This work was funded by the National Natural Science Foundation of China (21777145, 22076170), and the Program for Changjiang Scholars and Innovative Research Team in University (IRT_17R97).

Adult zebrafish are originated from the China Zebrafish Resource Center (Wuhan, China). The experiments were conducted in compliance with the national guide &#34;Laboratory Animal Guideline for Ethical Review of Animal Welfare (GB/T35892-2018).
1. Embryo collection
<ol>
	<li>Maintain fish in 20 L glass tanks with recirculating charcoal-filtered tap water system (pH 7.0 &#177; 0.2) at a constant temperature (28 &#177; 0.5 &#176;C) on a photoperiod of 14:10 h light: dark.</li>
	<li>Feed fish twice daily with Artemia nauplii. It is recommended that the food is given at max. 3% fish weight per day and should be eaten within 5 min every time14.</li>
	<li>Transfer well-developed adult zebrafish (with body length of 3-4 cm) into the spawning tank at a ratio of one male to two females the night before the breeding. 
	NOTE: The following morning, the fish start to spawn after the onset of the light cycle.</li>
	<li>Collect eggs using a Pasteur pipette. Rinse with 10% Hank&#39;s solution several times, and then check for fertilization using a microscope. Fertilized eggs undergo the cleavage period after approximately 2 h post fertilization (hpf) and can be clearly identified15.</li>
	<li>Incubate the fertilized embryos in a 500 mL beaker containing 200 mL of 10% Hank&#39;s solution with 1% methylene blue for disinfection at 28 &#176;C. Do not exceed a loading rate of 1 embryo/2 mL solution. 
	NOTE: 10% Hank&#39;s solution is made up of 137 mM NaCl, 5.4 mM KCl, 0.25 mM Na2HPO4, 0.44 mM KH2PO4, 1.3 mM CaCl2, 1.0 mM MgSO4 and 4.2 mM NaHCO3.</li>
</ol>
2. Preparation of microplastic suspensions
<ol>
	<li>Sonicate the stock solution of green fluorescently labeled polystyrene beads (10 mg/mL) with nominal diameter of 500 nm (excitation/emission: 460/500 nm) for 10 minutes.</li>
	<li>Dilute the stock solution with 10% Hank&#39;s solution to produce the desired exposure solutions (0.1, 1, and 10 mg/L).</li>
	<li>Always prepare the exposure solutions of microplastics before exposure. 
	&#8203;NOTE: Caution should be taken when assessing the toxic effects of microplastics, because the presence of preservatives, such as sodium azide, in the commercial particle formulations, can be toxic to different organisms 16. Therefore, these additives should be removed or accounted for in the controls before conducting toxicity experiment.</li>
</ol>
3. Microplastic exposure
<ol>
	<li>Randomly select 6 newly fertilized embryos (4 hpf), and then transfer into each well of 6-well plate containing 5 mL of microplastic solutions with different concentrations. Include the control groups containing 10% Hank&#39;s solution.
	<ol>
		<li>Use triplicate wells (with a total of 18 embryos) for each treatment.</li>
	</ol>
	</li>
	<li>Incubate the embryos under the same light: dark cycle and temperature as adults (see 1.2) and observe every 12 hours. Remove the dead immediately.</li>
	<li>Renew the microplastic solutions 90% every 24 h. During the exposure period, the fish are not fed. 
	NOTE: Generally, the hatching of embryo begins at 48 hpf and completes at about 72 hpf.</li>
</ol>
4. Assessment of microplastic distribution
<ol>
	<li>At 24, 48, 72, 96, and 120 h post fertilization, randomly select the embryos/larvae (one from each of the three replicates) and rinse with 10% Hank&#39;s solution.</li>
	<li>Transfer the larvae into a Petri dish and expose to 0.016% tricaine for anesthesia.
	<ol>
		<li>Prepare the stock solution of tricaine: 4 mg of tricaine powder is dissolved in 100 mL of double distilled water, and adjust the pH to 7.0 with Tris-HCl (pH 9.0). Store the stock solution in the freezer.</li>
		<li>Prepare the working solution. Dilute the stock solution to the desired concentration (0.016%) with 10% Hank&#39;s solution at room temperature14.</li>
	</ol>
	</li>
	<li>Arrange the embryos/larvae and prepare for observation.</li>
	<li>Observe the fish with a fluorescence microscope and image with imaging software.</li>
	<li>Quantify the fluorescence intensity in fish with ImageJ.</li>
</ol>

