We thank Ernst H. K. Stelzer for the opportunity to use his resources as well as his valuable comments regarding the manuscript, Anita Anderl for support with the Tribolium live imaging, Sven Plath for technical support as well as Ilan Davis, Nicole Grieder and Gerold Schubiger for sharing their transgenic Drosophila lines via the Bloomington Stock Center.

1. Preparatory work
<ol>
	<li>Choose an illumination lens/detection lens/camera combination for the LSFM that suits the scientific question and set up the microscope. The size of the field of view is the quotient of the camera chip size and the magnification of the detection lens. The illumination lens should be chosen so that the entire field of view is covered by a roughly planar light sheet34. Three recommended combinations are listed in Table 1.</li>
	<li>To prepare agarose aliquots and retrieval dishes, add 2 g low-melt agarose to 200 mL autoclaved tap water and heat the mixture in a microwave oven at 600-800 W until all agarose particles are dissolved. Prepare several 1 mL agarose aliquots in 1.5 mL or 2 mL reaction tubes, then fill several 90-mm &#216; Petri dishes 3-5 mm high with agarose. Store solidified aliquots and dishes at 4 &#176;C.</li>
	<li>For Drosophila: To prepare fresh rearing vials, cook an adequate amount of custom-made or commercially available Drosophila medium, transfer 5-15 mL into wide vials and store them at 4 &#176;C. To prepare egg-laying dishes, add 1 g of low-melt agarose to 50 mL of autoclaved tap water and heat the mixture in a microwave oven at 600-800 W until all agarose particles are dissolved. Allow the mixture to cool down to 45 &#176;C, then add 50 mL fruit juice (preferably apple or red grape) and mix thoroughly. Pour the mixture into 35 mm &#216; Petri dishes and store solidified egg-laying dishes at 4 &#176;C.</li>
	<li>For Tribolium: To prepare the growth medium, pass whole wheat flour as well as inactive dry yeast through a 710 &#181;m mesh size sieve, then supplement the sieved flour with 5% (w/w) sieved yeast. To prepare egg-laying medium, pass fine wheat flour as well as inactive dry yeast through a 250 &#181;m mesh size sieve, then supplement the sieved flour with 5% (w/w) sieved yeast.</li>
</ol>
2. Calibration of sample chamber based LSFMs using fluorescent microspheres
NOTE: The purpose of calibration is to align the focal points of the illumination and detection lenses (Figure 2A), as this is the premise for clear images. LSFMs should be calibrated regularly, at least once every 3-4 weeks.
<ol>
	<li>Re-liquefy an agarose aliquot in a dry block heater/mixer at 80 &#176;C, then allow the agarose aliquot to cool down to 35 &#176;C.</li>
	<li>Transfer 50 &#181;L of agarose to a 1.5 mL reaction tube and add 0.5 &#181;L of fluorescent microsphere solution. Mix at 1,400 rpm for 1 min.</li>
	<li>Fill the slotted hole of the cobweb holder with 10 &#181;L of agarose/fluorescent microsphere solution mixture, then aspirate as much agarose as possible until only a thin agarose film remains. Wait 30-60 s for solidification.</li>
	<li>Fill the sample chamber with autoclaved tap water. Insert the cobweb holder slowly into the sample chamber and move the slotted hole with the microtranslation stages in front of the detection lens.</li>
	<li>Rotate the cobweb holder with the rotation stage to a 45&#176; position relative to the illumination (x) and detection (z) axes. The cobweb holder should not be visible in the transmission light channel.</li>
	<li>Switch to the respective fluorescence channel and adjust the laser power as well as the exposure time so that the fluorescent microspheres provide proper signal.</li>
	<li>Specify a volume of view that covers the now transversely oriented agarose film completely. Define the z spacing by calculating the maximally&#160;possible axial resolution for the respective illumination lens/detection lens combination34. Alternatively, 4 times the lateral resolution can be used as a rough approximation.</li>
	<li>Record a three-dimensional test z stack of the fluorescent microspheres and compare the x, y and z maximum projections to the calibration chart (Figure 2). If the microspheres appear blurry, fuzzy, and/or distorted (Figure 2B,C), adjust the positions of the illumination and/or detection lens.</li>
</ol>
3. Collection of Drosophila embryos
<ol>
	<li>Transfer 100-200 adults of the Drosophila line of choice to a fresh rearing vial 2-3 days before the imaging assay to establish an egg collection culture. If not yet existent, consider using the old rearing vial to start a progeny culture. To ensure adults are no more than&#160;two weeks old, replace the embryo collection culture in time with the progeny culture.</li>
	<li>Warm an egg-laying dish&#160;to room temperature and add a drop of yeast paste on top of the agar.</li>
	<li>Transfer the adults from the egg collection culture to an empty narrow vial and place it on top of the egg-laying dish. Incubate the egg collection setup at room temperature for 15 min. Avoid anesthesia (cold, CO2) during this step if possible.</li>
