Estelle Hirsinger: Core funding from the Institut Pasteur and Agence Nationale de la Recherche (ANR-10-BLAN-121801 DEVPROCESS). Estelle Hirsinger is from the Centre National de la Recherche Scientifique (CNRS). Benjamin Steventon was funded by the Agence Nationale de la Recherche (ANR- 10-BLAN-121801 DEVPROCESS), then a Roux fellowship (Institut Pasteur) then an AFM-Te&#769;le&#769;thon fellowship (number 16829). He is now supported by a Wellcome Trust/Royal Society Sir Henry Dale Fellowship.

1. Preparation of Solutions and Pulled Glass Needle
<ol>
	<li>Make a 25x stock solution of Tricaine (3-amino benzoic acid ethyl ester, also called ethyl 3-aminobenzoate) at 4 mg/mL in 20 mM Tris pH 8.8 and bring that solution is at pH 7. Aliquot by 4 mL and store at -20 &#176;C. 
	NOTE: The anesthetic Tricaine acts preferentially on neural voltage-gated sodium channels thereby blocking muscle twitching and movement6.</li>
	<li>Make a working solution of Tricaine at a final concentration of 0.17 mg/ml in E3 embryo medium.7 
	NOTE: Make the Tricaine working solution extemporaneously as the pH of this solution drifts.</li>
	<li>Dissolve the low melting point agarose to a final concentration of 1.5% in E3 embryo medium in a 50 mL tube by heating the solution in a microwave. Let this solution equilibrate to 42 - 45 &#176;C in either a water bath or bench-top incubator. If preparing large dishes for vertical light-sheet imaging (step 4), make an additional 25 mL of 1% standard melting agarose in E3 embryo medium.</li>
	<li>Use borosilicate glass with filament capillaries with dimensions of OD 1.20 mm, ID 0.69 mm, length 10 mm.</li>
	<li>If capillaries are pulled on the type of heating-filament needle puller described in the Table of Equipment and Reagents, use the following settings: Heat 600, Pull 120, Velocity 50, Time 225, Pressure 500. With a pair of sharp forceps, break the needle just past the point at which it flexes to create a clean and sharp needle for orientating embryos and removing excess agarose. A micro-scalpel can also be used to remove excess agarose.</li>
</ol>
2. Embedding of Embryos for Inverted or Upright Microscopy
<ol>
	<li>Raise the embryos up to the appropriate stage in E3 embryo medium.7</li>
	<li>At the required stage, dechorionate embryos with a pair of sharp forceps underneath a binocular dissecting microscope.</li>
	<li>Incubate the dechorionated embryos for at least 5 min in the Tricaine working solution.</li>
	<li>Once the melted agarose solution has cooled to 45 &#176;C, use a glass Pasteur pipette to transfer the dechorionated embryo directly into the 50 ml tube with minimal transfer of E3 medium.</li>
	<li>Remove embryo together with approximately 1 ml of mounting medium and add approx. 100 &#181;l of mounting medium together with the embryo to the central circle of a 35 mm glass-bottomed petri dish with a 10 mm microwell.</li>
	<li>As the mounting medium is setting, move the embryo to the edge of the circle of agarose with the tail facing outwards (Figure 1A). Use the capillary needle to maintain the embryo in the desired lateral orientation until the gel is completely set. To image posterior body development, take care to orientate the embryo in as lateral an orientation as possible. Therefore, maintain the embryo in this position by careful adjustments with the capillary needle. 
	NOTE: Multiple embryos can be added to the mounting dish at this point, if required. If this is the case, transfer the low-melt agarose into the dish first and add the embryos to the center of the agarose drop. Then, push the embryos out to the edges of the glass ring and orientate into the desired position. Up to six embryos can be mounted this way before the agarose becomes solidified.</li>
</ol>
3. Removal of Excess Agarose Around the Posterior Body
NOTE: This section describes the procedure by which agarose is removed from the region surrounding the posterior body. In the case of posterior body elongation, it is important to ensure that the tail can grow-out normally. By removing the agarose after the embryo has been completely included by agarose, the embryo is left enclosed by the head region and approximately half of the yolk sac.