<ol><li>SAPEA (Science Advice for Policy by European Academies). <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=%22A+Scientific+Perspective+on+Microplastics+in+Nature+and+Society%22">A Scientific Perspective on Microplastics in Nature and Society</a>. , SAPEA. Berlin. (2019).</li>
<li>Plastics Europe. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=%22Plastics-the+facts+2019%22">Plastics-the facts 2019</a>. , Plastics Europe. Brussels. (2019).</li>
<li>Andrady, A. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Microplastics+in+the+marine+environment%5BTitle%5D)+AND+%22Marine+Pollution+Bulletin%22%5BJournal%5D)">Microplastics in the marine environment.</a> Marine Pollution Bulletin. 62, 1596-1605 (2011).</li>
<li>Arthur, C., Baker, J., Bamford, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Proceedings+of+the+International+Research+Workshop+on+the+Occurrence%2C+Effects+and+Fate+of+Microplastic+Marine+Debris%5BTitle%5D)+AND+%22National+Oceanic+and+Atmospheric+Administration+Technical+Memorandum%22%5BJournal%5D)">Proceedings of the International Research Workshop on the Occurrence, Effects and Fate of Microplastic Marine Debris.</a> National Oceanic and Atmospheric Administration Technical Memorandum. , (2009).</li>
<li>Ivleva, N. P., Wiesheu, A. C., Niessner, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Microplastic+in+aquatic+ecosystems%5BTitle%5D)+AND+%22Angewandte+Chemie+International+Edition%22%5BJournal%5D)">Microplastic in aquatic ecosystems.</a> Angewandte Chemie International Edition. 56, 1720-1739 (2017).</li>
<li>Lu, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Pollutant+toxicology+with+respect+to+microalgae+and+cyanobacteria%5BTitle%5D)+AND+%22Journal+of+Environmental+Sciences%22%5BJournal%5D)">Pollutant toxicology with respect to microalgae and cyanobacteria.</a> Journal of Environmental Sciences. 99, 175-186 (2021).</li>
<li>Huang, J. N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Exposure+to+microplastics+impairs+digestive+performance%2C+stimulates+immune+response+and+induces+microbiota+dysbiosis+in+the+gut+of+juvenile+guppy+%28Poecilia+reticulata%29%5BTitle%5D)+AND+%22Science+of+the+Total+Environment%22%5BJournal%5D)">Exposure to microplastics impairs digestive performance, stimulates immune response and induces microbiota dysbiosis in the gut of juvenile guppy (Poecilia reticulata).</a> Science of the Total Environment. 733, 138929(2020).</li>
<li>Prüst, M., Meijer, J., Westerink, R. H. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+plastic+brain%3A+neurotoxicity+of+micro-+and+nanoplastics%5BTitle%5D)+AND+%22Particle+and+Fibre+Toxicology%22%5BJournal%5D)">The plastic brain: neurotoxicity of micro- and nanoplastics.</a> Particle and Fibre Toxicology. 17, 24(2020).</li>
<li>Jakubowska, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Effects+of+chronic+exposure+to+microplastics+of+different+polymer+types+on+early+life+stages+of+sea+trout+Salmo+trutta%5BTitle%5D)+AND+%22Science+of+the+Total+Environment%22%5BJournal%5D)">Effects of chronic exposure to microplastics of different polymer types on early life stages of sea trout Salmo trutta.</a> Science of the Total Environment. 740, 139922(2020).</li>
<li>Qiang, L., Cheng, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Exposure+to+polystyrene+microplastics+impairs+gonads+of+zebrafish+%28Danio+rerio%29%5BTitle%5D)+AND+%22Chemosphere%22%5BJournal%5D)">Exposure to polystyrene microplastics impairs gonads of zebrafish (Danio rerio).</a> Chemosphere. 263, 128161(2021).</li>
