We thank John Murray for integrated strain, ujIs113, to generate lineaging strain BV514; Brandon Harvey (NIBIB) for help with testing the protocol; Jon Daniels and Gary Rondeau (Applied Scientific Instrumentation) for assistance with Micro-Manager and diSPIM instrument; and Andrew York and Hank Eden for their critical feedback on the diSPIM system. We also thank the Research Center for Minority Institutions program and the Instituto de Neurobiolog&#237;a Jose del Castillo (Universidad de Puerto Rico) for providing a meeting and brainstorming platform. Much of this work was conducted at the Marine Biological Laboratory at Woods Hole through the Whitman Program. This work was supported by the Intramural Research Programs of the NIH National Institute of Biomedical Imaging and Bioengineering and by NIH grant No. U01-HD075602 and No. R24-OD016474. Mark W. Moyle was supported by F32-NS098616 and Leighton H. Duncan was supported by a Diversity Supplement to R24-OD016474.

<ol>
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The authors have nothing to disclose.

C. elegans stands out as the only organism with the final positions and connectivity of each adult neuron known27. However, the developmental dynamics leading to organization of the working circuits and networks that makes up the C. elegans connectome remain unknown. Based on opportunities emerging from advances in light microscopy, we can now capture and analyze cell positions, morphogenesis, and neurogenesis throughout C. elegans embryonic development.
The procedure that we have described and that we routinely use in the lab yields 4D-isotropic images of labeled neurons and nuclei for cell-lineaging in C. elegans embryos. More importantly, we have optimized long-term imaging conditions with the diSPIM and coupled semi-automated lineaging capabilities with high-resolution images to improve the speed and precision of analyzing C. elegans embryogenesis. This integrated protocol will enable users to visualize and identify cells and quantitate three-dimensional features such as neurite migration and axon fasciculation through onset of early twitching. This procedure can be readily adapted into any facility with an ASI diSPIM system, and we recommend this system specifically for this protocol. Other SPIM formulations offered commercially may differ from the ASI configuration in the sample chamber and optical properties. However, data exported from other platforms can also be put through our data pipeline. Therefore, appraisal of their value in lineaging, a demanding test of image quality and instrument stability, is feasible. Even though we actively use the diSPIM to regularly image other specimens (such as drosophila and zebrafish embryos), the described and comprehensive lineaging analysis of embryos is still currently limited to the nematode species. For larger or thick samples, we opt to use stage-scanning approaches, which scan the samples through a stationary light sheet. Kumar et al. have previously demonstrated this improved diSPIM sectioning to yield high quality images from thick samples without additional modifications to the diSPIM10.
The critical steps within the protocol include mounting C. elegans embryos on the poly-L-lysine coated coverslip, data acquisition, and data processing. Harvesting and mounting C. elegans embryos on the glass coverslip can be challenging to inexperienced users, but here we provide a detailed protocol of key steps to facilitate learning. If long-term imaging is desired, we obtain best results harvesting four-cell or earlier embryos from 8-10 young adults28. Note that old adults are less desirable to harvest early stage embryos because they tend to contain older embryos in the uterus and unfertilized eggs. In regards to mounting embryos, problems such as blockage in the assembled aspirator (mouth pipette) or a too-large of an opening in the microcapillary pipette may prevent proper mounting and orientation of embryos. To prepare for optimal imaging, we perform pre-acquisition testing on early and late pre-twitching embryos to check the performance of the light sheets, cameras, objectives, and autofocus. We obtain best results when all of these operations are tested and yield high quality images during our pre-acquisition testing. This is particularly relevant for generating images with isotropic spatial resolution, for which raw images acquired from both views (objectives) must be of high-quality. After acquisition, the volumes acquired for each view are processed to yield isotropic images. It is important to use an appropriate graphics processing unit (GPU) card as described in this protocol (see below). This improves the processing speed at which the isotropically fused images are generated, shortening the time to data analyses. It is also imperative that users are running the latest version of CytoSHOW and are using the parameters provided with our download bundle for StarryNite auto-lineaging. If users are interested in using auto-lineaging for other samples (e.g., zebrafish, drosophila etc.) then additional optimization to the parameters used in StarryNite will be required (see references3,4).
