Name: multilayer mounting for long-term light sheet microscopy of zebrafish
Uploaded: 2014-02-27T14:15:00
Duration: 7 min 28 s
Description: Watch this Scientific Journal Video about Multilayer Mounting for Long-term Light Sheet Microscopy of Zebrafish at JoVE.com

We thank the Max Planck Society, the Human Frontier Science Program (HFSP) and the Boehringer Ingelheim Fonds for funding and support.

1. Material Preparation
<ol>
	<li>Preparation of FEP tubes
	<ol>
		<li>Clean the FEP tube (Figure 2A) by flushing liquids (in the steps noted below) through the tube and using a syringe attached with a blunt end cannula (Figure 2B). Note 1: Use gloves throughout the procedure. Note 2: We recommend cutting the tube in pieces of ca. 1 m in length and cleaning those in parallel.
		<ol>
			<li>First flush the tube with 1 M NaOH repeatedly. Then transfer the tube to a 50 ml centrifuge tube filled with 0.5 M NaOH and ultrasonicate it for 10 min.</li>
			<li>Transfer the polymer tube to a small basin with ddH2O. Flush the tube first with ddH2O and then with 70% EtOH.</li>
			<li>Subsequently transfer the FEP tube to a fresh centrifuge tube with 70% EtOH. Again ultrasonicate this centrifuge tube for 10 min.</li>
		</ol>
		</li>
		<li>Cut the cleaned FEP tube into pieces of about 3 cm (a typical length for use in zebrafish mounting and imaging) and straighten them if necessary. Store the cleaned FEP tubes in a fresh 50 ml centrifuge tube filled with ddH2O (Figure 2C).</li>
	</ol>
	</li>
	<li>Preparation of Tricaine stock solution
	<ol>
		<li>Prepare 0.4% Tricaine stock solution by dissolving 400 mg Tricaine (MS-222) in 97.9 ml ddH2O. After the powder has completely dissolved, adjust the pH to 7.0 using 2.1 ml 1M Tris (pH 9.0). Protect the solution from light. Store aliquots at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Preparation of 3% methylcellulose
	<ol>
		<li>Prepare 3% methylcellulose solution for the inner coating of the FEP tubes as described below. Heat up E311 in a glass bottle to approximately 60-70 &#176;C and add the appropriate amount of methylcellulose powder. Add a stirring bar and stir at 4 &#176;C. As soon as the solution is cooled down to below 30 &#176;C, add Tricaine solution to a final concentration of 100 mg/L. Keep stirring at 4 &#176;C overnight.</li>
	</ol>
	</li>
	<li>Preparation of 1.5% and 0.1% agarose in E3
	<ol>
		<li>Dissolve the appropriate amount of low melting point agarose powder in a defined volume of E3 in a glass flask. Heat up the mixture in a microwave and shake it once in a while, until the agarose solution appears homogeneous, without any remaining crystals.</li>
		<li>Prepare 1 ml aliquots of 0.1% agarose solution in 1.5 ml reaction tubes. The aliquots can be stored at 4 &#176;C.</li>
		<li>Use the 1.5% solution to coat a plastic Petri dish. The thickness of the agarose layer should be ca. 2 mm. After the agarose is solidified, add E3 medium on top of the layer to prevent it from drying. The coated dish can be stored at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Preparation of zebrafish
	<ol>
		<li>Keep Zebrafish (Danio rerio) adults and embryos at 28.5 &#176;C and handle them according to the established protocols11,12.</li>
		<li>Set up male and female fish in the afternoon and separate them using a divider. Remove the divider the following morning to time the mating of the fish.</li>
		<li>Collect the embryos in a dish filled with E3 and examine them using a stereomicroscope.</li>
		<li>At 24 hpf (or the developmental stage of your interest), select the zebrafish embryo for fluorescence expression and transfer the positive ones to a new dish with fresh E3.</li>
		<li>Carefully dechorionate the embryos using two sharp forceps (Figures 2D and 2E). Use the first forceps to grab and hold the chorion and the second forceps to tear it open and pull it off. Note 1: Perform this step with the help of a stereomicroscope.</li>
	</ol>
	</li>
</ol>
2. Multilayer Mounting
<ol>
	<li>Coating the FEP tube
	<ol>
		<li>Attach a blunt end cannula to a syringe. Insert the cannula carefully for about 3 mm into one end of a cleaned and cut piece of FEP tube (Figures 2F and 2G). Note 1: Use gloves and avoid any bending or squeezing of the tube.</li>
		<li>Dip the free end of the FEP tube into 3% methylcellulose and fill the tube with the solution.</li>
		<li>Slowly release the methylcellulose by pushing on the syringe. Discard the solution.</li>
		<li>Repeat steps 2.1.2 and 2.1.3 using E3. Note 1: The coated tubes need to be used for mounting immediately.</li>
	</ol>
	</li>
	<li>Transferring the zebrafish to the agarose
	<ol>
		<li>Heat up one aliquot of 0.1% agarose to 70&#176;C in a heat block. Vortex the reaction tube briefly and transfer it to a heat block set to 38 &#176;C (Figure 2H).</li>
		<li>Take the reaction tube out from the heat block and let it cool down briefly. Add Tricaine to a final concentration of 133-200 mg/L and vortex again (Figure 2I).</li>
		<li>Select one of the dechorionated zebrafish embryos and transfer it to the reaction tube with the preheated 0.1% agarose using a glass Pasteur pipette (Figures 2J and 2K). Note 1: Try to transfer as little additional E3 as possible.</li>
	</ol>
