The authors thank Meng-Chieh Shen for images, Jennifer Anderson for providing helpful comments on the manuscript, and members of the Farber laboratory for their contributions in developing these techniques. This study was funded by NIDDK-NIH award RO1DK093399 (S.A.F.), RO1GM63904 (The Zebrafish Functional Genomics Consortium: PI Stephen Ekker and Co-PI S.A.F), and F32DK096786 (J.P.O.). This content is solely the responsibility of the authors and does not necessarily represent the official views of NIH. Additional support was provided by the G. Harold and Leila Y. Mathers Charitable Foundation to the laboratory of S.A.F and the Carnegie Institution for Science endowment.

These protocols have been approved by the Carnegie Institution for Science Institutional Animal Care and Use Committee (protocol no. 139).
1. Animal Preparation
<ol>
	<li>Maintain adults and larvae at 28 &#176;C on a 14 hr:10 hr light:dark cycle. Feed adults twice daily with shell free Artemia (decapsulated, non-hatching, starting at 14 dpf) and commercial micropellets.</li>
	<li>These protocols are optimized for the use of 6-7 dpf larvae collected by natural spawning of the AB background. Protocols can be modified for larvae of other ages and backgrounds. Do not provide exogenous food prior to 6 dpf.</li>
	<li>Anesthetize larvae with Tricaine in embryo media (EM) (4.2 ml of 4 mg/ml Tricaine per 100 ml EM) at room temperature (RT).</li>
</ol>
2. Preparation of Lipid-rich Egg Yolk Feed
<ol>
	<li>Prepare and store the chicken egg yolks. Separate the yolk and white of 12 chicken eggs. Pool the yolks, aliquot 1 ml into 1.5 ml tubes, and store at -80 &#176;C up to 1 yr (12 yolks will make ~80 aliquots). Pool the whites, store, and use as a lipid-poor, protein-rich feed.</li>
	<li>Prepare fluorescent lipid analog(s) to be added to the egg yolk feed if desired.
	<ol>
		<li>Suspend the purchased, powdered fluorescent lipid analog in 100% ethanol or 100% chloroform at 0.1 &#181;g/&#181;l (see following note discussing solvents), aliquot in opaque glass tubes, seal with parafilm, and store according to manufacturer&#39;s instructions. Due to rapid evaporation of organic solvents do not use aliquot volumes less than ~400 &#181;l for long-term storage. 
		Note: If the lipid analog is suspended in ethanol, large volumes of the lipid analog will be used because it dries faster under N2 than chloroform. If the lipid analog is suspended in ethanol it does not have to be dried and resuspended before adding it to the liposome feed in step 2.2.4, but the total ethanol concentration should not exceed 0.1% in the liposome feed to avoid potentially confounding physiological effects of ethanol.</li>
		<li>For fluorescent fatty acid analogs: Prepare a total volume of 20 ml 5% egg yolk emulsion in EM. From that prepare 5 ml aliquots with a final concentration of 6.4 &#181;M 4,4-difluoro-4-bora-3a,4a-diaza-s-indacene (BODIPY C16) for live confocal imaging and food intake assays or 1 &#181;g/ml BODIPY C12 for stereomicroscope imaging. Transfer the desired amount of the fluorescent lipid analog into a 1.5 ml plastic tube. Dry the lipid under a stream of N2, taking care not to blow the lipid out of the tube.</li>
		<li>Immediately, resuspend in 5 &#181;l 100% ethanol by pipetting a stream down the sides of the tube, add 95 &#181;l EM, and mix well. 
		Note: The mixture should appear homogenous; if colored lipid remains on the sides of the tube or the liquid appears clumpy, the lipid has not fully solubilized and requires more ethanol. Protect the tube from light and store on ice.</li>
		<li>For fluorescent cholesterol analog: prepare a final concentration of 2.4 &#181;g/ml cholesterol in 5 ml 0.5-5% egg yolk emulsion for live imaging studies. Transfer the desired amount of cholesterol into a 1.5 ml plastic tube. Dry the lipid under a stream of N2, taking care not to blow the lipid out of the tube.</li>
		<li>Immediately, resuspend in 15 &#181;l of RT 100% ethanol by pipetting a stream down the sides of the tube and mix with 85 &#181;l of RT 1% fatty acid-free BSA in purified H2O. Protect the tube from light and store on ice.</li>
	</ol>
	</li>
	<li>Prepare Egg Yolk Liposome Feed.
	<ol>
		<li>Add a thawed aliquot of egg yolk to EM in a 50 ml conical tube. Vortex the diluted egg yolk mixture for 1-2 min; the mixture should appear to be a homogenous emulsion.</li>
		<li>Pour the mixture through a fine mesh strainer to remove protein conglomerates and collect in a new, fresh 50 ml conical tube.</li>
		<li>Pulse sonicate the egg mixture with a one-fourth inch tapered microtip (5 times (5 x 1 sec on, 1 sec off) pausing 5-10 sec between each set; output intensity: 6 W) at RT to create liposomes. Sonicator power settings are instrument dependent and therefore require optimization for each instrument and microtip. 