	<li>Return the adults to the rearing vial. Incubate the egg-laying dish at a convenient temperature/time combination, then transfer 10-20 embryos with a small paintbrush from the egg-laying dish to a 100 &#181;m mesh size cell strainer. Discard the egg-laying dish.</li>
	<li>Repeat steps 3.1 to 3.4 for each Drosophila line.</li>
</ol>
4. Collection of Tribolium embryos
NOTE: For convenience, a scheme of the Tribolium egg collection procedure is provided (Figure 3A) to which also the numbers within the brackets in this step refer.
<ol>
	<li>Transfer 200-300 adults (about 400-700 mg) of the Tribolium line of choice to an empty 1 L glass bottle 2-10 days before the imaging assay to establish an egg collection culture (Figure 3A_01). Fill the bottle with 50-100 g of fresh growth medium. If not yet existent, consider starting a progeny culture using available larvae and pupae. To ensure adults are no more than 3 months old, replace the egg collection culture in time with the progeny culture.</li>
	<li>Pass the egg collection culture through an 800 &#181;m mesh size sieve (Figure 3A_02). Return the growth medium, which contains non-staged embryos, to the initial bottle (Figure 3A_03) and transfer adults to an empty 1 L glass bottle (Figure 3A_04). Add 10 g of egg-laying medium (Figure 3A_05) and incubate the egg collection setup at room temperature for 1 h (Figure 3A_06,07).</li>
	<li>Pass the egg collection setup through the 800 &#181;m mesh size sieve (Figure 3A_08). Return adults to their initial bottle (Figure 3A_09). Depending on the developmental process that should be imaged, incubate the egg-laying medium, which now contains about 30-100 embryos, at a convenient temperature/time combination (Figure 3A_10).</li>
	<li>Pass the egg-laying medium through the 300 &#181;m mesh size sieve (Figure 3A_11) and transfer the embryos (Figure 3A_12) to a 100 &#181;m mesh size cell strainer. Discard the sieved egg-laying media (Figure 3A_13).</li>
	<li>Repeat steps 4.1 to 4.4 for each Tribolium line.</li>
</ol>
5. Sodium hypochlorite-based dechorionation
NOTE: Both Drosophila and Tribolium embryos are covered by a chorion, a protective and heavily light-scattering protein layer that is not essential for proper development as long as the embryos are kept moist after removal. The dechorionation protocol for the embryos of both species is identical.
<ol>
	<li>Prepare a 6-well plate&#160;by filling the A1, A2, A3 and B3 wells with 8 mL of autoclaved tap water and the B1 and B2 wells with 7 mL of autoclaved tap water and 1 mL of sodium hypochlorite (NaOCl) solution (Figure 3B). Observe the dechorionation process under a stereo microscope, ideally in transmission light. 
	CAUTION: Sodium hypochlorite is corrosive.</li>
	<li>Insert the embryo-containing cell strainer (Step 3.4 and/or 4.4) into the A1 well and wash the embryos about 30-60 s under gentle agitation.</li>
	<li>Move the cell strainer to the B1 well&#160;and shake the plates vigorously for 30 s, then transfer it to the A2 well and wash the embryos for 1 min under gentle agitation.</li>
	<li>Move the cell strainer to the B2 well and shake the plate vigorously until most embryos are completely dechorionated (Figure 3C), then transfer it to the A3&#160;well and wash the embryos for 1 min under gentle agitation.</li>
	<li>Store the cell strainer in the B3 well before proceeding with the mounting procedure.</li>
	<li>Repeat steps 5.1 to 5.5 for each line.</li>
</ol>
6. Mounting of multiple embryos using the cobweb holder
<ol>
	<li>Re-liquefy an agarose aliquot in a dry block heater/mixer at 80 &#176;C, then allow the agarose aliquot to cool down to 35 &#176;C.</li>
	<li>Pipet 10 &#181;L of agarose on top of the slotted hole of the cobweb holder. With the pipette tip, spread the agarose over the slotted hole, then aspirate as much agarose as possible until only a thin agarose film remains. Wait 30-60 s for solidification.</li>
	<li>For each line, carefully pick one or more embryos with a small paintbrush and place them on the agarose film.</li>
	<li>Arrange the embryos along the long axis of the slotted hole, then also align their anterior-posterior axis with the long axis of the slotted hole (Figure 3D).</li>
	<li>Stabilize the embryos carefully by pipetting 1-2 &#181;L of agarose into the gap between the embryos and the agarose film. Wait 30-60 s for solidification.</li>
	<li>Insert the cobweb holder with the mounted embryos slowly into the image buffer-filled sample chamber.</li>
</ol>
7. Comparative live imaging in sample chamber-based LSFMs
<ol>
	<li>Move one of the embryos with the microtranslation stages in front of the detection lens. Ensure that the cobweb holder is in a 45&#176; position relative to the illumination (x) and detection (z) axes (cf. Figure 1C).</li>
	<li>In the transmission light channel, move the embryo into the center of the field of view. The cobweb holder should not be visible.</li>
	<li>Move the embryo with the microtranslation stages in z until the midplane of the embryo overlaps with the focal plane, i.e. until the outline appears sharp. Without switching to the fluorescence channel, specify the volume of view by moving 250 &#181;m away from the midplane into both directions.</li>