	</li>
	<li>Mounting the zebrafish
	<ol>
		<li>Take up the embryo with the syringe-attached FEP tube (Figure 2L). The tube should be filled completely with 0.1% agarose , with the embryo positioned close to either end of the tube and with the head pointing towards the opening of the tube. Note 1: Make sure to take up some agarose before taking up the fish. Note 2: Pull very gently on the syringe to avoid bubbles. Note 3: Keep the tube in a horizontal orientation to avoid leakage of its content (Figures 2M and 2N).</li>
	</ol>
	</li>
	<li>Plugging the tube
	 <img alt="Figure 2" fo:content-width="5in" fo:src="/files/ftp_upload/51119/51119fig2highres.jpg" src="/files/ftp_upload/51119/51119fig2.jpg" /> 
	Figure 2. Multilayer mounting for zebrafish embryos. A) Fluorinated Ethylene Propylene (FEP) tubes on a cable drum before preparation. B) A syringe with a blunt end cannula attached. C) A cleaned and cut FEP tube. D, E) Dechorionation of 24 hpf zebrafish embryos. F) Attaching the tube to the blunt end cannula. G) The syringe with cannula and tube attached. H) An aliquot of 0.1% agarose inside a reaction tube while transferring it to a heat block. I) Vortexing of the melted agarose. J) Uptake of a dechorionated embryo using a glass pipette. K) Transfer of the embryo into the melted agarose. L) Intake of the embryo with surrounding agarose into the FEP tube. M, N) The zebrafish embryo inside the FEP tube. O) Cutting the tube between specimen and cannula. P) Plugging the tube by the use of an agarose-coated dish. Q) The embryo inside the plugged tube. R) The final sample preparation in a metal holder. <a href="https://www.jove.com/files/ftp_upload/51119/51119fig2highres.jpg" target="_blank">Click here to view larger image.</a> 
	<ol>
		<li>Use a razor blade to cut the tube off the cannula (Figure 2O).</li>
		<li>Slowly stick the end of the tube containing the fish through the entire layer of 1.5% agarose in the plastic dish to plug the tube (Figure 2P). Note 1: Avoid the formation of bubbles and try to obtain a straight plug surface by holding the tube exactly perpendicular to the agarose surface. Note 2: Rotate the tube slightly to obtain a nice plug.</li>
		<li>Keep the mounted specimen in a 1.5 ml reaction tube with E3 to avoid drying. The sample is now ready to be imaged. Before imaging, check whether the fish is sitting right on top of the plug (Figures 2Q and 2R). Make sure to also add Tricaine to the medium inside the imaging chamber at the same concentration as in the mounting medium.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Huisken, J., Stainier, D. Y. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Selective+plane+illumination+microscopy+techniques+in+developmental+biology.">Selective plane illumination microscopy techniques in developmental biology.</a> Development. 136 (12), 1963-1975 (2009).</li><li>Ahrens, M. B., Orger, M. B., Robson, D. N., Li, J. M., Keller, P. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whole-brain+functional+imaging+at+cellular+resolution+using+light-sheet+microscopy.">Whole-brain functional imaging at cellular resolution using light-sheet microscopy.</a> Nat. Methods. 10 (5), 413-420 (2013).</li><li>Swoger, J., Muzzopappa, M., L&#243;pez-Schier, H., Sharpe, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=4D+retrospective+lineage+tracing+using+SPIM+for+zebrafish+organogenesis+studies.">4D retrospective lineage tracing using SPIM for zebrafish organogenesis studies.</a> J. Biophoton. 4 (1-2), 122-134 (2011).</li><li>Jemielita, M., Taormina, M. J., DeLaurier, A., Kimmel, C. B., Parthasarathy, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparing+phototoxicity+during+the+development+of+a+zebrafish+craniofacial+bone+using+confocal+and+light+sheet+fluorescence+microscopy+techniques.">Comparing phototoxicity during the development of a zebrafish craniofacial bone using confocal and light sheet fluorescence microscopy techniques.</a> J. Biophoton. , 1-11 (2012).</li><li>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=Light+sheet+microscopy+for+real-time+developmental+biology.">Light sheet microscopy for real-time developmental biology.</a> Curr. Opin. Genet. Dev. 21 (5), 566-572 (2011).</li><li>Huisken, J., Swoger, J., Del Bene, F., Wittbrodt, J., Stelzer, E. H. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical+sectioning+deep+inside+live+embryos+by+selective+plane+illumination+microscopy.">Optical sectioning deep inside live embryos by selective plane illumination microscopy.</a> Science. 305 (5686), 1007-1009 (2004).</li><li>Keller, P. J., Stelzer, E. H. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Digital+scanned+laser+light+sheet+fluorescence+microscopy.">Digital scanned laser light sheet fluorescence microscopy.</a> Cold Spring Harb. Prot. 2010 (5), (2010).</li><li>Keller, P. J., Stelzer, E. H. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+in+vivo+imaging+of+entire+embryos+with+Digital+Scanned+Laser+Light+Sheet+Fluorescence+Microscopy.">Quantitative in vivo imaging of entire embryos with Digital Scanned Laser Light Sheet Fluorescence Microscopy.</a> Curr. Opin. 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The authors have nothing to disclose.

