We thank Albert Pan and Arndt Sieakmann for gifts of transgenic fish. We thank Koichi Kawakami at National Institute of Genetics, the National BioResource Project from the Ministry of Education, Culture, Sports, Science and Technology of Japan for the gift of the transgenic zebrafish HGn39b. We also thank Fatima Merchant and Kathleen Gajewski for assistance on confocal microscopy, and Tracey Theriault for photographs.
This work was supported by grants from the National Institute of Environmental Health Sciences of the National Institutes of Health (grant number P30ES023512 and contract number HHSN273201500010C). SU was supported by a fellowship from the Keck Computational Cancer Biology program (Gulf Coast Consortia CPRIT grant RP140113) and by Hugh Roy and Lillie Endowment Fund. J-&#197; G was supported by the Robert A. Welch Foundation (E-0004).

The animal work presented here was approved by the Institutional Animal Care and Use Committees (IACUCs) of the University of Houston and Indiana University.
1. Fish husbandry
NOTE: Work with vertebrate models requires an IACUC approved protocol. It should be conducted according to relevant national and international guidelines.
<ol>
	<li>Maintain adult zebrafish as described in previously published literature9.</li>
	<li>In the afternoon, place adult zebrafish in breeding tanks. Breed males to females at a ratio of 1:2.</li>
</ol>
2. Preparation of solutions
<ol>
	<li>Make a stock solution of 1% low melt agarose in embryo media (E3: 5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4, adjusted to pH 7.0). Aliquot the stock solution into 1.5 mL tubes and store them at 4 &#176;C.</li>
	<li>Make a stock solution of 4% (w/v) Tricaine (MS-222) in distilled water. Store at 4 &#176;C in a dark bottle. 
	CAUTION: Tricaine is toxic and should be weighed and dissolved in a fume hood.</li>
	<li>Make a stock solution of 20 mM N-phenylthiourea (PTU) in distilled water. Store at -20 &#176;C.</li>
</ol>
3. Preparation of embryos
<ol>
	<li>After mating, harvest embryos in E3 in a Petri dish and incubate them at 26.5 &#176;C for about 28 h before mounting. 
	NOTE: This slows down the development of the embryos so that the embryos are approximately at 30 somite stage at the beginning of imaging.</li>
	<li>Anesthetize embryos in 0.016-0.020% Tricaine in E3. To inhibit pigmentation, add PTU to a concentration of 200 &#181;M.</li>
	<li>Dechorionate the embryos using forceps under a dissecting microscope. Using two forceps, grip and gently pull the chorion apart to release the embryo.</li>
</ol>
4. Mounting in agarose
NOTE: The developed mounting method requires two different concentrations of low-melt agarose in E3 with 0.02% Tricaine and PTU as needed. The first agarose solution contains an optimal concentration of agarose at which the distortion and motility are at a minimum. The optimization is described in step 5 below.
<ol>
	<li>Heat the agarose solutions for the two layers (concentration defined in step 5 below and 1%) to 65 &#176;C. Let the agarose cool down to approximately 30 &#176;C just before mounting so that the embryo is not harmed by the heat. For mounting, use 35 mm glass bottom dishes with a No. 0 cover glass bottom. The cover glass attached to the bottom of the dish creates a 10 mm shallow (approx. 1.2 mm deep) well, in which the embryo is to be placed. 
	NOTE: In this case, the concentration with the least motility and distortions were between 0.025 to 0.040% agarose.</li>
	<li>Gently place a dechorionated embryo with one of its lateral sides toward the bottom of the dish using a glass pipette or micropipette. If using a micropipette, cut the outer part of the tip to increase the size of the opening to fit the embryo (Figure 1A). Carefully remove any remaining E3 with a micropipette.</li>
	<li>Add the first agarose solution to the small well created by the cover glass attached to the bottom of the dish to cover the embryo (Layer 1) (Figure 1B). Ensure that the agarose covers the small well but will not overflow it.</li>
	<li>Cover the small well with a cover glass (22 mm x 22 mm) (Figure 1C) to create a narrow agarose filled space with the embryo between the two cover glasses.</li>
	<li>Place a layer of 1% agarose solution on top of the cover glass all over the bottom of the dish (Layer 2) (Figure 1D). As this layer solidifies, it holds the cover glass in place.</li>
	<li>Fill the remaining portion of the dish with E3 containing 0.02% Tricaine to keep the system hydrated (Layer 3) (Figure 1E). 
	NOTE: In this setup, the cover glass and 1% agarose protect the bottom layer from getting diluted.</li>
</ol>
5. Optimization of agarose solution for layer 1
<ol>
	<li>To identify the optimal concentration of agarose for Layer 1, use a multiscale grid search approach. Mount embryos in increasing concentrations of agarose ranging from 0.01% to 1% followed by time-lapse imaging of embryo growth restriction and motility in the field of view. Identify the concentrations where both the distortion and motility are at a minimum.</li>
