We thank Amayra Hern&#225;ndez-Vega and Philippe-Alexandre Pouille for participating in setting up the basis for these protocols. We also thank the Molecular Imaging Platform from the IBMB-PCB, Xavier Esteban and members of the laboratory for continuous support. The Consolidated Groups Program of the Generalitat de Catalunya and DGI and Consolider Grants from the Ministry of Economy and Competitivity of Spain to EMB and DN supported this work.

All the protocol steps described below follow the animal care guidelines of our institutions.
1. Zebrafish Culture
<ol>
	<li>Breed and maintain adult zebrafish under standard conditions. 
	NOTE: AB and TL wild type embryos were used for this study.</li>
	<li>Collect embryos and grow them at 28.5 &#176;C in E3 embryo medium24. Stage them according to morphology as previously described19.</li>
</ol>
2. Atomic Force Microscopy
<ol>
	<li>Manually dechorionate staged zebrafish embryos (of different ages according to individual interests). Remove chorions with two thin and sharp forceps (see Table of Materials).
	<ol>
		<li>Grip the chorion using one forceps and make a tear in it with the other forceps. Then, holding the chorion in a region opposite to that of the tear, push the embryo gently through the opening.</li>
		<li>Perform the dechorionation under a dissecting stereomicroscope with an adjustable range of magnification between 8X and 50X.</li>
	</ol>
	</li>
	<li>Mount the embryos for AFM
	<ol>
		<li>Prepare a solution of 2% agarose in embryo medium and fill a 35-mm Petri dish with this. Let it solidify. Make small holes of 1.5 mm of diameter (twice the embryo size) and approximately 350 &#181;m depth (half the embryo size) with thin forceps in the agarose bed.</li>
		<li>Place the embryos in the pokes made in the agarose layer and assure they are in place by spreading around a solution of 0.5% low melting agarose in embryo medium. Pour this solution around just the embryos to secure them in the holes.</li>
		<li>Before the low melting point agarose solidifies at room temperature, rotate the embryos in such a way that the region of interest to be probed by AFM will face upwards (Figure 1A).</li>
	</ol>
	</li>
	<li>Examine the embryos by AFM
	<ol>
		<li>Locate and image the embryos in the petri dish employing a 20X objective in an Inverted Atomic Force Microscope (see Table of Materials25) at room temperature (23 - 24 &#176;C). 
		NOTE: The embryos are kept alive immersed in embryo medium and attached to the agarose bed.</li>
		<li>Probe the yolk cell surface of casted embryos employing spherical polystyrene beads of 4.5 &#181;m in diameter attached to a cantilever with a nominal spring constant of 0.01 N/m. Set the peak-to-peak amplitude of cantilever oscillation to 5 &#181;m and its frequency to 1 Hz.</li>
		<li>Collect data for each region to be tested in different positions and in several embryos, as a routine, for data averaging. 
		NOTE: A good starting point is to probe five positions located in the middle and at the corners of a 10 &#181;m x 10 &#181;m square by controlling the cantilever position with the XY piezo-actuators of the AFM microscope. Imaging and data collection took around 20 min per embryo. Recording data in different regions of the embryo yolk cell surface, e.g., the vegetal pole or different regions close to the EVL cells margin (Figure 1B and 1C), can only be done by employing distinct specimens oriented differently.</li>
	</ol>
	</li>
	<li>Calculate forces
	<ol>
		<li>Measure the vertical displacement of the AFM cantilever (z) with strain gauge sensors coupled to piezo-actuators, and its deflection (d) using a quadrant photodiode by the optical lever method with the microscope control software (see Table of Materials25).</li>
		<li>Use the slope of a d-z curve obtained from data collected from a bare region of a glass coverslip to calibrate the correlation between the photodiode signal and the cantilever deflection (d). 
		NOTE: The measured vertical displacement equals the cantilever deflection and the slope represents the deflection sensitivity of the optical lever26.</li>
		<li>Infer the cantilever spring constant (k) from the thermal fluctuations as previously described27.</li>
		<li>Compute the indentation of the sample (h) as: 
		h = (z - zc) - (d - doff)&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;(1) 
		NOTE: Here zc is the position of the contact point and doff is the offset of the photodiode.</li>
		<li>Calculate the force (F) on the cantilever as: 
		F = k &#183; d&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;(2)</li>
	</ol>
	</li>
	<li>Calculate the tension of the cortex in terms of a liquid-balloon model consisting of an elastic layer of cortical tension Tc enclosing a viscous liquid. In this model, for small indentations (in comparison to the size of the embryo), force increases proportionally4,28 as: 
	F = 4&#960; Tc [(Rb / Re) +1)] h&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;(3) 
	NOTE: Here Rb is the radius of the bead and Re is the radius of the embryo. For the zebrafish embryo, as the radius of the embryo (400 &#181;m) is two orders of magnitude larger than that of the bead (2.25 &#181;m), this equation can be approximated as: 
	F = 4&#960; Tc h&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;(4)
	<ol>
		<li>Calculate the tension force-indentation (F-h) curves in each embryo yolk cell region for five different point measurements.</li>