<ol>
	<li>Once the agarose drop has set, flood the petri dish with the Tricaine working solution.</li>
	<li>Under a dissecting binocular microscope with a transmitted-light base, adjust the mirror position and angle of incident light such that the strong contrast allows for the cuts in the agarose to be seen clearly through the operation.</li>
	<li>To perform cut 1 (Figure 1), use the capillary needle or the micro-scalpel to cut the agarose mid-way along the yolk, from a position just posterior to the forming heart field to the position of the somite 5 (the 5th anterior somite).</li>
	<li>Start the initial cut adjacent to the forming heart as shown, and the full way through the agarose to the glass.</li>
	<li>Keep the capillary or micro-scalpel deep within the agarose, and slowly saw up and down while beginning to make a long cut over the embryo as shown Figure 1. Cut as close to the embryo as possible without puncturing the embryo or yolk.</li>
	<li>Next, make cuts 2 and 3 (Figure 1), starting from the position against the embryo and with the cut going all the way down to the glass bottomed dish. Make cut 2 so that it is tangential to the first 5 somites, allowing for full dorsalward unfolding of the posterior body. Make cut 3 at a 90&#176; angle with the yolk surface and starting from the heartfield. 
	NOTE: Cutting the agarose until the edge of the glass circle (as shown in Figure 1) aids in removing the agarose as a complete block (step 3.6). However, this is not necessary.</li>
	<li>Starting at the intersection of cuts 1 and 3, make a slow diagonal cut towards the end of cut 2 whilst slowly lifting upwards in order to dislodge the square of agarose surrounding the posterior body. 
	Notes: In some cases, this will be released in one block and the operation is completed in one go. In others, it may take several attempts to dis-lodge all the agarose surrounding the embryo.</li>
	<li>With a pair of sharp forceps, remove the dislodged blocks of agarose from the embryo medium. To aid in this process, move the agarose pieces to the side of the dish and use the wall of the petri dish as support whilst lifting out the agarose pieces.</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/55210/55210fig1.jpg" /> 
Figure 1: Diagrams of Mounting Set-ups. (A) The diagram shows the position of the mounted embryo within the center glass ring of a petri dish. On the right is a zoom of the embryo with each successive cut through the surrounding agarose shown with dottted red lines. (B) The mounted embryos is diagrammed in lateral view displaying the ease of access for both inverted and upright objectives. (C) A similar digram showing how embryos can be mounted for vertical light sheet imaging set-ups. <a href="http://cloudfront.jove.com/files/ftp_upload/55210/55210fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="9">
	<li>If muscle twitching is not completely blocked at that point, add drops of Tricaine from the stock solution of 4 mg/ml, pH 7.</li>
</ol>
4. Mounting of Embryos for Vertical Light-sheet Microscopy
NOTE: This is a variation on the method outlined above that allows for the access of multiple objectives for imaging of samples by vertically-orientated SPIM. The idea behind this variation is to lift the sample slightly higher than the bottom of the dish, to allow for easy access of two imaging objectives.
<ol>
	<li>Prior to sample mounting, coat a 100 mm plastic petri dish with 1% agarose in E3 medium to a height of 5 mm and allow to set.</li>
	<li>Place a drop of 1 ml low-melting point agarose to the center of the dish and allow to set.</li>
	<li>Embed embryos in low-melting point agarose as in section 2. However, this time, remove the embryo from the 50 ml tube of agarose with a smaller drop of solution (0.5 ml) and place this small drop on top of the 1 ml drop in the center of the dish.</li>
	<li>Use the capillary needle to place the embryo in the center of the small drop and maintain its correct orientation until gel is set.</li>
	<li>Flood the dish with Tricaine working solution and remove the excess agarose as in section 3 but without cutting through the cushion of 1% standard agarose.</li>
</ol>