Although our integrated protocol provides images and lineaging results in the pre-twitching embryo, users should be aware that automated lineaging in the post-twitching embryo is currently not feasible: nuclear positions changes on the order of seconds in the post-twitching embryo, too rapidly to allow lineage tracking. However, the diSPIM has indeed demonstrated a promising capability to capture neurodevelopmental events and track some cell positions in the post-twitching stages of embryogenesis23,29. If users are interested in examining the post-twitching embryo, the diSPIM does provide the speed to obtain volumetric snapshots and track fine neurodevelopmental events, such as neurite outgrowth, in rapidly moving embryos.
This protocol will be foundational for the cell-by-cell completion of the WormGUIDES atlas30, as it will provide an integrated approach with high resolution isotropic images to identify and capture 3D morphologies of labeled neurons during the first 430 minutes of embryogenesis. As it stands, the prototype WormGUIDES atlas provides nuclear positions of cells in the developing embryo and aims to capture the developmental dynamics of a subset of embryonic neurons. This protocol will be a key for the integration of additional developing neurons into the WormGUIDES atlas30.
Our integrated protocol will also simplify exploring new gene expression profiles in the C. elegans embryo. In transgenic C. elegans, many cell-specific promoters spatially and temporally control transgene expression. While the expression patterns of most genes have been extensively characterized in the adult animal31,32,33,34, nearly all have yet to be characterized in the developing (especially late-stage) embryo. The C. elegans promoterome has been a useful resource to the worm community to drive cell-specific transgene expression, as well as determine whether gene function is cell-autonomous or non-autonomous. Capturing isotropic high-resolution and dynamic expression patterns of genes, and precisely identifying expressing cells via lineaging will be valuable to many in the scientific community.
Embryogenesis comprises two intertwined major processes, cellular differentiation and tissue morphogenesis. A great deal is known about the mechanisms and molecules that define distinct cell types during the development of C. elegans. However, little is known about the mechanisms important for cell migration, cell adhesion, and cell shape in the C. elegans embryo. With the C. elegans invariant cell lineage known, our protocol lets us readily discern the catalogued 3D-microanatomy of the embryo during morphogenesis at new levels of detail: e.g., axon fasciculation, synaptogenesis, and neuronal activity. Ardiel et al. have previously demonstrated the power of the diSPIM to capture calcium transients at the level of a single neurons in C. elegans embryos23. Many other aspects of developmental physiology are ripe for inquiry by these methods.
Finally, this protocol is largely automated and systematically reduces the time it takes to generate deconvolution images and perform cell-lineaging via StarryNite and Acetree. The software strategies used in this protocol can be applied to many questions of biology far-flung from the very specific fields in which we have demonstrated them here.
Details on software compatibility and download access
Information on Micro-Manager and plugins for diSPIM imaging are available at http://dispim.org/software/micro-manager and https://micro-manager.org/wiki/ASIdiSPIM_Plugin.
The data-processing pipeline currently requires a Windows operating system. We have bundled a single archive file to simplify installation of all required data-processing programs and support files. It is available for download at http://dispimlineage.wormguides.org.
CytoSHOW (http://run.cytoshow.org/) is based on the widely used and open source image analysis platform, ImageJ (v1). Java must be installed and up-to-date on the computer to use CytoSHOW, and updates to CytoSHOW are deployed automatically via Java Web Start. Many ImageJ-based functions of CytoSHOW are as described and illustrated at https://imagej.nih.gov/ij/docs/examples/index.html. CytoSHOW has been customized to display multidimensional raw data from the ASI diSPIM, as well as other imaging software that creates TIFF output. In principle, other multi-view SPIM imaging systems could be supported by minor modifications of CytoSHOW to allow this protocol to be carried out on different microscope systems.

SpimFusion was written in CUDA/C++ using Visual Studio 2013 with CUDA toolkit v7.5. Running SpimFusion requires specific computer hardware: a NVIDIA graphics processing unit (GPU) card with CUDA compute capability 1.0 or higher and a minimum of 2 GB graphics card memory. At the time of publication of our protocol, SpimFusion is unpublished (Min Guo and Hari Shroff) but available in the software bundle archive mentioned above.