	<li>Optionally, if imaging along multiple directions is required, rotate the embryo appropriately and repeat steps 7.2 and 7.3. The cobweb holder supports up to four orientations in steps of 90&#176;.</li>
	<li>Repeat steps 7.1 to 7.3 (or 7.4) for all other embryos mounted on the cobweb holder (Figure 3E). Ensure that the topmost embryo does not leave the imaging buffer when the bottommost embryo is in front of the detection lens.</li>
	<li>Define the fluorescence channel (laser power, exposure time, detection filter) and time lapse (interval, total duration) parameters and start the imaging process. For indicative values, consult the metadata table of the example datasets (Supplementary Table 1). For assays that last several days, consider covering the sample chamber opening at least partially to reduce evaporation.</li>
</ol>
8. Retrieval and further cultivation of imaged embryos
<ol>
	<li>When the imaging assay has ended, carefully remove the cobweb holder from the sample chamber.</li>
	<li>Detach the embryos from the agarose film with a small paintbrush and transfer them to an appropriately labeled microscope slide. Place the slide into a retrieval dish and incubate under the respective standard rearing conditions.</li>
	<li>Regarding the experimental modalities, estimate when embryogenesis is completed. As the hatching time point approaches, check the retrieval dishes frequently and transfer hatched Drosophila larvae to individual rearing vials and Tribolium larvae to individual wells of a 24-well plate. Fill the wells up to the half with growth medium. Incubate under the respective standard rearing conditions.</li>
	<li>Once the observed individuals are adults, provide them with a suitable mating partner and check for progeny after several days.</li>
</ol>
9. Image data processing and metadata documentation
NOTE: For image data processing, the ImageJ derivate FIJI35 is recommended (imagej.net/Fiji/Downloads). FIJI does not require installation and 32- as well as 64-bit versions are available. One of the most frequently used formats for LSFM data is the tagged image file format (TIFF), which allows the storage of&#160;image stacks in form of a TIFF container.
<ol>
	<li>Calculate the z maximum projections for all z stacks (Image | Stacks | Z Project, choose Max Intensity). Maximum projections are data simplification approaches that reduce the number of spatial dimensions&#160;from three to two. A FIJI script for batch processing is provided (Supplementary File 1).</li>
	<li>Concatenate the respective z maximum projections to create time stacks (t stacks). Save these in one TIFF container. Do this for all recorded embryos as well as the respective directions and&#160;fluorescence channels if applicable.</li>
	<li>Rotate the z and t stack&#160;around the z axis to align the anterior-posterior axis of the embryos with the x or y image axis (Image | Transform | Rotate). Crop the z stacks along all three image axes and the t stack in the x and y image axes so that only minimal buffer space (20-40 pixels along the x and y axes, 5-10 images along the z axis) around the embryo remains (Image | Adjust | Canvas Size for the x and y axes, Image | Stacks | Tools | Slice Keeper for the z axis).
	<ol>
		<li>Do this individually for all recorded embryos as well as the respective directions, if applicable. Fluorescence channels, if applicable, should be processed with identical rotation and cropping parameters.</li>
	</ol>
	</li>
	<li>Document the metadata as detailed as possible. As a guideline, the metadata table of the example datasets, which are featured in the Representative Results section, can be used (Supplementary Table 1).</li>
</ol>
10. Data visualization
NOTE: This data visualization guideline focuses primarily on the creation of z maximum projection image matrices that show several recorded embryos along multiple directions and/or over time. The following steps describe the data visualization procedure that was applied to the example datasets for the creation of the figures shown and videos linked in the Representative Results section.
<ol>
	<li>Combine t stacks of multiple directions (Image | Stacks | Tools | Combine) and/or merge multiple fluorescence channels (Image | Color | Merge Channels) to visualize the biological structure and/or process of interest.</li>
	<li>Adjust the intensities of the t stacks (Image | Adjust | Brightness/Contrast) as needed by using the &#8220;Set&#8221; function. The minimum displayed value should be set slightly above the background signal, the maximum displayed value should result in a convenient contrast. Document both values, as they can be used for a consistent adjustment of the respective z stacks. Imprint adjustments using the &#8220;Apply&#8221; function. Depending on the experimental modalities, consider processing all recorded embryos with identical values.</li>
	<li>Save intensity-adjusted t stacks as separate files, do no override the non-adjusted t stacks. Compilate suitable sub stacks from the adjusted t stacks with dedicated image selection functions (e.g., Image | Stacks | Tools | Slice Keeper) and use the montage tool (Image | Stacks | Make Montage) to create image grids that can be used for figure design.</li>
</ol>