		Note: Continuously rock liposome mixture at RT to maintain homogeneity until use.</li>
	</ol>
	</li>
	<li>Add the fluorescent lipid analog to the egg yolk liposomes, if desired. Immediately after sonication, pipet the prepared lipids into the liposome mixture and vortex at high speed for 30 sec to incorporate into liposomes. Protect the mixture from light by wrapping the tube in foil and continue rocking at RT.</li>
</ol>
3. Feeding Paradigm
<ol>
	<li>Prepare the Larvae for Feeding.
	<ol>
		<li>Prior to beginning the feed, screen larvae for obvious morphological defects that may impede feeding. Transfer larvae to 60 mm x 15 mm Petri dishes or 6- or 12-well plates, reduce EM to a minimum, and replace with 5 ml of the prepared liposome solution.</li>
		<li>If required, transfer unfed control larvae to 5 ml EM and/or 5 ml 5% egg white in EM and treat in parallel.</li>
	</ol>
	</li>
	<li>Gently rock the larvae in an incubated rocker (29-31 &#176;C at 30 rpm) throughout the feed to encourage food intake while continuing to shield from light if fluorescent lipid analogs are present. If larvae need to be exposed to light, minimize the exposure with a tinted glass incubator cover.</li>
	<li>At the end of the feeding period, rinse the freely swimming larvae in EM 3 times. 
	Note: Larvae may begin consuming liposomes at any point during the feed. If it is critical that larvae commence feeding at the same time, screen for food intake (see 3.4) after 1 hr of feeding and return only larvae that have commenced feeding to the egg yolk emulsion to complete the feed.</li>
	<li>Screen the larvae at this point for food intake as a small number may not eat (generally ~0-5%). Examine the intestines of larvae that have been anesthetized or slightly cooled on a metal block on ice to determine if they have fed: larvae that have consumed liposomes will have a darkened intestine (Figure 1).</li>
	<li>Image or process larvae for food intake immediately. Alternately, place larvae in fresh EM and incubate at 28 &#176;C to allow metabolic processing to occur.</li>
</ol>
4. Mounting Larvae for Live Imaging
<ol>
	<li>Prepare the Mounting Media.
	<ol>
		<li>For 3% methyl cellulose: add 0.75 g methyl cellulose to 25 ml of near boiling EM, vortex, rock at RT 2-3 days until completely dissolved, and store at 4 &#176;C for years at -20 &#176;C. Repeated cycles of freezing and thawing at 4 &#176;C will remove air bubbles. Store 3% methyl cellulose on ice while mounting larvae.</li>
		<li>For 1.2% low-mount agarose: add 0.3 g of low-melt agarose to 25 mL EM and dissolve by bringing to a boil, aliquot into 2 ml tubes, and maintain at 26-28 &#176;C on a heat block. Unused aliquots can be stored at RT or -20 &#176;C and boiled again at the time of use. Be sure that agarose is cool enough to handle before exposing to larvae or they may be over-heated.</li>
	</ol>
	</li>
	<li>For low-magnification, medium-throughput live imaging, place a glass slide on a tissue on a metal block on ice and wait for the slide to cool. Apply a generous amount of 3% methyl cellulose (which has been maintained at 4 &#176;C on ice) onto the slide, transfer a larva onto the 3% methyl cellulose, using a fishing-line poker (0.41 mm diameter fishing line glued to the end of a glass capillary tube) create a troth to drain excess EM (so that methyl cellulose will not be diluted) and position the larvae as desired.</li>
	<li>For short-term (&#60;20 min), high-magnification live imaging with an upright oil emersion objective, mount larvae with the coverslip lean-to method.
	<ol>
		<li>Use cyanoacrylate based glue (Superglue) to attach a a 22 mm x 30 mm coverslip to one end of a glass slide. Use a poker to put a thin line of 3% methyl cellulose on the slide adjacent to the coverslip and transfer larvae to the slide with a wide bore Pasteur pipet. Remove excess EM so it will not dilute the methyl cellulose.</li>
		<li>Using a poker, gently position larvae in the methyl cellulose on their sides (left side up to image more of the liver, right side up to image the gallbladder) with their heads next to the coverslip.</li>
		<li>Spot superglue in the corners of a second coverslip and gently place one edge of the coverslip on the mounted coverslip and the other on the slide, bridging the larvae. Take care not to apply excess pressure during this step or with the objective while imaging so that larvae remain uninjured.</li>
	</ol>
	</li>
	<li>For long-term, high-magnification imaging with an upright immersion objective, mount larvae in agarose. Add a larva to a small aliquot of 1.2% low melt agarose then transfer the larva in a small volume of agarose to a petri dish.