		<li>Compute the Tc for each F-h curve by non-linear least-squares fitting. For statistical analysis, the cortical tension Tc computed must be averaged from the different F-h curves.</li>
	</ol>
	</li>
	<li>Measure the viscoelastic properties (rheology) of the cortex by applying low amplitude (100 nm) multifrequency oscillations during AFM composed of sinusoidal waves of different frequencies25.
	<ol>
		<li>Compute an effective complex modulus g*(&#402;) in the frequency domain from the force indentation curves as: 
		g*(f) = [F(f) / h(f)] - ifb&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;(5) 
		here i is the imaginary unit and F(f) and h(f) are the frequency (f) spectra of force and indentation. b is the correction for the viscous drag on the cantilever extracted from the oscillations applied on the surface.</li>
		<li>Separate g*(&#402;) into real and imaginary parts as: 
		g*(&#402;) = g&#39;(f) + ig&#39;&#39;(f)&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;(6) 
		em&#62;g&#39;(f) is the elastic modulus and is a measure of the elastic energy stored and recovered per cycle of oscillation. g&#39;&#39;(f) is the viscous modulus that accounts for the dissipated energy.</li>
		<li>Calculate the loss tangent, which provides an index of the solid-like (&#60;1) or liquid-like (&#62;1) behavior of the material, as: 
		g&#39;&#39;(f) / g&#39;(f)&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;(7) 
		NOTE: With this type of measurement, it is possible to extract parameters indicating how viscous and how elastic is the probed material. Although b slightly depends on the distance of the end of the cantilever to the surface, given the very low values of g&#180;&#180; exhibited by the yolk cell (see Figure 2D below), small variations in b have a negligible impact in the measurements29.</li>
	</ol>
	</li>
</ol>
3. Particle Tracking Microrheology
<ol>
	<li>Perform particle microinjection into the yolk cell
	<ol>
		<li>Fabricate tailored microneedles using a horizontal micropipette puller and borosilicate capillary glass (see Table of Materials).
		<ol>
			<li>Prepare microneedles that have an outer diameter of 1.00 mm, an inner diameter of 0.58 mm and a length of 10 cm using puller settings: Pressure: 500; Heat: 510; Pull: 65; Velocity: 25; Time: 50.</li>
		</ol>
		</li>
		<li>Prepare an injection petri dish plate by creating straight indentation lanes in a 1% agarose in embryo medium bed employing custom made molds (see Table of Materials).
		<ol>
			<li>Turn the molds upside down and place them on top of the liquid agarose gel and remove the molds once the gel has solidified. Pipette the embryos into the grooves made by the mold in the agarose under a dissecting stereomicroscope at 1.2X magnification. 
			NOTE: The width and design of the molds enable the embryos to self-align.</li>
		</ol>
		</li>
		<li>Before injection, almost completely remove the medium to facilitate injection (the surface tension prevents the embryo/chorion from sticking to the needle when removing it after injection).</li>
		<li>Microinject fluorescent nanoparticles (radius a = 100 nm, see Table of Materials) diluted in water (1:1,000) in the vegetal part of the embryo yolk cell (Figure 3A).</li>
		<li>Adjust the micropipette with precision micromanipulators and inject the beads with an automatic microinjector with time and pressure controls (see Table of Materials). Set the pressure between 10 and 20 psi.</li>
		<li>Before injecting the bead solution, calibrate the volume to inject (0.5 nL) by measuring the droplet size delivered by the microinjector with a 1x 0.01 mm stage micrometer (see the Table of Materials).</li>
		<li>Employ a magnification of 1.6X in the dissecting stereomicroscope to visualize the embryos during the injections, which are performed at room temperature.</li>
	</ol>
	</li>
	<li>Assess the viscoelastic behavior of the yolk by recording thermal fluctuations
	<ol>
		<li>Dechorionate the microinjected embryos as in step 2.1 and embed them in a 0.5% low melting point agarose in embryo medium solution at 30 &#176;C. Afterwards, transfer them to glass bottom plates (see Table of Materials) and orient and push them towards the coverslip when the agarose is still liquid. 
		NOTE: When the agarose solidifies at room temperature, the embryos are ready to be imaged.</li>
		<li>Place the embryos mounted in the glass bottom plates on the stage of a confocal inverted microscope 2 h after microinjection. Capture images of the nanoparticles for 26 s at a sampling rate of 25 Hz with a 63X objective at room temperature in an inverted confocal microscope employing the standard commercial microscope software (pixel size = 166 nm, images captured every 40 ms).</li>
		<li>Compute the position of the particles&#39; centroids and define their trajectories over time with the TrackMate plugin of the open source ImageJ software (Figure 3B).</li>
		<li>Calculate the two-dimensional Mean Square Displacement (MSD) of each particle with custom-made software6,30 as: 
		&#60;&#916;r2 (&#964;)&#62; = &#60;[x(t + &#964;) - x(t)]2 + [y(t + &#964;) - y(t)]2&#62;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;(8) 
		NOTE: Here t is the elapsed time and the time lag. In a pure viscous liquid, the MSD increases inversely with viscosity (&#957;) according to the Stokes-Einstein relationship: 
		&#60;&#916;r2 (&#964;)&#62; = 4kB T&#964;/ 6&#960;&#957;a,&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;(9) 
		where T is the absolute temperature.</li>
	</ol>
	</li>
</ol>

