This work was partially supported by the National Institutes of Health (GM093282 to P.W.; 3U54AI057158-06S1 to R.D.S.) and startup funds from Albert Einstein College of Medicine.

1. Egg Collection and Dechorionation
<ol>
	<li>Collect and transfer zebrafish eggs to 35mm petri dish, remove as much water as possible and then add 1 mg/ml Pronease E in E3 embryo medium (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2&middot;2H2O, 0.33 mM MgSO4, pH = 7.4) to digest the chorion.</li>
	<li>After 3-5 minutes, merge the dish into a beaker filled with fish water (60 mg "instant Ocean" per liter distilled H2O), and gently transfer the eggs to the beaker and allow the eggs to "fall" into water.</li>
	<li>Rinse the eggs with fish water three times. Most eggs will be released from their chorion.</li>
	<li>Using a fire-polished glass Pasteur pipette, transfer the dechorionated eggs to agarose-coated petri dishes filled with E3 embryo medium.</li>
</ol>
2. Microinjection with GDP-FucAl
<ol>
	<li>Prepare the injection dishes following the reported protocol7.</li>
	<li>Transfer dechorionated eggs into injection dishes filled with E3 embryo medium.</li>
	<li>Prepare the needle, and load with 2 &mu;L injection solution. This solution contains 20 mM GDP-FucAl synthesized chemoenzymatically8 and either Alexa Fluor 594-dextran (5% w/v) as a tracer or phenol red loading dye (0.1% w/v) in 0.2 M KCl. As a negative control, replace GDP-FucAl with GDP-fucose in injection solution.</li>
	<li>Break the needle and adjust the injection pressure and duration to yield a 1 nL drop9.</li>
	<li>Inject the eggs with 1 nL of either solution.</li>
	<li>Transfer the eggs into agarose-coated petri dishes filled with E3 embryo medium.</li>
	<li>Incubate eggs at 28 &deg;C and remove the unfertilized eggs within three to four hours after fertilization.</li>
</ol>
3. BTTES-Cu(I)-catalyzed Click Chemistry Reaction
<ol>
	<li>When the embryos reach desired developmental stages (e.g. late gastrula, tissue segmentation and early larva), coat the base of a 96-well plate with agarose.</li>
	<li>Add 92 &mu;L E3 embryo medium to each well, followed by addition of 4 &mu;L Alexa Fluor-488 azide (from a 2.5 mM stock in H2O), 2 &mu;L BTTES-CuSO4 6:1 complex, and shake gently to mix.</li>
	<li>Transfer embryos into the well containing the click chemistry reagent using a fire-polished glass Pasteur pipette. Each well should contain less than five embryos.</li>
	<li>Add 2.5 &mu;L freshly prepared sodium ascorbate (from a 100 mM stock in H2O) to initiate the click reaction3. Final concentration of each reagent: Alexa Fluor-488 azide: 100 &mu;M; CuSO4: 50 &mu;M; BTTES: 300 &mu;M; sodium ascorbate: 2.5 mM.</li>
	<li>After 3 min, add 2 &mu;L bathocuproine sulphonate (50 mM stock in H2O), a biocompatible copper chelator, to quench the reaction then dilute immediately with 100 &mu;L E3 embryo medium.</li>
	<li>Transfer the embryos to a glass petri dish and wash the treated embryos 2 times with 15 mL E3 embryo medium.</li>
</ol>
4. Imaging
<ol>
	<li>Place a drop of ultralow melting point agarose (1.2% (w/v) in E3 embryo medium) on a MatTek glass bottom microwell dish, and place an embryo into the agarose drop.</li>
	<li>Position the embryos dorsally or laterally and place the dish on ice for 5 min to solidify the agarose drop. Add E3 embryo medium gently to the dish until it covers the agarose drop.</li>
	<li>Fluorescence and bright field images are acquired sequentially using a confocal microscope. All embryo images are acquired using a 5 &mu;m step interval. Composite figures are prepared using ImageJ software.</li>
</ol>
5. Representative Results
Figure 1 shows the workflow of our two-step labeling strategy. Figure 2 shows the labeling of zebrafish embryos via BTTES-mediated CuAAC at 9.5 hours post fertilization (hpf). BTTES is a tris (triazolylmethyl) amine-based ligand. It accelerates CuAAC dramatically when coordinating with the in situ generated Cu(I), and promotes the cycloaddition reaction rapidly in living systems without apparent toxicity. Immediately following a 3-min click reaction, we are able to detect robust labeling of the GDP-FucAl treated embryos (Figure 2, left panels). Only background fluorescence is detected for control embryos microinjected with GDP-fucose (Figure 2, right panels).
 <img src="/files/ftp_upload/2686/2686fig1.jpg" alt="Figure 1" /> 
Figure 1.The strategy of labeling fucosylated glycans in the enveloping layer of zebrafish embryos.
 <img src="/files/ftp_upload/2686/2686fig2.jpg" alt="Figure 2" /> 
Figure 2. In vivo imaging of fucosylated glycans during zebrafish embryogenesis via BTTES-Cu(I)-catalyzed click chemistry. One-cell stage zebrafish embryos are microinjected with a single dose of GDP-FucAl and allowed to develop to 9.5 hpf. The embryos are then reacted with Alexa Fluor 488-azide catalyzed by BTTES-Cu(I). Reacted embryos are imaged using confocal microscopy. Maximum intensity z-projection images of Alexa Fluor 488 fluorescence (upper panel); Bright field (lower panel). Scale bar: 100 &mu;m.

