The research presented in this article was supported by previous grants from National Institutes of Health/National Institute of Dental and Craniofacial Research (NIH/NIDCR) R01DE020884 to J.K.E. and National Institutes of Health/National Institute on Alcohol Abuse and Alcoholism (NIH/NIAAA) F32AA021320 to C.B.L. and by the current grant from National Institutes of Health/National Institute on Alcohol Abuse (NIH/NIAAA) R00AA023560 to C.B.L. We thank Rueben Gonzales for providing and assisting with gas chromatograph analysis. We thank Tiahna Ontiveros and Dr. Gina Nobles writing assistance.

All zebrafish embryos used in this procedure were raised and bred following established IACUC protocols27. These protocols were approved by the University of Texas at Austin and the University of Louisville.
NOTE: The zebrafish line Tg(fli1:EGFP)y1 was used in this study28. All water used in this procedure is sterile reverse osmosis water. All statistical analyses were performed using Graphpad Prism v8.2.1.
1. Making embryo media
<ol>
	<li>To make a 20x stock of embryo media, dissolve 17.5 g of NaCl, 0.75 g of KCl, 2.9 g of CaCl2, 0.41 g of K2HPO4, 0.142 g of NA2HPO4, and 4.9 g of MgSO4&#183;7H2O in 1 L of water. Ignore the white precipitate that forms; this will not impact the media. Filter sterilize the stock solution and store at 4 &#176;C.</li>
	<li>To create the working embryo media solution, dissolve 1.2 g of NaHCO3 in 1 L of 20x embryo media stock and add 19 L of water. Maintain the working embryo media solution at 28 &#176;C.</li>
</ol>
2. Measuring the embryonic volume using water displacement
NOTE: In this protocol, 24 h postfertilization (hpf) embryos (Figure 1) are used. The embryos used in the volume measurements are not used in the ethanol analysis.
<ol>
	<li>Place 10 embryos and extraembryonic fluid in 1.5 mL microcentrifuge tube marked at a volume of 250 &#956;L (Figure 2A). Add water to the 250 &#956;L fill line (see sample with dyed water in Figure 2B).</li>
	<li>Repeat step 2.1 to set up receptacles for embryos that have had their chorions removed.
	<ol>
		<li>To remove the chorion, place the embryos in their chorions in a 100 mm Petri dish with 2 mg/mL of protease cocktail in embryo media at room temperature (RT) for 10 min. Every few minutes, gently swirl the embryos to break the chorion.</li>
		<li>Once all the embryos are free of their chorion, remove the dechorionated embryos from protease cocktail/embryo media and place in a new 100 mm Petri dish with fresh embryo media to wash the embryos. Repeat this wash step 1 more time (for a total of 2 washes). Transfer the embryos to a fresh 100 mm Petri dish.</li>
	</ol>
	</li>
	<li>Using the smallest tip possible and without damaging the embryos, carefully remove all liquid from around the embryos using a p200 micropipettor (Figure 2C) and weigh the water with a scale with &#60;0.1 mg precision. To determine the volume of the sample of 10 embryos, subtract the weight/volume of the water (1 mL of water = 1 g of water.) removed from 250 &#956;L. To determine the volume of a single embryo, divide the difference between 250 &#956;L and the weight of the water removed by 10.</li>
</ol>
<img alt="Equation 1" src="/files/ftp_upload/60766/60766equ01.jpg" />
<img alt="Equation 2" src="/files/ftp_upload/60766/60766equ02.jpg" />
3. Treating embryos with ethanol
<ol>
	<li>Gather embryos from mating tanks and place in a standard 100 mm Petri dish. Count out no more than 100 embryos per single Petri dish and incubate at 28.5 &#176;C. 
	NOTE: If the chorion is to be removed (step 2.2), it needs to be removed before adding ethanol.</li>
	<li>At 6 hpf, add up to 100 embryos to a new standard 100 mm Petri dish either with embryo media or embryo media + 1% ethanol (v/v). Cover, but DO NOT seal the Petri dish. Place the embryos in a low temp incubator set at 28.5 &#176;C for 18 h, or until the embryos reach the developmental time point of 24 hpf.</li>
</ol>
4. Preparing workflow before processing the embryos for head space gas chromatography