Advanced microscopy techniques such as light sheet microscopy enable us to record image data with high temporal and spatial resolution over the course of several days, but also require sample mounting techniques to improve. We present a detailed protocol of multilayer mounting for light sheet microscopy of zebrafish development. The method is straightforward to implement and we use it in our lab on a daily basis.
The FEP tubes
Key components of our mounting strategy are tubes made of the polymer Fluorinated Ethylene Propylene (FEP). We decided to use FEP primarily because of its refractive index of 1.338, which is similar to water and therefore ideally suited for light sheet microscopy with water-dipping objectives and specimen requiring aqueous media. The tubes with 0.8 mm inner diameter and 0.4 mm wall thickness fit the developing zebrafish very well, can be used with typical sample holders suited for 1.6 mm capillaries and are our first choice for the presented mounting method. We tested them thoroughly in our home-built setups as well as in the commercial system &#34;Lightsheet Z.1&#34; by Carl Zeiss Microscopy and found that FEP tubes offer long-term stability, have only minimal impact on optical performance and are nonfluorescent10. They are, however, not suited for applications with samples that require a different refractive index, e.g. cleared specimen.
Tubes as well as foils made of FEP are available in a variety of diameters and thicknesses. In addition, FEP can be melted at 260 &#176;C and thus offers virtually endless possibilities for custom-made sample mounting supports. The polymer tubes tend to be supplied nonsterile, which is why the cleaning protocol incorporates several steps to ensure best optical quality. The coating with methylcellulose ensures that no part of the growing zebrafish embryo sticks to the polymer.
The mounting medium
The decision for 0.1% agarose is based on extensive tests of growth rate and immobility in different concentrations of agarose and methylcellulose10. Agarose concentrations above 0.1% already showed a severe impact on the growth rate of 24 hpf zebrafish. Additionally we use low doses of the anesthetizing drug Tricaine. Tricaine blocks action potentials, thus inhibits the signal transmission between brain and muscles and suppresses muscle contractions16. A concentration of 133-200 mg/L ensures sufficient immobilization of zebrafish between 24-72 hpf. However, Tricaine and other anesthetizing drugs we tested show dose dependent side effects such as cardiac edema. Additionally the required dose of Tricaine varies with the developmental stage of the embryo. We therefore recommend reducing the Tricaine concentration to the amount needed for the developmental stages that are imaged. To ensure best performance and prevent the formation of toxic by-products, freshly prepared or defrosted Tricaine stock solution should be used, protected from light and not heated above 30 &#176;C.
The embedding in FEP tubes also allows for easy modification of the mounting medium to account for specific needs. For time-lapse imaging of the heart, we recommend using 3% methylcellulose and 100 mg/L Tricaine instead of 0.1% agarose and 200 mg/L for mounting. Methylcellulose is more viscous than 0.1% agarose thus providing higher immobility on its own and less Tricaine needs to be used to ensure confinement of the embryo.
Tricaine is sensitive to temperatures above 30 &#176;C and can also be diluted by the medium in the imaging chamber, leading to decreased performance and possible twitching of the embryo. Therefore, make sure to use Tricaine also in the imaging medium in which the mounted specimen is immersed. If your experiments require the use of higher temperatures, we recommend exchanging the medium in the imaging chamber either constantly by using a perfusion system or manually with a tube and a syringe between two time-points.
Critical steps
The critical steps during the process of sample mounting are the intake of the specimen into the tube and the insertion of the agarose plug. The intake defines the initial position of the embryo, which is cumbersome to change afterwards. Multiple specimens should be available to mount and the mounting quality should be monitored with a stereomicroscope. If the sample orientation is not satisfactory, altering the speed of drawing up the specimen and avoiding subsequent movements of the plunger may help as this often twists the embryo. The 1.5% agarose plug is necessary to avoid leakage of the 0.1% agarose. The surface of the plug inside the tube should be flat to not influence the sample orientation during development. To achieve a good surface, different thicknesses of the agarose layers need to be tested and the tube needs to be put into the agarose normal to the surface. Rotating the tube a little bit and pulling out the tube carefully releases the plug from the dish. The quality of the mounting should be checked with a stereomicroscope before imaging.
Multiview imaging
One major advantage of light sheet microscopy is multiview imaging, the ability to rotate the specimen, acquire z-stacks from multiple angles and subsequently register and fuse them. An established method to register the z-stacks in 3D space is bead-based registration using fluorescent beads surrounding the sample as fiduciary markers 17. This method however relies on a solid mounting medium such as 1.5% agarose and thus seems to be incompatible with the multilayer mounting presented here. But several alternative solutions work depending on the application. First, the fluorescent beads can be incorporated into the agarose plug, which has to be in the field of view during the acquisition. Second, the stage positions can be calibrated before the actual experiments using a 1.5% agarose column with fluorescent beads. Third, alternative markers such as fluorescent nuclei can be used to identify the position of the z-stacks relative to each other.
Evaporation
Typical light sheet microscopy setups use an open sample chamber, which inevitable leads to evaporation of the medium. Since the medium is essential for the survival of the specimen, to restrict evaporation of the mounting medium and for the correct refractive index between optics and sample, we recommend taking one or more of the following measures to prevent evaporation. First, the medium in the chamber can be constantly exchanged by a perfusion system. Second, the medium can be manually refilled. Third, the chamber can be covered with a lid or a flexible foil.
Oxygen supply
The polymer FEP was developed to withstand a large variety of chemicals and is therefore robust, chemically inert and nonpermeable to gas. In the case of multilayer mounting oxygen can only penetrate through the agarose plug and the agarose-air interface, but it remains to be shown if the oxygen supply in multilayer mounting is severely lower when compared to mounting in 1.5% agarose without supporting polymer tube. However, as mounting in 1.5% agarose can only be used for about two hours before the zebrafish morphology is affected multilayer mounting seems to be the overall superior solution for long-term imaging.
Imaging earlier stages of development
Multilayer mounting has now become our standard embedding technique for time-lapse light sheet microscopy of zebrafish older than 24 hpf. Besides that, we also use the polymer tubes for imaging of earlier stage embryos. The embryos remain in their chorions and are mounted in a FEP tube with an inner diameter of 1 mm and filled with E3. The zebrafish can freely develop inside the chorion and can still be imaged well using light sheet microscopy.
Outlook
The presented multilayer sample mounting unleashes the full potential of light sheet microscopy for real-time developmental biology with zebrafish. It can easily be adopted for other microscopy techniques such as optical projection tomography (OPT) and other organisms. We believe that the standard-sized FEP tubes are just the first steps towards sample mounting techniques tailored for specific needs.