Troubleshooting
<table border="1">
	<tbody>
		<tr>
			<td>Problem</td>
			<td>Cause</td>
			<td>Remedy</td>
		</tr>
		<tr>
			<td>The embryos look unhealthy</td>
			<td>The microinjected solution contains contaminants</td>
			<td>The purity of the nucleotide sugars should be greater than 85%. Check your source.</td>
		</tr>
		<tr>
			<td>There is no labeling after the reaction</td>
			<td>Copper concentration is below 30 &mu;M</td>
			<td>Be careful not to dilute the reaction solution by adding excess E3 embryo medium when adding the embryos, as the reaction rate drops significantly when the copper concentration is below 30 &mu;M.</td>
		</tr>
		<tr>
			<td>The embryos die after the reaction</td>
			<td>Improper handling of the embryos</td>
			<td>Make sure the pipet is fire-polished and the reaction vessel is coated with a very thin layer of 0.5% agarose.</td>
		</tr>
		<tr>
			<td>The images look spotty</td>
			<td>The reagents are not properly mixed before adding the embryos</td>
			<td>Follow the recommended order of addition for the reagents, and make sure that the click chemistry reagents have been properly mixed before adding the embryos.</td>
		</tr>
		<tr>
			<td>The images show the embryos are damaged</td>
			<td>Vigorous shaking damages the embryos during the reaction</td>
			<td>Gently shake for less than 10 seconds once the embryos are in the solution.</td>
		</tr>
		<tr>
		 <td>&nbsp;</td>
		 <td>&nbsp;</td>
		 <td>&nbsp;</td>
	 </tr>
	</tbody>
</table>