<ol>
	<li>Make a solution of 5 M NaCl in water (450 &#956;L will be needed per sample tube) to denature all proteins and prevent ethanol metabolism. Make the protease cocktail (Table of Materials) at a concentration of 2 mg/mL in embryo media.</li>
	<li>Label a 1.5 mL microcentrifuge tube and a 2 mL gas chromatograph vial for each sample. Label additional 2 mL gas chromatograph vials for the ethanol standards described below as well as air, water, and the 5 M NaCl/protease cocktail blanks.</li>
	<li>Set up three p200 micropipettors two set to 50 &#956;L, and the third set to 200 &#956;L; a p1000 micropipettor set to 450 &#956;L and a p2 micropipettor set to 2 &#956;L. For the glass pipettes used to transfer embryos from the Petri dishes to the 1.5 mL microcentrifuge tubes, quickly pass the pipette tip through a flame to smooth the edges to not damage the embryos when drawing them into the pipette.</li>
</ol>
5. Processing embryos for head space gas chromatography
NOTE: Both embryos in their chorions and those previously removed from their chorions are treated the same for consistency in the calculation of dilution factors.
<ol>
	<li>Using the two p200 micropipettors set to 50 &#956;L, draw 50 &#956;L of the protease cocktail solution into one and draw 50 &#956;L of water into the second.</li>
	<li>Using the glass pipette and micropipettor, quickly place 10 embryos (from step 3.2) in a 1.5 mL microcentrifuge tube and close the cap (as described in Figure 2A). Repeat for all samples to be tested, controls, and ethanol treated embryos.</li>
	<li>Using the p200 micropipettor set to 200 &#956;L, quickly open the cap and remove all residual embryo media (as described in Figure 2C). Quickly place the pipette containing 50 &#956;L of water in the tube and rapidly but gently (to not damage the embryos) add, then remove the water. Quickly add 50 &#956;L of the protease cocktail solution from the waiting pipettor and close the tube cap (Figure 2D). 
	NOTE: Perform this process one sample at a time.</li>
	<li>Let the sample sit at RT for 10 min to allow the protease cocktail to degrade the chorion. Then, quickly add 450 &#956;L of 5 M NaCl and close the tube cap (Figure 2E). Vortex the samples for 10 min. To speed up the process, set up the mixer to vortex multiple samples at the same time. 
	NOTE: Add a small amount (~100 &#956;L) of a silica bead mixture (2 sizes, 0.5 mm and 1 mm bead) to any tube with embryos older than 24 hpf. In older embryos, the notochord will remain intact regardless of how long they are homogenized.</li>
	<li>After homogenizing for 10 min, quickly remove 2 &#956;L of homogenized embryo supernatant and add to a gas chromatograph vial. Quickly seal the vial with the polytetrafluoroethylene cap.</li>
</ol>
6. Preparing media and ethanol standards
<ol>
	<li>To prepare the media standards, dilute the media by a factor of 10 with a 5 M NaCl/protease cocktail solution. Add 2 &#956;L of each sample to a gas chromatograph vial and seal with a polytetrafluoroethylene cap.</li>
	<li>To prepare the ethanol standards, create a serial dilution of 100% ethanol in 5 M NaCl/protease cocktail solution to the following concentrations: 0.3125, 0.625, 1.25, 2.5, 5, 10, 20, and 40 mM. Add 2 &#956;L of each standard to a gas chromatograph vial and seal with a polytetrafluoroethylene cap.</li>
</ol>
7. Preparing the head space gas chromatograph
NOTE: This setup and protocol may need to be changed depending on the gas chromatograph used. Head space gas chromatography is used to quantify ethanol levels, not for separation.
<ol>
	<li>Set the heater for the autosampler to 58 &#176;C and turn on. Allow the heater to reach 58 &#176;C, and turn on the air and hydrogen gas lines feeding the gas chromatograph (for flame ionization used to quantify the ethanol). 
	NOTE: The psi should be set to properly operate the chromatograph according to the manufacturer&#39;s specifications. Make sure the helium line is on and set to the proper psi.</li>