<ol><li>Graeden, E., Sive, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Live+Imaging+of+the+Zebrafish+Embryonic+Brain+by+Confocal+Microscopy%5BTitle%5D)+AND+%22J+Vis+Exp%22%5BJournal%5D)">Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy.</a> J Vis Exp. (26), e1-e2 (2009).</li>
<li>Delaune, E. A., François, P., Shih, N. P., Amacher, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Single-Cell-Resolution+Imaging+of+the+Impact+of+Notch+Signaling+and+Mitosis+on+Segmentation+Clock+Dynamics%5BTitle%5D)+AND+%22Dev+Cell%22%5BJournal%5D)">Single-Cell-Resolution Imaging of the Impact of Notch Signaling and Mitosis on Segmentation Clock Dynamics.</a> Dev Cell. 23 (5), 995-1005 (2012).</li>
<li>Lawton, A. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Regulated+tissue+fluidity+steers+zebrafish+body+elongation%5BTitle%5D)+AND+%22Development%22%5BJournal%5D)">Regulated tissue fluidity steers zebrafish body elongation.</a> Development. 140 (3), 573-582 (2013).</li>
<li>Dray, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cell-Fibronectin+Interactions+Propel+Vertebrate+Trunk+Elongation+via+Tissue+Mechanics%5BTitle%5D)+AND+%22Curr+Biol%22%5BJournal%5D)">Cell-Fibronectin Interactions Propel Vertebrate Trunk Elongation via Tissue Mechanics.</a> Curr Biol. 23 (14), 1335-1341 (2013).</li>
<li>Steventon, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Species+tailoured+contribution+of+volumetric+growth+and+tissue+convergence+to+posterior+body+elongation+in+vertebrates%5BTitle%5D)+AND+%22Development%22%5BJournal%5D)">Species tailoured contribution of volumetric growth and tissue convergence to posterior body elongation in vertebrates.</a> Development. 143, 1732-1741 (2016).</li>
<li>Attili, S., Hughes, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Anaesthetic+Tricaine+Acts+Preferentially+on+Neural+Voltage-Gated+Sodium+Channels+and+Fails+to+Block+Directly+Evoked+Muscle+Contraction%5BTitle%5D)+AND+%22PLoS+ONE%22%5BJournal%5D)">Anaesthetic Tricaine Acts Preferentially on Neural Voltage-Gated Sodium Channels and Fails to Block Directly Evoked Muscle Contraction.</a> PLoS ONE. 9 (8), 103751-103756 (2014).</li>
<li>Westerfield, M. <a target="_blank" href="https://zfin.org/zf_info/zfbook/zfbk.html">The Zebrafish Book, 5th Edition; A guide for the laboratory use of zebrafish (Danio rerio)</a>. , University of Oregon Press. Eugene. Paperback (2007).</li>
<li>Schröter, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Dynamics+of+zebrafish+somitogenesis%5BTitle%5D)+AND+%22Dev+Dyn%22%5BJournal%5D)">Dynamics of zebrafish somitogenesis.</a> Dev Dyn. 237 (3), 545-553 (2008).</li>
<li>Megason, S. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((In+toto+imaging+of+embryogenesis+with+confocal+time-lapse+microscopy%5BTitle%5D)+AND+%22Methods+Mol+Biol+%28Clifton%2C+N.J.%29%22%5BJournal%5D)">In toto imaging of embryogenesis with confocal time-lapse microscopy.</a> Methods Mol Biol (Clifton, N.J.). 546, Chapter 19 317-332 (2009).</li>
<li>Sugiyama, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Illuminating+cell-cycle+progression+in+the+developing+zebrafish+embryo%5BTitle%5D)+AND+%22PNAS%22%5BJournal%5D)">Illuminating cell-cycle progression in the developing zebrafish embryo.</a> PNAS. 106 (49), 20812-20817 (2009).</li>
</ol>

The authors have nothing to disclose.

This mounting technique enables embryos to be kept still during transfer to the microscope and over long-term time-lapse imaging experiments aimed at following posterior body elongation at multiple length scales. Furthermore, it is versatile in that it allows for imaging on both upright and inverted microscopy set-ups, and a suggestion is made for how this can be further adapted to vertically orientated SPIM.
A critical step in this protocol is the careful removal of excess agarose surrounding...