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The authors declare no competing financial interests or other conflicts of interest.

Here we show that the material properties and some biomechanical parameters of the zebrafish yolk cell during epiboly can be readily estimated by AFM and nanoparticles microrheology.
While AFM has been employed to retrieve rheological features of cells and tissues in physiological conditions4,5,25,28, here we developed a protocol for applying AFM to intact developing...

The physical principles underlying the biomechanical control of morphogenetic processes are largely undefined. While biomechanical studies at the molecular and cellular level are gathering considerable momentum, the exploration of biomechanical parameters at the tissue/organism level is at its infancy. Hydrodynamic Regression1 or Video Force Microscopy2 allow researchers to distinguish active and passive forces, while laser microsurgery3, or the less intrusive atomic force microscopy (AFM) of dissociated cells4 or in vivo5 or nanoparticle microrheology6 allow the anticipation of a tissue&#39;s mechanical properties and responses.
AFM is a three-dimensional topographic technique with high atomic resolution resolving surface roughness and compliance from the deflection of cantilever tips upon contact with a probed surface7. Different methodologies employing AFM have been developed to investigate surface properties, including measuring friction, adhesion forces, and viscoelastic properties of diverse materials. Recently, AFM has emerged as a powerful tool to retrieve information on the mechanical properties of biological samples. In particular, AFM can non-invasively extract the complex shear modulus from single cells by applying oscillatory indentations with a micro tip attached to a cantilever of known bending constant8. This lets us infer resulting forces. Yet, AFM is not unique and different methodologies allow extracting rheological data and mechanical properties from individual cells (e.g., Micropipette aspiration9, magnetic twisting cytometry (MTC)10, or uniaxial tensile testing11).
Yet, the application of these novel methodologies to complex morphogenetic processes is not straightforward. The main challenges faced when determining the mechanical properties of embryonic tissues are the small (&#181;m to mm), soft (in the 102&#160;- 104 Pa range) and visco-elastic (giving rise to time-dependent phenomena) nature of the material12. It is therefore important to adapt the methods employed for the determination of mechanical properties (stiffness, viscosity, adhesion) to the specific case of embryos and developing organisms. Two important issues need to be taken into consideration when analyzing rheological approaches to study development: to avoid intrusive interference and to provide easy accessibility. In this scenario, micropipette aspiration, MTC, and tensile testing exhibit limitations. In the first case, the high deformation that a cell suffers might alter its physiological and mechanical properties9. In the second case, the need to firmly adhere the experimental tissue to a substrate to ensure that the stresses applied deform the cytoskeleton locally could also introduce effects on the cells&#39; activation state and, hence, in their mechanical features10. Last, uniaxial tensile approaches are limited by the geometry of developing organisms and accessibility11.
AFM seems to be better suited for the study of developmental processes as it allows the study of biological samples directly in their natural environment without any cumbersome sample preparation. Moreover, as dissociation of embryonic tissues is generally difficult, the reduced size of AFM probes provides a high degree of versatility with no limitation in the choice of the type of medium (either aqueous or non-aqueous), sample temperature, or chemical composition of the sample. AFM is versatile enough to be applied to large domains and time scales and has been employed to retrieve material properties of tissues in distinct developmental stages or physiological conditions. Whole native tissues such as