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 imaging, we see e.g., the following susceptibilities. The egg collection cultures may have different ages at the time of embryo collection. In Drosophila, it has been shown that this affects the qualities of the progeny45. Although working with appropriately desynchronized cultures is an option, simultaneous imaging with synchronized cultures is considerably more convenient. Both the Drosophila- and Tribolium-based representative results derive from synchronized egg collection cultures, and we can confidently exclude that the different fluorescence characteristics are an unintended effect of parental senescence.
Dechorionation is a crucial step as prolonged exposure to sodium hypochlorite decreases embryo viability. Inconveniently, the ionic strength of sodium hypochlorite decreases over time. Measurements of the ionic strength are possible46, but laborious and cumbersome. In simultaneous live imaging, a master mix can be used for all lines and/or conditions during the preparation of the dechorionation-associated 6-well plates.
Even in laboratories with efficient air condition, small but irregular temperature fluctuations are inevitable and also affect temperature-controlled9 sample chamber-based LSFMs. Theoretically, the temperature in the sample chamber can be logged, but an exact repetition is not possible. In simultaneous live imaging, these fluctuations also occur, but they affect all embryos likewise.
From the practical point of view, a protocol for simultaneous imaging of multiple embryos increases the throughput. In live imaging, the duration of the developmental process of interest determines how many assays can be conducted within a given period, which is an important consideration when access to LSFMs is limited. Prime examples for long-term recording are live imaging assays in which cells are tracked. Such experiments usually range over long development periods. While the embryonic development of Drosophila takes only about one day at room temperature47, Tribolium takes about seven days48. Hence, for this species, parallelization is essential to remain within a reasonable time frame, especially regarding&#160;assays where many conditions are compared.
The throughput, i.e., the number of embryos that are imaged simultaneously, is primarily limited by two factors: First is the space available along the y axis within the sample chamber. When the lowermost embryo is about to be imaged, the uppermost embryo should not have left the imaging buffer. Further, the cobweb holder and the associated support system should not be so lengthy that it wobbles and/or jitters noticeably during the z stack recording process. Second&#160;is the intended temporal resolution. The recording time per embryo can be approximated by multiplying the exposure time per two-dimensional image (usually 10-200 ms) with the number of planes per z stack (usually several hundred), the number of channels (usual between one and three), the number of directions (usually between one and eight) and&#160;a setup-dependent factor for the translation and rotation movements (estimated between 1.2 and 1.5). The ratio of the intended temporal resolution and the recording time per embryo indicates the maximum number of embryos that can be imaged simultaneously. In summary, our protocol is appropriate to advance from sequential live imaging to medium throughput, but not suitable for what is generally regarded as high throughput. For the latter, LSFMs suitable for microscope slides49,50,51 or well plates52 can be considered, but these implementations usually have other limitations.
Depending on the experimental question and hence the illumination lens/detection lens/camera combination in conjunction with the imaging parameters, simultaneously imaged LSFM datasets may require vast storage space in their raw state. This issue can best be illustrated by considering the provided Drosophila&#160;example&#160;datasets (Supplementary Table 1), i.e., DS0001-3. Each two-dimensional raw image had a data volume of about 2.8 Megabytes, each raw z stack consisted of 120 planes, and all three embryos were recorded along four directions a total of 143 times. In consequence, the raw datasets occupied about half a Terabyte. Through image data processing, i.e., cropping along all spatial dimensions (Step 9) in conjunction with ZIP-based compression, the data volume was reduced to 53.3 Gigabyte, which is about 10% of the initial amount. To process datasets in the Terabyte regime, the BigDataViewer53 plugin, which is pre-installed in FIJI (Plugins | BigDataViewer), is recommended.
In addition to the five scenarios outlined in the Representative Results section, our protocol can also be used to characterize the effect of RNAi knockdown effects, which is a popular approach in Tribolium54. However, it is only of limited use for the analysis of compounds (e.g., insecticides or other biochemical stressors) if these should be applied through the imaging buffer &#8211; which is the most straightforward approach &#8211; as simultaneous imaging of a control embryo is not possible.
We demonstrated our protocol using one of our custom-built microscopes8,55, but it is applicable to any sample chamber-based setup, e.g., OpenSPIM56 or most commercial LSFMs. This protocol complements the LSFM-associated resource establishment initiative that aims at encouraging novices to actively integrate light sheet-based technology&#160;into their research plans. Most universities and institutes nowadays operate well-equipped fluorescence microscopy&#160;facilities57. These usually provide access to one or more chamber-based LSFMs, but&#160;the staff cannot be expected to have the proper experimental solutions for every scientific question at hand.
Our results show that our experimental framework is suitable for Drosophila and Tribolium. We are confident that our comparison-focused protocol can be used to study the embryogenesis of other insect species. For instance,&#160;a LSFM-based approach has been applied to compare wild-type and knockdown development in the scuttle fly Megaselia abdita58, and follow-up&#160;studies would also benefit from our mounting method. In addition, transgenic lines of the two-spotted cricket Gryllus bimaculatus specifically designed for fluorescence live imaging59 are available, but the only currently available comparison-focused protocol60 is intended for conventional inverted microscopes and therefore does not address three-dimensional image acquisition or sample rotation. Additionally, it has already been shown that the cobweb holder is a convenient choice for&#160;zebrafish, whose embryos change their shape considerably during embryogenesis61. An adaptation of our protocol, which requires basically no optimization effort, would provide the above-described advantages also for this model organism.