<ol>
	<li>Laughlin, S. T., Bertozzi, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+the+glycome.">Imaging the glycome.</a> Proc. Natl. Acad. Sci. U. S. A. 106, 12-17 (2009).</li><li>Baskin, J. M., Bertozzi, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bioorthogonal+click+chemistry:+Covalent+labeling+in+living+systems.">Bioorthogonal click chemistry: Covalent labeling in living systems.</a> Qsar Comb. Sci. 26, 1211-1219 (2007).</li><li>Soriano del Amo, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biocompatible+copper(I)+catalysts+for+in+vivo+imaging+of+glycans.">Biocompatible copper(I) catalysts for in vivo imaging of glycans.</a> J. Am. Chem. Soc. 132, 16893-16899 (2010).</li><li>Chang, P. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Metabolic+labeling+of+sialic+acids+in+living+animals+with+alkynyl+sugars.">Metabolic labeling of sialic acids in living animals with alkynyl sugars.</a> Angew. Chem. Int. Ed. 48, 4030-4033 (2009).</li><li>Laughlin, S. T., Baskin, J. M., Amacher, S. L., Bertozzi, C. R. <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+imaging+of+membrane-associated+glycans+in+developing+zebrafish.">In vivo imaging of membrane-associated glycans in developing zebrafish.</a> Science. 320, 664-667 (2008).</li><li>Baskin, J. M., Dehnert, K. W., Laughlin, S. T., Amacher, S. L., Bertozzi, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualizing+enveloping+layer+glycans+during+zebrafish+early+embryogenesis.">Visualizing enveloping layer glycans during zebrafish early embryogenesis.</a> Proc. Natl. Acad. Sci. U. S. A. 107, 10360-10365 (2010).</li><li>Kemp, H. A., Carmany-Rampey, A., Moens, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generating+chimeric+zebrafish+embryos+by+transplantation.">Generating chimeric zebrafish embryos by transplantation.</a> J. Vis. Exp. , (2009).</li><li>Wang, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemoenzymatic+synthesis+of+GDP-L-fucose+and+the+Lewis+X+glycan+derivatives.">Chemoenzymatic synthesis of GDP-L-fucose and the Lewis X glycan derivatives.</a> Proc. Natl. Acad. Sci. U. S. A. 106, 16096-16101 (2009).</li><li>Rosen, J. N., Sweeney, M. F., Mably, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microinjection+of+zebrafish+embryos+to+analyze+gene+function.">Microinjection of zebrafish embryos to analyze gene function.</a> J. Vis. Exp. , (2009).</li><li>Westerfield, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> THE ZEBRAFISH BOOK, A guide for the laboratory use of zebrafish (Danio rerio). , (2007).</li><li>Kimmel, C. B., Warga, R. M., Schilling, T. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Origin+and+organization+of+the+zebrafish+fate+map.">Origin and organization of the zebrafish fate map.</a> Development. 108, 581-594 (1990).</li></ol>

No conflicts of interest declared.

Imaging biomolecules in vivo provides critical insights of their biological activities in their native environments. In this video, we demonstrate how the labeling of fucosylated glycans in the enveloping layer of zebrafish embryos is realized by microinjecting one-cell stage embryos with GDP-FucAl and a second-step fluorophore conjugation via BTTES-mediated biocompatible CuAAC3. Robust labeling can be achieved within 2-3 minutes, and the labeled glycans are detectable as early as 2.5 hpf. Importantly, a single dose of GDP-FucAl can yield a detectable alkyne-dependent fluorescent signal till 96 hpf. The same approach can be extended to multicolor time-lapse imaging experiments to monitor the trafficking of the alkyne-tagged fucosides in developing zebrafish via two sequential CuAAC. However, labeling is only achieved in the enveloping layer of the blastodisc due to the low tissue penetration depth of the click chemistry reagents. By fixing and permeabilizing the embryos metabolically treated with GDP-FucAl, we can achieve labeling of the interior body parts as well. Since transplantation can be easily performed from alkyne labeled donor embryos to unlabeled hosts between blastula stages (4 hpf) and the onset of gastrulation (5.3 hpf) following the zebrafish fate map10, 11, exploiting this manipulation may allow us to follow the allocation of these glycans in specific lineages of the embryo during development.
As many azide- and alkyne-functionalized reagents are commercially available, only metabolic manipulations are required to generate the tagged glycans in vivo for this procedure of click modification. Therefore, the chemical tools described here can be directly applied for studying other glycans, e.g. glycans containing sialic acid4 and N-acetylgalactosamine5, 6, in zebrafish embryos and in other living systems (the azide-tagged metabolic precursors of these monosaccharides are commercially available from Invitrogen).

<table border="1">
	<tbody>
		<tr>
			<td>Name of the reagent</td>
			<td>Company</td>
			<td>Catalog number</td>
		</tr>
		<tr>
			<td>Copper(II) sulfate pentahydrate</td>
			<td>Sigma-Aldrich</td>
			<td>203165</td>
		</tr>
		<tr>
			<td>Alexa Fluor 488 azide</td>
			<td>Invitrogen</td>
			<td>A10266</td>
		</tr>
		<tr>
			<td>dextran, Alexa Fluor 594</td>
			<td>Invitrogen</td>
			<td>D-22913</td>
		</tr>
		<tr>
			<td>(+)-Sodium L-ascorbate</td>
			<td>Sigma-Aldrich</td>
			<td>A7631</td>
		</tr>
		<tr>
			<td>Bathocuproinedisulfonic acid</td>
			<td>Acros Organics</td>
			<td>164060010</td>
		</tr>
		<tr>
			<td>Glass bottom microwell dish</td>
			<td>MatTek</td>
			<td>P35G-1.5-14-C</td>
		</tr>
	</tbody>
</table>

imaging glycans in zebrafish embryos by metabolic labeling and bioorthogonal click chemistry