arteries13 or bones14 have been studied from a topographical point of view and in some cases, like the sclera, mechanical properties have also been retrieved15. Topography has also been explored in live embryos allowing the visualization of, for instance, cell rearrangement during morphogenesis in Xenopus16. Last, comprehensive sample preparation protocols have been developed to determine mechanical parameters during different developmental processes. Nanoindentation maps have been generated on native unfixed tissue sections for the chick embryonic digestive tract11; adhesive and mechanical properties retrieved for individual ectoderm, mesoderm, and endoderm progenitor cells isolated from gastrulating zebrafish embryos4; stiffness measurements performed for the epiblast and primitive streak on explants from avian embryos17; and a distinct pattern of stiffness gradients directly defined in the embryonic brain of Xenopus5.
A significant qualitative leap for applying AFM for exploring embryonic development may come from approaching simple accessible morphogenetic processes. Zebrafish epiboly is an essential and conserved event, in which different tissues coordinate in a restricted spherical space to direct the expansion of the blastoderm in a few hours. At the onset of zebrafish epiboly (spherical stage), a superficial layer of cells, the enveloping layer (EVL), covers a semi-spherical cap of blastomeres centered on the animal pole of the embryo sitting on a massive yolk syncytial cell. Epiboly consists of the cortical vegetal ward expansion of the EVL, the deep cells (DCs) of the blastoderm, and the external layer of the syncytial yolk cell (E-YSL) around the yolk. Epiboly ends with the closure of the EVL and the DCs at the vegetal pole18,19,20. How and where forces are generated during epiboly and how they are globally coupled is not yet clear1,21.
Here we describe in detail how to apply AFM to infer passive mechanical tissue properties during epiboly progression in the zebrafish yolk cell in vivo. To do so, we employed small microspheres attached to AFM cantilevers as probes. This allows the retrieval of precise local information within the non-uniform yolk cell surface and to study tensional gradients and their dynamics over time. Alternative AFM approaches such as those using wedge cantilevers22, will not render local data that is precise enough. The wedge technique, which requires very careful manipulation skills to glue a wedge larger than the sample size at the end of the cantilever, is not well suited for probing embryo (~2-fold larger than the length of the cantilever) mechanics.
Modeling and characterizing the viscoelastic properties of cells in qualitative and quantitative ways allows for an improved understanding of their biomechanics. From a biomechanical point of view, it is essential to understand epiboly, and not just demand knowledge on cortical tension measurements and dynamics, which can be extracted by AFM; thus there is need for information on the biophysical properties of the tissue. To extract this information, a variety of cell rheology techniques have been developed over the years in order to characterize different cell types under different physiological conditions23. They include AFM, MTC, and Optical Tweezers (OT). These methods, however, have been proven inadequate for mesoscopic analyses at the scale of embryos or organs. As an alternative, we have successfully adapted nanoparticle-tracking microrheology6 for in vivo rheological measurements. This technique analyzes the Brownian displacements of individual particles and infers local micromechanical properties.
Together AFM and nanoparticle microrheology allow the definition of cortical tensional dynamics and internal mechanical properties of the yolk during epiboly. This information fully endorses a model in which anisotropic stress develops as a consequence of the differences in the deformation response of the EVL and the Yolk Cytoplasmic Layer (YCL) to the isotropic actomyosin contraction of the E-YSL cortex is essential for the directional net movement of the EVL and epiboly progression1.