To understand the complex events and interactions during embryogenesis, research in developmental biology is more and more moving from microscopy of single cells and organs to in vivo imaging of entire organisms. Traditional microscopy techniques typically fail to offer the spatial and temporal resolution to follow rapid events over a long period of time and often cause bleaching or phototoxic effects in the sample1. We and others found light sheet microscopy (Figure 1) to be the ideal technique for in vivo imaging of zebrafish2-4. The illumination of the sample with a thin sheet of laser light, orthogonal to the detection axis, offers fast optical sectioning with high spatial resolution as well as minimized photo-toxicity5. At the same time, due to this unique optical arrangement, traditional sample mounting techniques using Petri dishes or culture chambers are not suitable. In a typical light sheet microscope three translational motors and a rotational motor hold the sample, such that it can be precisely positioned, turned and viewed from any direction. In the past specimens for light sheet microscopy have often been embedded in 1-2% agarose inside a glass capillary6,7. In this case the solidified agarose cylinder with the embedded specimen is extruded out of the glass capillary into the sample chamber filled with water-like medium for imaging. Agarose has a refractive index close to water and is therefore perfectly suited for in vivo light sheet microscopy that facilitates water-corrected illumination- and detection optics. The sample is confined in the rigid agarose and can be imaged well for a short period of time, but morphogenetic changes and growth of the embryo are strongly impaired when imaging for several hours7,8. Although solutions for other applications have been developed9, the noninvasive long-term time-lapse zebrafish imaging potential of light sheet microscopy cannot be exploited using the conventional 1.5% agarose embedding.
We therefore developed a new mounting strategy for in vivo light sheet microscopy of zebrafish embryos10. The sample is mounted in 0.1% agarose with 200 mg/L Tricaine inside a tube made of the optically clear polymer Fluorinated Ethylene Propylene (FEP). The embedded embryo develops normally due to low viscosity of the medium. At the same time, the FEP tube confines the medium and the sample for the duration of the experiment. Here we provide a detailed protocol of the new mounting technique and present data reflecting its advantages over the conventional protocol. We demonstrate that with our method zebrafish development can be continuously imaged with high spatial and temporal resolution over a period of more than two days.
<img alt="Figure 1" fo:content-width="5in" fo:src="/files/ftp_upload/51119/51119fig1highres.jpg" src="/files/ftp_upload/51119/51119fig1.jpg" /> 
Figure 1. The principle of light sheet microscopy. A zebrafish mounted in a tube made of Fluorinated Ethylene Propylene (FEP) can be translated and rotated through the focal plane of the detection objective. A thin sheet of laser light (light sheet) is projected by the illumination objective into the focal plane of the detection objective and illuminates only a thin optical section of the specimen. The emitted fluorescence signal is collected by the detection objective and recorded on a camera chip.