	<ol>
		<li>Quickly position the larvae with a poker before the agarose solidifies. Allow the agarose to dry for 2-3 min and cover the agarose fully with fresh EM to prevent it from drying.</li>
	</ol>
	</li>
	<li>For long-term, high-magnification imaging with an inverted objective, mount larvae on a glass-bottom dish in agarose.
	<ol>
		<li>Cool a metal block on ice. Place the dish on the metal block separated by a tissue to prevent excessive cooling and larval freezing. Transfer a larva to the dish and remove the EM by wicking with a tissue.</li>
		<li>Place a drop of 28 &#176;C low-melt agarose (1.2%) on top of the larva and immediately position with a poker (left side down to best image the liver, right side down to image the gallbladder). Remove dish from the cold block, allow to dry for 2-3 min, and add a sufficient volume of fresh EM to cover the agarose (~2-5 ml).</li>
	</ol>
	</li>
</ol>
5. Food Intake Assay
<ol>
	<li>At the end of an egg yolk liposome feed containing 6.4 &#181;M BODIPY C16, pool a minimum of 10 washed larvae in a plastic 1.5 or 2 ml tube, remove EM, and snap freeze. The samples may be stored at -80 &#176;C protected from light for several months. 
	Note: If possible, collect multiple replicates. It is important to collect unfed larvae at this step to normalize for background lipid fluorescence (ideally larvae from the same clutch are used).</li>
	<li>Perform a modified Bligh-Dyer lipid extraction3,21.
	<ol>
		<li>Add 100 &#181;l homogenization buffer (20 mM Tris-Cl, 1 mM EDTA) to the sample on ice (alternatively purified H2O can be used). Homogenize on ice with a microtip sonicator (5 sec total: 1 sec on 1 sec off). Check that larvae are totally homogenized; if not, repeat. Transfer to a 13 ml disposable glass culture tube (other glass tubes may be used). 
		Note: Chloroform is hazardous; perform all subsequent lipid extraction steps at RT in a chemical hood.</li>
		<li>For each 100 &#181;l of homogenization buffer used, add 375 &#181;l of 1:2 (chloroform:methanol). Vortex 30-60 sec, incubate 10 min, add 125 &#181;l of chloroform per 100 &#181;l homogenization buffer used, and vortex 30 sec.</li>
		<li>Add 125 &#181;l of 200 mM Tris pH 7.5 per 100 &#181;l homogenization buffer used, vortex 30 sec, and centrifuge (2,000 x g, 5 min, RT, protected from light). Carefully remove the samples from the centrifuge. They will be separated into two phases: an upper phase is aqueous, an interface of larval debris, and a lower organic phase (contains lipids).</li>
		<li>With a clean glass pipette collect the bottom, organic phase and transfer it into a clean 13 ml glass tube. 
		Note: Take care to avoid the aqueous phase and larval debris at the interface; if contamination occurs, repeat centrifugation step and recollect. Discard the upper, aqueous phase; it may be helpful to remove and discard this phase before collecting the organic phase. The sample can be stored at -80 &#176;C for up to 1 month.</li>
		<li>Dry the organic phase under vacuum (0.12 atm) while protecting from light. Do not continue to dry the samples after the liquid has evaporated as it will reduce lipid solubility. Resuspend the lipids in 2:1 (chloroform:methanol) and store on ice. The optimal volume of the chloroform:methanol solution depends on the detection method used; start low and continue to dilute as needed. A general guideline is to use 10 &#181;l for 10 pooled larvae.</li>
	</ol>
	</li>
	<li>Spot the entire sample volumes in evenly spaced intervals onto a channeled TLC plate and scan the plate with a laser scanner routinely used for biomolecular imaging (e.g., a fluorescent plate reader). For the fluorescent lipid analogs included in the materials list that have an excitation maxima from 500-650 nm and emission maxima from 510-665 nm, use a 488 nm laser and with emission collection at 520 nm.
	<ol>
		<li>Quantify the total fluorescence of each spot with imaging software. 
		Note: Correct for naturally occurring background lipid fluorescence by subtracting the fluorescence of paired, unfed larval samples.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The techniques described here allow researchers to treat larval zebrafish with a lipid-rich feed, visualize dietary lipid processing in live larvae, and quantify larval food intake. To ensure success, special attention should be given to several critical steps. Commercial chicken eggs vary; to minimize potential variability we perform all assays on organic eggs from cage-free chickens that have not been enriched for omega-3 fatty acids. Lower feeding rates may be observed in fish younger than 6 dpf with remaining endogen...

