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. Neurobiol. 18 (6), 624-632 (2008).</li><li>Desmaison, A., Lorenzo, C., Rouquette, J., Ducommun, B., Lobjois, V. <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+sample+holder+for+single+plane+illumination+microscopy.">A versatile sample holder for single plane illumination microscopy.</a> J. Microsc. 10, (2013).</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>Nusslein-Volhard, C., Zebrafish Dahm, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> , (2002).</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> J. Vis. Exp. (69), (2012).</li><li>Jin, S. -. W., Beis, D., Mitchell, T., Chen, J. -. N., 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=Cellular+and+molecular+analyses+of+vascular+tube+and+lumen+formation+in+zebrafish.">Cellular and molecular analyses of vascular tube and lumen formation in zebrafish.</a> Development. 132 (23), 5199-5209 (2005).</li><li>Pauls, S., Geldmacher-Voss, B., Campos-Ortega, J. 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+zebrafish+histone+variant+H2A.F/Z+and+a+transgenic+H2A.F/Z:GFP+fusion+protein+for+in+vivo+studies+of+embryonic+development.">A zebrafish histone variant H2A.F/Z and a transgenic H2A.F/Z:GFP fusion protein for in vivo studies of embryonic development.</a> Dev. 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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">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</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

18.7K Views.  Max Planck Institute of Molecular Cell Biology and Genetics. 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.

