied to chemical screens to identify small molecules that augment or suppress liver regeneration.

To help assess the molecular mechanisms underlying zebrafish biliary-driven liver regeneration, we established a liver injury model in which the nitroreductase-expressing hepatocytes are genetically ablated upon metronidazole treatment. In this protocol, we describe how to adeptly manipulate, monitor and analyze hepatocyte ablation and biliary-driven liver regeneration.

The liver has a great capacity to regenerate. Hepatocytes, the parenchymal cells of the liver, can regenerate in one of two ways: hepatocyte- or ...

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method described here allows time-lapse analysis of normal and impaired kidney development in zebrafish embryos by using a fluorescence dissecting microscope equipped for structured illumination and z-stack acquisition. To visualize nephrogenesis, transgenic zebrafish (Tg(wt1b:GFP)) with fluorescently labeled kidney structures were used. Renal defects were triggered by injection of an antisense morpholino oligonucleotide against the Wilms tumor gene wt1a, a factor known to be crucial for kidney development.
The advantage of the experimental setup is the combination of a zoom microscope with simple strategies for re-adjusting movements in x, y or z direction without additional equipment. To circumvent focal drift that is induced by temperature variations and mechanical vibrations, an autofocus strategy was applied instead of utilizing a usually required environmental chamber. In order to re-adjust the positional changes due to a xy-drift, imaging chambers with imprinted relocation grids were employed.
In comparison to more complex setups for time-lapse recording with optical sectioning such as confocal laser scanning&#160;or light sheet microscopes, a zoom microscope is easy to handle. Besides, it offers dissecting microscope-specific benefits such as high depth of field and an extended working distance.
The method to study organogenesis presented here can also be used with fluorescence stereo microscopes not capable of optical sectioning. Although limited for high-throughput, this technique offers an alternative to more complex equipment that is normally used for time-lapse recording of developing tissues and organ dynamics.

The method described here allows time-lapse analysis of organ development in zebrafish embryos by using a fluorescence dissecting microscope capable of performing optical sectioning and simple strategies of readjustment to correct focal and planar drift.

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method ...

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 ...

Dissection of Larval Zebrafish Gonadal Tissue

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex determination system, where several genes distributed throughout the genome collectively determine the sex identities of individual fish. Currently, the genes involved in regulating gonad development and how they work remain elusive. Normally, isolating gonadal tissue is the first step to examine the sex developmental processes. Here, we present a procedure to isolate gonadal tissue from 17 dpf (days post fertilization) and 25 dpf zebrafish larvae. The isolated gonadal tissue may be subsequently examined by morphology and gene expression profiling.

Here, we present a protocol for isolating gonadal tissue of larval zebrafish, which will facilitate investigations of zebrafish sex differentiation and maintenance.

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex ...

Histological Analyses of Acute Alcoholic Liver Injury in Zebrafish

Alcoholic Liver Disease (ALD) refers to damage to the liver due to acute or chronic alcohol abuse. It is among the leading causes of alcohol-related morbidity and mortality and affects more than 2 million people in the United States. A better understanding of the cellular and molecular mechanisms underlying alcohol-induced liver injury is crucial for developing effective treatment for ALD. Zebrafish larvae exhibit hepatic steatosis and fibrogenesis after just 24 h of exposure to 2% ethanol, making them useful for the study of acute alcoholic liver injury. This work describes the procedure for acute ethanol treatment in zebrafish larvae and shows that it causes steatosis and swelling of the hepatic blood vessels. A detailed protocol for Hematoxylin and Eosin (H&#38;E) staining that is optimized for the histological analysis of the zebrafish larval liver, is also described. H&#38;E staining has several unique advantages over immunofluorescence, as it marks all liver cells and extracellular components simultaneously and can readily detect hepatic injury, such as steatosis and fibrosis. Given the increasing usage of zebrafish in modeling toxin and virus-induced liver injury, as well as inherited liver diseases, this protocol serves as a reference for the histological analyses performed in all these studies.

This protocol describes histological analyses of the livers from zebrafish larvae that have been treated with 2% ethanol for 24 h. Such acute ethanol treatment results in hepatic steatosis and swelling of the hepatic vasculature.

Alcoholic Liver Disease (ALD) refers to damage to the liver due to acute or chronic alcohol abuse. It is among the leading causes of alcohol-related ...