	<li>For the septum, make sure the number of samples injected is not &#62;100. For the injection fiber, make sure the number of samples injected is not &#62;500. 
	NOTE: If either is over the amount of injections, they will need to be changed before running the test samples.</li>
	<li>Turn on the analysis software and make sure the workstation is set up properly. Store all data according to lab/department standards. Create a new sample list for the samples to be run.</li>
	<li>Initiate the startup method and wait for the flame ionization detector to stabilize before running the samples. Once stable, run the software startup method to clean the fiber.</li>
</ol>
8. Sample measurements using head space gas chromatography
<ol>
	<li>Once the startup method is complete, create 1 new line per sample in the sample list. From the methods menu in the analysis software, load 436 Current spme ethanol 2013 3min absorb 2_5min rg run.METH for each sample on the sample list.
	<ol>
		<li>Fill out the sample list, starting with the air, water, 5 M NaCl/protease cocktail blanks. Then enter in the standards in order, from 0.3125 to 40 mM. Follow the standards with a second round of the air, water, and 5 M NaCl/protease cocktail blanks.</li>
		<li>Enter all homogenized embryo supernatant samples to be tested, from lowest to highest predicted ethanol concentration. End by entering a third and final round of the air, water, and 5 M NaCl/protease cocktail blanks.</li>
	</ol>
	</li>
	<li>Add the gas chromatograph vials to the autosampler in the order in which the samples were entered. Allow samples to warm for 10&#8211;15 min. Start the sample runs in the software.</li>
	<li>After all the samples and the final blanks have run, activate the shutdown method in the software by adding a final sample in the sample list and running standby.METH. Back up all data acquired during the sample runs. Turn off the equipment in the following order: autosampler heater, hydrogen tank and, only after the chromatograph temperature reaches 30 &#176;C, the air tank. Leave the helium tank on to preserve the wax column.</li>
</ol>
9. Sample ethanol peak integration and sample concentration analysis
NOTE: All values from 9.3 on were calculated in an excel file that all equations prefilled.
<ol>
	<li>Once the shutdown method is complete, click on Open chromatogram. Open the folder containing the results. Samples are automatically integrated in this program.</li>
	<li>In the results, make sure the correct peaks have been integrated (ethanol peaks between 2 and 2.5). Once all samples have been confirmed, print or export the results.</li>
	<li>Plot the peak height of the ethanol standards on a graph. Calculate the slope, Y intercept and R2 values for the ethanol standards (R2 should be &#62;0.99). 
	NOTE: These values will be used to determine the ethanol concentration from the sample peak height.</li>
	<li>For each sample, subtract the peak height of the sample from the Y intercept of the ethanol standards. Divide this value by the slope of the ethanol standards to obtain the ethanol value for each sample in the GC vials.</li>
</ol>
<img alt="Equation 3" src="/files/ftp_upload/60766/60766equ03.jpg" />
<ol start="5">
	<li>To calculate the ethanol concentration in the embryos, first calculate the dilution factor for each sample. Take the volume measure calculated in step 2.3 of this protocol and divide it by the volume measure plus 500 &#956;L. This represents the 5 M NaCl/protease cocktail solution added to each sample during embryo processing (steps 5.3 and 5.4).</li>
</ol>
<img alt="Equation 4" src="/files/ftp_upload/60766/60766equ04.jpg" />
<ol start="6">
	<li>Using this sample dilution factor, multiply by the sample ethanol value for each sample. The results will be in mM concentration. Calculate media reference samples by multiplying the media ethanol value by the dilution factor of 10.</li>
</ol>
<img alt="Equation 5" src="/files/ftp_upload/60766/60766equ05.jpg" />
<img alt="Equation 6" src="/files/ftp_upload/60766/60766equ06.jpg" />

