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="https://cloudfront-jove-com.remotexs.ntu.edu.sg/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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+Imaging+of+the+Zebrafish+Embryonic+Brain+by+Confocal+Microscopy.">Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy.</a> J Vis Exp. (26), e1-e2 (2009).</li><li>Delaune, E. A., Fran&#231;ois, P., Shih, N. P., Amacher, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-Cell-Resolution+Imaging+of+the+Impact+of+Notch+Signaling+and+Mitosis+on+Segmentation+Clock+Dynamics.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulated+tissue+fluidity+steers+zebrafish+body+elongation.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell-Fibronectin+Interactions+Propel+Vertebrate+Trunk+Elongation+via+Tissue+Mechanics.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Species+tailoured+contribution+of+volumetric+growth+and+tissue+convergence+to+posterior+body+elongation+in+vertebrates.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anaesthetic+Tricaine+Acts+Preferentially+on+Neural+Voltage-Gated+Sodium+Channels+and+Fails+to+Block+Directly+Evoked+Muscle+Contraction.">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="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Zebrafish Book, 5th Edition; A guide for the laboratory use of zebrafish (Danio rerio). , (2007).</li><li>Schr&#246;ter, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamics+of+zebrafish+somitogenesis.">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/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+toto+imaging+of+embryogenesis+with+confocal+time-lapse+microscopy.">In toto imaging of embryogenesis with confocal time-lapse microscopy.</a> Methods Mol Biol (Clifton, N.J.). 546, 317-332 (2009).</li><li>Sugiyama, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Illuminating+cell-cycle+progression+in+the+developing+zebrafish+embryo.">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 the posterior body that is important for allowing the normal development of this structure. It is important to take care here in not damaging the embryo, particularly when removing the agarose around the top of the yolk. In addition, care must be taken when removing the cut agarose block from around the embryo, as sometimes it is possible to lose the embryo from the mold during this process. For these reasons, this is a low-throughput method that is not suitable for the mounting of many zebrafish embryos simultaneously, in contrast to already published methods utilizing 3D printed plastic molds.8,9 However, embryos are highly stable by this mounting method and therefore can be transported to the microscope much easier, and retain their 3D orientation throughout posterior body elongation.
Another critical step is the accuracy of the initial lateral orientation of the embryo, avoiding any antero-posterior or dorso-ventral tilt. Either tilt will preclude the lateral vision of the posterior body extension, which is the most convenient angle of view for further analysis. The antero-posterior tilt will in addition result in the posterior body growing up or down with respect to the horizontal plane, which will mean the acquisition of higher stacks in z and thus time-lapses with a lower time resolution.
This method is versatile and has allowed for imaging of cell division rates with the use of the Fucci line5,10 as well as multi-scalar morphometric analyses.5 However, given the dramatic overall displacement of the tailbud during the elongation of the posterior body axis, only relatively low magnification objectives can be used to capture the entire process such as shown in Figure 2. This is because higher magnification results in the region of interest leaving the field of view within 3 - 4 hr of acquisition (Video Figure 1). Therefore, one way to further improve the imaging of the tailbud is to have an automated tracking algorithm that can track the overall movement of the tailbud and adjust the x,y,z position of the microscope stage during acquisition. Given the increased stability of the embryo in all axial dimensions with the mounting method described here, it should be amenable to such an approach.