Posterior body elongation is an essential process in embryonic development by which the embryo extends to form a large part of the body axis. It is an example of a complex morphogenetic process by which multiple cell behaviors act coordinately to generate the morphogenesis at the level of individual tissues. These differential tissue deformations then act together to generate the elongation of the posterior body at the whole structure level. To understand how these processes are controlled and coordinated during development, we must be able to follow these processes at multiple scales (i.e. at the level of molecules, cells, cell populations and tissues) and to relate this directly to the morphogenesis of the entire structure.
Zebrafish embryos are ideal for imaging posterior body elongation as their optical transparency and small size allows for the application of minimally invasive light imaging approaches well suited to live imaging.1 This has been evidenced by a series of recent publications that have shed light on posterior body development at the level of molecules,2 single cells,3 and inter-tissue behaviors,4 as well as at the level of cell population and whole organ.5
Advanced imaging techniques such as confocal, multi-photon microscopy and selective plane illumination microscopy (SPIM) are enabling the long term imaging of developmental processes with decreased effect of light toxicity and photo-bleaching. Robust techniques for the mounting of live samples are required to achieve three goals: 1) sufficient support to maintain samples in the correct orientation during transfer to the microscope and during acquisition, 2) sufficient freedom of movement of the sample to allow for the outgrowth of the posterior body region without affecting its normal development, and finally 3) a certain degree in versatility of the mounting method to allow imaging on diverse imaging set-ups.
This protocol introduces a mounting technique for imaging the development of the zebrafish D. rerio. This technique involves mounting embryos such that the head and yolk sac regions are almost entirely included in agarose, while leaving the posterior body region to elongate and develop normally. As such, it is also an appropriate method for the long-term imaging of other regions of the developing body as the agarose enables imaging by standard light imaging techniques. This protocol demonstrates mounting of embryos in a lateral orientation, although it is also possible to mount embryos in alterative orientations. It will further show how to adapt the method for upright, inverted and vertical light-sheet microscopy setups.

<table><tbody><tr><td>&lt;strong&gt;CONSUMABLES&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Glass-bottomed dishes</td><td>Mattek</td><td>P35-1.5-10-C</td><td>35 mm Petri dish, 10 mm microwell. No. 1.5 cover glass</td></tr><tr><td>Capillaries for injection needles</td><td>&amp;nbsp;Sutter&amp;nbsp;</td><td>BF 120-94-10</td><td>We use orosilicate glass with filament, OD 1.20 mm, ID 0.94 mm, length 10 cm. However, filament needles are not necessary and most injection standard needles should work.</td></tr><tr><td>Micro-scalpel</td><td>Feather</td><td>P-715</td><td>Micro Feather disposable opthalmic scalpel with plastic handle</td></tr><tr><td>Pasteur Pipettes</td><td></td><td></td><td>230 mm long</td></tr><tr><td>&lt;strong&gt;REAGENTS&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Tricaine</td><td>Sigma-Aldrich</td><td>A5040</td><td></td></tr><tr><td>Low-melting point agarose</td><td>Sigma-Aldrich</td><td>A9414</td><td></td></tr><tr><td>&lt;strong&gt;EQUIPMENT&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Fine forceps&amp;nbsp;</td><td>FINE SCIENCE TOOLS GMBH&amp;nbsp;</td><td>11252-30&amp;nbsp;</td><td>Dumont #5</td></tr><tr><td>Needle puller&amp;nbsp;</td><td>Sutter&amp;nbsp;</td><td>P97</td><td>Heating-filament needle puller</td></tr><tr><td>Binocular dissecting microscope</td><td>Leica</td><td>S8 Apo</td><td></td></tr></tbody></table>