	<li>To optimize the concentration of agarose further, mount the embryos using a finer range of concentrations of agarose (e.g., between 0.025 and 0.040% agarose) depending on the concentration found to be best in step 5.1 (e.g., 0.025%, 0.028%, 0.031%, etc.). 
	NOTE: In our laboratory, the optimal agarose concentration was around 0.03%.</li>
</ol>
6. Time-lapse imaging
NOTE: This mounting method works for any inverted microscope with time-lapse functionality for fluorescence and bright field imaging.
<ol>
	<li>Perform time-lapse imaging for the whole embryo or parts of it for up to 55 h. For imaging of several embryos, use a stage adaptor that holds several dishes with one embryo mounted per dish. Rotate the dishes one by one so that the embryos are approximately horizontally positioned as determined by eye. For optimal embryo growth and development, use a microscope stage with an incubator set at 28.5 &#176;C.</li>
	<li>Select a low magnification objective in the microscope software. Under the eye piece, locate and finely align the embryos horizontally by rotating the dishes using bright field illumination and record their locations in the software. Create a filename for automatic saving of the data. 
	NOTE: In this experiment, a plan apo &#955; 4x 0.2 NA objective and the XY tab within the ND acquisition window were used to record the locations of the embryos. Data were saved in .nd2 format. The objective was selected by clicking on its icons in the Ti Pad tab.</li>
	<li>Set the pinhole size, scan speed, image size, and zoom. Next, select a higher magnification objective for capturing the images. 
	NOTE: In this experiment, a plan-apo 10x 0.45 NA objective was used for imaging of whole embryos, and a super-fluor 20x 0.75 NA objective was used for the higher magnification imaging of parts of the embryo. The pinhole was set to 1.2 AU (19.2 &#181;m for the 10x lens), the scan speed was set for a pixel dwell time of 2.4 &#181;s and the image size was set to 1024 x 1024 pixels with a scan zoom of one, giving a pixel size of 1.24 &#181;m for the 10x lens and 0.62 &#181;m for the 20x lens.</li>
	<li>Select the channels for the fluorescence to be imaged. Adjust the image capture settings one channel at a time. For each fluorescence channel adjust the laser power and the detector high voltage, making sure to collect the best dynamic range possible while avoiding the saturation and limiting photobleaching. 
	NOTE: In this experiment, GFP was imaged with the 488 green channel, (488 nm laser and emission between 500 and 550 nm), and RFP with the 561 red channel (561 nm laser and emission between 570 and 600 nm), and the transmission image was collected using the 561 nm laser and the transmitted light detector (TD channel).</li>
	<li>To image the whole embryo with the 10x objective, image several fields of view with overlap and stitch them together using the microscope software. 
	NOTE: As the embryos will grow substantially in size during the imaging, make sure that there is space in the field of view anterior and posterior to the embryo. Use the Scan Large Image tab of the ND acquisition window and select a 4 x 1 pattern with 10% overlap to capture four adjacent fields of view.</li>
	<li>Configure the settings for capturing the Z-stacks using the microscope software. 
	NOTE: Because the embryo is mounted close to the bottom of the glass dish, its center will move away from the bottom as it grows. Use the Z tab of the ND acquisition window and select the Asymmetrical relative range. With this method, the current focus plane is used as the Reference plane and the rest of the planes are asymmetrically distributed above and below to include the whole embryo within the Z-stack volume, with enough space to account for the specimen&#39;s growth. In all experiments, the interval was set to 11 &#181;m and the total range to about 45 planes.</li>
	<li>In order to maintain the focus of multiple embryos during long-term time-lapse imaging, use an automated focus. If imaging several embryos, adjust the individual offset levels for each embryo to focus at the Reference plane. 
	NOTE: In this experiment, a laser-based focus stabilization system was used.</li>
	<li>Set up time-lapse parameters in the microscope software and image for a duration of 55 h at about 1 h intervals. At each cycle, capture two embryo datasets sequentially and save data automatically after each cycle.</li>
	<li>Process images using an on-line or off-line version of the microscope software. If more than one embryo was imaged, create independent files for each embryo by splitting the dataset based on the location of imaging. Use maximum intensity projections or a similar tool to convert 3D time data sets into 2D time data sets. Create movie files by using the Save as menu option, selecting .avi as the file format. Select the No compression option with 200 ms intervals for a playback speed of 5 frames /s. 
	NOTE: The original dataset of two embryos in this experiment was split using the Split locations function in the software (File | Import/Export | Split Multipoints). 3D time datasets were converted to 2D time data sets using the Maximum intensity projection function (Image | ND Processing| Create Maximum Intensity Projection Image in Z).</li>
</ol>