<table border="0" cellpadding="0" cellspacing="0" width="1097">
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	</colgroup>
	<tbody>
		<tr height="52">
			<td height="52" style="height:52px;width:268px;">Fluorinated Ethylene Propylene (FEP) tubes</td>
			<td style="width:143px;">Bola</td>
			<td style="width:113px;">S 1815-04</td>
			<td style="width:573px;">0.8 / 1.6 mm inner / outer diameter, other sizes available</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Omnifix-F Solo 1 ml Syringe</td>
			<td style="width:143px;">B. Braun Melsungen AG</td>
			<td style="width:113px;">9161406 V</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;">Sterican Single-Use Cannula, blunt</td>
			<td style="width:143px;">B. Braun Melsungen AG</td>
			<td style="width:113px;">9180109</td>
			<td style="width:573px;">0.8 x 22 mm, other sizes available, outer diameter of cannula has to fit inner diameter of FEP tube</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">50 ml Polypropylene Centrifuge Tube</td>
			<td style="width:143px;">Corning</td>
			<td>430829</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">1 M NaOH</td>
			<td style="width:143px;"></td>
			<td style="width:113px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">0.5 M NaOH&#160;</td>
			<td style="width:143px;"></td>
			<td style="width:113px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">70% EtOH</td>
			<td style="width:143px;"></td>
			<td style="width:113px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:268px;">Methylcellulose</td>
			<td style="width:143px;">Sigma</td>
			<td>M0387</td>
			<td>supplied as powder</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:268px;">E3 medium (for zebrafish embryos)</td>
			<td style="width:143px;"></td>
			<td style="width:113px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:268px;">1.5 ml Reaction Tube</td>
			<td style="width:143px;">Eppendorf</td>
			<td style="width:113px;">3810X</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:268px;">Agarose, low gelling temperature</td>
			<td style="width:143px;">Sigma</td>
			<td>A9414</td>
			<td>supplied as powder</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:268px;">Petri Dish</td>
			<td style="width:143px;">Greiner</td>
			<td>633180</td>
			<td>plastic, 94 x 16 mm</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:268px;">Tricaine</td>
			<td style="width:143px;">Sigma</td>
			<td>E10521</td>
			<td style="width:573px;">synonyms:&#160; Ethyl 3-aminobenzoate methanesulfonate, MS-222, TS 222, Tricaine methanesulfonate&#160;</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:268px;">10x/0.3 W Microscope Objective Lens</td>
			<td style="width:143px;">Leica</td>
			<td style="width:113px;">HCX APO L</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:268px;">EMCCD Camera</td>
			<td style="width:143px;">Andor Technology</td>
			<td style="width:113px;">iXon 885 EMCCD</td>
			<td></td>
		</tr>
	</tbody>
</table>