<li>Hamed, M., Soliman, H. A. M., Osman, A. G. M., Sayed, A. E. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Antioxidants+and+molecular+damage+in+Nile+Tilapia+%28Oreochromis+niloticus%29+after+exposure+to+microplastics%5BTitle%5D)+AND+%22Environmental+Science+and+Pollution+Research%22%5BJournal%5D)">Antioxidants and molecular damage in Nile Tilapia (Oreochromis niloticus) after exposure to microplastics.</a> Environmental Science and Pollution Research. 27, 14581-14588 (2020).</li>
<li>Burns, E. E., Boxall, A. B. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Microplastics+in+the+aquatic+environment%3A+Evidence+for+or+against+adverse+impacts+and+major+knowledge+gaps%5BTitle%5D)+AND+%22Environmental+Toxicology+and+Chemistry%22%5BJournal%5D)">Microplastics in the aquatic environment: Evidence for or against adverse impacts and major knowledge gaps.</a> Environmental Toxicology and Chemistry. 37, 2776-2796 (2018).</li>
<li>Ma, H., Pu, S., Liu, S., Bai, Y., Mandal, S., Xing, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Microplastics+in+aquatic+environments%3A+Toxicity+to+trigger+ecological+consequences%5BTitle%5D)+AND+%22Environmental+Pollution%22%5BJournal%5D)">Microplastics in aquatic environments: Toxicity to trigger ecological consequences.</a> Environmental Pollution. 261, 114089(2020).</li>
<li>Westerfield, M. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aWesterfield%2C+M+%22The+Zebrafish+Book%3A+A+Guide+for+the+Laboratory+Use+of+Zebrafish+%28Danio+reio%29.+4th+ed%22">The Zebrafish Book: A Guide for the Laboratory Use of Zebrafish (Danio reio). 4th ed</a>. , University of Oregon Press. Eugene. (2000).</li>
<li>Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B., Schilling, T. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Stages+of+embryonic+development+of+the+zebrafish%5BTitle%5D)+AND+%22Developmental+Dynamics%22%5BJournal%5D)">Stages of embryonic development of the zebrafish.</a> Developmental Dynamics. 203, 253-310 (1995).</li>
<li>Pikuda, O., Xu, E. G., Berk, D., Tufenkji, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Toxicity+assessments+of+micro-+and+nanoplastics+can+be+confounded+by+preservatives+in+commercial+formulations%5BTitle%5D)+AND+%22Environmental+Science+%26+Technology+Letters%22%5BJournal%5D)">Toxicity assessments of micro- and nanoplastics can be confounded by preservatives in commercial formulations.</a> Environmental Science & Technology Letters. 6, 21-25 (2019).</li>
<li>Lidster, K., Readman, G. D., Prescott, M. J., Owen, S. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((International+survey+on+the+use+and+welfare+of+zebrafish+Danio+rerio+in+research%5BTitle%5D)+AND+%22Journal+of+Fish+Biology%22%5BJournal%5D)">International survey on the use and welfare of zebrafish Danio rerio in research.</a> Journal of Fish Biology. 90, 1891-1905 (2017).</li>
<li>Pitt, J. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Uptake%2C+tissue+distribution%2C+and+toxicity+of+polystyrene+nanoparticles+in+developing+zebrafish+%28Danio+rerio%29%5BTitle%5D)+AND+%22Aquatic+Toxicology%22%5BJournal%5D)">Uptake, tissue distribution, and toxicity of polystyrene nanoparticles in developing zebrafish (Danio rerio).</a> Aquatic Toxicology. 194, 185-194 (2018).</li>
<li>Lin, S. J., Zhao, Y., Nel, A. E., Lin, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Zebrafish%3A+An+in+vivo+model+for+nano+EHS+studies%5BTitle%5D)+AND+%22Small%22%5BJournal%5D)">Zebrafish: An in vivo model for nano EHS studies.</a> Small. 9, 1608-1618 (2013).</li>
</ol>