Imaging glycans in vivo has recently been enabled using a bioorthogonal chemical reporter strategy by treating cells or organisms with azide- or alkyne-tagged monosaccharides1, 2. The modified monosaccharides, processed by the glycan biosynthetic machinery, are incorporated into cell surface glycoconjugates. The bioorthogonal azide or alkyne tags then allow covalent conjugation with fluorescent probes for visualization, or with affinity probes for enrichment and glycoproteomic analysis. This protocol describes the procedures typically used for noninvasive imaging of fucosylated glycans in zebrafish embryos, including: 1) microinjection of one-cell stage embryos with GDP-5-alkynylfucose (GDP-FucAl), 2) labeling fucosylated glycans in the enveloping layer of zebrafish embryos with azide-conjugated fluorophores via biocompatible Cu(I)-catalyzed azide-alkyne cycloaddition (CuAAC), and 3) imaging by confocal microscopy3. The method described here can be readily extended to visualize other classes of glycans, e.g. glycans containing sialic acid4 and N-acetylgalactosamine5, 6, in developing zebrafish and in other living organisms.

Watch this Scientific Journal Video about Imaging Glycans in Zebrafish Embryos by Metabolic Labeling and Bioorthogonal Click Chemistry at JoVE.com

Imaging Glycans in Zebrafish Embryos by Metabolic Labeling and Bioorthogonal Click Chemistry

A click-chemistry based method that allows for the rapid, noninvasive, and robust labeling of alkyne-tagged glycans in zebrafish embryos is described. Fucosylated glycans in the enveloping layer of zebrafish embryos in the late gastrulation stage were imaged in this study.

Imaging glycans in vivo has recently been enabled using a bioorthogonal chemical reporter strategy by treating cells or organisms with azide- or ...

imaging-glycans-zebrafish-embryos-metabolic-labeling-bioorthogonal

Nanyang Technological University

Research

JoVE Journal

Biology

13.6K Views.  Albert Einstein College of Medicine, Yeshiva University. A click-chemistry based method that allows for the rapid, noninvasive, and robust labeling of alkyne-tagged glycans in zebrafish embryos is described. Fucosylated glycans in the enveloping layer of zebrafish embryos in the late gastrulation stage were imaged in this study.

Video: Imaging Glycans in Zebrafish Embryos by Metabolic Labeling and Bioorthogonal Click Chemistry

Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo. Here we describe a method to mount zebrafish embryos for long-term imaging and demonstrate how to automate the capture of time-lapse images using a confocal microscope. We also describe a method to create controlled, precise damage to individual branches of peripheral sensory axons in zebrafish using the focused power of a femtosecond laser mounted on a two-photon microscope. The parameters for successful two-photon axotomy must be optimized for each microscope. We will demonstrate two-photon axotomy on both a custom built two-photon microscope and a Zeiss 510 confocal/two-photon to provide two examples.

Zebrafish trigeminal sensory neurons can be visualized in a transgenic line expressing GFP driven by a sensory neuron specific promoter 1. We have adapted this zebrafish trigeminal model to directly observe sensory axon regeneration in living zebrafish embryos. Embryos are anesthetized with tricaine and positioned within a drop of agarose as it solidifies. Immobilized embryos are sealed within an imaging chamber filled with phenylthiourea (PTU) Ringers. We have found that embryos can be continuously imaged in these chambers for 12-48 hours. A single confocal image is then captured to determine the desired site of axotomy. The region of interest is located on the two-photon microscope by imaging the sensory axons under low, non-damaging power. After zooming in on the desired site of axotomy, the power is increased and a single scan of that defined region is sufficient to sever the axon. Multiple location time-lapse imaging is then set up on a confocal microscope to directly observe axonal recovery from injury.