<table>
	<tbody>
		<tr>
			<td>Inverted optical microscope</td>
			<td>Nikon</td>
			<td>TE2000</td>
			<td>Visualization of the sample and AFM cantilever</td>
		</tr>
		<tr>
			<td>Atomic force microscope (AFM)</td>
			<td>Custom made</td>
			<td>-</td>
			<td>Any commercialized AFM could be employed</td>
		</tr>
		<tr>
			<td>Inverted Confocal microscope</td>
			<td>Zeiss</td>
			<td>LSM780</td>
			<td>Nanorheology data collection</td>
		</tr>
		<tr>
			<td>Agarose D1 Low EEO</td>
			<td>Conda</td>
			<td>8016.00</td>
			<td>Casting embryos</td>
		</tr>
		<tr>
			<td>Ultrapure LMP Agarose (low melting)</td>
			<td>Invitrogen</td>
			<td>16520-100</td>
			<td>Securing embryos</td>
		</tr>
		<tr>
			<td>Zebrafish Microinjection and Transplantation Molds</td>
			<td>World Precision Instruments</td>
			<td>Z-MOLDS</td>
			<td>Casting embryos. Custom made with the original dimensions can also be employed</td>
		</tr>
		<tr>
			<td>Dumont 5 Forceps</td>
			<td>FST</td>
			<td>11251-20</td>
			<td>1.5 mm diameter</td>
		</tr>
		<tr>
			<td>Si3N4 cantilever</td>
			<td>Novascan</td>
			<td>PT.GS</td>
			<td>Measuring the embryo by AFM. Spherical tip: 4.5 &#181;m diameter and k=0.01 N/m</td>
		</tr>
		<tr>
			<td>Fluorescent nanoparticles</td>
			<td>Life Technologies</td>
			<td>FluoSpheres F8811,</td>
			<td>For tracking nanorheology</td>
		</tr>
		<tr>
			<td>Micropipette puller</td>
			<td>Sutter Instruments</td>
			<td>P-2000</td>
			<td>Fabricating tailored microneedles</td>
		</tr>
		<tr>
			<td>Borosilicate capillary glass</td>
			<td>Warner Instruments</td>
			<td>G100TF-4</td>
			<td>Fabricating tailored microneedles</td>
		</tr>
		<tr>
			<td>Micromanipulator</td>
			<td>Narishige</td>
			<td>MN-153</td>
			<td>Manipulating the micropipette</td>
		</tr>
		<tr>
			<td>Microinjector</td>
			<td>Eppendorf</td>
			<td>FemtoJet Express</td>
			<td>Controlling injection time and pressure</td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>Leica</td>
			<td>DFC365FX</td>
			<td>Visualization of the embryos during injection</td>
		</tr>
		<tr>
			<td>Analysis Software</td>
			<td>MathWorks</td>
			<td>Matlab</td>
			<td>Analyzing AFM and particle tracking data</td>
		</tr>
		<tr>
			<td>Zebrafish</td>
			<td>AB and TL wild type</td>
			<td>-</td>
			<td>Strains employed for embryo collection</td>
		</tr>
		<tr>
			<td>Glass Bottom Plates</td>
			<td>Mat Tek</td>
			<td>P35G-0.170-14-C</td>
			<td>Mounting embryos for nanorheology data collection</td>
		</tr>
		<tr>
			<td>Stage micrometer</td>
			<td>FST</td>
			<td>29025-02</td>
			<td></td>
		</tr>
	</tbody>
</table>

afm and microrheology in the zebrafish embryo yolk cell

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. Various propositions claim to have been important contributions to the understanding of the mechanical properties of cells and tissues in their spatiotemporal organization in different developmental and morphogenetic processes. However, due to the lack of reliable and accessible tools to measure material properties and tensional parameters in vivo, validating these hypotheses has been difficult. Here we present methods employing atomic force microscopy (AFM) and particle tracking with the aim of quantifying the mechanical properties of the intact zebrafish embryo yolk cell during epiboly. Epiboly is an early conserved developmental process whose study is facilitated by the transparency of the embryo. These methods are simple to implement, reliable, and widely applicable since they overcome intrusive interventions that could affect tissue mechanics. A simple strategy was applied for the mounting of specimens, AFM recording, and nanoparticle injections and tracking. This approach makes these methods easily adaptable to other developmental times or organisms.

Cortical tension measurements 
For each measurement point, five force-displacement (F-z) curves were acquired by AFM by ramping the cantilever at 1 Hz with a peak-to-peak amplitude of 5 &#181;m (velocity = 10 &#181;m/s) up to a maximum indentation of ~2 &#181;m. This procedure was taking less than 20 min and was not affected by epiboly progression. Each experimental condition was tested in at least 5 embryos.
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Watch this Scientific Journal Video about AFM and Microrheology in the Zebrafish Embryo Yolk Cell at JoVE.com

AFM and Microrheology in the Zebrafish Embryo Yolk Cell

The lack of tools to measure material properties and tensional parameters in vivo prevents validating their roles during development. We employed atomic force microscopy (AFM) and nanoparticle-tracking to quantify mechanical features on the intact zebrafish embryo yolk cell during epiboly. These methods are reliable and widely applicable avoiding intrusive interventions.

Elucidating the factors that direct the spatio-temporal organization of evolving tissues is one of the primary purposes in the study of development. ...

afm-and-microrheology-in-the-zebrafish-embryo-yolk-cell

Research

JoVE Journal

Developmental Biology

8.2K Views.  Consejo Superior de Investigaciones Científicas. The lack of tools to measure material properties and tensional parameters in vivo prevents validating their roles during development. We employed atomic force microscopy (AFM) and nanoparticle-tracking to quantify mechanical features on the intact zebrafish embryo yolk cell during epiboly. These methods are reliable and widely applicable avoiding intrusive interventions.