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 border="0" cellpadding="0" cellspacing="0" width="727">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:231px;">CONSUMABLES</td>
			<td style="width:236px;"></td>
			<td style="width:135px;"></td>
			<td style="width:125px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Glass-bottomed dishes</td>
			<td>Mattek</td>
			<td>P35-1.5-10-C</td>
			<td>35mm petri dish, 10mm microwell. No. 1.5 cover glass</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Capillaries for injection needles</td>
			<td>&#160;Sutter&#160;</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 height="21">
			<td height="21" style="height:21px;">Micro-scalpel</td>
			<td>Feather</td>
			<td>P-715</td>
			<td>Micro Feather disposable opthalmic scalpel with plastic handle</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Pasteur Pipettes</td>
			<td></td>
			<td></td>
			<td>230 mm long</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">REAGENTS</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Tricaine</td>
			<td>Sigma-Aldrich</td>
			<td>A5040</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Low-melting point agarose</td>
			<td>Sigma-Aldrich</td>
			<td>A9414</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">EQUIPMENT</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Fine forceps&#160;</td>
			<td>FINE SCIENCE TOOLS GMBH&#160;</td>
			<td>11252-30&#160;</td>
			<td>Dumont #5</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Needle puller&#160;</td>
			<td>Sutter&#160;</td>
			<td>P97</td>
			<td>Heating-filament needle puller</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">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 resulting confocal stacks were maximally projected to be displayed as shown. This movie clearly shows that the posterior body is allowed to move freely throughout the duration of the movie and shows similar changes in morphology as seen in embryos that are allowed to develop freely of their chorion in normal culture conditions.
An additional example is shown in Figure 2B and animated/video Figure 2. Here, embryos were injected at the 16 cell stage with a combination of mRNAs encoding histone 2B-mCherry protein (that labels the nuclei) and eGFP:CAAX box (that labels membranes; for details see reference5). They were then imaged on an upright multiphoton microscope with a water immersion 25X objective as diagrammed in Figure 1B. Images were taken for 3 hr from the 10 somite stage in order to visualize cellular behaviors during tailbud formation. Cells can be seen to be generating active protrusions and directional movements as the tailbud forms normally.
<img alt="Figure 2" src="/files/ftp_upload/55210/55210fig2.jpg" /> 
Figure 2: Examples of Time-lapse Imaging of Posterior Body Elongation at 10X Magnification. (A) Consecutive frames of a representative movie covering the process of axial elongation. Embryos are shown in lateral view with posterior to the right side of the image. (B) Consecutive frames of a higher magnification movie showing individual cell behaviors during tailbud formation. Embryos are shown in lateral view with posterior to the right. Dotted lines outline the forming tailbud. Small colored lines show tracks of nuclei to follow individual cell movements. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55210/55210fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Video 1" src="/files/ftp_upload/55210/55210video1.jpg" /> 
Animated/Video Figure 1: Time-lapse Movie of Posterior Body Elongation of a KikumeGR mRNA Injected Zebrafish Embryo by Confocal Microscopy. Embryo is shown with anterior to the left and posterior to the right. Images were taken at 10 min intervals at 10X magnification. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55210/Video_Figure_1.mov" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Video 2" src="/files/ftp_upload/55210/55210video2.jpg" /> 
Animated/Video Figure 2: Time-lapse Movie of Posterior Body Elongation of a Nuclear-mCherry and Membrane-GFP Labelled Zebrafish Embryo Imaged by Two-photon Microscopy. Embryo is shown with anterior to the right and posterior to the left. Images were taken at 2 min intervals at 25X magnification. Colored lines display tracks of cells for the last ten time-steps for each tracked cell. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/55210/video_figure_2.mov" target="_blank">Please click here to view a larger version of this figure.</a>

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

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

11.3K 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 ...

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

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

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

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

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

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method described here allows time-lapse analysis of normal and impaired kidney development in zebrafish embryos by using a fluorescence dissecting microscope equipped for structured illumination and z-stack acquisition. To visualize nephrogenesis, transgenic zebrafish (Tg(wt1b:GFP)) with fluorescently labeled kidney structures were used. Renal defects were triggered by injection of an antisense morpholino oligonucleotide against the Wilms tumor gene wt1a, a factor known to be crucial for kidney development.
The advantage of the experimental setup is the combination of a zoom microscope with simple strategies for re-adjusting movements in x, y or z direction without additional equipment. To circumvent focal drift that is induced by temperature variations and mechanical vibrations, an autofocus strategy was applied instead of utilizing a usually required environmental chamber. In order to re-adjust the positional changes due to a xy-drift, imaging chambers with imprinted relocation grids were employed.
In comparison to more complex setups for time-lapse recording with optical sectioning such as confocal laser scanning&#160;or light sheet microscopes, a zoom microscope is easy to handle. Besides, it offers dissecting microscope-specific benefits such as high depth of field and an extended working distance.
The method to study organogenesis presented here can also be used with fluorescence stereo microscopes not capable of optical sectioning. Although limited for high-throughput, this technique offers an alternative to more complex equipment that is normally used for time-lapse recording of developing tissues and organ dynamics.

The method described here allows time-lapse analysis of organ development in zebrafish embryos by using a fluorescence dissecting microscope capable of performing optical sectioning and simple strategies of readjustment to correct focal and planar drift.

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method ...