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	<li>Kimmel, C. B., Ballard, W. W., Kimmel, S. R., Ullmann, B., Schilling, T. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stages+of+embryonic+development+of+the+zebrafish.">Stages of embryonic development of the zebrafish.</a> Developmental Dynamics. 203 (3), 253-310 (1995).</li><li>Kaufmann, A., Mickoleit, M., Weber, M., Huisken, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multilayer+mounting+enables+long-term+imaging+of+zebrafish+development+in+a+light+sheet+microscope.">Multilayer mounting enables long-term imaging of zebrafish development in a light sheet microscope.</a> Development. 139 (17), 3242-3247 (2012).</li><li>Schmid, 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=High-speed+panoramic+light-sheet+microscopy+reveals+global+endodermal+cell+dynamics.">High-speed panoramic light-sheet microscopy reveals global endodermal cell dynamics.</a> Nature Communications. 4, 2207 (2013).</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 in Molecular Biology. 546, 317-332 (2009).</li><li>Masselink, W., Wong, J. C., Liu, B., Fu, J., Currie, P. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Low-cost+silicone+imaging+casts+for+zebrafish+embryos+and+larvae.">Low-cost silicone imaging casts for zebrafish embryos and larvae.</a> Zebrafish. 11 (1), 26-31 (2014).</li><li>Wittbrodt, J. N., Liebel, U., Gehrig, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generation+of+orientation+tools+for+automated+zebrafish+screening+assays+using+desktop+3D+printing.">Generation of orientation tools for automated zebrafish screening assays using desktop 3D printing.</a> BMC Biotechnology. 14, 36 (2014).</li><li>Weijts, B., Tkachenko, E., Traver, D., Groisman, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Four-Well+Dish+for+High-Resolution+Longitudinal+Imaging+of+the+Tail+and+Posterior+Trunk+of+Larval+Zebrafish.">A Four-Well Dish for High-Resolution Longitudinal Imaging of the Tail and Posterior Trunk of Larval Zebrafish.</a> Zebrafish. 14 (5), 489-491 (2017).</li><li>Hirsinger, E., Steventon, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+Versatile+Mounting+Method+for+Long+Term+Imaging+of+Zebrafish+Development.">A Versatile Mounting Method for Long Term Imaging of Zebrafish Development.</a> Journal of Visualized Experiment. (119), (2017).</li><li>Avdesh, A., 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=Regular+care+and+maintenance+of+a+zebrafish+(Danio+rerio)+laboratory:+an+introduction.">Regular care and maintenance of a zebrafish (Danio rerio) laboratory: an introduction.</a> Journal of Visualized Experiment. (69), e4196 (2012).</li><li>McCollum, C. W., 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=Embryonic+exposure+to+sodium+arsenite+perturbs+vascular+development+in+zebrafish.">Embryonic exposure to sodium arsenite perturbs vascular development in zebrafish.</a> Aquatic Toxicology. 152, 152-163 (2014).</li><li>Pan, Y. A., 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=Zebrabow:+multispectral+cell+labeling+for+cell+tracing+and+lineage+analysis+in+zebrafish.">Zebrabow: multispectral cell labeling for cell tracing and lineage analysis in zebrafish.</a> Development. 140 (13), 2835-2846 (2013).</li><li>Asakawa, 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=Genetic+dissection+of+neural+circuits+by+Tol2+transposon-mediated+Gal4+gene+and+enhancer+trapping+in+zebrafish.">Genetic dissection of neural circuits by Tol2 transposon-mediated Gal4 gene and enhancer trapping in zebrafish.</a> Proceedings of the National Academy of Science U. S. A. 105 (4), 1255-1260 (2008).</li><li>Nagayoshi, S., 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=Insertional+mutagenesis+by+the+Tol2+transposon-mediated+enhancer+trap+approach+generated+mutations+in+two+developmental+genes:+tcf7+and+synembryn-like.">Insertional mutagenesis by the Tol2 transposon-mediated enhancer trap approach generated mutations in two developmental genes: tcf7 and synembryn-like.</a> Development. 135 (1), 159-169 (2008).</li><li>Huang, W. 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=Combined+use+of+MS-222+(tricaine)+and+isoflurane+extends+anesthesia+time+and+minimizes+cardiac+rhythm+side+effects+in+adult+zebrafish.">Combined use of MS-222 (tricaine) and isoflurane extends anesthesia time and minimizes cardiac rhythm side effects in adult zebrafish.</a> Zebrafish. 7 (3), 297-304 (2010).</li><li>Craig, M. P., Gilday, S. D., Hove, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dose-dependent+effects+of+chemical+immobilization+on+the+heart+rate+of+embryonic+zebrafish.">Dose-dependent effects of chemical immobilization on the heart rate of embryonic zebrafish.</a> Laboratory Animal (NY). 35 (9), 41-47 (2006).</li><li>Swinburne, I. A., Mosaliganti, K. R., Green, A. A., Megason, S. 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The authors have nothing to disclose.