Video: Multilayer Mounting for Long-term Light Sheet Microscopy of Zebrafish

Long-term Imaging Mammalian Cells using Wide-Field Microscopy

Actually what I'm gonna do, mount it in here first and then swap out You. So now I'm gonna aspirate try and be as gentle as possible.Okay. A few seconds just holding.Okay.Alright.So the important features of our lifestyle imaging grid are that, so it's an inverted scope, which means the objective, the samples here in the objective is underneath. So the objective is pointing up and the whole scope, or practically the whole, practically the whole scope, including the objectives and the sample are at 37 degrees. So everything in this plastic box is at 37 degrees.And right on top of the stage we have an air and CO2 mix. That is 5%CO2. So the idea is that the sample is actually in a cell, it's like a cell incubator.And so we have, this is our mixer for the air and the CO2 and, and it gets humidified and heated and then piped onto the, the sample on the stage.Okay. And today we're just gonna be doing fluorescence one channel GFP fluorescence. This is the hose that the CO2 and 5%CO2 goes in and it just fits into the stage holder.That's important. And then we can close these, take these down, and then we go to our first position. And what I'm gonna do is go to the first field, which is my control sample, and I'll pick positions based on what I see on the screen.So I just make sure I actually look into the microscope in each new, well, because the focal plane for each of the wells is different, the, the dish is not perfectly flat on the bottom. So you will need to make fine adjustments when you move from well to the well. So right now I'm on the first, well, what I'll do is I'll focus, so I just hit, I just told, I just turned on the light, the transmitted light for the microscope.And I have make sure all the light is going to the eye pieces. And so I just look in, make sure I get a nice clean field. And then what I'll do is turn that light off, turn the fluorescence light on, make sure the cells that I'm looking at look healthy and they're not apoptotic.And, and I turn that off. And what I do next is send, so now I'm gonna start picking positions, which means I have to te send the light path to the camera. This is the camera right here.So I tell the microscope to send all the camera, they don't see anything in the eye pieces. What I'll do is in the software controlling the microscope, I'll choose the right filter set, which for our purposes is psy. And then set an exposure time.And I know these cells very well and I know that they take a 0.2 second exposure time. So I tell, that's what I tell the software to do. And what I'll do is hit focus.This is, and it's basically just alive. The, A light will turn on and you can, oh, I don't know if you can see cells, but the reason this is a good field is because there's, all the cells are, even in terms of their fluorescence, There are no apoptotic cells. Apoptotic cells tend to be very bright and there's no dirt or schmutz in the field.And you can see that they're actually, this is a metaphase cell and that's a metaphase cell, that's a cell that's probably just exited mitosis. And I can just tell that because it looks like those are, those two sets of chromatin are very nicely oriented to each other. Like they just exited an face.So that's a good field. So, and now that I know that the exposure time and the filter set is correct, and this is gonna be software specific, I'll go and I'll take positions and now the software has memorized the X, Y, Z position of that field. And then when I do pick another field, it's, and I'll, so when I'm picking, when I'm picking new fields, but in the same, well, I'll use the, I'll move the i'll, I'll turn the shutter, I'll open the shutter and I'll look to see what I see on the screen and I'll pick fields based on that versus looking in the field.So I don't see any of my tonics in there. But again, even nicely centered, these cells will go through my PS I continue, here's a good example. Here's a good example of what I would avoid that, see how bright that is?That's very bright. That's probably remnants of an apoptotic cell. And the reason I would avoid that is because since it's so bright, when the software auto focuses, it will autofocus just on that and it won't be in the same plane as all these other cells.And so I won't get any information from that field. That looks nice. There's a, this is a pro metaphase right here, That's A metaphase.So, and I'm gonna take three positions per well. So that was three positions and now I'm going to go to my next Well, so you can, so what I will probably do is take images every 10 minutes. So it's a compromise.Lifestyle imaging is always a compromise, right? You have to image, you have to keep the cells healthy, especially if you're interested in mitosis. So you have to minimize their exposure to the fluorescent line.And, but you wanna get information. So you have to take images, you have to, so you have to compromise, you have to make a choice about what you want to capture, what kind of information you wanna capture. For our purposes, mitosis tends to take about an hour.So if we're taking 10 minute intervals, that's pretty good. We see prophase, we see metaphase, we see anaphase, and we can, we can see when there are, there are treatments or drugs or siRNAs that affect those specific cell cycle stages. So for our purposes, that's enough, but we always want more.We always want more positions or finer time and interval or something like that. So it's a compromise. So, and then I'll hit start.And so our software, it does an autofocusing pass on the first pass. So once I hit start, it should start auto autofocusing and you'll see the shutter open, open and close between auto focusing. So I'll hit start.So that's the shutter opening and closing. The Z motor is engaging and it's actually trying to find the plane with the most contrast, which will be the most in focus. You can actually see on the screen going in and out of finding the best plane of focus that might be worrisome.So what it does is it, it drives the Z motor to find the best, the most in focus plane focus, well the most in image. And then it acquires picture there and then it learns that Z position. And then we'll use that Z position for the next 12 passes.And then we'll do the auto focusing pass again. What Kind of experiments do they do? Okay, so our lab is mostly interested in understanding what regulates mitic progression, or at least I'm most interested in understanding that.So with live cell imaging and especially our multi position, long-term imaging scope, you can take, you can film cells for two or three days entering anding mitosis, and you can image multiple rounds of mitosis so you can understand. And then you can put drugs or RNAs or any kind of perturbations on the cells that you would like to study. So you can understand how, how those drugs or, or genes are, are important in regulating mitotic progression.Because it's not fixed cell work and live cell work, you can understand what, when you need to have a perturbation to affect mitosis or what stage of mitosis is affected. So it's really powerful in that respect. What, what are the main technical elements of this experiments?What are the possible problems? Well, the biggest problem besides getting the rig set up, getting the rig set up is not trivial. A lot of people have the rig, but just making sure it works well for your application.The financial element is not, you know, is not small. So there's the setting up of the apparatus and then, and then once you gather the data, there's a real data analysis issue. It takes these movie, if, so, if you're gathering 40 movies each with 30 cells, so you have 40 in 40 fields that you just filmed and each of those fields has 30 cells, then you have to sit down with each one of those movies and look at each one of those cells in that movie.So it takes a really long time to do data analysis. And that's, so it takes a week to analyze data that it took you two days to collect. So there's a, that's a very big obstacle in terms of gathering lots of data.So it's not, while it's high throughput data collection, it's not high throughput analysis. So what would you make to make this experiment really high throughput? So our lab is in the process of developing a program that does the data analysis automatically.And it's a pretty decent program. It's not perfect. The human eye is always better, but it's getting there.So What are the interesting application of this experiment? Maybe other fields you can mention? Besides cell division?You mean for the, oh, for the live cell imaging? Yes.Well, you can do cell motility assays. You can, depending on the, the microscopy that's involved, you can, you know, look at how cells, you could look at tumor development if you really wanted to.But there, you, you're, if as long as you have the right imaging set up, you can, you're pretty much not limited in terms of anything. Our scope happens to be just a wide field and we use 20 by 20 x objective. So it's a pretty low resolution set up, but that's sufficient for what we need to see.So there are, but there definitely aren't, you wouldn't be limited. Anything that has any dynamic kind of experiment, which is biology, it would be, would really benefit from lifestyle imaging. So calcium imaging, although you couldn't do it on our apparatus, that's a live cell.You definitely need live cell imaging for that. Like I said, patient assays, motility assays, those kinds of things would really benefit from live cell imaging. Would you like to tell a few words about your previous experience?How long do you work on these techniques? What do you you work on before? Wow.So in my PhD I didn't work.I did a lot of confocal microscopy, but no lifestyle imaging. And so during my postdoc I was, I helped Dr.King get this apparatus that, that we just used to set up these fragment, get that up and running. And, and so during my postdoc, I had a lot of experience with both widefield with TimeLapse imaging on our scope, which is the widefield and also TimeLapse imaging with confocal microscopy.And I would say that it's really powerful. You get a lot of information, but it's not easy to get working perfectly. So it really requires a lot of attention and a lot of attention to details at every little step of the process.And, and then you still have problems. So it's a tough application, but you can really get, if, if it's what you need to have to, to study your problem, your scientific problem, then it, it's invaluable in that respect, but it's tough. So find someone who knows what they're doing and use them.That's my advice. So good.