Here we describe a method for mounting zebrafish embryos for long-term imaging, two-photon imaging and tissue-damage techniques, and time-lapse confocal imaging.

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo.  ...

Generating Chimeric Zebrafish Embryos by Transplantation

One of the most powerful tools used to gain insight into complex developmental processes is the analysis of chimeric embryos. A chimera is defined as an organism that contains cells from more than one animal; mosaics are one type of chimera in which cells from more than one genotype are mixed, usually wild-type and mutant. In the zebrafish, chimeras can be readily made by transplantation of cells from a donor embryo into a host embryo at the appropriate embryonic stage. Labeled donor cells are generated by injection of a lineage marker, such as a fluorescent dye, into the one-cell stage embryo. Labeled donor cells are removed from donor embryos and introduced into unlabeled host embryos using an oil-controlled glass pipette mounted on either a compound or dissecting microscope. Donor cells can in some cases be targeted to a specific region or tissue of the developing blastula or gastrula stage host embryo by choosing a transplantation site in the host embryo based on well-established fate maps.

A step-by-step guide to generating targeted chimeric zebrafish embryos by transplantation at the blastula or gastrula stage.

One of the most powerful tools used to gain insight into complex developmental processes is the analysis of chimeric embryos. A chimera is defined as an ...

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating cells. Cell cycle division is an integral part of growth and reproduction and deregulation of key cell cycle components have been implicated in the precipitating events of carcinogenesis 1,2. Molecular agents in anti-cancer therapies frequently target biological pathways responsible for the regulation and coordination of cell cycle division 3. Although cell cycle kinetics tend to vary according to cell type, the distribution of cells amongst the four stages of the cell cycle is rather consistent within a particular cell line due to the consistent pattern of mitogen and growth factor expression. Genotoxic events and other cellular stressors can result in a temporary block of cell cycle progression, resulting in arrest or a temporary pause in a particular cell cycle phase to allow for instigation of the appropriate response mechanism. 
The ability to experimentally observe the behavior of a cell population with reference to their cell cycle progression stage is an important advance in cell biology. Common procedures such as mitotic shake off, differential centrifugation or flow cytometry-based sorting are used to isolate cells at specific stages of the cell cycle 4-6. These fractionated, cell cycle phase-enriched populations are then subjected to experimental treatments. Yield, purity and viability of the separated fractions can often be compromised using these physical separation methods. As well, the time lapse between separation of the cell populations and the start of experimental treatment, whereby the fractionated cells can progress from the selected cell cycle stage, can pose significant challenges in the successful implementation and interpretation of these experiments.
Other approaches to study cell cycle stages include the use of chemicals to synchronize cells. Treatment of cells with chemical inhibitors of key metabolic processes for each cell cycle stage are useful in blocking the progression of the cell cycle to the next stage. For example, the ribonucleotide reductase inhibitor hydroxyurea halts cells at the G1/S juncture by limiting the supply of deoxynucleotides, the building blocks of DNA. Other notable chemicals include treatment with aphidicolin, a polymerase alpha inhibitor for G1 arrest, treatment with colchicine and nocodazole, both of which interfere with mitotic spindle formation to halt cells in M phase and finally, treatment with the DNA chain terminator 5-fluorodeoxyridine to initiate S phase arrest 7-9. Treatment with these chemicals is an effective means of synchronizing an entire population of cells at a particular phase. With removal of the chemical, cells rejoin the cell cycle in unison. Treatment of the test agent following release from the cell cycle blocking chemical ensures that the drug response elicited is from a uniform, cell cycle stage-specific population. However, since many of the chemical synchronizers are known genotoxic compounds, teasing apart the participation of various response pathways (to the synchronizers vs. the test agents) is challenging. 
Here we describe a metabolic labeling method for following a subpopulation of actively cycling cells through their progression from the DNA replication phase, through to the division and separation of their daughter cells. Coupled with flow cytometry quantification, this protocol enables for measurement of kinetic progression of the cell cycle in the absence of either mechanically- or chemically- induced cellular stresses commonly associated with other cell cycle synchronization methodologies 10. In the following sections we will discuss the methodology, as well as some of its applications in biomedical research.