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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+and+Validation+of+a+Method+for+Alcohol+Analysis+in+Brain+Tissue+by+Headspace+Gas+Chromatography+with+Flame+Ionization+Detector.">Development and Validation of a Method for Alcohol Analysis in Brain Tissue by Headspace Gas Chromatography with Flame Ionization Detector.</a> Journal of Analytical Toxicology. 40 (8), 653-658 (2016).</li><li>Schlatter, J., Chiadmi, F., Gandon, V., Chariot, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+determination+of+methanol,+acetaldehyde,+acetone,+and+ethanol+in+human+blood+by+gas+chromatography+with+flame+ionization+detection.">Simultaneous determination of methanol, acetaldehyde, acetone, and ethanol in human blood by gas chromatography with flame ionization detection.</a> Human and Experimental Toxicology. 33 (1), 74-80 (2013).</li><li>Schier, C. J., Mangieri, R. A., Dilly, G. A., Gonzales, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microdialysis+of+ethanol+during+operant+ethanol+self-administration+and+ethanol+determination+by+gas+chromatography.">Microdialysis of ethanol during operant ethanol self-administration and ethanol determination by gas chromatography.</a> Journal of Visualized Experiments. (67), e4142 (2012).</li><li>Adalsteinsson, E., Sullivan, E. V., Mayer, D., Pfefferbaum, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+quantification+of+ethanol+kinetics+in+rat+brain.">In vivo quantification of ethanol kinetics in rat brain.</a> Neuropsychopharmacology. 31 (12), 2683-2691 (2006).</li><li>Quertemont, E., Green, H. L., Grant, K. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Brain+ethanol+concentrations+and+ethanol+discrimination+in+rats:+effects+of+dose+and+time.">Brain ethanol concentrations and ethanol discrimination in rats: effects of dose and time.</a> Psychopharmacology. 168 (3), 262-270 (2003).</li><li>Flentke, G. R., Klinger, R. H., Tanguay, R. L., Carvan, M. J., Smith, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+evolutionarily-conserved+mechanism+of+calcium-dependent+neurotoxicity.">An evolutionarily-conserved mechanism of calcium-dependent neurotoxicity.</a> Alcoholism: Clinical and Experimental Research. 38 (5), 1255-1265 (2014).</li><li>Reimers, M. J., Flockton, A. R., Tanguay, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ethanol-+and+acetaldehyde-mediated+developmental+toxicity+in+zebrafish.">Ethanol- and acetaldehyde-mediated developmental toxicity in zebrafish.</a> Neurotoxicology and Teratology. 26 (6), 769-781 (2004).</li><li>Zhang, C., Ojiaku, P., Cole, G. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Forebrain+and+hindbrain+development+in+zebrafish+is+sensitive+to+ethanol+exposure+involving+agrin,+Fgf,+and+sonic+hedgehog+function.">Forebrain and hindbrain development in zebrafish is sensitive to ethanol exposure involving agrin, Fgf, and sonic hedgehog function.</a> Birth Defects Research Part A: Clinical and Molecular Teratology. 97 (1), 8-27 (2013).</li></ol>