multilayer mounting for long-term light sheet microscopy of zebrafish

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

Multiday Imaging of Zebrafish Vasculature Development
We use 1 dpf Tg(kdrl:GFP)13 zebrafish to demonstrate the advantages of the multilayer mounting strategy for long-term light sheet microscopy. The aim is to image the development of the vasculature in the entire zebrafish over a course of 2 days. The zebrafish embryo was mounted in 0.1% agarose inside a methylcellulose coated polymer tube. Here, the zebrafish is not restricted by the mounting medium and can develop normally. Its head raises, the tail extends and vascular sprouting appears unhindered (Figure 3).
<img alt="Figure 3" fo:content-width="5in" fo:src="/files/ftp_upload/51119/51119fig3highres.jpg" src="/files/ftp_upload/51119/51119fig3.jpg" /> 
Figure 3. Multiday imaging of zebrafish vasculature development. The vasculature of a multilayer mounted 1 dpf Tg(kdrl:GFP) 13 zebrafish develops unhindered in a light sheet microscope over the course of two days. A 488 nm laser light sheet excited the fluorescence and the signal was detected using 10X/0.3 W objective and an EMCCD camera. Z-stacks were acquired on four adjacent positions every 10 min and subsequently stitched with Fiji 15. Maximum intensity projections of a subset of time-points are shown. Scale bar: 500 &#956;m. <a href="https://www.jove.com/files/ftp_upload/51119/51119fig3highres.jpg" target="_blank">Click here to view larger image.</a>
Imaging of Early Zebrafish Embryogenesis
Here, we demonstrate the capability of FEP tubes as a mounting support for light sheet microscopy of early zebrafish embryogenesis during the first day after fertilization. The 8 hpf Tg(H2A:GFP)14 zebrafish embryo was mounted with its intact chorion inside an E3-filled tube with an inner diameter of 1 mm. That results in a slight clamping of the chorion. All developmental processes during embryogenesis are unaffected by the mounting and can be optimally visualized using light sheet microscopy. In this example, we imaged the specimen over a course of 12 hr (Figure 4). Typical shape changes during early embryogenesis, such as the increasing elongation of the yolk and thickening of tissue along the midline, would be oppressed when using the traditional mounting without chorion in 1.5% agarose.
<img alt="Figure 4" fo:content-width="5in" fo:src="/files/ftp_upload/51119/51119fig4highres.jpg" src="/files/ftp_upload/51119/51119fig4.jpg" /> 
Figure 4. Imaging of early zebrafish embryogenesis. A Tg(H2A:GFP)14 zebrafish passes through early embryogenesis. A 488 nm laser light sheet excited GFP and the fluorescence signal was detected with a 10X/0.3 W objective and an EMCCD camera. Z-stacks from two angles were acquired every 3 min over a course of 12 hr. Normalized maximum intensity projections of a subset of the time-lapse are shown. Visible motion blur at 19 hpf results from the embryo twitching during acquisition, as no anesthetic were used. Scale bar: 150 &#956;m. <a href="https://www.jove.com/files/ftp_upload/51119/51119fig4highres.jpg" target="_blank">Click here to view larger image.</a>

Watch this Scientific Journal Video about Multilayer Mounting for Long-term Light Sheet Microscopy of Zebrafish at JoVE.com

Multilayer Mounting for Long-term Light Sheet Microscopy of Zebrafish

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

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

multilayer-mounting-for-long-term-light-sheet-microscopy-of-zebrafish

Research

JoVE Journal

Biology

Long-term Imaging Mammalian Cells using Wide-Field Microscopy

Application of Light-cured Dental Adhesive Resin for Mounting Electrodes or Microdialysis Probes in Chronic Experiments

In chronic recording experiments, self-curing dental acrylic resins have been used as a mounting base of electrodes or microdialysis-probes. Since these acrylics do not bond to the bone, screws have been used as anchors. However, in small experimental animals like finches or mouse, their craniums are very fragile and can not successfully hold the anchors. In this report, we propose a new application of light-curing dental resins for mounting base of electrodes or microdialysis probes in chronic experiments. This material allows direct bonding to the cranium. Therefore, anchor screws are not required and surgical field can be reduced considerably. Past experiences show that the bonding effect maintains more than 2 months. Conventional resin's window of time when the materials are pliable and workable is a few minutes. However, the window of working time for these dental adhesives is significantly wider and adjustable.

In this report, we propose a new application of light-curing dental resins for mounting base of electrodes or microdialysis probes in chronic experiments. This material allows direct bonding to the cranium.

In chronic recording experiments, self-curing dental acrylic resins have been used as a mounting base of electrodes or microdialysis-probes. Since these ...