Tracking subtle changes in the progression and kinetics of cell cycle stages can be accomplished by use of a combination of metabolic labeling of nucleic acids with BrdU and total genomic DNA staining via Propidium Iodide. This method avoids the need of chemical synchronization of cycling cells, thereby preventing the introduction of non-specific DNA damage, which in turn affects cell cycle progression.

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating ...

Metabolic Profile Analysis of Zebrafish Embryos

A growing goal in the field of metabolism is to determine the impact of genetics on different aspects of mitochondrial function. Understanding these relationships will help to understand the underlying etiology for a range of diseases linked with mitochondrial dysfunction, such as diabetes and obesity. Recent advances in instrumentation, has enabled the monitoring of distinct parameters of mitochondrial function in cell lines or tissue explants. Here we present a method for a rapid and sensitive analysis of mitochondrial function parameters in vivo during zebrafish embryonic development using the Seahorse bioscience XF 24 extracellular flux analyser. This protocol utilizes the Islet Capture microplates where a single embryo is placed in each well, allowing measurement of bioenergetics, including: (i) basal respiration; (ii) basal mitochondrial respiration (iii) mitochondrial respiration due to ATP turnover; (iv) mitochondrial uncoupled respiration or proton leak and (iv) maximum respiration. Using this approach embryonic zebrafish respiration parameters can be compared between wild type and genetically altered embryos (mutant, gene over-expression or gene knockdown) or those manipulated pharmacologically. It is anticipated that dissemination of this protocol will provide researchers with new tools to analyse the genetic basis of metabolic disorders in vivo in this relevant vertebrate animal model.

Zebrafish represent a powerful vertebrate model that has been under-utilised for metabolic studies. Here we describe a rapid way to measure the in vivo metabolic profile of developing zebrafish that allows the comparison of different mitochondrial function parameters between genetically or pharmacologically manipulated embryos, thereby increasing the applicability of this organism.

A growing goal in the field of metabolism is to determine the impact of genetics on different aspects of mitochondrial function. Understanding these ...

Metabolic Labeling of Leucine Rich Repeat Kinases 1 and 2 with Radioactive Phosphate

Leucine rich repeat kinases 1 and 2 (LRRK1 and LRRK2) are paralogs which share a similar domain organization, including a serine-threonine kinase domain, a Ras of complex proteins domain (ROC), a C-terminal of ROC domain (COR), and leucine-rich and ankyrin-like repeats at the N-terminus. The precise cellular roles of LRRK1 and LRRK2 have yet to be elucidated, however LRRK1 has been implicated in tyrosine kinase receptor signaling1,2, while LRRK2 is implicated in the pathogenesis of Parkinson's disease3,4. In this report, we present a protocol to label the LRRK1 and LRRK2 proteins in cells with 32P orthophosphate, thereby providing a means to measure the overall phosphorylation levels of these 2 proteins in cells. In brief, affinity tagged LRRK proteins are expressed in HEK293T cells which are exposed to medium containing 32P-orthophosphate. The 32P-orthophosphate is assimilated by the cells after only a few hours of incubation and all molecules in the cell containing phosphates are thereby radioactively labeled. Via the affinity tag (3xflag) the LRRK proteins are isolated from other cellular components by immunoprecipitation. Immunoprecipitates are then separated via SDS-PAGE, blotted to PVDF membranes and analysis of the incorporated phosphates is performed by autoradiography (32P signal) and western detection (protein signal) of the proteins on the blots. The protocol can readily be adapted to monitor phosphorylation of any other protein that can be expressed in cells and isolated by immunoprecipitation.

Leucine rich repeat kinases 1 and 2 (LRRK1 and LRRK2) are multidomain proteins which encode both GTPase and kinase domains and which are phosphorylated in cells. Here, we present a protocol to label LRRK1 and LRRK2 in cells with 32P orthophosphate, thereby providing a means to measure their overall cellular phophorylation levels.

Leucine rich repeat kinases 1 and 2 (LRRK1 and LRRK2) are paralogs which share a similar domain organization, including a serine-threonine kinase domain, ...