The authors have nothing to disclose.

As a developmental model system, zebrafish are ideally suited to study the impact of environmental factors on development. They produce large numbers of externally fertilized embryos, which allows for precise timing and dosage paradigms in ethanol studies. This, combined with the live imaging capabilities and the genetic and developmental conservation with humans, make zebrafish a powerful model system for teratology studies. Described is a protocol for measuring embryonic ethanol concentrations in developing zebrafish e...

Fetal Alcohol Spectrum Disorders (FASD) describes a wide array of neurological impairments and craniofacial dysmorphologies associated with embryonic ethanol exposure1. Multiple factors, including timing and dosage of ethanol exposure and genetic background, contribute to the variation of FASD2,3. In humans, the complex relationship of these variables makes studying and understanding the etiology of FASD challenging. Animal models have proven crucial in developing our understanding of the mechanistic basis of ethanol teratogenicity. A wide variety of animal model systems has been used to study multiple aspects of FASD and results have been remarkably consistent with what is found in exposure in humans4. Rodent model systems are used to examine many aspects of FASD, with mice being the most common5,6,7. The majority of this work has focused on developmental defects to early ethanol exposure8, though later exposure to ethanol has been shown to cause developmental anomalies as well9. Moreover, the genetic capabilities of mice have greatly aided in our ability to probe the genetic underpinnings of FASD10,11. These studies in mice strongly suggest that there are gene-ethanol interactions with the sonic hedgehog pathway, retinoic acid signaling, Superoxide dismutase, Nitric oxide synthase I, Aldh2 and Fancd28,10,11,12,13,14,15,16,17,18,19,20,21. These studies show that animal models are critical to advancing our understanding of FASD and its underlying mechanisms.
The zebrafish has emerged as a powerful model system to examine many aspects of ethanol teratogenesis22,23. Due to their external fertilization, high fecundity, genetic tractability, and live imaging capabilities, zebrafish are ideally suited to study factors such as timing, dosage, and genetics of ethanol teratogenesis. Ethanol can be administered to precisely staged embryos and the embryos can then be imaged to examine the direct impact of ethanol during developmental processes. This work can be related directly to humans, because the genetic programs of development are highly conserved between zebrafish and humans and can therefore help guide FASD human studies24. While zebrafish have been used to examine ethanol teratogenesis, a lack of consensus in reporting embryonic ethanol concentrations makes comparison to humans difficult25. In mammalian systems, blood-alcohol levels correlate directly to tissue ethanol levels26. Many of the zebrafish studies treat embryos before complete formation of their circulatory system. With no maternal sample to examine, a process to assess ethanol concentrations is required to quantify ethanol levels within the embryo. Here we describe a process to quantify ethanol concentrations in a developing zebrafish embryo using head space gas chromatography.

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quantification of ethanol levels in zebrafish embryos using head space gas chromatography

Fetal Alcohol Spectrum Disorders (FASD) describe a highly variable continuum of ethanol-induced developmental defects, including facial dysmorphologies and neurological impairments. With a complex pathology, FASD affects approximately 1 in 100 children born in the United States each year. Due to the highly variable nature of FASD, animal models have proven critical in our current mechanistic understanding of ethanol-induced development defects. An increasing number of laboratories has focused on using zebrafish to examine ethanol-induced developmental defects. Zebrafish produce large numbers of externally fertilized, genetically tractable, translucent embryos. This allows researchers to precisely control timing and dosage of ethanol exposure in multiple genetic contexts and quantify the impact of embryonic ethanol exposure through live imaging techniques. This, combined with the high degree of conservation of both genetics and development with humans, has proven zebrafish to be a powerful model in which to study the mechanistic basis of ethanol teratogenicity. However, ethanol exposure regimens have varied between different zebrafish studies, which has confounded the interpretation of zebrafish data across these studies. Here is a protocol to quantify ethanol concentrations in zebrafish embryos using head space gas chromatography.

Blood ethanol levels cannot be determined in early embryonic zebrafish, because they lack a fully formed circulatory system. To determine the level of ethanol concentration in the zebrafish embryos, the ethanol levels are measured directly from homogenized embryonic tissue. To properly measure the embryonic ethanol concentrations, the embryonic volume has to be taken into account. The embryo (yolk attached) sits inside the chorion (eggshell) surrounded by extraembryonic fluid (Figure 1). Any...

Watch this Scientific Journal Video about Quantification of Ethanol Levels in Zebrafish Embryos Using Head Space Gas Chromatography at JoVE.com

Quantification of Ethanol Levels in Zebrafish Embryos Using Head Space Gas Chromatography

This work describes a protocol to quantify ethanol levels in a zebrafish embryo using head space gas chromatography from proper exposure methods to embryo processing and ethanol analysis.

Fetal Alcohol Spectrum Disorders (FASD) describe a highly variable continuum of ethanol-induced developmental defects, including facial dysmorphologies ...

quantification-ethanol-levels-zebrafish-embryos-using-head-space-gas

Universidad de Cartagena UDEC

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Developmental Biology

5.6K Views.  University of Louisville. This work describes a protocol to quantify ethanol levels in a zebrafish embryo using head space gas chromatography from proper exposure methods to embryo processing and ethanol analysis.