The authors have nothing to disclose.

One of the exclusive application areas of LSFMs is developmental biology. In this discipline, it is of importance to look at living specimens, otherwise morphogenetic processes cannot be described in a dynamic manner. An experimental framework for the simultaneous live imaging in sample chamber-based LSFMs, as described here, is convenient for two major reasons.
Ambient variance, which is unavoidable in sequential live imaging, can be addressed pro-actively. In insect embryo-associated live im...

Fluorescence microscopy is one of the most essential imaging techniques in the life sciences, especially in cell and developmental biology. In confocal fluorescence microscopes1, which are state-of-the-art for three-dimensional fluorescence imaging since the mid-1990s, the same lens is used for fluorophore excitation and emission light detection. The illumination laser beam excites all fluorophores along the illumination/detection axis and the respective out-of-focus signal is discriminated prior to&#160;detection by a pinhole. Hence, for each two-dimensional image, the entire specimen is illuminated. Consequently, for each three-dimensional image, i.e., a stack of spatially consecutive two-dimensional images, the entire specimen is illuminated several dozen to a few hundred times2, which promotes photobleaching and phototoxicity3.
Almost twenty years ago, light sheet-based technology4 emerged as a promising alternative for three-dimensional fluorescence imaging and thus became a valuable tool in developmental biology5. In this approach, illumination and detection are decoupled. The illumination lens is used to generate a light sheet with a depth&#160;of only a few micrometers within the focal plane of the perpendicularly arranged detection lens. Hence, for each two-dimensional image, only a thin planar volume around the focal plane is illuminated. Consequently, for each three-dimensional image, the entire specimen is illuminated only once, which strongly decreases photobleaching and phototoxicity6. For this reason, light sheet fluorescence microscopes (LSFMs) offer&#160;efficient solutions to study complex processes on multiple biologically relevant scales and are, therefore, of particular value in developmental biology, where specimens as large as several millimeters have to be analyzed at the subcellular level.
Historically, LSFMs have been sample chamber-based7,8. In these setups, the illumination (x) and detection (z) axes are usually arranged perpendicularly to the gravity axis (y). Sample chambers offer ample experimental freedom. Firstly, they provide large imaging buffer capacities, which in turn eases the use of a perfusion system to control the environment, e.g., to maintain a specific temperature9 or to apply biochemical stressors. Further, they support customized mounting methods10 that are tailored to the respective experimental needs while preserving the three-dimensional, in some instances dynamic11, integrity of the specimen. Additionally, sample chamber-based setups are usually equipped with a rotation function that is used to revolve the specimens around the y axis and thus image them along two, four or even more directions. Since&#160;embryos of commonly used model organisms are, in the context of microscopy, relatively large, successive imaging along the ventral-dorsal, lateral, and/or anterior-posterior body axes provides a more comprehensive representation. This allows e.g., long-term tracking of cells that move along complex three-dimensional migration paths12,13.
Light sheet-based fluorescence microscopy has been applied extensively to study the embryonic morphogenesis of Drosophila melanogaster, both systematically14,15 as well as with a specific focus on the biophysical aspects of development. For instance, it was used to gather high-resolution morphogenetic data in order to detect a biomechanical link between endoderm invagination and axis extension during germband elongation16 and further to relate the complex cellular flow with force generation patterns during gastrulation17. It has also been combined with other state-of-the-art techniques, e.g., optogenetics to investigate the regulation of Wnt signaling during anterior-posterior patterning in the epidermis18.
However, studying only one species does not provide insights into the evolution of development. To understand embryogenesis within the phylogenetic context, intensive research has been conducted with alternative insect model organisms. One of the most comprehensively investigated species is the red flour beetle Tribolium castaneum, an economically relevant stored grain pest19, whose embryonic morphogenesis has also already been systematically imaged with LSFM20. The embryonic morphogenesis of these two species differs remarkably in several aspects, e.g., the segmentation mode21, as well as the formation and degradation of extra-embryonic membranes22. The latter aspect has already been extensively analyzed using LSFMs. For instance, it has been shown that the serosa, an extra-embryonic tissue that envelops and protects the Tribolium embryo from various hazards for the better part of its embryogenesis23,24, also&#160;acts as the morphogenetic &#8220;driver&#8221; for its own withdrawal process during dorsal closure25. Further, it has been demonstrated that during gastrulation, a particular region of the blastoderm remains anchored to the vitelline membrane in order to create asymmetric tissue movements26 and, following this observation, that regionalized tissue fluidization allows cells to sequentially leave the serosa edge during serosa window closure27.
In all Drosophila- and Tribolium-associated studies cited above, sample chamber-based LSFMs have been used. In most, the embryos were recorded along multiple directions using the sample rotation function. Although not stated explicitly, it can be assumed that they have been recorded individually and thus independent of each other in sequential live imaging assays, similar to our previous work on Tribolium20,28. In certain scenarios, such an approach is acceptable, but especially in quantitative comparative approaches, ambient variance can distort the results. For instance, it has long been known that the developmental speed of insects is temperature-dependent29, but a more recent study further suggests that in Drosophila, temperature may also affect the concentration of morphogens30. Consequently, if certain characteristics of embryogenesis, e.g., the dynamic proportions, division rates and migration velocities of cells, should be precisely quantified, sufficient repetitions without ambient variance are required. This minimizes standard deviations and standard errors, which in turn facilitates juxtaposition with other, even just marginally divergent experimental conditions.
However, sample chamber-based LSFMs are primarily designed for high content rather than high throughput assays. Unlike confocal microscopes, which are typically equipped with standardized clamp mechanisms for microscopy slides, Petri dishes and well plates, nearly all sample chamber-based LSFMs use cylinder-based clamp mechanisms. These mechanisms are intended for custom-made sample holders that are rotation-compatible as well as non-invasive10, but usually not designed for more than one specimen20,31,32. A framework for simultaneous live imaging of two or more embryos, in which the advantages of sample chamber-based setups are not compromised, addresses the ambient variance issue thereby increasing the value of LSFMs for comparative studies.
In our protocol, we present an experimental framework for comparative live imaging in sample chamber-based LSFMs (Figure 1A) in which the y axis is used as an option to &#8220;stack&#8221; embryos. Firstly, we provide a fluorescent microsphere-based calibration guideline for sample chamber-based LSFMs, which is especially important for instruments that lack a calibration assistant. Secondly, we describe a mounting method for multiple embryos based on the cobweb holder28 (Figure 1B) that is compatible with sample rotation and thus allows simultaneous imaging of multiple specimens along multiple directions (Figure 1C). Several embryos are aligned on top of a thin agarose film and, after insertion into the sample chamber, moved successively through the light sheet to acquire three-dimensional images. Thirdly, we provide three exemplary live imaging datasets for Drosophila as well as for Tribolium. For the former, we juxtapose transgenic lines with fluorescently labeled nuclei. For the latter, we compare the performance of transgenic sublines that carry the same transgene, but at different genomic locations. Finally, we discuss the importance of parallelization with regard to comparative live imaging and ambient variance33, debate the throughput limit of our experimental framework and evaluate adaption of our approach to other model organisms.