A specially built command-line driven version of StarryNite requires that the freely available MATLAB Compiler Runtime is installed, but does not require a license for commercial MATLAB software. The MATLAB Compiler Runtime is included in the software bundle archive mentioned above. The code for StarryNite as used in this protocol is essentially unchanged from that used for confocal images6. However, several operational matters in the creation of input images for StarryNite processing and the handling of StarryNite results have been addressed here by methods in CytoSHOW that enable a continuous data processing pipeline for fused isotropic diSPIM volumes. These changes are automated by CytoSHOW code that handles these pre-and post-processing steps. CytoSHOW also edits a pre-optimized diSPIM-specific template StarryNite parameter set to automatically tune the segmentation algorithm to the fluorescence intensity of nuclei in the imaged data. The unique parameters used by StarryNite on each diSPIM data set are then saved in a file along with the output image and lineaging data.
A custom version of AceTree that works with 16-bit images and maintains compatibility with Java3D rendering is best suited for this protocol. It is also included in the software bundle archive mentioned above.

C. elegans stands out as the only organism in which every cell in the embryo can be observed throughout neurodevelopment. With the entire cell-lineage known and invariant1, and with the development of new tools that allow labeling and continuous imaging of single cells in embryos, biologists can now begin examining different steps in the development of the nematode nervous system from all angles - cell birth; migration and differentiation; neurite formation, targeted outgrowth and fasciculation; synapse formation; and tuning of functional circuits. Capturing neuronal outgrowth dynamics in the C. elegans embryo, by combining stably expressed reporters and fluorescence microscopy, is valuable to the scientific community.
Developmental studies in C. elegans often leverage the invariant cell-lineage and cell-fate maps of this species to augment contextual understanding at the single-cell level within the intact organism1. Auto-lineaging analysis - using StarryNite2,3,4 and AceTree5,6,7,8 software - benefits from high contrast, high resolution images of fluorescent nuclei. To work optimally, StarryNite/AceTree also depends upon predictable constrained orientation of imaged embryos during development. Confocal microscopy, done in C. elegans embryos compressed between two coverslips, has been the standard auto-lineaging microscopy method for more than a decade because it provides both high contrast/high resolution and a predictable constrained orientation of the embryo7,8. We previously described the construction and use of a novel light-sheet-based dual-view inverted selective plane illumination microscope (diSPIM) for live sample imaging such as C. elegans embryogenesis9,10,11,12,13. Light-sheet microscopy, in general, provides low phototoxicity, high speed, and long-term imaging of live 3D specimens14,15. The diSPIM method, specifically, produces four-dimensional (4D) images with nearly isotropic spatial resolution of approximately 330 nm9.
Compared with confocal microscopy, diSPIM offers higher signal-to-noise and speed, more isotropic spatial resolution, and is more suitable for long-term in vivo imaging16. We therefore worked to adapt diSPIM data for input into StarryNite/AceTree and investigated whether this would enhance the lineaging analyses. A major hurdle is that diSPIM specimens cannot be easily constrained by eggshell-compression to adopt expected orientations for StarryNite/ AceTree. Random orientation of cell positions in the volume being analyzed degrades the accuracy of auto-lineaging analysis.
We therefore employed CytoSHOW, a viewer-guided user interface which allows users to select precise 3D orientation of embryos during pre-processing of diSPIM images, yielding image data that is both quality-optimized and context-aware for input into StarryNite/AceTree. Upon user-selection of imaged embryos, CytoSHOW orchestrates an automated data processing pipeline. Cropped and background-subtracted embryo images are saved within TIFF stack files for each position, timepoint and view. CytoSHOW then iteratively calls the program SpimFusion to co-register and jointly deconvolve the two pre-processed views, using the Richardson-Lucy17,18 algorithm to yield isotropic high-resolution volumetric images. A diSPIM-specific set of parameters has been optimized for StarryNite to govern its behavior during image-segmentation and nucleus-tracking in isotropically fused images. Fused images and lineaging results are then edited using AceTree, which allows users to identify and fix any errors in the auto-lineage trace generated by StarryNite. AceTree can also can present lineage-tree and 3D modeled renderings of tracked nuclei in the embryo. We find that auto-lineaging speed and accuracy are markedly enhanced using isotropically fused images, when compared to raw images from either SPIM camera. Our protocol, while optimized for the C. elegans application described here, could be generally adapted for auto-lineaging of diSPIM data produced for other species or specimens. If this is the intended use of the protocol, please note that additional tuning of the StarryNite parameters will likely be required for new specimens, as described3,4.
Successful implementation of this protocol results in images with 4D-isotropic resolution and enables biologists to trace cell lineages, while simultaneously identifying and analyzing neurons in the developing C. elegans embryo. Moreover, by merging several post-processing algorithms - with hardware acceleration being the most time-consuming of these - we can now analyze both fine subcellular details and the cell-lineages and cell-fates of live embryos in essentially real time. This new protocol allows precise, informed manipulation and observation of cell behavior during probative studies of differentiation and morphogenesis in vivo. In this manuscript, we present a detailed explanation of the improved protocols we have developed for lineaging and cell tracking in developing C. elegans embryos, to enhance studies of embryogenesis, gene expression or neurodevelopment.