Preparation of Drosophila S2 cells for Light Microscopy

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature and genome sequence database. Cultured Drosophila S2 cells, the product of disassociated 20-24 hour old embryos1, possess all these properties. Consequently, S2 cells are extremely well-suited for the analysis of cellular processes, including the discovery of the genes encoding the molecular components of the process or mechanism of interest. The features of S2 cells that are most responsible for their utility are the ease with which they are maintained, their exquisite sensitivity to double-stranded (ds)RNA-mediated interference (RNAi), and their tractability to fluorescence microscopy as either live or fixed cells.
S2 cells can be grown in a variety of media, including a number of inexpensive, commercially-available, fully-defined, serum-free media2. In addition, they grow optimally and quickly at 21-24&deg;C and can be cultured in a variety of containers. Unlike mammalian cells, S2 cells do not require a regulated atmosphere, but instead do well with normal air and can even be maintained in sealed flasks.
Complementing the ease of RNAi in S2 cells is the ability to readily analyze experimentally-induced phenotypes by phase or fluorescence microscopy of fixed or live cells. S2 cells grow in culture as a single monolayer but do not display contact inhibition. Instead, cells tend to grow in colonies in dense cultures. At low density, S2 cultures grown on glass or tissue culture-treated plastic are round and loosely-attached. However, the cytology of S2 cells can be greatly improved by inducing them to flatten extensively by briefly culturing them on a surface coated with the lectin, concanavalin A (ConA)3. S2 cells can also be stably transfected with fluorescently-tagged markers to label structures or organelles of interest in live or fixed cells. Therefore, the usual scenario for the microscopic analysis of cells is this: first, S2 cells (which can possess transgenes to express tagged markers) are treated by RNAi to eliminate a target protein(s). RNAi treatment time can be adjusted to allow for differences in protein turn-over kinetics and to minimize cell trauma/death if the target protein is important for viability. Next, the treated cells are transferred to a dish containing a coverslip pre-coated with conA to induce cells to spread and tightly adhere to the glass. Finally, cells are imaged with the researcher's choice of microscopy modes. S2 cells are particularly good for studies requiring extended visualization of live cells since these cells stay healthy at room temperature and normal atmosphere.

Preparation of Drosophila S2 cells for Light Microscopy

Drosophila Schneider (S2) cells are an increasingly popular system for the discovery and functional analysis of genes. Our goal is to describe some of the microscopic techniques that make S2 cells such an increasingly important experimental system.

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature ...

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

Our research activities target the use of biological methods for the evaluation of environmental quality, with particular reference to saltwater/brackish water and sediment. The choice of biological indicators must be based on reliable scientific knowledge and, possibly, on the availability of standardized procedures. In this article, we present a standardized protocol that used the marine crustacean Artemia to evaluate the toxicity of chemicals and/or of marine environmental matrices. Scientists propose that the brine shrimp (Artemia) is a suitable candidate for the development of a standard bioassay for worldwide utilization. A number of papers have been published on the toxic effects of various chemicals and toxicants on brine shrimp (Artemia). The major advantage of this crustacean for toxicity studies is the overall availability of the dry cysts; these can be immediately used in testing and difficult cultivation is not demanded1,2. Cyst-based toxicity assays are cheap, continuously available, simple and reliable and are thus an important answer to routine needs of toxicity screening, for industrial monitoring requirements or for regulatory purposes3. The proposed method involves the mortality as an endpoint. The numbers of survivors were counted and percentage of deaths were calculated. Larvae were considered dead if they did not exhibit any internal or external movement during several seconds of observation4. This procedure was standardized testing a reference substance (Sodium Dodecyl Sulfate); some results are reported in this work. This article accompanies a video that describes the performance of procedural toxicity testing, showing all the steps related to the protocol.

Long-term Lethal Toxicity Test with the Crustacean Artemia franciscana

This study concerns the development and standardization of a valuable methodological protocol to determine long-term (14 days) lethal toxicity exerted by chemical substances, industrial wastewater or sewage and liquid environmental samples on the saltwater crustacean, Artemia franciscana.

Our research activities target the use of biological methods for the evaluation of environmental quality, with particular reference to saltwater/brackish ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Light Sheet Fluorescence Microscopy of Plant Roots Growing on the Surface of a Gel

One of the key questions in understanding plant development is how single cells behave in a larger context of the tissue. Therefore, it requires the observation of the whole organ with a high spatial- as well as temporal resolution over prolonged periods of time, which may cause photo-toxic effects. This protocol shows a plant sample preparation method for light-sheet microscopy, which is characterized by mounting the plant vertically on the surface of a gel. The plant is mounted in such a way that the roots are submerged in a liquid medium while the leaves remain in the air. In order to ensure photosynthetic activity of the plant, a custom-made lighting system illuminates the leaves. To keep the roots in darkness the water surface is covered with sheets of black plastic foil. This method allows long-term imaging of plant organ development in standardized conditions.

This protocol shows a plant sample preparation method for light-sheet microscopy. The setup is characterized by mounting the plant vertically on the surface of a gel and letting it grow in controlled bright conditions. This allows long-term observation of plant organ development in standardized conditions.

One of the key questions in understanding plant development is how single cells behave in a larger context of the tissue. Therefore, it requires the ...