A mounting method for extended time-lapse confocal microscopy of whole zebrafish embryos is described here. The most critical step for the mounting method is to identify the optimal concentration of agarose that will allow for unrestricted zebrafish embryo growth, and at the same time keep the embryos in a completely fixed position for confocal imaging. Because the optimal concentration of agarose is very narrow, this value is very sensitive to the errors in measurement of the weight of agarose and the volume of E3 durin...

The zebrafish has long been a model organism for developmental biology, and microscopy is the major method to visualize embryonic development. The advantages of using zebrafish embryos for developmental studies include small size, optical clarity, rapid development, and high fecundity of the adult fish. The generation of transgenic zebrafish lines expressing fluorescence in certain tissues or cells have allowed for a direct visualization of tissue development in ways not possible with larger vertebrate animals. In combination with time-lapse microscopy, details and dynamics of the tissue development can be readily studied.
A difficulty with imaging zebrafish development is that the embryos grow substantially in length; the embryo extends its length four times within the first 3 days of life1. Also, the body of the early embryo is soft, and easily becomes distorted if growth is restricted. Yet, to perform confocal microscopy, the embryo must be immobilized. To keep embryos in a fixed position for confocal time-lapse imaging, they are regularly anesthetized and mounted in 0.3-1% low-melt agarose. This has the advantage of allowing for some growth during imaging for a certain period of time, while restricting movements of the embryo. Parts of the embryo can efficiently be imaged like this. However, when using this method for imaging of the whole embryo for extended time periods, distortions are observed because of restricted growth caused by the agarose. Thus, other mounting methods are required. Kaufmann and colleagues have described an alternative mounting of zebrafish embryos for light sheet microscopy, such as selective plane illumination microscopy (SPIM), by mounting the embryos in fluorinated ethylene propylene (FEP) tubes containing low concentrations of agarose or methylcellulose2. This technique produces a superb visualization of embryogenesis over time. Schmid et al. describe mounting of up to six embryos in agarose in FEB tubes for light-sheet microscopy3 providing visualization of several embryos in one imaging session. Molds have been used to create embryo arrays for efficient mounting of larger numbers of embryos4. Masselkink et al. have constructed 3D printed plastic molds that can be used to make silicon casts that zebrafish embryos at different stages can be placed in, enabling mounting in a constant position for imaging, including confocal imaging5. 3D printing has also been used to make molds for consistent positioning of zebrafish embryos in 96-well format6. Some molds are customized for certain stages and may not permit unrestricted growth for long time periods, whereas other molds are more flexible. Recently, Weijts et al. published the design and fabrication of a four-well dish for live imaging of zebrafish embryos7. In this dish, the tail and trunk of anesthetized fish embryos are placed manually under a clear silicone roof attached just above a cover glass to form a pocket. The embryo is then fixed in this position by the addition of 0.4% agarose. This mounting allows for the imaging of the about 2 mm long posterior part (trunk and tail) of the embryo, and as up to 12 embryos can be mounted per well, the method allows for the imaging of multiple samples. Similarly, Hirsinger and Steventon recently presented a method where the head of the fish is mounted in agarose, while the tail can freely grow, and this method also efficiently facilitates imaging of the trunk and tail region of the embryo8.