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>&#160;</td>
		</tr>
		<tr>
			<td>24-well plate</td>
			<td>Orange Scientific</td>
			<td>4430300</td>
			<td>Only for live imaging involving Tribolium</td>
		</tr>
		<tr>
			<td>35-mm &#216; Petri dish</td>
			<td>Fisher Scientific</td>
			<td>153066</td>
			<td>Only for live imaging involving Drosophila.</td>
		</tr>
		<tr>
			<td>90-mm &#216; Petri dish</td>
			<td>Fisher Scientific</td>
			<td>L9004575</td>
			<td></td>
		</tr>
		<tr>
			<td>100-&#181;m mesh size cell strainer</td>
			<td>BD Biosciences</td>
			<td>352360</td>
			<td></td>
		</tr>
		<tr>
			<td>250-&#181;m mesh size sieve</td>
			<td>VWR International</td>
			<td>200.025.222-038</td>
			<td>Only for live imaging involving Tribolium</td>
		</tr>
		<tr>
			<td>300-&#181;m mesh size sieve</td>
			<td>VWR International</td>
			<td>200.025.222-040</td>
			<td>Only for live imaging involving Tribolium</td>
		</tr>
		<tr>
			<td>710-&#181;m mesh size sieve</td>
			<td>VWR International</td>
			<td>200.025.222-050</td>
			<td>Only for live imaging involving Tribolium</td>
		</tr>
		<tr>
			<td>800-&#181;m mesh size sieve</td>
			<td>VWR International</td>
			<td>200.025.222-051</td>
			<td>Only for live imaging involving Tribolium</td>
		</tr>
		<tr>
			<td>405 fine wheat flour</td>
			<td>Demeter e.V.</td>
			<td>SP061006</td>
			<td>Only for live imaging involving Tribolium</td>
		</tr>
		<tr>
			<td>commercially available Drosophila medium</td>
			<td>Genesee Scientific</td>
			<td>66-115</td>
			<td>Only for live imaging involving Drosophila / Custom-made Drosophila medium may also be used</td>
		</tr>
		<tr>
			<td>fluorescent microspheres, 1.0 &#181;m &#216;</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>Only for live imaging involving Tribolium</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 Drosophila</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 Tribolium / 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 Drosophila</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 examples for the last two scenarios.
In our first exemplary application, we show the fluorescence signal dynamics of three embryos that derive from different transgenic Drosophila lines (Table 2) over a period of about 1 day (Figure 4, Supplementary Movie 1). For these lines, we expected rather similar fluorescence patterns since all of them express nuclear-localized EGFP/GFP under control of different presumably ubiquitous and constitutively active promoters. However, our comparative live imaging results show that there are strong spatiotemporal differences in the expression patterns that are certainly not secondary effects due to ambient variance.
In our second application example, we compare&#160;the performance of three embryos that derive from the AGOC{Zen1&#8217;#O(LA)-mEmerald} #1, #2 and #3 (Zen1&#8217;LA-mE #1, #2 and #3, respectively) transgenic Tribolium sublines36. All of these carry the same piggyBac-based transgene that leads to expression of mEmerald-linked37 Lifeact38 under control of the zerkn&#252;llt 139 promoter, a transcription factor involved in serosa specification. Our comparative live imaging results, which illustrate the serosa window closure process during gastrulation, suggest that subline #2 provides a remarkably stronger overall signal than the other two sublines (Figure 5, Supplementary Movie 2). This indicates that also in Tribolium, the genomic context may have a strong influence on the expression level of transgenes that carry an expression cassette.
Successful retrieval of all six embryos that are shown in this section was possible as described in Step 8 of our protocol. All of them developed into fully functional and fertile adults (Supplementary Table 1, &#8220;Retrieval&#8221; row), indicating that the overall procedure, from dechorionation over mounting to recording, was non-invasive. This level of quality control is essential whenever wild-type development is expected, e.g., regarding control embryos that are imaged simultaneously with embryos in which one or more genes are knocked down or knocked out.
All datasets that were used to create the representative results are provided as resources that will help especially LSFM novices to evaluate the quality of their own work. The digital object identifier-based download links can be found in the metadata table (Supplementary Table 1, &#8220;Data Access&#8221; row).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61713/61713fig01.jpg" /> 