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 ...

Setting Up a Simple Light Sheet Microscope for In Toto Imaging of C. elegans Development

Fast and low phototoxic imaging techniques are pre-requisite to study the development of organisms in toto. Light sheet based microscopy reduces photo-bleaching and phototoxic effects compared to confocal microscopy, while providing 3D images with subcellular resolution. Here we present the setup of a light sheet based microscope, which is composed of an upright microscope and a small set of opto-mechanical elements for the generation of the light sheet. The protocol describes how to build, align the microscope and characterize the light sheet. In addition, it details how to implement the method for in toto imaging of C. elegans embryos using a simple observation chamber. The method allows the capture of 3D two-colors time-lapse movies over few hours of development. This should ease the tracking of cell shape, cell divisions and tagged proteins over long periods of time.

Setting Up a Simple Light Sheet Microscope for In Toto Imaging of C. elegans Development

This protocol describes the setup of a light sheet microscope and its implementation for in vivo imaging of C. elegans embryos.

Fast and low phototoxic imaging techniques are pre-requisite to study the development of organisms in toto. Light sheet based microscopy reduces ...

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 ...

Analysis of the Gap Junction-dependent Transfer of miRNA with 3D-FRAP Microscopy

Small antisense RNAs, like miRNA and siRNA, play an important role in cellular physiology and pathology and, moreover, can be used as therapeutic agents in the treatment of several diseases. The development of new, innovative strategies for miRNA/siRNA therapy is based on an extensive knowledge of the underlying mechanisms. Recent data suggest that small RNAs are exchanged between cells in a gap junction-dependent manner, thereby inducing gene regulatory effects in the recipient cell. Molecular biological techniques and flow cytometric analysis are commonly used to study the intercellular exchange of miRNA. However, these methods do not provide high temporal resolution, which is necessary when studying the gap junctional flux of molecules. Therefore, to investigate the impact of miRNA/siRNA as intercellular signaling molecules, novel tools are needed that will allow for the analysis of these small RNAs at the cellular level. The present protocol describes the application of three-dimensional fluorescence recovery after photobleaching (3D-FRAP) microscopy to elucidating the gap junction-dependent exchange of miRNA molecules between cardiac cells. Importantly, this straightforward and non-invasive live-cell imaging approach allows for the visualization and quantification of the gap junctional shuttling of fluorescently labeled small RNAs in real time, with high spatio-temporal resolution. The data obtained by 3D-FRAP confirm a novel pathway of intercellular gene regulation, where small RNAs act as signaling molecules within the intercellular network.

Here, we describe the application of three-dimensional fluorescence recovery after photobleaching (3D-FRAP) for the analysis of the gap junction-dependent shuttling of miRNA. In contrast to commonly applied methods, 3D-FRAP allows for the quantification of the intercellular transfer of small RNAs in real time, with high spatio-temporal resolution.

Small antisense RNAs, like miRNA and siRNA, play an important role in cellular physiology and pathology and, moreover, can be used as therapeutic agents ...

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 ...