<table><tbody><tr><td>6-well plate</td><td>Orange Scientific</td><td>4430500</td><td>&lt;strong&gt;&amp;nbsp;&lt;/strong&gt;</td></tr><tr><td>24-well plate</td><td>Orange Scientific</td><td>4430300</td><td>Only for live imaging involving &lt;em&gt;Tribolium&lt;/em&gt;</td></tr><tr><td>35-mm &amp;Oslash; Petri dish</td><td>Fisher Scientific</td><td>153066</td><td>Only for live imaging involving &lt;em&gt;Drosophila&lt;/em&gt;.</td></tr><tr><td>90-mm &amp;Oslash; Petri dish</td><td>Fisher Scientific</td><td>L9004575</td><td></td></tr><tr><td>100 &amp;micro;m mesh size cell strainer</td><td>BD Biosciences</td><td>352360</td><td></td></tr><tr><td>250 &amp;micro;m mesh size sieve</td><td>VWR International</td><td>200.025.222-038</td><td>Only for live imaging involving &lt;em&gt;Tribolium&lt;/em&gt;</td></tr><tr><td>300 &amp;micro;m mesh size sieve</td><td>VWR International</td><td>200.025.222-040</td><td>Only for live imaging involving &lt;em&gt;Tribolium&lt;/em&gt;</td></tr><tr><td>710 &amp;micro;m mesh size sieve</td><td>VWR International</td><td>200.025.222-050</td><td>Only for live imaging involving &lt;em&gt;Tribolium&lt;/em&gt;</td></tr><tr><td>800 &amp;micro;m mesh size sieve</td><td>VWR International</td><td>200.025.222-051</td><td>Only for live imaging involving &lt;em&gt;Tribolium&lt;/em&gt;</td></tr><tr><td>405 fine wheat flour</td><td>Demeter e.V.</td><td>SP061006</td><td>Only for live imaging involving &lt;em&gt;Tribolium&lt;/em&gt;</td></tr><tr><td>commercially available Drosophila medium</td><td>Genesee Scientific</td><td>66-115</td><td>Only for live imaging involving &lt;em&gt;Drosophila&lt;/em&gt; / Custom-made &lt;em&gt;Drosophila&lt;/em&gt; medium may also be used</td></tr><tr><td>fluorescent microspheres, 1.0 &amp;micro;m &amp;Oslash;</td><td>Thermo Fisher Scientific</td><td>T7282</td><td></td></tr><tr><td>inactive dry yeast</td><td>Genesee Scientific</td><td>62-108</td><td></td></tr><tr><td>low-melt agarose</td><td>Carl Roth</td><td>6351.2</td><td></td></tr><tr><td>narrow vials</td><td>Genesee Scientific</td><td>32-109</td><td>Only for live imaging involving &lt;em&gt;Drosophila&lt;/em&gt;</td></tr><tr><td>small paint brush</td><td>VWR International</td><td>149-2121</td><td></td></tr><tr><td>sodium hypochlorite (NaOCl), ~12% active Cl</td><td>Carl Roth</td><td>9062.3</td><td>Caution: sodium hypochlorite is corrosive</td></tr><tr><td>whole wheat flour</td><td>Demeter e.V.</td><td>SP061036</td><td>Only for live imaging involving &lt;em&gt;Tribolium&lt;/em&gt; / United Kingdom: wholemeal flour</td></tr><tr><td>wide vials</td><td>Genesee Scientific</td><td>32-110</td><td>Only for live imaging involving &lt;em&gt;Drosophila&lt;/em&gt;</td></tr></tbody></table>