<table>
	<tbody>
		<tr>
			<td>Steps 1-4</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Concavity slides</td>
			<td>ThermoFisher Scientific</td>
			<td>1519006</td>
			<td>5-18mm diameter, 0.6-0.8mm deep, 1.2-1.5mm</td>
		</tr>
		<tr>
			<td>Dissecting microscope with 10&#215;&#8211;50&#215; zoom range</td>
			<td>Motic</td>
			<td>SMZ-171</td>
			<td></td>
		</tr>
		<tr>
			<td>E. coli (OP50)</td>
			<td>Caenorhabditis Genetics Center (CGC)</td>
			<td>OP50</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass coverslips, no. 1.5, 24 &#215; 50 mm</td>
			<td>VWR International</td>
			<td>48393-241</td>
			<td></td>
		</tr>
		<tr>
			<td>M9 Buffer</td>
			<td></td>
			<td></td>
			<td>Stiernagle, T. Maintenance of C. elegans. WormBook. 1-11, doi:10.1895/wormbook.1.101.1, (2006).</td>
		</tr>
		<tr>
			<td>Methyl cellulose &#160;</td>
			<td>Sigma-Aldrich</td>
			<td>H7509-25G</td>
			<td></td>
		</tr>
		<tr>
			<td>Microcapillary pipette aspirator tube</td>
			<td>Sigma-Aldrich</td>
			<td>A5177</td>
			<td></td>
		</tr>
		<tr>
			<td>Microcapillary pipettes, 0.4-mm i.d</td>
			<td>Drummond Scientific</td>
			<td>1-000-800</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle, no. 18G x 1 &#189; (1.2mm x 40mm)</td>
			<td>BD Precision Glide</td>
			<td>305196</td>
			<td></td>
		</tr>
		<tr>
			<td>NGM plates</td>
			<td></td>
			<td></td>
			<td>prepared as described by Brenner (1974)</td>
		</tr>
		<tr>
			<td>O-ring for imaging chamber</td>
			<td>O-Rings West</td>
			<td>M1.5X40</td>
			<td></td>
		</tr>
		<tr>
			<td>Pasteur pipette</td>
			<td>Corning/Sigma-Aldrich</td>
			<td>CLS7095D5X</td>
			<td></td>
		</tr>
		<tr>
			<td>Platinum wire, 0.5-mm diameter</td>
			<td>Sigma-Aldrich</td>
			<td>267201</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly-L-lysine</td>
			<td>Sigma-Aldrich</td>
			<td>P1524</td>
			<td></td>
		</tr>
		<tr>
			<td>Stainless steel rectangular chamber (76.0 mm x 50.5 mm)</td>
			<td>Applied Scientifics Instrumentations (ASI)</td>
			<td>I2450</td>
			<td></td>
		</tr>
		<tr>
			<td>Worm Eyelash Pick</td>
			<td></td>
			<td></td>
			<td>Hart, A. C. Behavior. WormBook.&#160; (2006).</td>
		</tr>
		<tr>
			<td>Worm Pick</td>
			<td></td>
			<td></td>
			<td>Stiernagle, T. Maintenance of C. elegans. WormBook. 1-11, doi:10.1895/wormbook.1.101.1, (2006).</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Steps 5-6</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>488 nm long-pass filter</td>
			<td>Semrock</td>
			<td>LP02-488 RU-2</td>
			<td></td>
		</tr>
		<tr>
			<td>561-nm notch filter</td>
			<td>Semrock</td>
			<td>NF03-561E-25</td>
			<td></td>
		</tr>