a versatile mounting method for long term imaging of zebrafish development

Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. In particular, posterior body elongation is an essential process in embryonic development by which multiple tissue deformations act together to direct the formation of a large part of the body axis. In order to observe this process by long-term time-lapse imaging it is necessary to utilize a mounting technique that allows sufficient support to maintain samples in the correct orientation during transfer to the microscope and acquisition. In addition, the mounting must also provide sufficient freedom of movement for the outgrowth of the posterior body region without affecting its normal development. Finally, there must be a certain degree in versatility of the mounting method to allow imaging on diverse imaging set-ups. Here, we present a mounting technique for imaging the development of posterior body elongation in the zebrafish D. rerio. This technique involves mounting embryos such that the head and yolk sac regions are almost entirely included in agarose, while leaving out the posterior body region to elongate and develop normally. We will show how this can be adapted for upright, inverted and vertical light-sheet microscopy set-ups. While this protocol focuses on mounting embryos for imaging for the posterior body, it could easily be adapted for the live imaging of multiple aspects of zebrafish development.

The protocol outlined above details a versatile technique for the mounting of zebrafish embryos for long-term time lapse imaging. An example of this is shown in Figure 2A and in animated/video Figure 1. Embryos were injected at the 1 cell stage with mRNA encoding the photoconvertible fluorescent protein kikumeGR. At the 15 somite stage they were mounted as described above and imaged for 12 hr on an inverted confocal microscope with a 10X objective. The re...

Watch this Scientific Journal Video about A Versatile Mounting Method for Long Term Imaging of Zebrafish Development at JoVE.com

A Versatile Mounting Method for Long Term Imaging of Zebrafish Development

Here, we present a versatile mounting method that allows for the long-term time-lapse imaging of the posterior body development of live zebrafish embryos without perturbing normal development.

Zebrafish embryos offer an ideal experimental system to study complex morphogenetic processes due to their ease of accessibility and optical transparency. ...

a-versatile-mounting-method-for-long-term-imaging-zebrafish

Research

JoVE Journal

Developmental Biology

11.4K Views.  Institut Pasteur. Here, we present a versatile mounting method that allows for the long-term time-lapse imaging of the posterior body development of live zebrafish embryos without perturbing normal development.

Video: A Versatile Mounting Method for Long Term Imaging of Zebrafish Development

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 ...

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.  ...

Biology

A Layered Mounting Method for Extended Time-Lapse Confocal Microscopy of Whole Zebrafish Embryos

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues or cells. A difficulty with imaging whole embryo development is that zebrafish embryos grow substantially in length. When mounted as regularly done in 0.3-1% low melt agarose, the agarose imposes growth restriction, leading to distortions in the soft embryo body. Yet, to perform confocal time-lapse microscopy, the embryo must be immobilized. This article describes a layered mounting method for zebrafish embryos that restrict the motility of the embryos while allowing for the unrestricted growth. The mounting is performed in layers of agarose at different concentrations. To demonstrate the usability of this method, whole embryo vascular, neuronal and muscle development was imaged in transgenic fish for 55 consecutive hours. This mounting method can be used for easy, low-cost imaging of whole zebrafish embryos using inverted microscopes without requirements of molds or special equipment.

This article describes a method to mount fragile zebrafish embryos for extended time-lapse confocal microscopy. This low-cost method is easy to perform using regular glass-bottom microscopy dishes for imaging on any inverted microscope. The mounting is performed in layers of agarose at different concentrations.

Dynamics of development can be followed by confocal time-lapse microscopy of live transgenic zebrafish embryos expressing fluorescence in specific tissues ...

Visualizing the Developing Brain in Living Zebrafish using Brainbow and Time-lapse Confocal Imaging

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in vivo imaging techniques can provide a clear window into these processes in the living organism. For example, dividing cells and their progeny can be followed in real time as the nervous system forms. In recent years, technical advances in multicolor techniques have expanded the types of questions that can be investigated. The multicolor Brainbow approach can be used to not only distinguish among like cells, but also to color-code multiple different clones of related cells that each derive from one progenitor cell. This allows for a multiplex lineage analysis of many different clones and their behaviors simultaneously during development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells over real time within the developing zebrafish nervous system. This is particularly useful for following cellular interactions among like cells, which are difficult to label differentially using traditional promoter-driven colors. Our approach can be used for tracking lineage relationships among multiple different clones simultaneously. The large datasets generated using this technique provide rich information that can be compared quantitatively across genetic or pharmacological manipulations. Ultimately the results generated can help to answer systematic questions about how the nervous system develops.