Correlative Super-resolution and Electron Microscopy to Resolve Protein Localization in Zebrafish Retina

We present a method to investigate the subcellular protein localization in the larval zebrafish retina by combining super-resolution light microscopy and scanning electron microscopy. The sub-diffraction limit resolution capabilities of super-resolution light microscopes allow improving the accuracy of the correlated data. Briefly, 110 nanometer thick cryo-sections are transferred to a silicon wafer and, after immunofluorescence staining, are imaged by super-resolution light microscopy. Subsequently, the sections are preserved in methylcellulose and platinum shadowed prior to imaging in a scanning electron microscope (SEM). The images from these two microscopy modalities are easily merged using tissue landmarks with open source software. Here we describe the adapted method for the larval zebrafish retina. However, this method is also applicable to other types of tissues and organisms. We demonstrate that the complementary information obtained by this correlation is able to resolve the expression of mitochondrial proteins in relation with the membranes and cristae of mitochondria as well as to other compartments of the cell.

This protocol describes the necessary steps to obtain subcellular protein localization results on zebrafish retina by correlating super-resolution light microscopy and scanning electron microscopy images.

We present a method to investigate the subcellular protein localization in the larval zebrafish retina by combining super-resolution light microscopy and ...

Establishment of Larval Zebrafish as an Animal Model to Investigate Trypanosoma cruzi Motility In Vivo

Chagas disease is a parasitic infection caused by Trypanosoma cruzi, whose motility is not only important for localization, but also for cellular binding and invasion. Current animal models for the study of T. cruzi allow limited observation of parasites in vivo, representing a challenge for understanding parasite behavior during the initial stages of infection in humans. This protozoan has a flagellar stage in both vector and mammalian hosts, but there are no studies describing its motility in vivo.The objective of this project was to establish a live vertebrate zebrafish model to evaluate T. cruzi motility in the vascular system. Transparent zebrafish larvae were injected with fluorescently labeled trypomastigotes and observed using light sheet fluorescence microscopy (LSFM), a noninvasive method to visualize live organisms with high optical resolution. The parasites could be visualized for extended periods of time due to this technique&#39;s relatively low risk of photodamage compared to confocal or epifluorescence microscopy. T. cruzi parasites were observed traveling in the circulatory system of live zebrafish in different-sized blood vessels and the yolk. They could also be seen attached to the yolk sac wall and to the atrioventricular valve despite the strong forces associated with heart contractions. LSFM of T. cruzi-inoculated zebrafish larvae is a valuable method that can be used to visualize circulating parasites and evaluate their tropism, migration patterns, and motility in the dynamic environment of the cardiovascular system of a live animal.

Establishment of Larval Zebrafish as an Animal Model to Investigate Trypanosoma cruzi Motility In Vivo

In this protocol, fluorescently labeled T. cruzi&#160;were injected into transparent zebrafish larvae, and parasite motility was observed in vivo using light sheet fluorescence microscopy.

Chagas disease is a parasitic infection caused by Trypanosoma cruzi, whose motility is not only important for localization, but also for cellular binding ...

Visualizing the Node and Notochordal Plate In Gastrulating Mouse Embryos Using Scanning Electron Microscopy and Whole Mount Immunofluorescence

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is the formation of the transient organizers, the node and notochordal plate, from cells that have passed through the primitive streak. The proper formation of these signaling centers is essential for the development of the body plan and techniques to visualize them are of high interest to mouse developmental biologists. The node and notochordal plate lie on the ventral surface of gastrulating mouse embryos around embryonic day (E) 7.5 of development. The node is a cup-shaped structure whose cells possess a single slender cilium each. The proper subcellular localization and rotation of the cilia in the node pit determines left-right asymmetry. The notochordal plate cells also possess single cilia albeit shorter than those of the node cells. The notochordal plate forms the notochord which acts as an important signaling organizer for somitogenesis and neural patterning. Because the cells of the node and notochordal plate are transiently present on the surface and possess cilia, they can be visualized using scanning electron microscopy (SEM). Among other techniques used to visualize these structures at the cellular level is whole mount immunofluorescence (WMIF) using the antibodies against the proteins that are highly expressed in the node and notochordal plate. In this report, we describe our optimized protocols to perform SEM and WMIF of the node and notochordal plate in developing mouse embryos to help in the assessment of tissue shape and cellular organization in wild-type and gastrulation mutant embryos.

The node and notochordal plate are transient signaling organizers in developing mouse embryos that can be visualized using several techniques. Here, we describe in detail how to perform two of the techniques to study their structure and morphogenesis: 1) scanning electron microscopy (SEM); and 2) whole mount immunofluorescence (WMIF).

The post-implantation mouse embryo undergoes major shape changes after the initiation of gastrulation and morphogenesis. A hallmark of morphogenesis is ...

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

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

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

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