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.

Our protocol describes an experimental framework for comparative fluorescence live imaging in sample chamber based LSFMs. For instance, the framework can be used to juxtapose (i) embryos of two or more species, (ii) embryos of lines in which one or more genes are knocked out plus wild-type controls, (iii) multiple embryos of the same transgenic line, (iv) embryos from different transgenic lines, or (v) embryos from sublines that carry the same transgene, but at different genomic locations. In this section, we provide exa...

Watch this Scientific Journal Video about Simultaneous Live Imaging of Multiple Insect Embryos in Sample Chamber-Based Light Sheet Fluorescence Microscopes at JoVE.com

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

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 ...

simultaneous-live-imaging-multiple-insect-embryos-sample-chamber

Research

JoVE Journal

Biology

3.0K Views.  Goethe-Universität. Op lichtplaten gebaseerde fluorescentiemicroscopie is het meest waardevolle hulpmiddel in de ontwikkelingsbiologie. Een belangrijk probleem in vergelijkende studies is de variantie van de omgeving. Ons protocol beschrijft een experimenteel kader voor gelijktijdige live beeldvorming van meerdere specimens en pakt dit probleem daarom proactief aan.

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

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Developmental Biology

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM compared to confocal systems are its low phototoxicity, gentle mounting strategies, fast acquisition with high signal to noise ratio and the possibility of imaging samples from various angles (views) for long periods of time. Imaging from multiple views unleashes the full potential of LSFM, but at the same time it can create terabyte-sized datasets. Processing such datasets is the biggest challenge of using LSFM. In this protocol we outline some solutions to this problem. Until recently, LSFM was mostly performed in laboratories that had the expertise to build and operate their own light sheet microscopes. However, in the last three years several commercial implementations of LSFM became available, which are multipurpose and easy to use for any developmental biologist. This article is primarily directed to those researchers, who are not LSFM technology developers, but want to employ LSFM as a tool to answer specific developmental biology questions. 
Here, we use imaging of zebrafish eye development as an example to introduce the reader to LSFM technology and we demonstrate applications of LSFM across multiple spatial and temporal scales. This article describes a complete experimental protocol starting with the mounting of zebrafish embryos for LSFM. We then outline the options for imaging using the commercially available light sheet microscope. Importantly, we also explain a pipeline for subsequent registration and fusion of multiview datasets using an open source solution implemented as a Fiji plugin. While this protocol focuses on imaging the developing zebrafish eye and processing data from a particular imaging setup, most of the insights and troubleshooting suggestions presented here are of general use and the protocol can be adapted to a variety of light sheet microscopy experiments.

Light sheet fluorescence microscopy is an excellent tool for imaging embryonic development. It allows recording of long time-lapse movies of live embryos in near physiological conditions. We demonstrate its application for imaging zebrafish eye development across wide spatio-temporal scales and present a pipeline for fusion and deconvolution of multiview datasets.