The mechanisms by which the intestine regulates dietary lipid processing, the liver controls complex lipid synthesis and lipoprotein metabolism, and how these organs work with the central nervous system to control food intake are incompletely understood. It is of biomedical interest to elucidate this biology in light of the current epidemics of obesity, cardiovascular disease, diabetes, and non-alcoholic fatty liver disease. Studies in cell culture and mice have provided the majority of our understanding of the mechanistic relationships between dietary lipids and disease, and zebrafish (Danio rerio) are emerging as an ideal model to complement this work.
Zebrafish have similar gastrointestinal (GI) organs, lipid metabolism, and lipoprotein transport to higher vertebrates 1,2, develop rapidly, and are genetically tractable. The optical clarity of the larval zebrafish facilitates in vivo studies, a particular advantage for study of the GI system as its extracellular milieu (i.e., bile, microbiota, endocrine signaling) is virtually impossible to model ex vivo. In accordance, a body of research combining the genetic tractability and conduciveness to live imaging of zebrafish larvae with a variety of dietary manipulations (high-fat3,4, -cholesterol5, and -carbohydrate diets6,7), and models of cardiovascular disease8, diabetes9,10, hepatic steatosis11-13, and obesity14-16, are emerging to provide a host of metabolic insights.
An essential aspect of transitioning the larval zebrafish into metabolic research is the optimization of techniques developed in other model animals to the zebrafish and the development of novel assays that exploit the unique strengths of the zebrafish. This protocol presents techniques developed and optimized to feed larval zebrafish a lipid-rich meal, visualize dietary lipid processing from whole body to subcellular resolution, and measure food intake. Chicken egg yolk was chosen to compose the lipid-rich meal as it contains high levels of fats and cholesterol (lipids compose ~58% of chicken egg yolk, of which ~5% is cholesterol, 60% are triglycerides, and 35% are phospholipids). Chicken egg yolk provides more fat than typical commercial zebrafish micropellet foods (~15% lipids) and the advantage that it is a standardized feed with known percentages of specific fatty acids species, as zebrafish diets and feeding regiments have not been standardized across labs17. Moreover, fluorescent lipid analogs provided in the egg yolk visualize transport and accumulation of dietary lipids18, image cellular components including lipid droplets by acting both as vital dyes3 and through covalent incorporation into complex lipids, investigate metabolism through thin layer chromatography (TLC)19 and high performance liquid chromatography (HPLC) (S.A.F. unpublished data), and provide a quantitative assay for total food intake20.