A Submerged Filter Paper Sandwich for Long-term Ex Ovo Time-lapse Imaging of Early Chick Embryos

Due to its availability, low cost, flat geometry, and transparency, the ex ovo chick embryo has become a major vertebrate animal model for the study of morphogenetic events, such as gastrulation2, neurulation3-5, somitogenesis6, heart bending7,8, and brain formation9-13, during early embryogenesis. Key to understanding morphogenetic processes is to follow them dynamically by time-lapse imaging. The acquisition of time-lapse movies of chick embryogenesis ex ovo has been limited either to short time windows or to the need for an incubator to control temperature and humidity around the embryo14. Here, we present a new technique to culture chick embryos ex ovo for high-resolution time-lapse imaging using transmitted light microscopy. The submerged filter paper sandwich is a variant of the well-established filter paper carrier technique (EC-culture)1 and allows for the culturing of chick embryos without the need for a climate chamber. The embryo is sandwiched between two identical filter paper carriers and is kept fully submerged in a simple, temperature-controlled medium covered by a layer of light mineral oil. Starting from the primitive streak stage (Hamburger-Hamilton stage 5, HH5)15 up to at least the 28-somite stage (HH16)15, embryos can be cultured with either their ventral or dorsal side up. This allows the acquisition of time-lapse movies covering about 30 hr of embryonic development. Representative time-lapse frames and movies are shown. Embryos are compared morphologically to an embryo cultured in the standard EC-culture.
The submerged filter paper sandwich provides a stable environment to study early dorsal and ventral morphogenetic processes. It also allows for live fluorescence imaging and micromanipulations, such as microsurgery, bead implantation, microinjection, gene silencing, and electroporation, and has a strong potential to be combined with immersion objectives for laser-based imaging (including light-sheet microscopy).

A Submerged Filter Paper Sandwich for Long-term Ex Ovo Time-lapse Imaging of Early Chick Embryos

Here, we present a new technique to culture chick embryos ex ovo for time-lapse imaging using transmitted light and fluorescence microscopy. The submerged filter paper sandwich is a variant of the well-established filter paper carrier technique1 that allows for the culturing of chick embryos fully-submerged in a liquid culture medium.

Due to its availability, low cost, flat geometry, and transparency, the ex ovo chick embryo has become a major vertebrate animal model for the study of ...

A Simple Chamber for Long-term Confocal Imaging of Root and Hypocotyl Development

Several aspects of plant development, such as lateral root morphogenesis, occur on time spans of several days. To study underlying cellular and subcellular processes, high resolution time-lapse microscopy strategies that preserve physiological conditions are required. Plant tissues must have adequate nutrient and water supply with sustained gaseous exchange but, when submerged and immobilized under a coverslip, they are particularly susceptible to anoxia. One strategy that has been successfully employed is the use of a perfusion system to maintain a constant supply of oxygen and nutrients. However, such arrangements can be complicated, cumbersome, and require specialized equipment. Presented here is an alternative strategy for a simple imaging system using perfluorodecalin as an immersion medium. This system is easy to set up, requires minimal equipment, and is easily mounted on a microscope stage, allowing several imaging chambers to be set up and imaged in parallel. In this system, lateral root growth rates are indistinguishable from growth rates under standard conditions on agar plates for the first two days, and lateral root growth continues at reduced rates for at least another day. Plant tissues are supplied with nutrients via an agar slab that can be used also to administer a range of pharmacological compounds. The system was established to monitor lateral root development but is readily adaptable to image other plant organs such as hypocotyls and primary roots.

Presented here is a simple technique for high-resolution confocal time-lapse imaging of root and hypocotyl development for up to 3 days using high numerical-aperture objectives and perfluorodecalin as an immersion medium.

Several aspects of plant development, such as lateral root morphogenesis, occur on time spans of several days. To study underlying cellular and ...

Induction of Hypoxia in Living Frog and Zebrafish Embryos

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

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

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

Long-term Live Imaging of Drosophila Eye Disc

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have been used to study many biological processes, such as cell proliferation, differentiation, growth, migration, apoptosis, competition, cell-cell signaling, and compartmental boundary formation. However, methods for the long-term ex vivo culture and live imaging of the imaginal discs have not been satisfactory, despite many efforts. Recently, we developed a method for the long-term ex vivo culture and live imaging of imaginal discs for up to 18 h. In addition to using a high insulin concentration in the culture medium, a low-melting agarose was also used to embed the disc to prevent it from drifting during the imaging period. This report uses the eye-antennal discs as an example. Photoreceptor R3/4-specific m&#948;0.5-Ga4 expression was followed to demonstrate that photoreceptor differentiation and ommatidial rotation can be observed during a 10 h live imaging period. This is a detailed protocol describing this simple method.