In vivo imaging is a powerful tool that can be used to investigate the cellular mechanisms underlying nervous system development. Here we describe a technique for using time-lapse confocal microscopy to visualize large numbers of multicolor Brainbow-labeled cells in real time within the developing zebrafish nervous system.

Development of the vertebrate nervous system requires a precise coordination of complex cellular behaviors and interactions. The use of high resolution in ...

Multicolor Time-lapse Imaging of Transgenic Zebrafish: Visualizing Retinal Stem Cells Activated by Targeted Neuronal Cell Ablation

High-resolution time-lapse imaging of living zebrafish larvae can be utilized to visualize how biological processes unfold (for review see 1). Compound transgenic fish which express different fluorescent reporters in neighboring cell types provide a means of following cellular interactions 2 and/or tissue-level responses to experimental manipulations over time. In this video, we demonstrate methods that can be used for imaging multiple transgenically labeled cell types serially in individual fish over time courses that can span from minutes to several days. The techniques described are applicable to any study seeking to correlate the "behavior" of neighboring cells types over time, including: 1) serial 'catch and release' methods for imaging a large number of fish over successive days, 2) simplified approaches for separating fluorophores with overlapping excitation/emission profiles (e.g., GFP and YFP), 3) use of hypopigmented mutant lines to extend the time window available for high-resolution imaging into late larval stages of development, 4) use of membrane targeted fluorescent reporters to reveal fine morphological detail of individual cells as well as cellular details in larger populations of cells, and 5) a previously described method for chemically-induced ablation of transgenically targeted cell types; i.e., nitroreductase (NTR) mediated conversion of prodrug substrates, such as metronidazole (MTZ), to cytotoxic derivatives 3,5.
As an example of these approaches, we will visualize the ablation and regeneration of a subtype of retinal bipolar neuron within individual fish over several days. Simultaneously we will monitor several other retinal cell types, including neighboring non-targeted bipolar cells and potential degeneration-stimulated retinal stem cells (i.e., M&#971;ller glia). This strategy is being applied in our lab to characterize cell- and tissue-level (e.g., stem cell niche) responses to the selective loss and regeneration of targeted neuronal cell types.

In this video, techniques for multicolor confocal time-lapse imaging and targeted cell ablation are provided. Time-lapse imaging is used to monitor the behavior of multiple cell types of interest in vivo. Targeted cell ablation facilitates the study neural circuit function and cell-specific neuronal regeneration paradigms.

High-resolution time-lapse imaging of living zebrafish larvae can be utilized to visualize how biological processes unfold (for review see 1). Compound ...

Analysis of Zebrafish Cardiac Contraction with Virtual Reality Interaction

4D Light-sheet Imaging of Zebrafish Cardiac Contraction

Zebrafish is an intriguing model organism known for its remarkable cardiac regeneration capacity. Studying the contracting heart in vivo is essential for gaining insights into structural and functional changes in response to injuries. However, obtaining high-resolution and high-speed 4-dimensional (4D, 3D spatial + 1D temporal) images of the zebrafish heart to assess cardiac architecture and contractility remains challenging. In this context, an in-house light-sheet microscope (LSM) and customized computational analysis are&#160;used to overcome these technical limitations. This strategy, involving LSM system construction, retrospective synchronization, single cell tracking, and user-directed analysis, enables one to investigate the micro-structure and contractile function across the entire heart at the single-cell resolution in the transgenic Tg(myl7:nucGFP) zebrafish larvae. Additionally, we are able to further incorporate microinjection of small molecule compounds to induce cardiac injury in a precise and controlled manner. Overall, this framework allows one to track physiological and pathophysiological changes, as well as the regional mechanics at the single-cell level during cardiac morphogenesis and regeneration.