The author declares no competing or financial interests.

According to the guideline on the protection of animals used for scientific purposes, such as EU Directive 2010/63/EU, animal ethics permission is not mandatory for an experiment with early life-stages of zebrafish until the stage of being capable of independent feeding (5 days post fertilization)17. However, best welfare practice is important for optimizing the use of zebrafish, and, for example, the humane methods of anesthesia and euthanasia should be of concern. Ethyl 3-aminobenzoate methanesulphate (MS-222, or tricaine), the routinely used agent in the most laboratories, is employed here for anesthesia and euthanasia.
Before observation under the microscope, the embryos and larvae should be rinsed since the microplastics adsorbed on the external surface might interfere with the results. In addition, the autofluorescence in the embryos/larvae, especially around the yolk sac, which has been reported occasionally, might be problematic. The presence of many biomacromolecules, such as flavins, nicotinamide-adenine dinucleotide (NAD), aromatic amino acids, lipofuscins, advanced glycation end products, and collage, will emit light when excited at the appropriate wavelength. 
It is important to note that, as the particulate pollutant, the size of microplastic is considered as one of the determining factors of bioavailability, and toxicity18. The nominal diameter of microplastic used here is 500 nm, which is comparative to the pore size of the embryo chorion (within the range of 300 nm to 1 &#181;m)19. Therefore, these microplastics are not expected to easily pass through the zebrafish chorion. Consistently, there is little fluorescence visible in the embryos before hatching (Figure 1). Since the chorion will act as an effective barrier against the particles with large size, the dechorionation process before exposure may be needed. Chorion can be removed easily using the forceps, but enzymatic dechorionation with pronase is preferred when the embryos are handled in bulk. However, although dechorionation will increase the bioavailability and facilitate the high-throughput screening for the toxicity of substances, the embryo with chorion intact is more recommended to assess the ecotoxicity of pollutants when considering the condition of exposure in the "real" world. 
Although considerable efforts have been devoted to investigating the adverse effects of microplastics on fish, the current knowledge, including that of bioaccumulation, remain limited or even conflicting. These across-study inconsistencies are mainly attributed to the differences of properties of particles, including size, density, and surface characteristics (for example, surface charge). The behavior of microplastics in the solution is critical to the bioavailability as well. The physicochemical characteristics of microplastics should be tracked over the exposure duration, and the aggregation phenomenon that may occur should be recorded. In fact, for the exposures that require the microplastics to be suspended for an extended period, sonication or stirring with a magnetic bar is recommended.

Since first synthesized in the 1900s, plastics are widely used in various fields, resulting in rapid growth of global production1. In 2018, approximately 360 million tons of plastics were produced worldwide2. The plastics in the natural environment will degrade to fine particles due to chemical, physical or biological processes3. Generally, fine plastic particles &lt;5 mm in size are defined as microplastics4. Microplastics are also engineered for specific applications, such as microbeads from cosmetic products5. As near-permanent contaminants, microplastics are accumulated in the environment, and have attracted increasing attention from scientists, policymakers and the public1,6. Previous studies documented that microplastics could cause adverse effects in fish, such as gastrointestinal damage7, neurotoxicity8, endocrine disruption9, oxidative stress10 and DNA damage11. However, the toxicity of microplastics has not been fully revealed so far12,13.
Zebrafish embryos offer a lot of experimental advantages, including small size, external fertilization, optical transparency and large clutches, and is considered as an ideal model organism for in vivo studying the effects of pollutants on fish at early life stages. In addition, only limited amounts of test substances are needed for the evaluation of biological responses. Here, zebrafish embryos are exposed to different concentrations of microplastics (0.1, 1, 10 mg/L) for 5 days, and the bioaccumulation and distribution of microplastics in zebrafish embryos/larvae are evaluated. This result will advance our understanding about the toxicity of microplastics to fish, and the method described here can potentially be generalized to determine the accumulation and distribution of other types of fluorescent materials in the early life stages of zebrafish.

<table><tbody><tr><td>Fluorescent microscope</td><td>Nikon, Japan</td><td>Eclipse Ti-S</td><td></td></tr><tr><td>Green fluorescently labeled polystyrene beads</td><td>Phosphorex, USA</td><td>2103A</td><td></td></tr><tr><td>Tricaine</td><td>Sigma-Aldrich, USA</td><td>A5040</td><td></td></tr></tbody></table>

accumulation and distribution of fluorescent microplastics in the early life stages of zebrafish

As a new type of environmental pollutant, microplastic has been widely found in the aquatic environment and poses a high threat to aquatic organisms. The bioaccumulation of microplastics plays a key role in their toxic effects; however, as a particulate, their bioaccumulations are different from many other pollutants. Described here is a feasible method to visually determine the accumulation and distribution of microplastics in zebrafish embryos or larvae using fluorescent microplastics. Embryos are exposed to different concentrations (0.1, 1, and 10 mg/L) of fluorescent microplastics with a diameter of 500 nm for 120 h. It is shown in the results that microplastics can bioaccumulate in zebrafish embryos/larvae in a concentration-dependent manner. Before hatching, strong fluorescence is found around the embryonic chorion; while in zebrafish larvae, the yolk sac, pericardium, and gastrointestinal tract are the main accumulated sites of microplastics. The results demonstrate the uptake and internalization of microplastics in zebrafish at early life stages, which will provide basis for better understanding the impact of microplastics on aquatic animals.