		<tr>
			<td>BLP02-561R-25, quantity 2</td>
			<td>Semrock</td>
			<td></td>
			<td>561 nm EdgeBasic best-value long-pass edge filter</td>
		</tr>
		<tr>
			<td>Control software for bottom camera</td>
			<td>Jenoptik</td>
			<td></td>
			<td>ProgRes CapturePro</td>
		</tr>
		<tr>
			<td>diSIPM assembly video</td>
			<td>Applied Scientifics Instrumentations (ASI)</td>
			<td></td>
			<td>https://youtu.be/TAgbr6IrTqw ; http://www.asiimaging.com</td>
		</tr>
		<tr>
			<td>diSPIM alignment video</td>
			<td>Applied Scientifics Instrumentations (ASI)</td>
			<td></td>
			<td>https://youtu.be/qnOrg30NNuE</td>
		</tr>
		<tr>
			<td>diSPIM imaging PC</td>
			<td>Intel</td>
			<td></td>
			<td>Intel Xeon CPU E5-2630 2.6GHz, 12 cores in total,&#160;64&#160;GB memory, Windows 7</td>
		</tr>
		<tr>
			<td>FF01-525/45-25, quantity 2</td>
			<td>Semrock</td>
			<td></td>
			<td>525/45 nm BrightLine single-band bandpass filter</td>
		</tr>
		<tr>
			<td>FF555-DI03-25X36, quantity 2</td>
			<td>Semrock</td>
			<td></td>
			<td>555 nm edge BrightLine single-edge dichroic beamsplitter</td>
		</tr>
		<tr>
			<td>Imaging PC Graphics Card</td>
			<td>NVIDIDA</td>
			<td></td>
			<td>NVIDIA GeForce GTX 1080 Ti graphics cards</td>
		</tr>
		<tr>
			<td>Kumar et al diSPIM Setup</td>
			<td>Applied Scientifics Instrumentations (ASI)</td>
			<td></td>
			<td>Instrument setup for this protocol is identical to Kumar et al 10,11 &#160;diSPIM, which makes use of 40x 0.8NA water immersion lenses for imaging. (See steps 5.1 and note)</td>
		</tr>
		<tr>
			<td>Micro Manager</td>
			<td>Micro-Manager</td>
			<td></td>
			<td>https://micro-manager.org/</td>
		</tr>
		<tr>
			<td>Modifications to Kumar et al diSPIM Setup (see below)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Optical table with isolators, 4 feet &#215; 6 feet &#215; 12 inches</td>
			<td>TMC</td>
			<td>784-651-02DR and 14-416-34</td>
			<td></td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Steps 7-10</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Analysis PC</td>
			<td>Intel</td>
			<td></td>
			<td>Intel Core i7-8700K CPU 3.70GHz, 6 cores in total, 64 GB memory, Windows 10</td>
		</tr>
		<tr>
			<td>Analysis PC Graphics Card</td>
			<td>NVIDIDA</td>
			<td></td>
			<td>NVIDIA GeForce GTX 1080 Ti graphics cards</td>
		</tr>
		<tr>
			<td>Installation instructions</td>
			<td>Software bundle</td>
			<td></td>
			<td>http://dispimlineage.wormguides.org/diSPIMlineaging_InstallationInstructions.htm</td>
		</tr>
		<tr>
			<td>Software bundle</td>
			<td>Software bundle</td>
			<td></td>
			<td>http://dispimlineage.wormguides.org</td>
		</tr>
	</tbody>
</table>