Author Spotlight: High-Resolution 4D Light-Sheet Imaging and Virtual Reality in Zebrafish for Single-Cell Analysis of Heart Function

This protocol utilizes light-sheet imaging to investigate cardiac contractile function in zebrafish larvae and gain insights into cardiac mechanics through cell tracking and interactive analysis.

Zebrafish is an intriguing model organism known for its remarkable cardiac regeneration capacity. Studying the contracting heart in vivo is essential for ...

Bioengineering

Multilayer Mounting for Long-term Light Sheet Microscopy of Zebrafish

Light sheet microscopy is the ideal imaging technique to study zebrafish embryonic development. Due to minimal photo-toxicity and bleaching, it is particularly suited for long-term time-lapse imaging over many hours up to several days. However, an appropriate sample mounting strategy is needed that offers both confinement and normal development of the sample. Multilayer mounting, a new embedding technique using low-concentration agarose in optically clear tubes, now overcomes this limitation and unleashes the full potential of light sheet microscopy for real-time developmental biology.

The development of zebrafish can be followed over days with light sheet microscopy when embryos are embedded in optically clear polymer tubes with low-concentration agarose.

Light sheet microscopy is the ideal imaging technique to study zebrafish embryonic development. Due to minimal photo-toxicity and bleaching, it is ...

4-Dimensional Imaging of Zebrafish Optic Cup Morphogenesis

Visual system function requires the establishment of precise tissue and organ structures. In the vertebrate eye, structural defects are a common cause of visual impairment, yet mechanisms of eye morphogenesis are still poorly understood. The basic organization of the embryonic eye is conserved throughout vertebrates, thus live imaging of zebrafish embryos has become a powerful approach to directly observe eye development at real time under normal and pathological conditions. Dynamic cell processes including movements, morphologies, interactions, division, and death can be visualized in the embryo. We have developed methods for uniform labeling of subcellular structures and timelapse confocal microscopy of early eye development in zebrafish. This protocol outlines the method of generating capped mRNA for injection into the 1-cell zebrafish embryo, mounting embryos at optic vesicle stage (~12 hours post fertilization, hpf), and performing multi-dimensional timelapse imaging of optic cup morphogenesis on a laser scanning confocal microscope, such that multiple datasets are acquired sequentially in the same imaging session. Such an approach yields data that can be used for a variety of purposes, including cell tracking, volume measurements, three-dimensional (3D) rendering, and visualization. Our approaches allow us to pinpoint the cellular and molecular mechanisms driving optic cup development, in both wild type and genetic mutant conditions. These methods can be employed directly by other groups or adapted to visualize many additional aspects of zebrafish eye development.

This protocol describes an approach for in toto labeling and multidimensional imaging of zebrafish early eye development. We describe labeling, embedding, and four dimensional (4D) imaging using laser scanning confocal microscopy, and considerations for optimizing acquisition of datasets for dissecting mechanisms of optic cup morphogenesis.

Visual system function requires the establishment of precise tissue and organ structures. In the vertebrate eye, structural defects are a common cause of ...

Neuroscience

Layered Agar Mounting: Preparing Live Zebrafish Embryos for Long-Term Imaging with an Inverted Microscope

Source: Upadhyay S., et.al. <a href="https://www.jove.com/v/60321/" target="_blank">A Layered Mounting Method for Extended Time-Lapse Confocal Microscopy of Whole Zebrafish Embryos</a>. J. Vis. Exp. (2020).
This article describes a method to mount live zebrafish embryos for long term imaging. This method is cost effective and easy to perform using regular glass-bottom microscopy dishes for imaging on Inverted Microscope. The mounting is performed in layers of agarose at different concentrations.

Source: Upadhyay S., et.al. A Layered Mounting Method for Extended Time-Lapse Confocal Microscopy of Whole Zebrafish Embryos. J. Vis. Exp. (2020).
This ...

A Versatile Mounting Method for Long Term Imaging of Zebrafish Development

Samenvatting

Meer video's verkennen