<table><tbody><tr><td>Tricaine (ethyl 3-aminobenzoate methanesulofnate salt)</td><td>Sigma-Aldrich</td><td>A5040-25G</td><td>Anesthesia for larval zebrafish</td></tr><tr><td>Chicken eggs</td><td>N/A</td><td>N/A</td><td>Organic, cage-free eggs, not enriched for omege-3 fatty acids</td></tr><tr><td>Ultrasonic processor 3000 sonicator</td><td>Misonix, Inc.</td><td>S-3000</td><td>To make egg yolk liposomes</td></tr><tr><td>Sonabox acoustic enclosure</td><td>Misonix, Inc.</td><td>432B</td><td>To make egg yolk liposomes</td></tr><tr><td>1/8&amp;rdquo; tapered microtip</td><td>Misonix, Inc.</td><td>419</td><td>To make egg yolk liposomes</td></tr><tr><td>Amber vials (4 ml, glass)</td><td>National Scientific</td><td>13-425</td><td>Lipid storage; includes vials, open-top caps, and cap septa</td></tr><tr><td>Incu-Shaker Mini&amp;nbsp;</td><td>Benchmark</td><td>1222U12</td><td>Incubated shaker for feeds</td></tr><tr><td>BODIPY FL C16&amp;nbsp;</td><td>Thermo Fisher Scientific</td><td>D3821</td><td>Fluorescent lipid analog; (4,4-Difluoro-5,7-Dimethyl-4-Bora-3a,4a-Diaza-s-Indacene-3-Hexadecanoic Acid)</td></tr><tr><td>BODIPY FL C12&amp;nbsp;</td><td>Thermo Fisher Scientific</td><td>D3822</td><td>Fluorescent lipid analog; (4,4-Difluoro-5,7-Dimethyl-4-Bora-3a,4a-Diaza-s-Indacene-3-Dodecanoic Acid)</td></tr><tr><td>BODIPY FL C5&amp;nbsp;</td><td>Thermo Fisher Scientific</td><td>D3834</td><td>Fluorescent lipid analog; (4,4-Difluoro-5,7-Dimethyl-4-Bora-3a,4a-Diaza-s-Indacene-3-Pentanoic Acid)</td></tr><tr><td>BODIPY FL C5</td><td>Thermo Fisher Scientific</td><td>D2183</td><td>Fluorescent lipid analog; (4,4-Difluoro-5,7-Dimethyl-4-Bora-3a,4a-Diaza-s-Indacene-3-Propionic Acid)</td></tr><tr><td>TopFluor cholesterol&amp;nbsp;</td><td>Avanti Polar Lipids Inc.</td><td>810255</td><td>Fluorescent lipid analog; 23-(dipyrrometheneboron difluoride)-24-norcholesterol</td></tr><tr><td>Fatty acid-free BSA</td><td>Sigma-Aldrich</td><td>A0281-1G</td><td>For TopFluor cholesterol solubilization</td></tr><tr><td>Methyl cellulose</td><td>Sigma-Aldrich</td><td>M0387</td><td>Mounting media for live larval imaging; 75 x 25 x 1 mm</td></tr><tr><td>Low melt agarose</td><td>Thermo Fisher Scientific</td><td>BP165-25</td><td>Mounting media for live larval imaging; 22 x 30</td></tr><tr><td>VWR microscope slides&amp;nbsp;</td><td>VWR&amp;nbsp;</td><td>16004-422</td><td>Mounting larvae for live imaging</td></tr><tr><td>Coverslips&amp;nbsp;</td><td>Cover Glass</td><td>12-544A</td><td>Mounting larvae for live imaging</td></tr><tr><td>Super glue</td><td>Loctite</td><td>LOC01-30379</td><td>Mounting larvae for live imaging</td></tr><tr><td>FluoroDish (glass bottom dish)</td><td>World Precision Instruments, Inc.&amp;nbsp;</td><td>FD35-100</td><td>Mounting larvae for live imaging; 35 mm dish, 23 mm glass, 0.17 mm glass thickness&amp;nbsp;&amp;nbsp;</td></tr><tr><td>Confocal microscope</td><td>Leica Microsytems</td><td>SP-2, SP-5</td><td>Microscope for high magnification live imaging</td></tr><tr><td>Stereoscope</td><td>Nikon</td><td>SM21500</td><td>Microscope for low magnification live imaging</td></tr><tr><td>Glass culture tubes&amp;nbsp;</td><td>Kimble</td><td>73500-13100</td><td>Lipid extraction; (13 x 100 mm; 13 ml)</td></tr><tr><td>Savant SpeedVac Plus&amp;nbsp;</td><td>ThermoQuest</td><td>SC210A</td><td>Lipid extraction</td></tr><tr><td>Channeled TLC plates</td><td>Whatman Scientific</td><td>WC4855-821</td><td>Food intake assay; LK5D Silica Gel 150 A, 20 x 20 cm, 250 um thick; Discontinued</td></tr><tr><td>Channeled TLC plates</td><td>Analtech, Inc.</td><td>66911</td><td>Food intake assay; Direct replacement for Whatman Scientific TLC plates</td></tr><tr><td>Typhoon 9410 Variable Mode Imager</td><td>GE Healthcare</td><td>9410</td><td>Fluorescent plate reader for food intake assay</td></tr><tr><td>ImageQuant software</td><td>GE Healthcare</td><td>29000605</td><td>Analysis of food intake assay</td></tr><tr><td>5 3/4&amp;rsquo; Wide bore, borosilicate disposable pasteur pipets&amp;nbsp;&amp;nbsp;&amp;nbsp;</td><td>Kimble</td><td>63A53WT</td><td>Transfering larvae</td></tr></tbody></table>