Video: Quantification of Ethanol Levels in Zebrafish Embryos Using Head Space Gas Chromatography

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion (FDAP)

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological systems. In FDAP experiments, the clearance of proteins within living organisms can be measured. A protein of interest is tagged with a photoconvertible fluorescent protein, expressed in vivo and photoconverted, and the decrease in the photoconverted signal over time is monitored. The data is then fitted with an appropriate clearance model to determine the protein half-life. Importantly, the clearance kinetics of protein populations in different compartments of the organism can be examined separately by applying compartmental masks. This approach has been used to determine the intra- and extracellular half-lives of secreted signaling proteins during zebrafish development. Here, we describe a protocol for FDAP experiments in zebrafish embryos. It should be possible to use FDAP to determine the clearance kinetics of any taggable protein in any optically accessible organism.

Measuring Protein Stability in Living Zebrafish Embryos Using Fluorescence Decay After Photoconversion FDAP

Protein levels in cells and tissues are often tightly regulated by the balance of protein production and clearance. Using Fluorescence Decay After Photoconversion (FDAP), the clearance kinetics of proteins can be experimentally measured in vivo.

Protein stability influences many aspects of biology, and measuring the clearance kinetics of proteins can provide important insights into biological ...

Mechanical Vessel Injury in Zebrafish Embryos

Zebrafish (Danio rerio) embryos have proven to be a powerful model for studying a variety of developmental and disease processes. External development and optical transparency make these embryos especially amenable to microscopy, and numerous transgenic lines that label specific cell types with fluorescent proteins are available, making the zebrafish embryo an ideal system for visualizing the interaction of vascular, hematopoietic, and other cell types during injury and repair in vivo. Forward and reverse genetics in zebrafish are well developed, and pharmacological manipulation is possible. We describe a mechanical vascular injury model using micromanipulation techniques that exploits several of these features to study responses to vascular injury including hemostasis and blood vessel repair. Using a combination of video and timelapse microscopy, we demonstrate that this method of vascular injury results in measurable and reproducible responses during hemostasis and wound repair. This method provides a system for studying vascular injury and repair in detail in a whole animal model.

This article describes a method for creating a mechanical vessel injury in zebrafish embryos. This injury model provides a platform for studying hemostasis, injury-related inflammation, and wound healing in an organism ideally suited for real-time microscopy.

Zebrafish (Danio rerio) embryos have proven to be a powerful model for studying a variety of developmental and disease processes.  External development ...

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration have been thoroughly analyzed in vitro. However, in vivo cell migration somehow differs from in vitro migration, and has proven more difficult to analyze, being less accessible to direct observation and manipulation. This protocol uses the migration of the prospective prechordal plate in the early zebrafish embryo as a model system to study the function of candidate genes in cell migration. Prechordal plate progenitors form a group of cells which, during gastrulation, undergoes a directed migration from the embryonic organizer to the animal pole of the embryo. The proposed protocol uses cell transplantation to create mosaic embryos. This offers the combined advantages of labeling isolated cells, which is key to good imaging, and of limiting gain/loss of function effects to the observed cells, hence ensuring cell-autonomous effects. We describe here how we assessed the function of the TORC2 component Sin1 in cell migration, but the protocol can be used to analyze the function of any candidate gene in controlling cell migration in vivo.

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Combining cell transplantation, cytoskeletal labeling and loss/gain of function approaches, this protocol describes how the migrating zebrafish prospective prechordal plate can be used to analyze the function of a candidate gene in in vivo cell migration.

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration ...

Isolation and Characterization of Single Cells from Zebrafish Embryos

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been described at the morphological level, little is known about the molecular basis of cellular changes that occur as the organism develops. With recent advancements in microfluidics and multiplexing technologies, it is now possible to characterize gene expression in single cells. This allows for investigation of heterogeneity between individual cells of specific cell populations to identify and classify cell subtypes, characterize intermediate states that occur during cell differentiation, and explore differential cellular responses to stimuli. This study describes a protocol to isolate viable, single cells from zebrafish embryos for high throughput multiplexing assays. This method may be rapidly applied to any zebrafish embryonic cell type with fluorescent markers. An extension of this method may also be used in combination with high throughput sequencing technologies to fully characterize the transcriptome of single cells. As proof of principle, the relative abundance of cardiac differentiation markers was assessed in isolated, single cells derived from nkx2.5 positive cardiac progenitors. By evaluation of gene expression at the single cell level and at a single time point, the data support a model in which cardiac progenitors coexist with differentiating progeny. The method and work flow described here is broadly applicable to the zebrafish research community, requiring only a labeled transgenic fish line and access to microfluidics technologies.