Video: AFM and Microrheology in the Zebrafish Embryo Yolk Cell

Microbead Implantation in the Zebrafish Embryo

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic conservation, as well as similarities in cellular, molecular, and physiological processes, with other vertebrates including humans. During early ontogeny, zebrafish embryos are optically transparent, allowing researchers to visualize the dynamics of organogenesis using a simple stereomicroscope. Microbead implantation is a method that enables tissue manipulation through the alteration of factors in local environments. This allows researchers to assay the effects of any number of signaling molecules of interest, such as secreted peptides, at specific spatial and temporal points within the developing embryo. Here, we detail a protocol for how to manipulate and implant beads during early zebrafish development.

The zebrafish is an excellent model system for genetic and developmental studies. Bead implantation is a valuable tissue manipulation technique that can be used to interrogate developmental mechanisms by introducing alterations in local cellular environments. This protocol describes how to perform microbead implantation in the zebrafish embryo.

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic ...

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Primary Endodermal Epithelial Cell Culture from the Yolk Sac Membrane of Japanese Quail Embryos

We established an endodermal epithelial cell culture model (EEC) for studying the function of certain enzymes and proteins in mediating nutrient utilization by avian embryos during development. Fertilized Japanese quail eggs were incubated at 37 &#176;C for 5 days and then yolk sac membranes (YSM) were collected to establish the EEC culture system. We isolated the embryonic endoderm layer from YSM, and sliced the membrane into 2 - 3 mm pieces and partially digested with collagenase before seeding in 24-well culture plates. The EECs proliferate out of the tissue and are ready for cell culture studies. We found that the EECs had typical characteristics of YSM in vivo, for example, accumulation of lipid droplets, expression of sterol O-acyltransferase and lipoprotein lipase. The partial digestion treatment significantly increased the successful rate of EEC culture. Utilizing the EECs, we demonstrated that the expression of SOAT1 was regulated by the cAMP dependent protein kinase A related pathway. This primary Japanese quail EEC culture system is a useful tool to study embryonic lipid transportation and to clarify the role of genes involved in mediating nutrient utilization in YSM during avian embryonic development.

To study the mechanism of lipid utilization in yolk sac membranes during the late stages of avian embryonic development, we established a primary Japanese quail embryonic endodermal epithelial cell culture system.

We established an endodermal epithelial cell culture model (EEC) for studying the function of certain enzymes and proteins in mediating nutrient ...

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin condensation, microtubule-kinetochore attachment, chromosome segregation, and cytokinesis in a small time frame. Errors in the delicate process can result in human disease, including birth defects and cancer. Traditional approaches investigating human mitotic disease states often rely on cell culture systems, which lack the natural physiology and developmental/tissue-specific context advantageous when studying human disease. This protocol overcomes many obstacles by providing a way to visualize, with high resolution, chromosome dynamics in a vertebrate system, the zebrafish. This protocol will detail an approach that can be used to obtain dynamic images of dividing cells, which include: in vitro transcription, zebrafish breeding/collecting, embryo embedding, and time-lapse imaging. Optimization and modifications of this protocol are also explored. Using H2A.F/Z-EGFP (labels chromatin) and mCherry-CAAX (labels cell membrane) mRNA-injected embryos, mitosis in AB wild-type, auroraBhi1045, and esco2hi2865 mutant zebrafish is visualized. High resolution live imaging in zebrafish allows one to observe multiple mitoses to statistically quantify mitotic defects and timing of mitotic progression. In addition, observation of qualitative aspects that define improper mitotic processes (i.e., congression defects, missegregation of chromosomes, etc.) and improper chromosomal outcomes (i.e., aneuploidy, polyploidy, micronuclei, etc.) are observed. This assay can be applied to the observation of tissue differentiation/development and is amenable to the use of mutant zebrafish and pharmacological agents. Visualization of how defects in mitosis lead to cancer and developmental disorders will greatly enhance understanding of the pathogenesis of disease.

Mitosis is critical to every living organism and defects often lead to cancer and developmental disorders. Using this imaging protocol and zebrafish as a model system, researchers can visualize mitosis in a live vertebrate organism and the multitude of defects that arise when mitotic processes are defective.

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin ...