Light sheet fluorescence microscopy (LSFM) is gaining more and more popularity as a method to image embryonic development. The main advantages of LSFM ...

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. The observation of Tribolium embryos with light sheet-based fluorescence microscopy has multiple advantages over conventional widefield and confocal fluorescence microscopy. Due to the unique properties of a light sheet-based microscope, three dimensional images of living specimens can be recorded with high signal-to-noise ratios and significantly reduced photo-bleaching as well as photo-toxicity along multiple directions over periods that last several days. With more than four years of methodological development and a continuous increase of data, the time seems appropriate to establish standard operating procedures for the usage of light sheet technology in the Tribolium community as well as in the insect community at large. This protocol describes three mounting techniques suitable for different purposes, presents two novel custom-made transgenic Tribolium lines appropriate for long-term live imaging, suggests five fluorescent dyes to label intracellular structures of fixed embryos and provides information on data post-processing for the timely evaluation of the recorded data. Representative results concentrate on long-term live imaging, optical sectioning and the observation of the same embryo along multiple directions. The respective datasets are provided as a downloadable resource. Finally, the protocol discusses quality controls for live imaging assays, current limitations and the applicability of the outlined procedures to other insect species.
This protocol is primarily intended for developmental biologists who seek imaging solutions that outperform standard laboratory equipment. It promotes the continuous attempt to close the gap between the technically orientated laboratories/communities, which develop and refine microscopy methodologically, and the life science laboratories/communities, which require 'plug-and-play' solutions to technical challenges. Furthermore, it supports an axiomatic approach that moves the biological questions into the center of attention.

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

Imaging the morphogenesis of insect embryos with light sheet-based fluorescence microscopy has become state of the art. This protocol outlines and compares three mounting techniques appropriate for Tribolium castaneum embryos, introduces two novel custom-made transgenic lines well suited for live imaging, discusses essential quality controls and indicates current experimental limitations.

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and computational modeling to classify and compare biological processes. Although cell culture model systems are great for high content imaging, high throughput studies of cell morphology suggest that ex vivo cultures are limited in recapitulating the morphological complexity found in cells within living organisms. As such, there is a need for a scalable high throughput model system to image living cells within an intact organism. Described here is a protocol for using a high content image analyzer to simultaneously acquire multiple time-lapse videos of embryonic Drosophila melanogaster development during the syncytial blastoderm stage. The syncytial blastoderm has traditionally served as a great in vivo model for imaging biological events; however, obtaining a significant number of experimental replicates for quantitative and high-throughput approaches has been labor intensive and limited by the imaging of a single embryo per experimental repeat. Presented here is a method to adapt imaging and microinjection approaches to suit a high content imaging system, or any inverted microscope capable of automated multipoint acquisition. This approach enables the simultaneous acquisition of 6-12 embryos, depending on desired acquisition factors, within a single imaging session.

Presented here is a protocol to microinject and simultaneously image multiple Drosophila embryos during embryonic development using a plate-based, high content imager.

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and ...

Visualizing Ocular Morphogenesis by Lightsheet Microscopy Using rx3:GFP Transgenic Zebrafish

Vertebrate eye development is a complex process that begins near the end of embryo gastrulation and requires the precise coordination of cell migration, proliferation, and differentiation. Time-lapse imagining offers unique insight to the behavior of cells during eye development because it allows us to visualize oculogenesis in vivo. Zebrafish are an excellent model to visualize this process due to their highly conserved vertebrate eye and their ability to develop rapidly and externally while remaining optically transparent. Time-lapse imaging studies of zebrafish eye development are greatly facilitated by use of the transgenic zebrafish line Tg(rx3:GFP). In the developing forebrain, rx3:GFP expression marks the cells of the single eye field, and GFP continues to be expressed as the eye field evaginates to form an optic vesicle, which then invaginates to form an optic cup. High resolution time lapse imaging of rx3:GFP expression, therefore, allows us to track the eye primordium through time as it develops into the retina. Lightsheet microscopy is an ideal method to image ocular morphogenesis over time due to its ability to penetrate thicker samples for fluorescent imaging, minimize photobleaching and phototoxicity, and image at a high speed. Here, a&#160;protocol is&#160;provided&#160;for time-lapse imaging of ocular morphogenesis using a commercially available lightsheet microscope and an image processing workstation to analyze the resulting data. This protocol details the procedures for embryo anesthesia, embedding in low melting temperature agarose, suspension in the imaging chamber, setting up the imaging parameters, and finally analyzing the imaging data using image analysis software. The resulting dataset can provide valuable insights into the process of ocular morphogenesis, as well as perturbations to this process as a result of genetic mutation, exposure to pharmacological agents, or other experimental manipulations.