This article describes a layered mounting method for zebrafish embryos that restrict the movements of the embryos while allowing for unrestricted growth. The advantages of this mounting method are that it is a low-cost, fast and easy method to mount embryos of various stages for imaging using any inverted microscope. The mounting permits long-term imaging of the whole body (head, trunk and tail) during embryo development. To showcase the usability of this method, whole embryo vascular, neuronal and muscle development was imaged in transgenic fish. Two embryos per session, at two wavelengths in 3D were imaged by time-lapse microscopy for 55 consecutive hours to render movies of tissue development.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Low melting agarose</td>
			<td>Sigma-Aldrich, MO</td>
			<td>A9414</td>
			<td>Store dissolved solution at 4 &#176;C</td>
		</tr>
		<tr>
			<td>35 mm glass bottom dishes with No. 0 coverslip and 10 mm diameter of glass bottom</td>
			<td>MatTek Corporation, MA</td>
			<td>P35GCOL-0-10-C</td>
			<td></td>
		</tr>
		<tr>
			<td>Tricaine (MS-222)</td>
			<td>Sigma-Aldrich, MO</td>
			<td>E10521</td>
			<td>Store dissolved solution at 4 &#176;C</td>
		</tr>
		<tr>
			<td>N-phenylthiourea (PTU)</td>
			<td>Sigma-Aldrich, MO</td>
			<td>P7629</td>
			<td>Store dissolved solution at -20 &#176;C</td>
		</tr>
		<tr>
			<td>Micro cover glass 22x22 mm</td>
			<td>VWR</td>
			<td>48366 067</td>
			<td></td>
		</tr>
		<tr>
			<td>Leica DMi8 fluorescence microscope</td>
			<td>Leica</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>LAS X software</td>
			<td>Leica</td>
			<td>NA</td>
			<td>Microscope software</td>
		</tr>
		<tr>
			<td>DMC4500 digital microscope camera</td>
			<td>Leica</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Nikon A1S confocal microscope</td>
			<td>Nikon Instruments Inc.</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Nikon NIS AR Version 4.40</td>
			<td>Nikon Instruments Inc.</td>
			<td>NA</td>
			<td>Microscope software</td>
		</tr>
	</tbody>
</table>

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.

Development of the mounting method 
The main aim of this work was to develop a low-cost mounting technique for time-lapse imaging of zebrafish development for extended periods of time. The layered mounting method was developed to allow for full growth of the fragile zebrafish embryo body, while restricting its movements. If the agarose concentration of layer 1 is too high, the embryos will become distorted and curved (Figure 2). Embryos grown at 0.1% and 0.5% agarose ha...

Watch this Scientific Journal Video about A Layered Mounting Method for Extended Time-Lapse Confocal Microscopy of Whole Zebrafish Embryos at JoVE.com

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

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 ...

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Developmental Biology

8.8K Views.  University of Houston. 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.