Figure 1: Experimental framework and mounting method overview.&#160;(A) Flowchart of the ten protocol steps with short reminders and an estimation of the required time. Tasks marked with asterisks do not have to be performed during every assay but depending on the circumstances. Green boxes indicate steps associated with Drosophila and/or Tribolium, blue boxes indicate steps associated with microscopy. (B) Detail drawing of the cobweb holder. The presented design is suitable for cylinder-based clamp mechanisms, which are commonly used in sample chamber-based LSFMs. Specifications are in millimeters. The holder can be scaled as long as the working distance of the detection lens is respected. (C) Recording sequence for live imaging of multiple embryos along multiple directions. At first, z stacks of the uppermost embryo are recorded along up to four orientations, i.e. 0&#176;, 90&#176;, 180&#176; and 270&#176;. Subsequently, the embryo below is moved in front of the detection lens and the recording/rotation sequence begins anew. <a href="https://www.jove.com/files/ftp_upload/61713/61713fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61713/61713fig02v2.jpg" /> 
Figure 2: Fluorescent microsphere-based calibration chart for LSFMs.&#160;The chart illustrates how fluorescent microspheres appear&#160;in the x, y and z maximum projections if the LSFM is correctly calibrated (A), or if there is an illumination (B) or detection offset (C). <a href="https://www.jove.com/files/ftp_upload/61713/61713fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61713/61713fig03v2.jpg" /> 
Figure 3: Collection, dechorionation, mounting and imaging of multiple embryos.&#160;(A) Tribolium embryo collection overview. The illustrations are cited throughout Step 4 according to the numbers in the upper left. (B) Preparation and transfer scheme for the 6-well plate used during Step 5. The B1 and B2 wells contained diluted sodium hypochlorite (NaOCl), which induces dechorionation. (C) The 100 &#181;m cell strainer, which was used to transfer the embryos from one well to another, within the B2 well. The upper detail image shows a close-up of several Tribolium embryos on top of the cell strainer mesh, the lower detail image shows a transmission light stereo microscope image of several Tribolium embryos with partially detached chorion. (D) Cobweb holder with three mounted Tribolium embryos. The embryos, as well as their anterior-posterior axes, were aligned along the long axis of the slotted hole. (E) Movement of the cobweb holder within the sample chamber of the LSFM during the recording of three embryos. <a href="https://www.jove.com/files/ftp_upload/61713/61713fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61713/61713fig04.jpg" /> 
Figure 4: Comparative fluorescence live imaging of one embryo each from the #01691, #29724 and #24163 transgenic Drosophila lines.&#160;Embryos are shown along two (of four recorded) directions over a period of 20:00 h (from a total imaging period of 23:40 h). No dynamic intensity correction over time was performed. Metadata for the Drosophila datasets can be found in Supplementary Table 1. ZA, Z maximum projection with image adjustment. Scale bar, 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/61713/61713fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61713/61713fig05.jpg" /> 
Figure 5: Comparative fluorescence live imaging of one embryo each from the Zen1&#8217;LA-mE #1, #2 and #3 transgenic Tribolium sublines.&#160;(A) Embryos are shown along four directions during gastrulation, just prior to serosa window closure. (B) Embryos are shown ventrally over a period of 01:30 h (from a total imaging period of 118:00 h). Image adjustment was performed with identical minimum and maximum displayed values, no dynamic intensity correction over time was performed. Metadata for the Tribolium datasets can be found in Supplementary Table 1. Datasets were synchronized to the stage shown in (A). ZA, Z maximum projection with image adjustment. Scale bar, 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/61713/61713fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Combination</td>
			<td>Rationale</td>
			<td>Illumination lens</td>
			<td>Detection lens</td>
			<td>Camera</td>
			<td>Reference</td>
		</tr>
		<tr>
			<td>#1</td>
			<td>entire embryo on the cellular level,&#160;large pixel spacing&#160;(small data volume)</td>
			<td>2.5&#215; magnification numerical aperture 0.06&#160;(air)</td>
			<td>10&#215; magnification numerical aperture 0.3&#160;(water-dipping)</td>
			<td>CCD with 1.4 megapixel (1392&#215;1040, 6.45 &#181;m pitch)</td>