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.

When fed on a rocker at 29-31 &#176;C, the majority of healthy larvae (&#8805;95%) will eat within 1 hr. Upon consuming the egg yolk emulsion, the larval intestine darkens in color. Very dark intestines can be observed at 2 hr (Figure 1). If larvae are unfed or fail to feed, the intestine remains clear. Larvae fed egg white exhibit a distended intestinal lumen that does not darken in color.
<img alt="Figure 1" src="/files/ftp_upload/54735/54735...

Watch this Scientific Journal Video about High-fat Feeding Paradigm for Larval Zebrafish: Feeding, Live Imaging, and Quantification of Food Intake at JoVE.com

High-fat Feeding Paradigm for Larval Zebrafish: Feeding, Live Imaging, and Quantification 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 ...

high-fat-feeding-paradigm-for-larval-zebrafish-feeding-live-imaging

Research

JoVE Journal

Biology

10.4K Views.  Carnegie Institution for Science. 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. 

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

Experimental Protocol for Examining Behavioral Response Profiles in Larval Fish: Application to the Neuro-stimulant Caffeine

Fish models and behaviors are increasingly used in the biomedical sciences; however, fish have long been the subject of ecological, physiological and toxicological studies. Using automated digital tracking platforms, recent efforts in neuropharmacology are leveraging larval fish locomotor behaviors to identify potential therapeutic targets for novel small molecules. Similar to these efforts, research in the environmental sciences and comparative pharmacology and toxicology is examining various behaviors of fish models as diagnostic tools in tiered evaluation of contaminants and real-time monitoring of surface waters for contaminant threats. Whereas the zebrafish is a popular larval fish model in the biomedical sciences, the fathead minnow is a common larval fish model in ecotoxicology. Unfortunately, fathead minnow larvae have received considerably less attention in behavioral studies. Here, we develop and demonstrate a behavioral profile protocol using caffeine as a model neurostimulant. Though photomotor responses of fathead minnows were occasionally affected by caffeine, zebrafish&#160;were markedly more sensitive for photomotor and locomotor endpoints, which responded at environmentally relevant levels. Future studies are needed to understand comparative behavioral sensitivity differences among fish with age and time of day, and to determine whether similar behavioral effects would occur in nature and be indicative of adverse outcomes at the individual or population levels of biological organization.

