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>Si&lt;sub&gt;3&lt;/sub&gt;N&lt;sub&gt;4&lt;/sub&gt; cantilever</td><td>Novascan</td><td>PT.GS</td><td>Measuring the embryo by AFM. Spherical tip: 4.5 &amp;micro;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>Measuring injection volume</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

Universidad de Cartagena UDEC

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

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 ...

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM compared to confocal systems are its low phototoxicity, gentle mounting strategies, fast acquisition with high signal to noise ratio and the possibility of imaging samples from various angles (views) for long periods of time. Imaging from multiple views unleashes the full potential of LSFM, but at the same time it can create terabyte-sized datasets. Processing such datasets is the biggest challenge of using LSFM. In this protocol we outline some solutions to this problem. Until recently, LSFM was mostly performed in laboratories that had the expertise to build and operate their own light sheet microscopes. However, in the last three years several commercial implementations of LSFM became available, which are multipurpose and easy to use for any developmental biologist. This article is primarily directed to those researchers, who are not LSFM technology developers, but want to employ LSFM as a tool to answer specific developmental biology questions. 
Here, we use imaging of zebrafish eye development as an example to introduce the reader to LSFM technology and we demonstrate applications of LSFM across multiple spatial and temporal scales. This article describes a complete experimental protocol starting with the mounting of zebrafish embryos for LSFM. We then outline the options for imaging using the commercially available light sheet microscope. Importantly, we also explain a pipeline for subsequent registration and fusion of multiview datasets using an open source solution implemented as a Fiji plugin. While this protocol focuses on imaging the developing zebrafish eye and processing data from a particular imaging setup, most of the insights and troubleshooting suggestions presented here are of general use and the protocol can be adapted to a variety of light sheet microscopy experiments.

Light sheet fluorescence microscopy is an excellent tool for imaging embryonic development. It allows recording of long time-lapse movies of live embryos in near physiological conditions. We demonstrate its application for imaging zebrafish eye development across wide spatio-temporal scales and present a pipeline for fusion and deconvolution of multiview datasets.

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM ...

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 ...

Microstructured Devices for Optimized Microinjection and Imaging of Zebrafish Larvae

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning fundamental developmental biology through to large-scale genetic and chemical screens. However, many experiments, especially those related to infection and xenograft models, rely on microinjection and imaging of embryos and larvae, which are laborious techniques that require skill and expertise. To improve the precision and throughput of current microinjection techniques, we developed a series of microstructured devices to orient and stabilize zebrafish embryos at 2 days post fertilization (dpf) in ventral, dorsal, or lateral orientation prior to the procedure. To aid in the imaging of embryos, we also designed a simple device with channels that orient 4 zebrafish laterally in parallel against a glass cover slip. Together, the tools that we present here demonstrate the effectiveness of photolithographic approaches to generate useful devices for the optimization of zebrafish techniques.

Microinjection of zebrafish embryos and larvae is a crucial but challenging technique used in many zebrafish models. Here, we present a range of microscale tools to aid in the stabilization and orientation of zebrafish for both microinjection and imaging.

Zebrafish have emerged as a powerful model of various human diseases and a useful tool for an increasing range of experimental studies, spanning ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Biology

Dissection and Lateral Mounting of Zebrafish Embryos: Analysis of Spinal Cord Development

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene knockdown approaches can be utilized to examine the molecular mechanisms underlying nervous system development. Second, large clutches of developmentally synchronized embryos provide large experimental sample sizes. Third, the optical clarity of the zebrafish embryo permits researchers to visualize progenitor, glial, and neuronal populations. Although zebrafish embryos are transparent, specimen thickness can impede effective microscopic visualization. One reason for this is the tandem development of the spinal cord and overlying somite tissue. Another reason is the large yolk ball, which is still present during periods of early neurogenesis. In this article, we demonstrate microdissection and removal of the yolk in fixed embryos, which allows microscopic visualization while preserving surrounding somite tissue. We also demonstrate semipermanent mounting of zebrafish embryos. This permits observation of neurodevelopment in the dorso-ventral and anterior-posterior axes, as it preserves the three-dimensionality of the tissue.

Developmental processes such as proliferation, patterning, differentiation, and axon guidance can be readily modeled in the zebrafish spinal cord. In this article, we describe a mounting procedure for zebrafish embryos, which optimizes visualization of these events.

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene ...

Neuroscience

Direct Force Measurements of Subcellular Mechanics in Confinement using Optical Tweezers

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue morphogenesis goes in hand with molecular and structural changes at the single cell level that result in variations of subcellular mechanical properties. As a consequence, even within the same cell, different organelles and compartments resist differently to mechanical stresses; and mechanotransduction pathways can actively regulate their mechanical properties. The ability of a cell to adapt to the microenvironment of the tissue niche thus is in part due to the ability to sense and respond to mechanical stresses. We recently proposed a new mechanosensation paradigm in which nuclear deformation and positioning enables a cell to gauge the physical 3D environment and endows the cell with a sense of proprioception to decode changes in cell shape. In this article, we describe a new method to measure the forces and material properties that shape the cell nucleus inside living cells, exemplified on adherent cells and mechanically confined cells. The measurements can be performed non-invasively with optical traps inside cells, and the forces are directly accessible through calibration-free detection of light momentum. This allows measuring the mechanics of the nucleus independently from cell surface deformations and allowing dissection of exteroceptive and interoceptive mechanotransduction pathways. Importantly, the trapping experiment can be combined with optical microscopy to investigate the cellular response and subcellular dynamics using fluorescence imaging of the cytoskeleton, calcium ions, or nuclear morphology. The presented method is straightforward to apply, compatible with commercial solutions for force measurements, and can easily be extended to investigate the mechanics of other subcellular compartments, e.g., mitochondria, stress-fibers, and endosomes.