			<td>this study, Drosophila41, Tribolium20,41</td>
		</tr>
		<tr>
			<td>#2</td>
			<td>entire embryo on the cellular/subcellular level,&#160;small pixel spacing&#160;(large data volume)</td>
			<td>2.5&#215; magnification numerical aperture 0.06&#160;(air)</td>
			<td>20&#215; magnification numerical aperture 0.5&#160;(water-dipping)</td>
			<td>sCMOS with 5.5 megapixel (2560&#215;2160, 6.5 &#181;m pitch)</td>
			<td>Tribolium9</td>
		</tr>
		<tr>
			<td>#3</td>
			<td>specific regions of the embryo on the subcellular level or specialized applications e.g. live imaging of dissected germbands42,43</td>
			<td>5.0&#215; magnification numerical aperture 0.16&#160;(air)</td>
			<td>40&#215; magnification numerical aperture 0.75&#160;(water-dipping)</td>
			<td>CCD with 1.4 megapixel (1392&#215;1040, 6.45 &#181;m pitch)</td>
			<td>Tribolium44</td>
		</tr>
	</tbody>
</table>
Table 1: Recommended illumination lens/detection lens/camera combinations for live imaging of Drosophila and Tribolium embryos using LSFM. All combinations have been successfully employed in previous studies (see Reference row). Please note that most sample chamber-based LSFMs use water-dipping lenses for detection, which are primarily designed for imaging buffers with a refractive index of 1.33 (such as autoclaved tap water or other aqueous media). Imaging buffers with other refractive indices can be used40, but this changes certain properties of the optical system, e.g. the working distance.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Line</td>
			<td>Genotype</td>
			<td>Fluorophore expression</td>
		</tr>
		<tr>
			<td>#01691</td>
			<td>y[1] w[67c23]; P{w[+mC]=Ubi-GFP.nls}ID-2; P{Ubi-GFP.nls}ID-3</td>
			<td>NLS-GFP under control of the polyubiquitin promoter</td>
		</tr>
		<tr>
			<td>#29724</td>
			<td>w[*]; P{w[+mC]=Tub84B-EGFP.NLS}3</td>
			<td>NLS-GFP under control of the alpha-tubulin 84B promoter</td>
		</tr>
		<tr>
			<td>#24163</td>
			<td>w[*]; P{w[+mC]=His2Av-EGFP.C}2/SM6a</td>
			<td>EGFP-labeled histone 2A variant in all cells</td>
		</tr>
	</tbody>
</table>
Table 2: The three transgenic Drosophila lines used in the first application example. The &#8220;line&#8221; column refers to the Bloomington Drosophila Stock Center (bdsc.indiana.edu) stock number.
Supplementary Video 1: Comparative fluorescence live imaging of one embryo each from the #01691, #29724 and #24163 transgenic Drosophila lines. Each transgenic line expresses nuclear-localized EGFP/GFP under control of a presumably ubiquitous and constitutively active promoter. ZA, Z maximum projection with image adjustment. No dynamic intensity correction over time was performed. ZA, Z maximum projection with image adjustment. <a href="https://cloudflare.jove.com/files/ftp_upload/61713/Supplementary_Video_1_JQ.avi" target="_blank">Please click here to download this video.</a>
Supplementary Video 2: Comparative fluorescence live imaging of one embryo each from the Zen1&#8217;LA-mE #1, #2 and #3 transgenic Tribolium sublines. Each transgenic subline expresses mEmerald-labeled Lifeact under control of the zerkn&#252;llt 1 promoter, a transcription factor involved in serosa specification. Please note that the Zen1&#8217;LA-mE #1 embryo turns approximately 90&#176; within the serosa right after serosa window closure. Datasets were synchronized to the stage shown in Figure 4A. Image adjustment was performed with identical minimum and maximum displayed values, no dynamic intensity correction over time was performed. Embryos are shown ventrally and laterally over a period of 48:00 h. ZA, Z maximum projection with image adjustment. <a href="https://cloudflare.jove.com/files/ftp_upload/61713/Supplementary_Video_2_JQ.avi" target="_blank">Please click here to download this video.</a>
Supplementary File 1: FIJI batch processing script for the automated calculation of z maximum projections from all z stacks within one folder. The script can be opened in FIJI via drag-and-drop. At the start (&#8220;Run&#8221;) only the input folder must be specified. The output subfolder (&#34;Z Maximum Projections&#34;) is created automatically. <a href="https://cloudflare.jove.com/files/ftp_upload/61713/Supplementary_File_1_FIJI_Batch_Processing_Script.zip" target="_blank">Please click here to download this file.</a>
Supplementary Table 1: Metadata and parameter for the long-term live imaging datasets DS0001-6. <a href="https://cloudflare.jove.com/files/ftp_upload/61713/20_07_11_JoVE_LiveImaging_SupplementaryTable1_Revision.docx" target="_blank">Please click here to download this table.</a>