Here, a&#160;protocol is&#160;provided&#160;for time-lapse imaging of ocular morphogenesis using a commercially available lightsheet microscope and an image processing workstation to analyze the resulting data. This protocol details the procedures for embryo anesthesia, embedding in low melting temperature agarose, suspension in the imaging chamber, setting up the imaging parameters, and finally analyzing the imaging data using image analysis software.&#160;

Vertebrate eye development is a complex process that begins near the end of embryo gastrulation and requires the precise coordination of cell migration, ...

Imaging C. elegans Embryos using an Epifluorescent Microscope and Open Source Software

Cellular processes, such as chromosome assembly, segregation and cytokinesis,are inherently dynamic. Time-lapse imaging of living cells, using fluorescent-labeled reporter proteins or differential interference contrast (DIC) microscopy, allows for the examination of the temporal progression of these dynamic events which is otherwise inferred from analysis of fixed samples1,2. Moreover, the study of the developmental regulations of cellular processes necessitates conducting time-lapse experiments on an intact organism during development. The Caenorhabiditis elegans embryo is light-transparent and has a rapid, invariant developmental program with a known cell lineage3, thus providing an ideal experiment model for studying questions in cell biology4,5and development6-9. C. elegans is amendable to genetic manipulation by forward genetics (based on random mutagenesis10,11) and reverse genetics to target specific genes (based on RNAi-mediated interference and targeted mutagenesis12-15). In addition, transgenic animals can be readily created to express fluorescently tagged proteins or reporters16,17. These traits combine to make it easy to identify the genetic pathways regulating fundamental cellular and developmental processes in vivo18-21. In this protocol we present methods for live imaging of C. elegans embryos using DIC optics or GFP fluorescence on a compound epifluorescent microscope. We demonstrate the ease with which readily available microscopes, typically used for fixed sample imaging, can also be applied for time-lapse analysis using open-source software to automate the imaging process.

Imaging C. elegans Embryos using an Epifluorescent Microscope and Open Source Software

The C. elegans embryo is a powerful system for studying cell biology and development. We present a protocol for live imaging of C. elegans embryos utilizing DIC optics or fluorescence using readily available epifluorescent microscopes and open-source software.

Cellular processes, such as chromosome assembly, segregation and cytokinesis,are inherently dynamic. Time-lapse imaging of living cells, using ...

Refined Multiview Light Sheet Microscopy Protocol for Zebrafish Embryos

Light Sheet Microscopy Imaging and Mounting Strategies for Early Zebrafish Embryos

Light sheet microscopy has become the methodology of choice for live imaging of zebrafish embryos over long time scales with minimal phototoxicity. In particular, a multiview system, which allows sample rotation, enables imaging of entire embryos from different angles. However, in most imaging sessions with a multiview system, sample mounting is a troublesome process as samples are usually prepared in a polymer tube. To aid in this process, this protocol describes basic mounting strategies for imaging early zebrafish development between the 70% epiboly and early somite stages. Specifically, the study provides statistics on the various positions the embryos default to at the 70% epiboly and bud stages within the chorion. Furthermore, it discusses the optimum number of angles and the interval between angles required for imaging whole zebrafish embryos at the early stages of development so that cellular-scale information can be extracted by fusing the different views. Finally, since the embryo covers the entire field of view of the camera, which is required to obtain a cellular-scale resolution, this protocol details the process of using bead information from above or below the embryo for the registration of the different views.

Author Spotlight: Strategies for Mounting Zebrafish Embryos for High-Resolution Multiview Light-Sheet Microscopy &#8212; Techniques for Imaging and Image Reconstruction

A sample preparation strategy for imaging early zebrafish embryos within an intact chorion using a light-sheet microscope is described. It analyzes the different orientations that embryos acquire within the chorion at the 70% epiboly and bud stages and details imaging strategies for obtaining cellular-scale resolution throughout the embryo using the light-sheet system.

Light sheet microscopy has become the methodology of choice for live imaging of zebrafish embryos over long time scales with minimal phototoxicity. In ...

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 ...

Light Sheet Microscopy Sample Preparation: Mounting Live Zebrafish Embryos for Long-Term Imaging

Source: Icha J., et al. <a href="https://www.jove.com/v/53966/" target="_blank">Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development</a>. J. Vis. Exp. (2016).
This article describes sample preparation for light-sheet fluorescence microscopy. It also demonstrates a method to mount live zebrafish embryos for long term imaging, which allows recording of long time-lapse movies of live embryos in near-physiological conditions.

Source: Icha J., et al. Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development. J. Vis. Exp. (2016).
This article describes sample ...

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

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