The distribution and accumulation of fluorescent microplastics are shown in Figure 1 and Table 1. No visible fluorescence is observed in the unexposed group (control). However, an accumulation of fluorescence is found surrounding the chorion after exposure to different concentrations of microplastics (24 hpf). Green fluorescence is also detected in larvae, and the fluorescence levels appear to increase in a concentration- and time-dependent manner. The yolk sac, pericardium, and gastrointestinal tract are the main accumulated sites of microplastics (Figure 2).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62117/62117fig01.jpg" /> 
Figure 1:&#160;Distribution of fluorescent polystyrene microplastics in embryos/larvae of zebrafish (40&#215;). The fish are sampled from the control group, or the groups exposed to 500-nm microplastics at 0.1, 1 and 10 mg/L.&#160;Scale bar 100 &#181;m <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62117/62117fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62117/62117fig02.jpg" /> 
Figure 2:&#160;The sites of microplastic accumulation in zebrafish larvae (40&#215;). This larva is sampled from the group exposed to 500-nm microplastics at 10 mg/L for 120 hours. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62117/62117fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Concentration</td>
			<td>Embryo</td>
			<td></td>
			<td>Larva</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>(mg/L)</td>
			<td>24 hpf</td>
			<td>48 hpf&#160;a</td>
			<td>48 hpf b</td>
			<td>72 hpf</td>
			<td>96 hpf</td>
			<td>120 hpf</td>
		</tr>
		<tr>
			<td>Cont.</td>
			<td>1.2&#177;0.1</td>
			<td>2.6&#177;0.3</td>
			<td>2.2</td>
			<td>3.0&#177;0.2</td>
			<td>2.6&#177;0.7</td>
			<td>3.3&#177;0.3</td>
		</tr>
		<tr>
			<td>1</td>
			<td>1.2&#177;0.2</td>
			<td>5.0&#177;0.1</td>
			<td>5.3</td>
			<td>7.5&#177;0.5</td>
			<td>8.7&#177;0.5</td>
			<td>10.0&#177;1.9</td>
		</tr>
		<tr>
			<td>0.1</td>
			<td>7.0&#177;0.9</td>
			<td>26.1&#177;2.9</td>
			<td>8.9</td>
			<td>18.4&#177;0.7</td>
			<td>16.3&#177;2.8</td>
			<td>25.7&#177;2.7</td>
		</tr>
		<tr>
			<td>10</td>
			<td>9.1&#177;1.1</td>
			<td>82.3&#177;5.3</td>
			<td>30.4</td>
			<td>32.7&#177;3.2</td>
			<td>41.6&#177;0.4</td>
			<td>44.1&#177;0.9</td>
		</tr>
		<tr>
			<td colspan="7">a: only two embryos were assessed; b: only one larva was assessed.</td>
		</tr>
	</tbody>
</table>
Table 1: The change of fluorescence level in zebrafish following exposure to fluorescent microplastics (n=3). Due to the influence of chorion on the absorption of fluorescent microplastics, the data are divided into two parts, that of the embryos (before hatching) and larvae (after hatching).

Watch this Scientific Journal Video about Accumulation and Distribution of Fluorescent Microplastics in the Early Life Stages of Zebrafish at JoVE.com

Accumulation and Distribution of Fluorescent Microplastics in the Early Life Stages of Zebrafish

Zebrafish embryos/larvae develop externally and are optically transparent. The bioaccumulation of microplastics in fish at early life stages is readily assessed with fluorescently labeled microbeads.

As a new type of environmental pollutant, microplastic has been widely found in the aquatic environment and poses a high threat to aquatic organisms. The ...

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Nanyang Technological University

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Video: Accumulation and Distribution of Fluorescent Microplastics in the Early Life Stages of Zebrafish

Microbead Implantation in the Zebrafish Embryo

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Developmental Biology

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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.