isotropic light-sheet microscopy and automated cell lineage analyses to catalogue caenorhabditis elegans embryogenesis with subcellular resolution

Caenorhabditis elegans (C. elegans) stands out as the only organism in which the challenge of understanding the cellular origins of an entire nervous system can be observed, with single cell resolution, in vivo. Here, we present an integrated protocol for the examination of neurodevelopment in C. elegans embryos. Our protocol combines imaging, lineaging and neuroanatomical tracing of single cells in developing embryos. We achieve long-term, four-dimensional (4D) imaging of living C. elegans&#160;embryos with nearly isotropic spatial resolution through the use of Dual-view Inverted Selective Plane Illumination Microscopy (diSPIM). Nuclei and neuronal structures in the nematode embryos are imaged and isotropically fused to yield images with resolution of ~330 nm in all three dimensions. These minute-by-minute high-resolution 4D data sets are then analyzed to correlate definitive cell-lineage identities with gene expression and morphological dynamics at single-cell and subcellular levels of detail. Our protocol is structured to enable modular implementation of each of the described steps and enhance studies on embryogenesis, gene expression, or neurodevelopment.

We first validated the viability of embryos imaged using the protocol&#39;s parameters for diSPIM acquisition (sections 1-6). Ten embryos were simultaneously imaged at 20 &#176;C, one volume/embryo/minute, from the 2-cell stage to the 2-fold stage (7.5 hours, 451 volumes/embryo). To monitor cell divisions throughout embryogenesis, we used strain BV514, which ubiquitously expresses the mCherry::Histone reporter constructs from the integrated transgene array ujIs11324. Figure 6 shows a timeline of this first half of embryonic development for one of the imaged embryos. Each image represents a single-view maximum-intensity projection (produced by steps 7-8) of the imaged embryo. We found that the optimized protocols did not induce any detectable phototoxicity to the embryos, as assessed by timing of cell divisions (not shown), time of hatching, and timing related to developmental milestones (Figure 6 and references1,25,26).
We then applied the protocol to analyze outgrowth dynamics of single neurons in developing embryos. We imaged DCR7692 (olaex4655), a transgenic nematode strain that expresses GFP off the neuropeptide flp-19 promoter in a subset of unidentified cells (DACR2819, Pflp-19 (3.6kb)::Syn21::GFP::CAAX::p10 3&#8217;UTR). Following the steps of the protocol outlined here, we determined that the unidentified cells correspond to motor neurons RMDDL and RMDDR, to the excretory canal cell, and to two muscle cells (Figure 7). We then examined and quantified the outgrowth dynamics of the RMDDL and RMDDR neurons. We observed that the RMDDL and RMDDR neurons are obliquely shaped as early as 360 minutes post fertilization, with the longer cellular axis representing the subsequent axis for neurite outgrowth (Figure 7 and Movie S1). &#173;&#173;&#173;Using the &#8220;simple neurite tracing&#8221; plugin in FIJI and applying it to 3D reconstructions of isotropically fused volumes, we then quantified the stereotypic outgrowth of the RMDDL and RMDDR neurites for six embryos. We determined that the outgrowth dynamics were stereotyped for RMDDL and RMDDR across embryos (herein called RMDDs). From 385-410 minutes post fertilization, the RMDDs neurites extended 6.0&#177;0.5&#181;m (mean &#177; SEM; n = 12 neurites) anterior of the cell bodies (Figure 7B,C,I). From 415-445 minutes post fertilization, both neurites extend dorsally into and around the presumptive nerve ring (asterisk in Figure 7D). On average, each RMDD neurite extended 11.0 &#177; 0.6 &#181;m (mean &#177; SEM; n = 12 neurites) from the cell body before synchronously meeting its contralateral counterpart at the apex of the ring (Figure 7I). Importantly, our representative results demonstrate that we are able to examine, compare and quantify neuronal developmental features for single identifiable cells by using our integrated protocol (Figure 7 and Figure 8).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59533/59533fig1.jpg" /> 