Identification Of Erythromyeloid Progenitors And Their Progeny In The Mouse Embryo By Flow Cytometry

Macrophages are professional phagocytes from the innate arm of the immune system. In steady-state, sessile macrophages are found in adult tissues where they act as front line sentinels of infection and tissue damage. While other immune cells are continuously renewed from hematopoietic stem and progenitor cells (HSPC) located in the bone marrow, a lineage of macrophages, known as resident macrophages, have been shown to be self-maintained in tissues without input from bone marrow HSPCs. This lineage is exemplified by microglia in the brain, Kupffer cells in the liver and Langerhans cells in the epidermis among others. The intestinal and colon lamina propria are the only adult tissues devoid of HSPC-independent resident macrophages. Recent investigations have identified that resident macrophages originate from the extra-embryonic yolk sac hematopoiesis from progenitor(s) distinct from fetal hematopoietic stem cells (HSC). Among yolk sac definitive hematopoiesis, erythromyeloid progenitors (EMP) give rise both to erythroid and myeloid cells, in particular resident macrophages. EMP are only generated within the yolk sac between E8.5 and E10.5 days of development and they migrate to the fetal liver as early as circulation is connected, where they expand and differentiate until at least E16.5. Their progeny includes erythrocytes, macrophages, neutrophils and mast cells but only EMP-derived macrophages persist until adulthood in tissues. The transient nature of EMP emergence and the temporal overlap with HSC generation renders the analysis of these progenitors difficult. We have established a tamoxifen-inducible fate mapping protocol based on expression of the macrophage cytokine receptor Csf1r promoter to characterize EMP and EMP-derived cells in vivo by flow cytometry.

While infiltrating macrophages are continuously recruited to adult tissues from circulating precursors, resident macrophages seed their tissue during development, where they are maintained without further input from progenitors. The progenitors for resident macrophages were recently identified. Here, we present methods for the genetic fate mapping of the resident macrophage progenitors.

Macrophages are professional phagocytes from the innate arm of the immune system. In steady-state, sessile macrophages are found in adult tissues where ...

Use of Immunolabeling to Analyze Stable, Dynamic, and Nascent Microtubules in the Zebrafish Embryo

Microtubules (MTs) are dynamic and fragile structures that are challenging to image in vivo, particularly in vertebrate embryos. Immunolabeling methods are described here to analyze distinct populations of MTs in the developing neural tube of the zebrafish embryo. While the focus is on neural tissue, this methodology is broadly applicable to other tissues. The procedures are optimized for early to mid-somitogenesis-stage embryos (1 somite to 12 somites), however they can be adapted to a range of other stages with relatively minor adjustments. The first protocol provides a method to assess the spatial distribution of stable and dynamic MTs and perform a quantitative analysis of these populations with image-processing software. This approach complements existing tools to image microtubule dynamics and distribution in real-time, using transgenic lines or transient expression of tagged constructs. Indeed, such tools are very useful, however they do not readily distinguish between dynamic and stable MTs. The ability to image and analyze these distinct microtubule populations has important implications for understanding mechanisms underlying cell polarization and morphogenesis. The second protocol outlines a technique to analyze nascent MTs specifically. This is accomplished by capturing the de novo growth properties of MTs over time, following microtubule depolymerization with the drug nocodazole and a recovery period after drug washout. This technique has not yet been applied to the study of MTs in zebrafish embryos, but is a valuable assay for investigating the in vivo function of proteins implicated in microtubule assembly.

Immunolabeling methods to analyze distinct populations of microtubules in the developing zebrafish brain are described here, which are broadly applicable to other tissues. The first protocol outlines an optimized method for immunolabeling stable and dynamic microtubules. The second protocol provides a method to image and quantify nascent microtubules specifically.

Microtubules (MTs) are dynamic and fragile structures that are challenging to image in vivo, particularly in vertebrate embryos. Immunolabeling methods ...

Effectiveness of the Air Stripping in Two Salmonid Fish, Rainbow Trout (Oncorhynchus Mykiss) and Brown Trout (Salmo Trutta Morpha fario)

Egg collection is one of the most crucial procedures during fish reproduction in salmonid hatcheries. Classic methods involve the use of hand massage on fish abdomens to expel the eggs. An alternative method uses the pressure of gas injected into the body cavity, which causes the subsequent release of the eggs. This method is believed to have less negative effects on both the welfare and egg quality of the broodstocks. Herein, we compare the results of air and hand stripping methods with respect to one-year survival and egg quantity and quality in two salmonid fish, rainbow trout (Oncorhynchus mykiss) and brown trout (Salmo trutta morpha fario). Our results indicate that air stripping yielded a better quality of eggs and higher one-year survival rate in rainbow trout. In addition, air stripping resulted in lower mortality rate than the group subjected to hand stripping (25% vs. 35%). The pH and hatching rate of the hand stripped group was lower than those of the air stripped group. In the case of brown trout, the quality of eggs obtained by both hand and air-stripping methods was similar; however, the one-year losses in fish were higher in air stripped group (15% compared to 0% in hand stripped fish). Although the advantages of air stripping method over hand stripping in terms of egg quality might not be observed in all salmonid species, the air-stripping procedure might be a promising option to be adopted in hatcheries as it ensures a high level of reproducibility and efficiency.