Analysis of Oxidative Stress in Zebrafish Embryos

High levels of reactive oxygen species (ROS) may cause a change of cellular redox state towards oxidative stress condition. This situation causes oxidation of molecules (lipid, DNA, protein) and leads to cell death. Oxidative stress also impacts the progression of several pathological conditions such as diabetes, retinopathies, neurodegeneration, and cancer. Thus, it is important to define tools to investigate oxidative stress conditions not only at the level of single cells but also in the context of whole organisms. Here, we consider the zebrafish embryo as a useful in vivo system to perform such studies and present a protocol to measure in vivo oxidative stress. Taking advantage of fluorescent ROS probes and zebrafish transgenic fluorescent lines, we develop two different methods to measure oxidative stress in vivo: i) a &#8220;whole embryo ROS-detection method&#8221; for qualitative measurement of oxidative stress and ii) a &#8220;single-cell&#160;ROS detection method&#8221; for quantitative measurements of oxidative stress. Herein, we demonstrate the efficacy of these procedures by increasing oxidative stress in tissues by oxidant agents and physiological or genetic methods. This protocol is amenable for forward genetic screens and it will help address cause-effect relationships of ROS in animal models of oxidative stress-related pathologies such as neurological disorders and cancer.

Here we report a protocol to measure oxidative stress in living zebrafish embryos. This procedure allows reactive oxygen species (ROS) detection in both whole embryo tissues and single-cell populations. This protocol will accomplish both qualitative and quantitative analyses.

High levels of reactive oxygen species (ROS) may cause a change of cellular redox state towards oxidative stress condition. This situation causes ...

Semi-automated Imaging of Tissue-specific Fluorescence in Zebrafish Embryos

Zebrafish embryos are a powerful tool for large-scale screening of small molecules. Transgenic zebrafish that express fluorescent reporter proteins are frequently used to identify chemicals that modulate gene expression. Chemical screens that assay fluorescence in live zebrafish often rely on expensive, specialized equipment for high content screening. We describe a procedure using a standard epifluorescence microscope with a motorized stage to automatically image zebrafish embryos and detect tissue-specific fluorescence. Using transgenic zebrafish that report estrogen receptor activity via expression of GFP, we developed a semi-automated procedure to screen for estrogen receptor ligands that activate the reporter in a tissue-specific manner. In this video we describe procedures for arraying zebrafish embryos at 24-48 hours post fertilization (hpf) in a 96-well plate and adding small molecules that bind estrogen receptors. At 72-96 hpf, images of each well from the entire plate are automatically collected and manually inspected for tissue-specific fluorescence. This protocol demonstrates the ability to detect estrogens that activate receptors in heart valves but not in liver.

Described here is a protocol for semi-automated imaging of tissue-specific fluorescence in zebrafish embryos.

Zebrafish embryos are a powerful tool for large-scale screening of small molecules. Transgenic zebrafish that express fluorescent reporter proteins are ...

Measurement of Metabolic Rate in Drosophila using Respirometry

Metabolic disorders are a frequent problem affecting human health. Therefore, understanding the mechanisms that regulate metabolism is a crucial scientific task. Many disease causing genes in humans have a fly homologue, making Drosophila a good model to study signaling pathways involved in the development of different disorders. Additionally, the tractability of Drosophila simplifies genetic screens to aid in identifying novel therapeutic targets that may regulate metabolism. In order to perform such a screen a simple and fast method to identify changes in the metabolic state of flies is necessary. In general, carbon dioxide production is a good indicator of substrate oxidation and energy expenditure providing information about metabolic state. In this protocol we introduce a simple method to measure CO2 output from flies. This technique can potentially aid in the identification of genetic perturbations affecting metabolic rate.

Measurement of Metabolic Rate in Drosophila using Respirometry

Metabolic disorders are among one of the most common diseases in humans. The genetically tractable model organism D. melanogaster can be used to identify novel genes that regulate metabolism. This paper describes a relatively simple method which allows studying the metabolic rate in flies by measuring their CO2 production.

Metabolic disorders are a frequent problem affecting human health. Therefore, understanding the mechanisms that regulate metabolism is a crucial ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...