This protocol describes a method for isolating single cells from zebrafish embryos, enriching for cells of interest, capturing zebrafish cells in microfluidic based single cell multiplex systems, and assessing gene expression from single cells.

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been ...

Development of an Ethanol-induced Fibrotic Liver Model in Zebrafish to Study Progenitor Cell-mediated Hepatocyte Regeneration

Sustained liver fibrosis with continuation of extracellular matrix (ECM) protein build-up results in the loss of cellular competency of the liver, leading to cirrhosis with hepatocellular dysfunction. Among multiple hepatic insults, alcohol abuse can lead to significant health problems including liver failure and hepatocellular carcinoma. Nonetheless, the identity of endogenous cellular sources that regenerate hepatocytes in response to alcohol has not been properly investigated. Moreover, few studies have effectively modeled hepatocyte regeneration upon alcohol-induced injury. We recently reported on establishing an ethanol (EtOH)-induced fibrotic liver model in zebrafish in which hepatic progenitor cells (HPCs) gave rise to hepatocytes upon near-complete hepatocyte loss in the presence of fibrogenic stimulus. Furthermore, through chemical screens using this model, we identified multiple small molecules that enhance hepatocyte regeneration. Here we describe in detail the procedures to develop an EtOH-induced fibrotic liver model and to perform chemical screens using this model in zebrafish. This protocol will be a critical tool to delineate the molecular and cellular mechanisms of how hepatocyte regenerates in the fibrotic liver. Furthermore, these methods will facilitate potential discovery of novel therapeutic strategies for chronic liver disease in vivo.

Sustained fibrosis with deposition of excessive extracellular matrix proteins leads to cirrhosis. Alcohol abuse is one of the main causes of severe liver disease. We established an ethanol-induced zebrafish fibrotic liver model to study the mechanisms and strategies of promoting hepatocyte regeneration upon alcohol-induced injury.

Sustained liver fibrosis with continuation of extracellular matrix (ECM) protein build-up results in the loss of cellular competency of the liver, leading ...

Biosensing Motor Neuron Membrane Potential in Live Zebrafish Embryos

The protocols described here are designed to allow researchers to study cell communication without altering the integrity of the environment in which the cells are located. Specifically, they have been developed to analyze the electrical activity of excitable cells, such as spinal neurons. In such a scenario, it is crucial to preserve the integrity of the spinal cell, but it is also important to preserve the anatomy and physiological shape of the systems involved. Indeed, the comprehension of the manner in which the nervous system-and other complex systems-works must be based on a systemic approach. For this reason, the live zebrafish embryo was chosen as a model system, and the spinal neuron membrane voltage changes were evaluated without interfering with the physiological conditions of the embryos. 
Here, an approach combining the employment of zebrafish embryos with a FRET-based biosensor is described. Zebrafish embryos are characterized by a very simplified nervous system and are particularly suited for imaging applications thanks to their transparency, allowing for the employment of fluorescence-based voltage indicators at the plasma membrane during zebrafish development. The synergy between these two components makes it possible to analyze the electrical activity of the cells in intact living organisms, without perturbing the physiological state. Finally, this non-invasive approach can co-exist with other analyses (e.g., spontaneous movement recordings, as shown here).

Protocols described here allow for the study of the electrical properties of excitable cells in the most non-invasive physiological conditions by employing zebrafish embryos in an in vivo system together with a fluorescence resonance energy transfer (FRET)-based genetically encoded voltage indicator (GEVI) selectively expressed in the cell type of interest.

The protocols described here are designed to allow researchers to study cell communication without altering the integrity of the environment in which the ...