Effectiveness of the Air Stripping in Two Salmonid Fish, Rainbow Trout Oncorhynchus Mykiss and Brown Trout Salmo Trutta Morpha fario

The principal aim of this study is to standardize and test the pneumatic method (air stripping) of collecting eggs in rainbow trout and brown trout. This method allows effective and simple collection of the eggs without the necessity of fish abdomen massage.

Egg collection is one of the most crucial procedures during fish reproduction in salmonid hatcheries. Classic methods involve the use of hand massage on ...

Following Endocardial Tissue Movements via Cell Photoconversion in the Zebrafish Embryo

During embryogenesis, cells undergo dynamic changes in cell behavior, and deciphering the cellular logic behind these changes is a fundamental goal in the field of developmental biology. The discovery and development of photoconvertible proteins have greatly aided our understanding of these dynamic changes by providing a method to optically highlight cells and tissues. However, while photoconversion, time-lapse microscopy, and subsequent image analysis have proven to be very successful in uncovering cellular dynamics in organs such as the brain or the eye, this approach is generally not used in the developing heart due to challenges posed by the rapid movement of the heart during the cardiac cycle. This protocol consists of two parts. The first part describes a method for photoconverting and subsequently tracking endocardial cells (EdCs) during zebrafish atrioventricular canal (AVC) and atrioventricular heart valve development. The method involves temporally stopping the heart with a drug in order for accurate photoconversion to take place. Hearts are allowed to resume beating upon removal of the drug and embryonic development continues normally until the heart is stopped again for high-resolution imaging of photoconverted EdCs at a later developmental time point. The second part of the protocol describes an image analysis method to quantify the length of a photoconverted or non-photoconverted region in the AVC in young embryos by mapping the fluorescent signal from the three-dimensional structure onto a two-dimensional map. Together, the two parts of the protocol allows one to examine the origin and behavior of cells that make up the zebrafish AVC and atrioventricular heart valve, and can potentially be applied for studying mutants, morphants, or embryos that have been treated with reagents that disrupt AVC and/or valve development.

This protocol describes a method for the photoconversion of Kaede fluorescent protein in endocardial cells of the living zebrafish embryo that enables the tracking of endocardial cells during atrioventricular canal and atrioventricular heart valve development.

During embryogenesis, cells undergo dynamic changes in cell behavior, and deciphering the cellular logic behind these changes is a fundamental goal in the ...

Minimally Invasive Embryo Transfer and Embryo Vitrification at the Optimal Embryo Stage in Rabbit Model

Assisted reproductive techniques (ARTs), such as in vitro embryo culture or embryo cryopreservation, affect natural development patterns with perinatal and postnatal consequences. To ensure the innocuousness of ART applications, studies in animal models are necessary. In addition, as a last step, embryo development studies require evaluation of their capacity to develop full-term healthy offspring. Here, embryo transfer to the uterus is indispensable to perform any ARTs-related experiment.
The rabbit has been used as a model organism to study mammalian reproduction for over a century. In addition to its phylogenetic proximity to the human species and its small size and low maintenance cost, it has important reproductive characteristics such as induced ovulation, a chronology of early embryonic development similar to humans and a short gestation that allow us to study the consequences of ART application easily. Moreover, ARTs (such as intracytoplasmic sperm injection, embryo culture, or cryopreservation) are applied with suitable efficiency in this species.
Using the laparoscopic embryo transfer technique and the cryopreservation protocol presented in this article, we describe 1) how to transfer embryos through an easy, minimally invasive technique and 2) an effective protocol for long-term storage of rabbit embryos to provide time-flexible logistical capacities and the ability to transport the sample. The outcomes obtained after transferring rabbit embryos at different developmental stages indicate that morula is the ideal stage for rabbit embryo recovery and transfer. Thus, an oviductal embryo transfer is required, justifying the surgical procedure. Furthermore, rabbit morulae are successfully vitrified and laparoscopically transferred, proving the effectiveness of the described techniques.

Assisted reproductive techniques (ARTs) are in continuous evaluation to improve outcomes and reduce the associated risks. This manuscript describes a minimally invasive embryo transfer procedure with an efficient cryopreservation protocol that allows the use of rabbits as an ideal animal model of human reproduction.

Assisted reproductive techniques (ARTs), such as in vitro embryo culture or embryo cryopreservation, affect natural development patterns with perinatal ...