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

Universidad San Sebastian

Research

JoVE Journal

Biology

2.9K Views.  Goethe-Universität. 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.

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

Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo. Here we describe a method to mount zebrafish embryos for long-term imaging and demonstrate how to automate the capture of time-lapse images using a confocal microscope. We also describe a method to create controlled, precise damage to individual branches of peripheral sensory axons in zebrafish using the focused power of a femtosecond laser mounted on a two-photon microscope. The parameters for successful two-photon axotomy must be optimized for each microscope. We will demonstrate two-photon axotomy on both a custom built two-photon microscope and a Zeiss 510 confocal/two-photon to provide two examples.

Zebrafish trigeminal sensory neurons can be visualized in a transgenic line expressing GFP driven by a sensory neuron specific promoter 1. We have adapted this zebrafish trigeminal model to directly observe sensory axon regeneration in living zebrafish embryos. Embryos are anesthetized with tricaine and positioned within a drop of agarose as it solidifies. Immobilized embryos are sealed within an imaging chamber filled with phenylthiourea (PTU) Ringers. We have found that embryos can be continuously imaged in these chambers for 12-48 hours. A single confocal image is then captured to determine the desired site of axotomy. The region of interest is located on the two-photon microscope by imaging the sensory axons under low, non-damaging power. After zooming in on the desired site of axotomy, the power is increased and a single scan of that defined region is sufficient to sever the axon. Multiple location time-lapse imaging is then set up on a confocal microscope to directly observe axonal recovery from injury.

Here we describe a method for mounting zebrafish embryos for long-term imaging, two-photon imaging and tissue-damage techniques, and time-lapse confocal imaging.

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo.  ...

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Monitoring Plasmid Replication in Live Mammalian Cells over Multiple Generations by Fluorescence Microscopy

Few naturally-occurring plasmids are maintained in mammalian cells. Among these are genomes of gamma-herpesviruses, including Epstein-Barr virus (EBV) and Kaposi's Sarcoma-associated herpesvirus (KSHV), which cause multiple human malignancies 1-3. These two genomes are replicated in a licensed manner, each using a single viral protein and cellular replication machinery, and are passed to daughter cells during cell division despite their lacking traditional centromeres 4-8.
Much work has been done to characterize the replications of these plasmid genomes using methods such as Southern blotting and fluorescence in situ hybridization (FISH). These methods are limited, though. Quantitative PCR and Southern blots provide information about the average number of plasmids per cell in a population of cells. FISH is a single-cell assay that reveals both the average number and the distribution of plasmids per cell in the population of cells but is static, allowing no information about the parent or progeny of the examined cell.
Here, we describe a method for visualizing plasmids in live cells. This method is based on the binding of a fluorescently tagged lactose repressor protein to multiple sites in the plasmid of interest 9. The DNA of interest is engineered to include approximately 250 tandem repeats of the lactose operator (LacO) sequence. LacO is specifically bound by the lactose repressor protein (LacI), which can be fused to a fluorescent protein. The fusion protein can either be expressed from the engineered plasmid or introduced by a retroviral vector. In this way, the DNA molecules are fluorescently tagged and therefore become visible via fluorescence microscopy. The fusion protein is blocked from binding the plasmid DNA by culturing cells in the presence of IPTG until the plasmids are ready to be viewed.
This system allows the plasmids to be monitored in living cells through several generations, revealing properties of their synthesis and partitioning to daughter cells. Ideal cells are adherent, easily transfected, and have large nuclei. This technique has been used to determine that 84% of EBV-derived plasmids are synthesized each generation and 88% of the newly synthesized plasmids partition faithfully to daughter cells in HeLa cells. Pairs of these EBV plasmids were seen to be tethered to or associated with sister chromatids after their synthesis in S-phase until they were seen to separate as the sister chromatids separated in Anaphase10. The method is currently being used to study replication of KSHV genomes in HeLa cells and SLK cells. HeLa cells are immortalized human epithelial cells, and SLK cells are immortalized human endothelial cells. Though SLK cells were originally derived from a KSHV lesion, neither the HeLa nor SLK cell line naturally harbors KSHV genomes11. In addition to studying viral replication, this visualization technique can be used to investigate the effects of the addition, removal, or mutation of various DNA sequence elements on synthesis, localization, and partitioning of other recombinant plasmid DNAs.