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Biology

Elucidating Microglia Behavior in Zebrafish Optic Tectum Development

Live Imaging and Characterization of Microglia Dynamics in the Zebrafish Embryo

Microglia are highly dynamic cells and their migration and colonization of the brain parenchyma is a crucial step for proper brain development and function. Externally developing zebrafish embryos possess optical transparency, which along with well-characterized transgenic reporter lines that fluorescently label microglia, make zebrafish an ideal vertebrate model for such studies. In this paper, we take advantage of the unique features of the zebrafish model to visualize the dynamics of microglia cells in vivo and under physiological conditions. We use confocal microscopy to record a timelapse of microglia cells in the optic tectum of the zebrafish embryo and then, extract tracking data using the IMARIS 10.0 software to obtain the cells' migration path, mean speed, and distribution in the optic tectum at different developmental stages. This protocol can be a useful tool to elucidate the physiological significance of microglia behavior in various contexts, contributing to a deeper characterization of these highly motile cells.

Author Spotlight: Using Zebrafish to Explore Microglia Migration During Brain Development

We demonstrate a method that takes advantage of fast scanning confocal microscopy to perform live imaging of microglia cells in the developing zebrafish optic tectum, allowing for the analysis of the dynamics of these cells in vivo.

Microglia are highly dynamic cells and their migration and colonization of the brain parenchyma is a crucial step for proper brain development and ...

Immunology and Infection

Evaluation of Zebrafish Kidney Function Using a Fluorescent Clearance Assay

The zebrafish embryo offers a tractable model to study organogenesis and model human genetic disease. Despite its relative simplicity, the zebrafish kidney develops and functions in almost the same way as humans. A major difference in the construction of the human kidney is the presence of millions of nephrons compared to the zebrafish that has only two. However, simplifying such a complex system into basic functional units has aided our understanding of how the kidney develops and operates. In zebrafish, the midline located glomerulus is responsible for the initial blood filtration into two pronephric tubules that diverge to run bilaterally down the embryonic axis before fusing to each other at the cloaca. The pronephric tubules are heavily populated by motile cilia that facilitate the movement of filtrate along the segmented tubule, allowing the exchange of various solutes before finally exiting via the cloaca2-4. Many genes responsible for CKD, including those related to ciliogenesis, have been studied in zebrafish5. However, a major draw back has been the difficulty in evaluating zebrafish kidney function after genetic manipulation. Traditional assays to measure kidney dysfunction in humans have proved non translational to zebrafish, mainly due to their aquatic environment and small size. For example, it is not physically possible to extract blood from embryonic staged fish for analysis of urea and creatinine content, as they are too small. In addition, zebrafish do not produce enough urine for testing on a simple proteinuria &#8216;dipstick&#8217;, which is often performed during initial patient examinations. We describe a fluorescent assay that utilizes the optical transparency of the zebrafish to quantitatively monitor the clearance of a fluorescent dye, over time, from the vasculature and out through the kidney, to give a read out of renal function1,6-9.

The zebrafish is a popular tool to model chronic kidney disease (CKD). However, their small size makes it impossible to evaluate renal function using traditional methods. We describe a fluorescent dye kidney clearance assay1 that allows quantitative analysis of zebrafish kidney function in CKD.

The zebrafish embryo offers a tractable model to study organogenesis and model human genetic disease. Despite its relative simplicity, the zebrafish ...

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 ...

Environmental Screening of Aeromonas hydrophila, Mycobacterium spp., and Pseudocapillaria tomentosa in Zebrafish Systems