Here, we present a protocol to investigate the intracellular mechanical properties of isolated embryonic zebrafish cells in three-dimensional confinement with direct force measurement by an optical trap.

During the development of a multicellular organism, a single fertilized cell divides and gives rise to multiple tissues with diverse functions. Tissue ...

Refined Multiview Light Sheet Microscopy Protocol for Zebrafish Embryos

Light Sheet Microscopy Imaging and Mounting Strategies for Early Zebrafish Embryos

Light sheet microscopy has become the methodology of choice for live imaging of zebrafish embryos over long time scales with minimal phototoxicity. In particular, a multiview system, which allows sample rotation, enables imaging of entire embryos from different angles. However, in most imaging sessions with a multiview system, sample mounting is a troublesome process as samples are usually prepared in a polymer tube. To aid in this process, this protocol describes basic mounting strategies for imaging early zebrafish development between the 70% epiboly and early somite stages. Specifically, the study provides statistics on the various positions the embryos default to at the 70% epiboly and bud stages within the chorion. Furthermore, it discusses the optimum number of angles and the interval between angles required for imaging whole zebrafish embryos at the early stages of development so that cellular-scale information can be extracted by fusing the different views. Finally, since the embryo covers the entire field of view of the camera, which is required to obtain a cellular-scale resolution, this protocol details the process of using bead information from above or below the embryo for the registration of the different views.

Author Spotlight: Strategies for Mounting Zebrafish Embryos for High-Resolution Multiview Light-Sheet Microscopy &#8212; Techniques for Imaging and Image Reconstruction

A sample preparation strategy for imaging early zebrafish embryos within an intact chorion using a light-sheet microscope is described. It analyzes the different orientations that embryos acquire within the chorion at the 70% epiboly and bud stages and details imaging strategies for obtaining cellular-scale resolution throughout the embryo using the light-sheet system.

Light sheet microscopy has become the methodology of choice for live imaging of zebrafish embryos over long time scales with minimal phototoxicity. In ...

An Efficient Protocol to Assess ERK Activity Modulation in Early Zebrafish Noonan Syndrome Models via Live FRET Microscopy and Immunofluorescence

RASopathies are genetic syndromes caused by ERK hyperactivation and resulting in multisystemic diseases that can also lead to cancer predisposition. Despite a broad genetic heterogeneity, germline gain-of-function mutations in key regulators of the RAS-MAPK pathway underlie the majority of the cases, and, thanks to advanced sequencing techniques, potentially pathogenic variants affecting the RAS-MAPK pathway continue to be identified. Functional validation of the pathogenicity of these variants, essential for accurate diagnosis, requires fast and reliable protocols, preferably in vivo. Given the scarcity of effective treatments in early childhood, such protocols, especially if scalable in cost-effective animal models, can be instrumental in offering a preclinical ground for drug repositioning/repurposing. 
Here we describe step-by-step the protocol for rapid generation of transient RASopathy models in zebrafish embryos and direct inspection of live disease-associated ERK activity changes occurring already during gastrulation through real-time multispectral F&#246;rster resonance energy transfer (FRET) imaging. The protocol uses a transgenic ERK reporter recently established and integrated with the hardware of commercial microscopes. We provide an example application for Noonan syndrome (NS) zebrafish models obtained by expression of the Shp2D61G. We describe a straightforward method that enables registration of ERK signal change in the NS fish model before and after pharmacological signal modulation by available low-dose MEK inhibitors. We detail how to generate, retrieve, and assess ratiometric FRET signals from multispectral acquisitions before and after treatment and how to cross-validate the results via classical immunofluorescence on whole embryos at early stages. We then describe how, via examining standard morphometric parameters, to query late changes in embryo shape, indicative of a resulting impairment of gastrulation, in the same embryos whose ERK activity is assessed by live FRET at 6 h post fertilization.

RASopathies are multisystem genetic syndromes caused by RAS-MAPK pathway hyperactivation. Potentially pathogenic variants awaiting validation emerge continuously while poor preclinical evidence limits therapy. Here, we describe our in vivo protocol to test and cross-validate RASopathy-associated ERK activation levels and its pharmacological modulation during embryogenesis by live FRET imaging in Teen-reporter zebrafish.

RASopathies are genetic syndromes caused by ERK hyperactivation and resulting in multisystemic diseases that can also lead to cancer predisposition. ...

Biochemistry

Elucidating Microglia Behavior in Zebrafish Optic Tectum Development

Live Imaging and Characterization of Microglia Dynamics in the Zebrafish Embryo

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

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

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