A method of observing individual DNA molecules in live cells is described. The technique is based on the binding of a fluorescently tagged lac repressor protein to binding sites engineered into the DNA of interest. This method can be adapted to follow many recombinant DNAs in live cells over time.

Few naturally-occurring plasmids are maintained in mammalian cells.  Among these are genomes of gamma-herpesviruses, including Epstein-Barr virus  (EBV) ...

Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or&#160;photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.

We demonstrate the use of fluorescence photo activation localization microscopy (FPALM) to simultaneously image multiple types of fluorescently labeled molecules&#160;within cells. The techniques described yield the localization of thousands to hundreds of thousands of individual fluorescent labeled proteins, with a precision of tens of nanometers&#160;within single cells.

Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled ...

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.

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 ...

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 ...

Sample Preparation for Single Virion Atomic Force Microscopy and Super-resolution Fluorescence Imaging

Immobilization of virions to glass surfaces is a critical step in single virion imaging. Here we present a technique&#160;adopted from single molecule imaging assays which allows adhesion of single virions to glass surfaces with specificity. This preparation is based on grafting the surface of the glass with a mixture of PLL-g-PEG and PLL-g-PEG-Biotin, adding a layer of avidin, and finally creating virion anchors through attachment of biotinylated virus specific antibodies. We have applied this technique across a range of experiments including atomic force microscopy (AFM) and super-resolution fluorescence imaging. This sample preparation method results in a control adhesion of the virions to the surface.

The attachment of virions to a surface is a requirement for single virion imaging by Super-resolution fluorescence imaging or atomic force microscopy (AFM). Here we demonstrate a sample preparation method for controlled adhesion of virions to glass surfaces suitable for use in AFM and super-resolution fluorescence imaging.

Immobilization of virions to glass surfaces is a critical step in single virion imaging. Here we present a technique&#160;adopted from single molecule ...

Semi-automated Imaging of Tissue-specific Fluorescence in Zebrafish Embryos

Zebrafish embryos are a powerful tool for large-scale screening of small molecules. Transgenic zebrafish that express fluorescent reporter proteins are frequently used to identify chemicals that modulate gene expression. Chemical screens that assay fluorescence in live zebrafish often rely on expensive, specialized equipment for high content screening. We describe a procedure using a standard epifluorescence microscope with a motorized stage to automatically image zebrafish embryos and detect tissue-specific fluorescence. Using transgenic zebrafish that report estrogen receptor activity via expression of GFP, we developed a semi-automated procedure to screen for estrogen receptor ligands that activate the reporter in a tissue-specific manner. In this video we describe procedures for arraying zebrafish embryos at 24-48 hours post fertilization (hpf) in a 96-well plate and adding small molecules that bind estrogen receptors. At 72-96 hpf, images of each well from the entire plate are automatically collected and manually inspected for tissue-specific fluorescence. This protocol demonstrates the ability to detect estrogens that activate receptors in heart valves but not in liver.

Described here is a protocol for semi-automated imaging of tissue-specific fluorescence in zebrafish embryos.

Zebrafish embryos are a powerful tool for large-scale screening of small molecules. Transgenic zebrafish that express fluorescent reporter proteins are ...

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) ...