Figure 1: diSPIM sample mounting setup. (A) Preparation of coverslip with Poly-L-lysine. In the inset, 10 &#181;L of Poly-L lysine was used to coat the coverslip for 5 minutes. Poly-L-lysine allows the embryo eggshell to stick firmly to the coverslip in the rectangle. (B) Schematic of the steel imaging chamber and the assembled chamber. In the inset, representative embryos are oriented with the anterior-posterior axis perpendicular to the long axis on the coverslip. (C) Assembled aspirator tube with microcapillary pipette. (D) Steel imaging chamber mounted in sample holder under diSPIM 40x objectives. <a href="https://www.jove.com/files/ftp_upload/59533/59533fig1large.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/59533/59533fig2.jpg" /> 
Figure 2: Long-term diSPIM imaging setup in Micro-Manager. (A) Recommended diSPIM laser power parameters (red rectangle) optimized for prolonged imaging while reducing phototoxicity (as assessed by higher hatching rate of C. elegans embryos). Set 561 nm laser to 79 &#181;W (0.25) and 488 nm laser to 179 &#181;W (0.5). Note that exact calibration of software settings to laser power varies among diSPIM installations. It is recommended that users measure and calibrate the laser power in order to achieve 79 &#181;W (561 nm) and 179 &#181;W (488 nm) laser power. (B) diSPIM parameters for data saving (green rectangle), volume settings (blue square), and slice settings (orange square). (C) diSPIM autofocus parameters for long-term imaging of C. elegans embryogenesis (see steps 6.1-6.6). <a href="https://www.jove.com/files/ftp_upload/59533/59533fig2large.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/59533/59533fig3.jpg" /> 
Figure 3: Image visualization and data-processing setup using CytoSHOW. (A) Raw diSPIM images opened by CytoSHOW. CytoSHOW is able to open images captured by both camera paths (SPIM A and B). These raw images are opened in multidimensional windows called diSPIM monitor. In diSPIM monitor, a &#8220;bowtie pattern&#8221; is generated to select the embryo&#8217;s anterior, posterior, dorsal and ventral edges (see step 9.1). Bow-tie selections indicate embryo orientation for deconvolution and StarryNite-assisted lineaging tracing. (B) Optimized parameters used to generate isotropic images. In the Deconvolved while acquiring window, set the parameters specified in steps 9.5.1-9.5.8. <a href="https://www.jove.com/files/ftp_upload/59533/59533fig3large.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/59533/59533fig4.jpg" /> 
Figure 4: diSPIM camera configuration. (A) Photograph of diSPIM camera placements and orientations. (B) Depiction of +90-degree rotations of SPIM A to match SPIM B images collected. (C) Input volumes relative to orientation index +1 based on our diSPIM&#8217;s camera configuration (see step 9.5.2). We rotate SPIM A image(s) +90 degrees around Y-Axis before registration to match SPIM B image(s). Scale bars = 10 &#181;m. Images are representative single-view, maximum-intensity projections and deconvolution images of 1.5-fold embryo with labeled nuclei (561-nm, red) and neurons (488-nm, green). <a href="https://www.jove.com/files/ftp_upload/59533/59533fig4large.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/59533/59533fig5.jpg" /> 
Figure 5: Curation and editing C. elegans embryonic lineage in AceTree. (A) We use AceTree to edit StarryNite&#8217;s lineage traces (see references5,6,8; manuscripts are also included in our download bundle). AceTree will display C. elegans systematic names for each nucleus (green rectangle) upon completion of steps 10.1-10.2. This window (A) provides information (black rectangle) about each cell in the lineage trace (ABa, highlighted in blue) that help guide users when tracking and editing the lineage traces. It is recommended that users verify and compare lineaged cells and their positions to the C. elegans embryonic cell lineage previously reported by Sulston et al.1 In addition, if users are interested in locating specific cells in the deconvolved data series (see below, B) enter the C. elegans systematic name in the search bar (orange rectangle). (B) The user&#8217;s deconvolved data series also opens automatically upon completion of steps 10.1-10.2. Shown here an isotropically fused image of a four-cell stage embryo with nuclei labeled in red. During tracking of a nuclei, users should change the intensity of the image (red square) and navigate through time and z using the arrow keys on their keyboard (time-left/right, z-up/down). (C) 3D cartoon of the timepoint in (B) with certain functionalities (purple rectangle) that enables rotatable 3D-visualization. For an overview of AceTree and its 3D functionality, see references5,6,8. <a href="https://www.jove.com/files/ftp_upload/59533/59533fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/59533/59533fig6.jpg" /> 