Long-term Live Imaging of Drosophila Eye Disc

This protocol describes a detailed method for the long-term ex vivo culture and live imaging of a Drosophila imaginal disc. It demonstrates photoreceptor differentiation and ommatidial rotation within the 10 h period of live imaging of the eye disc. The protocol is simple and does not require expensive setup.

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have ...

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 ...

A Semi-high-throughput Imaging Method and Data Visualization Toolkit to Analyze C. elegans Embryonic Development

C. elegans is the premier system for the systematic analysis of cell fate specification and morphogenetic events during embryonic development. One challenge is that embryogenesis dynamically unfolds over a period of about 13 h; this half day-long timescale has constrained the scope of experiments by limiting the number of embryos that can be imaged. Here, we describe a semi-high-throughput protocol that allows for the simultaneous 3D time-lapse imaging of development in 80&#8211;100 embryos at moderate time resolution, from up to 14 different conditions, in a single overnight run. The protocol is straightforward and can be implemented by any laboratory with access to a microscope with point visiting capacity. The utility of this protocol is demonstrated by using it to image two custom-built strains expressing fluorescent markers optimized to visualize key aspects of germ-layer specification and morphogenesis. To analyze the data, a custom program that crops individual embryos out of a broader field of view in all channels, z-steps, and timepoints and saves the sequences for each embryo into a separate tiff stack was built. The program, which includes a user-friendly graphical user interface (GUI), streamlines data processing by isolating, pre-processing, and uniformly orienting individual embryos in preparation for visualization or automated analysis. Also supplied is an ImageJ macro that compiles individual embryo data into a multi-panel file that displays maximum intensity fluorescence projection and brightfield images for each embryo at each time point. The protocols and tools described herein were validated by using them to characterize embryonic development following knock-down of 40 previously described developmental genes; this analysis visualized previously annotated developmental phenotypes and revealed new ones. In summary, this work details a semi-high-throughput imaging method coupled with a cropping program and ImageJ visualization tool that, when combined with strains expressing informative fluorescent markers, greatly accelerates experiments to analyze embryonic development.

A Semi-high-throughput Imaging Method and Data Visualization Toolkit to Analyze C. elegans Embryonic Development

This work describes a semi-high-throughput protocol that allows simultaneous 3D time-lapse imaging of embryogenesis in 80&#8211;100 C. elegans embryos in a single overnight run. Additionally, image processing and visualization tools are included to streamline data analysis. The combination of these methods with custom reporter strains enables detailed monitoring of embryogenesis.

C. elegans is the premier system for the systematic analysis of cell fate specification and morphogenetic events during embryonic development. One ...

Development of a Larval Zebrafish Infection Model for Clostridioides difficile

Clostridioides difficile infection (CDI) is considered to be one of the most common healthcare-associated gastrointestinal infections in the United States. The innate immune response against C. difficile has been described, but the exact roles of neutrophils and macrophages in CDI are less understood. In the current study, Danio rerio (zebrafish) larvae are used to establish a C. difficile infection model for imaging the behavior and cooperation of these innate immune cells in vivo. To monitor C. difficile, a labeling protocol using a fluorescent dye has been established. A localized infection is achieved by microinjecting labeled C. difficile, which actively grows in the zebrafish intestinal tract and mimics the intestinal epithelial damage in CDI. However, this direct infection protocol is invasive and causes microscopic wounds, which can affect experimental results. Hence, a more noninvasive microgavage protocol is described here. The method involves delivery of C. difficile cells directly into the intestine of zebrafish larvae by intubation through the open mouth. This infection method closely mimics the natural infection route of C. difficile.

Development of a Larval Zebrafish Infection Model for Clostridioides difficile

Presented here is a safe and effective method to infect zebrafish larvae with fluorescently labeled anaerobic C. difficile by microinjection and noninvasive microgavage.

Clostridioides difficile infection (CDI) is considered to be one of the most common healthcare-associated gastrointestinal infections in the United ...

A Versatile Mounting Method for Long Term Imaging of Zebrafish Development

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Chapters in this video

Imaging Subcellular Structures in the Living Zebrafish Embryo

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

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

A Submerged Filter Paper Sandwich for Long-term Ex Ovo Time-lapse Imaging of Early Chick Embryos

A Simple Chamber for Long-term Confocal Imaging of Root and Hypocotyl Development

Induction of Hypoxia in Living Frog and Zebrafish Embryos

Long-term Live Imaging of Drosophila Eye Disc

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

A Semi-high-throughput Imaging Method and Data Visualization Toolkit to Analyze C. elegans Embryonic Development

Development of a Larval Zebrafish Infection Model for Clostridioides difficile