Here, we present a protocol to examine larval zebrafish and fathead minnow locomotor activities and photomotor responses (PMR) using an automated tracking software. When incorporated in common toxicology bioassays, analyses of these behaviors provide a diagnostic tool to examine chemical bioactivity. This protocol is described using caffeine, a model neurostimulant.

Fish models and behaviors are increasingly used in the biomedical sciences; however, fish have long been the subject of ecological, physiological and ...

Environment

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 ...

Cancer Research

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

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.

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 ...

Developmental Biology

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 ...

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

High Throughput Danio Rerio Energy Expenditure Assay

Zebrafish are an important model organism with inherent advantages that have the potential to make zebrafish a widely applied model for the study of energy homeostasis and obesity. The small size of zebrafish allows for assays on embryos to be conducted in a 96- or 384-well plate format, Morpholino and CRISPR based technologies promote ease of genetic manipulation, and drug treatment by bath application is viable. Moreover, zebrafish are ideal for forward genetic screens allowing for novel gene discovery. Given the relative novelty of zebrafish as a model for obesity, it is necessary to develop tools that fully exploit these benefits. Herein, we describe a method to measure energy expenditure in thousands of embryonic zebrafish simultaneously. We have developed a whole animal microplate platform in which we use 96-well plates to isolate individual fish and we assess cumulative NADH2 production using the commercially available cell culture viability reagent alamarBlue. In poikilotherms the relationship between NADH2 production and energy expenditure is tightly linked. This energy expenditure assay creates the potential to rapidly screen pharmacological or genetic manipulations that directly alter energy expenditure or alter the response to an applied drug (e.g. insulin sensitizers).

High Throughput Danio Rerio Energy Expenditure Assay

Zebrafish are an important model organism for the study of energy homeostasis. By utilizing a NADH2 sensitive redox indicator, alamar Blue, we have developed an assay that measures the metabolic rate of zebrafish larvae in a 96 well plate format and can be applied to drug or gene discovery.

Zebrafish are an important model organism with inherent advantages that have the potential to make zebrafish a widely applied model for the study of ...

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-throughput Measurement of Gut Transit Time Using Larval Zebrafish

Zebrafish are used as alternative model organisms for drug safety testing. The gastrointestinal (GI) tract of zebrafish has genetic, neuronal, and pharmacological similarities to that of the mammals. GI intolerance during clinical testing of drug candidates is common and may pose a serious threat to human health. Testing for GI toxicity in preclinical mammalian models can be expensive in terms of time, test compound, and labor. The high-throughput method presented here may be used to predict GI safety issues. Compared to mammalian models, this method allows for more expedient assessment of test compound effects on GI transit while using low quantities of compound. In this method, larval zebrafish (7 days post fertilization) are fed food containing a fluorescent label. After feeding, each larval fish is placed into a well of a 96-conical-bottom-well plate and dosed with test compound (dissolved in water) or the vehicle. As gut transit occurs, fecal matter accumulates on the bottom of the wells, and the rate at which this happens is monitored by measuring fluorescence from the bottom of the well repeatedly over time using a plate spectrophotometer. The fluorescence from larvae in a given treatment group are averaged and these values are graphed along with standard error, for each measurement time, yielding a curve representing average transit of food over time. Effects on gut transit time are identified by comparing the area under the curve for each treatment group to that of the vehicle-treated group. This method detected changes in zebrafish GI transit time induced by drugs with known clinical GI effects; it can be employed to interrogate dozens of treatments for GI effects per day. As such, safer compounds can be quickly prioritized for mammalian testing, which expedites discovery and proffers 3Rs advancement.

The goal of this protocol is to measure the transit time of fluorescently labeled food through the gut of larval zebrafish in a high throughput fashion.

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

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