Induction of Hypoxia in Living Frog and Zebrafish Embryos

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and zebrafish embryos. Our system comprises a chamber featuring a simple setup that is nevertheless robust to induce and maintain a specific oxygen concentration and temperature in any experimental solution of choice. The presented system is very cost-effective but highly functional, it allows induction and sustainment of hypoxia for direct experiments in vivo and for various time periods up to 48 h.
To monitor and study the effects of hypoxia, we have employed two methods - measurement of levels of hypoxia-inducible factor 1alpha (HIF-1&#945;) in whole embryos or specific tissues and determination of retinal stem cell proliferation by 5-ethynyl-2&#8242;-deoxyuridine (EdU) incorporation into the DNA. HIF-1&#945; levels can serve as a general hypoxia marker in the whole embryo or tissue of choice, here embryonic retina. EdU incorporation into the proliferating cells of embryonic retina is a specific output of hypoxia induction. Thus, we have shown that hypoxic embryonic retinal progenitors decrease proliferation within 1 h of incubation under 5% oxygen of both frog and zebrafish embryos.
Once mastered, our setup can be employed for use with small aquatic model organisms, for direct in vivo experiments, any given time period and under normal, hypoxic or hyperoxic oxygen concentration or under any other given gas mixture.

We introduce a novel hypoxic chamber system for use with aquatic organisms such as frog and zebrafish embryos. Our system is simple, robust, cost-effective and allows the induction and sustainment of hypoxia in vivo and for up to 48 h. We present 2 reproducible methods to monitor the effectiveness of hypoxia.

Here, we introduce a novel system for hypoxia induction, which we developed to study the effects of hypoxia in aquatic organisms such as frog and ...

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling processes of excitable neuronal and muscular cells and many other biological processes, such as embryonic developmental patterning. However, there is a need for in vivo electrical activity monitoring in vertebrate embryogenesis. The advances of genetically encoded fluorescent voltage indicators (GEVIs) have made it possible to provide a solution for this challenge. Here, we describe how to create a transgenic voltage indicator zebrafish using the established voltage indicator, ASAP1 (Accelerated Sensor of Action Potentials 1), as an example. The Tol2 kit and a ubiquitous zebrafish promoter, ubi, were chosen in this study. We also explain&#160;the processes of Gateway site-specific cloning, Tol2 transposon-based zebrafish transgenesis, and the imaging process for early-stage fish embryos and fish tumors using regular epifluorescent microscopes. Using this fish line, we found that there are cellular electric voltage changes during zebrafish embryogenesis, and fish larval movement. Furthermore, it was observed that in a few zebrafish malignant peripheral nerve sheath tumors, the tumor cells were generally polarized compared to the surrounding normal tissues.

Here, we show the process of creating a cellular electric voltage reporter zebrafish line to visualize embryonic development, movement, and fish tumor cells in vivo.

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling ...

Preparation of Drosophila Larval Samples for Gas Chromatography-Mass Spectrometry (GC-MS)-based Metabolomics

Recent advances in the field of metabolomics have established the fruit fly Drosophila melanogaster as a powerful genetic model for studying animal metabolism. By combining the vast array of Drosophila genetic tools with the ability to survey large swaths of intermediary metabolism, a metabolomics approach can reveal complex interactions between diet, genotype, life-history events, and environmental cues. In addition, metabolomics studies can discover novel enzymatic mechanisms and uncover previously unknown connections between seemingly disparate metabolic pathways. In order to facilitate more widespread use of this technology among the Drosophila community, here we provide a detailed protocol that describes how to prepare Drosophila larval samples for gas chromatography-mass spectrometry (GC-MS)-based metabolomic analysis. Our protocol includes descriptions of larval sample collection, metabolite extraction, chemical derivatization, and GC-MS analysis. Successful completion of this protocol will allow users to measure the relative abundance of small polar metabolites, including amino acids, sugars, and organic acids involved in glycolysis and the TCA cycles.

Preparation of Drosophila Larval Samples for Gas Chromatography-Mass Spectrometry GC-MS-based Metabolomics

This protocol describes how to prepare Drosophila larvae for GC-MS-based metabolomic analysis.

Recent advances in the field of metabolomics have established the fruit fly Drosophila melanogaster as a powerful genetic model for studying animal ...

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 ...

Quantification of Ethanol Levels in Zebrafish Embryos Using Head Space Gas Chromatography

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Preparation of Drosophila Larval Samples for Gas Chromatography-Mass Spectrometry (GC-MS)-based Metabolomics

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