LarvaSPA, A Method for Mounting Drosophila Larva for Long-Term Time-Lapse Imaging

Live imaging is a valuable approach for investigating cell biology questions. The Drosophila larva is particularly suited for in vivo live imaging because the larval body wall and most internal organs are transparent. However, continuous live imaging of intact Drosophila larvae for longer than 30 min has been challenging because it is difficult to noninvasively immobilizeimmobilizing larvae for a long time. Here we present a larval mounting method called LarvaSPA that allows for continuous imaging of live Drosophila larvae with high temporal and spatial resolution for longer than 10 hours. This method involves partially attaching larvae to the coverslip using a UV-reactive glue and additionally restraining larval movement using a polydimethylsiloxane (PDMS) block. This method is compatible with larvae at developmental stages from second instar to wandering third instar. We demonstrate applications of this method in studying dynamic processes of Drosophila somatosensory neurons, including dendrite growth and injury-induced dendrite degeneration. This method can also be applied to study many other cellular processes that happen near the larval body wall.

LarvaSPA, A Method for Mounting Drosophila Larva for Long-Term Time-Lapse Imaging

This protocol describes a method for mounting Drosophila larvae to achieve longer than 10 h of uninterrupted time-lapse imaging in intact live animals. This method can be used to image many biological processes close to the larval body wall.

Live imaging is a valuable approach for investigating cell biology questions. The Drosophila larva is particularly suited for in vivo live imaging because ...

Simultaneous Live Imaging of Multiple Insect Embryos in Sample Chamber-Based Light Sheet Fluorescence Microscopes

Light sheet-based fluorescence microscopy offers efficient solutions to study complex processes on multiple biologically relevant scales. Sample chamber-based setups, which are specifically designed to preserve the three-dimensional integrity of the specimen and usually feature sample rotation, are the best choice in developmental biology. For instance, they have been used to document the entire embryonic morphogenesis of the fruit fly Drosophila melanogaster and the red flour beetle Tribolium castaneum. However, many available live imaging protocols provide only experimental frameworks for single embryos. Especially for comparative studies, such approaches are inconvenient, since sequentially imaged specimens are affected by ambient variance. Further, this limits the number of specimens that can be assayed within a given time. We provide an experimental framework for simultaneous live imaging that increases the throughput in sample chamber-based setups and thus ensures&#160;similar ambient conditions for all specimens. Firstly, we provide a calibration guideline for light sheet fluorescence microscopes. Secondly, we propose a mounting method for multiple embryos that is compatible with sample rotation. Thirdly, we provide exemplary three-dimensional live imaging datasets of Drosophila, for which we juxtapose three transgenic lines with fluorescently labeled nuclei, as well as of Tribolium, for which we compare the performance of three transgenic sublines that carry the same transgene, but at different genomic locations. Our protocol is specifically designed for comparative studies as it pro-actively addresses ambient variance, which is always present in sequential live imaging. This is especially important for quantitative analyses and characterization of aberrational phenotypes, which result e.g., from knockout experiments. Further, it increases the overall throughput, which is highly convenient when access to light sheet&#160;fluorescence microscopes is limited. Finally, the proposed mounting method can be adapted for other insect species and further model organisms, e.g., zebrafish, with basically no optimization effort.

Light sheet-based fluorescence microscopy is the most valuable tool in developmental biology. A major issue in comparative studies is ambient variance. Our protocol describes an experimental framework for simultaneous live imaging of multiple specimens and, therefore, addresses this issue pro-actively.

Light sheet-based fluorescence microscopy offers efficient solutions to study complex processes on multiple biologically relevant scales. Sample ...

Light Sheet Microscopy of Fast Cardiac Dynamics in Zebrafish Embryos

Embryonic cardiac research has greatly benefited from advances in fast in vivo light sheet fluorescence microscopy (LSFM). Combined with the rapid external development, tractable genetics, and translucency of the zebrafish, Danio rerio, LSFM has delivered insights into cardiac form and function at high spatial and temporal resolution without significant phototoxicity or photobleaching. Imaging of beating hearts challenges existing sample preparation and microscopy techniques. One needs to maintain a healthy sample in a constricted field of view and acquire ultrafast images to resolve the heartbeat. Here we describe optimized tools and solutions to study the zebrafish heart in vivo. We demonstrate the applications of bright transgenic lines for labeling the cardiac constituents and present novel gentle embedding and immobilization techniques that avoid developmental defects and changes in heart rate. We also propose a data acquisition and analysis pipeline adapted to cardiac imaging. The entire workflow presented here focuses on zebrafish embryonic heart imaging but can also be applied to various other samples and experiments.

We describe optimized tools to study the zebrafish heart in vivo with light sheet fluorescence microscopy. Specifically, we suggest bright cardiac transgenic lines and present new gentle embedding and immobilization techniques that avoid developmental and heart defects. A possible data acquisition and analysis pipeline adapted to cardiac imaging is also provided.