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

Capturing Tissue Repair in Zebrafish Larvae with Time-lapse Brightfield Stereomicroscopy

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two layers of skin that enclose undifferentiated mesenchyme, and because the larval tail fin regenerates rapidly within 2-3 days. Using this system, we demonstrate a method for capturing the repair dynamics of the amputated tail fin with time-lapse video brightfield stereomicroscopy. We demonstrate that fin amputation triggers a contraction of the amputation wound and extrusion of cells around the wound margin, leading to their subsequent clearance. Fin regeneration proceeds from proximal to distal direction after a short delay. In addition, developmental growth of the larva can be observed during all stages. The presented method provides an opportunity for observing and analyzing whole tissue-scale behaviors such as fin development and growth in a simple microscope setting, which is easily adaptable to any stereomicroscope with time-lapse capabilities.

We present a protocol for capturing the dynamics of zebrafish larval tail fin regeneration on a whole-tissue scale using brightfield-based stereomicroscopy. This technique enables capturing the regeneration dynamics with single cell resolution. This methodology can be adapted to any stereomicroscope equipped with a CCD camera and time-lapse software.

The zebrafish larval tail fin is ideal for studying tissue regeneration due to the simple architecture of the larval fin-fold, which comprises of two ...

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 ...

An Ex Vivo Method for Time-Lapse Imaging of Cultured Rat Mesenteric Microvascular Networks

Angiogenesis, defined as the growth of new blood vessels from pre-existing vessels, involves endothelial cells, pericytes, smooth muscle cells, immune cells, and the coordination with lymphatic vessels and nerves. The multi-cell, multi-system interactions necessitate the investigation of angiogenesis in a physiologically relevant environment. Thus, while the use of in vitro cell-culture models have provided mechanistic insights, a common critique is that they do not recapitulate the complexity associated with a microvascular network. The objective of this protocol is to demonstrate the ability to make time-lapse comparisons of intact microvascular networks before and after angiogenesis stimulation in cultured rat mesentery tissues. Cultured tissues contain microvascular networks that maintain their hierarchy. Immunohistochemical labeling confirms the presence of endothelial cells, smooth muscle cells, pericytes, blood vessels and lymphatic vessels. In addition, labeling tissues with BSI-lectin enables time-lapse comparison of local network regions before and after serum or growth factor stimulation characterized by increased capillary sprouting and vessel density. In comparison to common cell culture models, this method provides a tool for endothelial cell lineage studies and tissue specific angiogenic drug evaluation in physiologically relevant microvascular networks.

An Ex Vivo Method for Time-Lapse Imaging of Cultured Rat Mesenteric Microvascular Networks

Angiogenesis involves multi-cell, multi-system interactions that need to be investigated in a physiologically relevant environment. The objective of this study is to demonstrate the ability of the rat mesentery culture model to make time-lapse comparisons of intact microvascular networks during angiogenesis.

Angiogenesis, defined as the growth of new blood vessels from pre-existing vessels, involves endothelial cells, pericytes, smooth muscle cells, immune ...

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.

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. ...

Multi-Photon Time Lapse Imaging to Visualize Development in Real-time: Visualization of Migrating Neural Crest Cells in Zebrafish Embryos

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to numerous cell types throughout the body. Understanding the biology of the neural crest has been limited, reflecting a lack of genetically tractable models that can be studied in vivo and in real-time. Zebrafish is a particularly important developmental model for studying migratory cell populations, such as the neural crest. To examine neural crest migration into the developing eye, a combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time videos of the developing eye in transgenic zebrafish embryos, namely Tg(sox10:EGFP) and Tg(foxd3:GFP), as sox10 and foxd3 have been shown in numerous animal models to regulate early neural crest differentiation and likely represent markers for neural crest cells. Multi-photon time-lapse imaging was used to discern the behavior and migratory patterns of two neural crest cell populations contributing to early eye development. This protocol provides information for generating time-lapse videos during zebrafish neural crest migration, as an example, and can be further applied to visualize the early development of many structures in the zebrafish and other model organisms.

A combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time imaging of neural crest migration in Tg(sox10:EGFP) and Tg(foxd3:GFP) zebrafish embryos.

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to ...