Health monitoring systems are developed and used in zebrafish research facilities because pathogens of Danio rerio such as Aeromonas hydrophila, Mycobacterium spp., and Pseudocapillaria tomentosa have the potential to impair animal welfare and research. The fish are typically analyzed post mortem to detect microbes. The use of sentinels is a suggested way to improve the sensitivity of the surveillance and to reduce the number of animals to sample. The setting of a pre-filtration sentinel tank out of a recirculating system is described. The technique is developed to prevent water pollution and to represent the fish population by a careful selection of age, gender, and strains. In order to use the minimum number of animals, techniques to screen the environment are also detailed. Polymerase Chain Reaction (PCR) on surface sump swabs is used to significantly improve the detection of some prevalent and pathogenic mycobacterial species such as Mycobacterium fortuitum, Mycobacterium haemophilum, and Mycobacterium chelonae. Another environmental method consists of processing the sludge at the bottom of a holding tank or sump to look for P. tomentosa eggs. This is a cheap and fast technique that can be applied in quarantine where a breeding device is submerged into the holding tank of imported animals. Finally, PCR is applied to the sludge sample and A. hydrophila is detected at the sump&#39;s bottom and surface. Generally, these environmental screening techniques applied to these specific pathogens have led to an increased sensitivity compared to the testing of pre-filtration sentinels.

Environmental Screening of Aeromonas hydrophila, Mycobacterium spp., and Pseudocapillaria tomentosa in Zebrafish Systems

This protocol describes the use of sump swabs and sludge analysis of zebrafish systems, which leads to increased detection compared to the sole use of sentinels to detect pathogens such as Aeromonas hydrophila, Mycobacterium spp., and Pseudocapillaria tomentosa. A system to monitor P. tomentosa eggs in quarantine is also proposed.

Health monitoring systems are developed and used in zebrafish research facilities because pathogens of Danio rerio such as Aeromonas hydrophila, ...

Simple and Effective Administration and Visualization of Microparticles in the Circulatory System of Small Fishes Using Kidney Injection

The systemic administration of micro-size particles into a living organism can be applied for vasculature visualization, drug and vaccine delivery, implantation of transgenic cells and tiny optical sensors. However, intravenous microinjections into small animals, which are mostly used in biological and veterinary laboratories, are very difficult and require trained personnel. Herein, we demonstrate a robust and efficient method for the introduction of microparticles into the circulatory system of adult zebrafish (Danio rerio) by injection into the fish kidney. To visualize the introduced microparticles in the vasculature, we propose a simple intravital imaging technique in fish gills. In vivo monitoring of the zebrafish blood pH was accomplished using an injected microencapsulated fluorescent probe, SNARF-1, to demonstrate one of the possible applications of the described technique. This article provides a detailed description of the encapsulation of pH-sensitive dye and demonstrates the principles of the quick injection and visualization of the obtained microcapsules for in vivo recording of the fluorescent signal. The proposed method of injection is characterized by a low mortality rate (0-20%) and high efficiency (70-90% success), and it is easy to institute using commonly available equipment. All described procedures can be performed on other small fish species, such as guppies and medaka.

This article demonstrates the principles of a quick, minimally invasive injection of fluorescent microparticles into the circulatory system of small fishes and the in vivo visualization of the microparticles in fish blood.

The systemic administration of micro-size particles into a living organism can be applied for vasculature visualization, drug and vaccine delivery, ...

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.

Zebrafish are used as alternative model organisms for drug safety testing. The gastrointestinal (GI) tract of zebrafish has genetic, neuronal, and ...

Small Molecule Screening and Toxicity Testing in Early-stage Zebrafish Larvae

Zebrafish have become a prominent model organism for human translational research, toxicity testing of small molecules and environmental pollutants, therapeutic drug discovery, and other biomedical research applications. The presented protocol aims to provide a clear guide for drug screening and toxicity testing in early-stage zebrafish larvae (aged 3-7 days post fertilization, dpf). Several methodologies describing toxicity testing in zebrafish larvae have been published, albeit lacking a standardized protocol for exposure durations, rate of media changes, and morphology scoring, amongst others. We experimentally determined the ideal larvae ages for conducting the experiments, the plate sizes that work best for those ages, the rate of media changes that provide reliable results while limiting wastage&#160;of test compound, and a morphology scoring system that is easily quantifiable in a statistical manner. The manuscript describes in detail a protocol for toxicity testing in zebrafish larvae aged between&#160;3 to 7 dpf, aiming to target the gap in drug toxicity testing and taking the first step towards a standardized method similar to the Fish Embryo Acute Toxicity test.

This protocol describes a drug screening procedure in zebrafish larvae 3-7 days post fertilization followed by morphological assessment.

Zebrafish have become a prominent model organism for human translational research, toxicity testing of small molecules and environmental pollutants, ...

Accumulation and Distribution of Fluorescent Microplastics in the Early Life Stages of Zebrafish

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