Quantitative Whole-mount Immunofluorescence Analysis of Cardiac Progenitor Populations in Mouse Embryos

The use of ever-advancing imaging techniques has contributed broadly to our increased understanding of embryonic development. Pre-implantation development and organogenesis are two areas of research that have benefitted greatly from these advances, due to the high quality of data that can be obtained directly from imaging pre-implantation embryos or ex vivo organs. While pre-implantation embryos have yielded data with especially high spatial resolution, later stages have been less amenable to three-dimensional reconstruction. Obtaining high-quality 3D or volumetric data for known embryonic structures in combination with fate mapping or genetic lineage tracing will allow for a more comprehensive analysis of the morphogenetic events taking place during embryogenesis.
This protocol describes a whole-mount immunofluorescence approach that allows for the labeling, visualization, and quantification of progenitor cell populations within the developing cardiac crescent, a key structure formed during heart development. The approach is designed in such a way that both cell- and tissue-level information can be obtained. Using confocal microscopy and image processing, this protocol allows for three-dimensional spatial reconstruction of the cardiac crescent, thereby providing the ability to analyze the localization and organization of specific progenitor populations during this critical phase of heart development. Importantly, the use of reference antibodies allows for successive masking of the cardiac crescent and subsequent quantitative measurements of areas within the crescent. This protocol will not only enable a detailed examination of early heart development, but with adaptations should be applicable to most organ systems in the gastrula to early somite stage mouse embryo.

Here, we describe a protocol for whole mount immunofluorescence and image-based quantitative volumetric analysis of early stage mouse embryos. We present this technique as a powerful approach to qualitatively and quantitatively assess cardiac structures during development, and propose that it may be widely adaptable to other organ systems.

The use of ever-advancing imaging techniques has contributed broadly to our increased understanding of embryonic development. Pre-implantation development ...

Whole-mount Confocal Microscopy for Adult Ear Skin: A Model System to Study Neuro-vascular Branching Morphogenesis and Immune Cell Distribution

Here, we present a protocol of a whole-mount adult ear skin imaging technique to study comprehensive three-dimensional neuro-vascular branching morphogenesis and patterning, as well as immune cell distribution at a cellular level. The analysis of peripheral nerve and blood vessel anatomical structures in adult tissues provides some insights into the understanding of functional neuro-vascular wiring and neuro-vascular degeneration in pathological conditions such as wound healing. As a highly informative model system, we have focused our studies on adult ear skin, which is readily accessible for dissection. Our simple and reproducible protocol provides an accurate depiction of the cellular components in the entire skin, such as peripheral nerves (sensory axons, sympathetic axons, and Schwann cells), blood vessels (endothelial cells and vascular smooth muscle cells), and inflammatory cells. We believe this protocol will pave the way to investigate morphological abnormalities in peripheral nerves and blood vessels as well as the inflammation in the adult ear skin under different pathological conditions.

Here, we describe a high resolution whole-mount imaging method in the entire adult mouse ear skin, which enables us to visualize branching morphogenesis and patterning of peripheral nerves and blood vessels, as well as immune cell distribution.

Here, we present a protocol of a whole-mount adult ear skin imaging technique to study comprehensive three-dimensional neuro-vascular branching ...

Whole Mount Immunohistochemistry in Zebrafish Embryos and Larvae

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. Although many immunohistochemistry protocols have been optimized for mammalian tissue and tissue sections, these protocols often require modification and optimization for non-mammalian model organisms. Zebrafish are increasingly used as a model system in basic, biomedical, and translational research to investigate the molecular, genetic, and cell biological mechanisms of developmental processes. Zebrafish offer many advantages as a model system but also require modified techniques for optimal protein detection. Here, we provide our protocol for whole-mount fluorescence immunohistochemistry in zebrafish embryos and larvae. This protocol additionally describes several different mounting strategies that can be employed and an overview of the advantages and disadvantages each strategy provides. We also describe modifications to this protocol to allow detection of chromogenic substrates in whole mount tissue and fluorescence detection in sectioned larval tissue. This protocol is broadly applicable to the study of many developmental stages and embryonic structures.

Here, we present a protocol for fluorescent antibody-mediated detection of proteins in whole preparations of zebrafish embryos and larvae.

Immunohistochemistry is a widely used technique to explore protein expression and localization during both normal developmental and disease states. ...