Figure 6: Timed developmental dynamics of C. elegans embryos on the diSPIM. Top panel, diSPIM images showing the first half of embryonic development for one of the imaged embryos (Strain BV514 ujIs11324). Embryos were imaged continuously, every minute for 7.5 hours (at 20 &#176;C). The first two images of top panel represent 4- and 8-cell embryos with nuclei (red) and positions of polar bodies (dense red spheres, marked with blue asterisks). Each image represents a single-view maximum-intensity projection of the imaged embryo. Scale bars = 10 &#181;m. The timeline (horizontal bar) represents minutes post fertilization (m.p.f.) of the development of C. elegans embryos. We validated that our protocol&#39;s parameters for diSPIM acquisition did not induce any detectable phototoxicity to the imaged embryos as assessed by viability, timing of cell divisions, timing of hatching and timing of developmental milestones (see references1,25,26). We note that the timing of developmental milestones was reproducible across embryos with our imaging parameters (SEM &#177; 8.174 minutes for 6.4 hour long imaging sessions; n = 10). <a href="https://www.jove.com/files/ftp_upload/59533/59533fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/59533/59533fig7_v2.jpg" /> 
Figure 7. Cellular identification and single cell characterization of neurite outgrowth dynamics in developing C. elegans embryos. Dual-color imaging of a strain made by crossing BV514 ujIs11324 (for lineaging) and DCR7692 (olaex4655), a transgenic nematode strain that expresses GFP off the neuropeptide flp-19 promoter in a subset of unidentified cells. (A-H) Following the steps of the protocol outlined here, we determined that the unidentified cells correspond to motor neurons RMDDL and RMDDR (yellow arrows), to the excretory canal cell (blue arrows), and to two muscle cells (white arrows). (I) Quantification of the outgrowth dynamics of the RMDDL and RMDDR neurons by using FIJI plugin &#8220;simple neurite tracing&#8221; and applying it to 3D reconstructions of isotropically fused volumes. Note how both RMDDL and RMDDR show stereotypic outgrowth dynamics, each extending synchronously for a total length of 11.0&#177;0.6&#181;m (mean &#177; SEM; n = 12 neurites) and meeting at the dorsal apex of nerve ring (see also Movie S1). <a href="https://www.jove.com/files/ftp_upload/59533/59533fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/59533/59533fig8.jpg" /> 
Figure 8: Examination of Isotropic diSPIM images of neuronal morphologies in C. elegans embryos. Isotropic visualization of AVHL and AVHR neurons (yellow arrows). Using the diSPIM, neuronal morphologies can be captured yielding four-dimensional (4D) images with isotropic spatial resolution of approximately 330 nm. The diSPIM allows users to virtually rotate image volumes with identical resolution in all direction. Images in A-D are maximum-intensity projections of the same isotropically fused diSPIM image volume from distinct rotations around the embryo&#39;s long axis. Scale bars = 5 &#181;m. <a href="https://www.jove.com/files/ftp_upload/59533/59533fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Movie S1: C. elegans embryo developing from 280 to 434 minutes post fertilization. Isotropic movie of strain DCR7692 (olaex4655) expressing ujIs113 ubiquitously with DACR2819 sparsely labeling RMDD neurites (Figure 7A-D, yellow arrows). DACR2819 also labels two muscle cells (Figure 7A-D, white arrows) and excretory canal cell (Figure 7A-D, blue arrow) during embryonic development (Figure 7A-D). Scale bars = 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/59533/Movie_S1.mov">Please click here to download this file.</a>

Watch this Scientific Journal Video about Isotropic Light-Sheet Microscopy and Automated Cell Lineage Analyses to Catalogue Caenorhabditis elegans Embryogenesis with Subcellular Resolution at JoVE.com

Isotropic Light-Sheet Microscopy and Automated Cell Lineage Analyses to Catalogue Caenorhabditis elegans Embryogenesis with Subcellular Resolution

Here, we present a combinatorial approach using high-resolution microscopy, computational tools, and single-cell labeling in living C. elegans embryos to understand single cell dynamics during neurodevelopment.

Caenorhabditis elegans (C. elegans) stands out as the only organism in which the challenge of understanding the cellular origins of an entire nervous ...

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Developmental Biology

8.1K Views.  Yale University School of Medicine. Here, we present a combinatorial approach using high-resolution microscopy, computational tools, and single-cell labeling in living C. elegans embryos to understand single cell dynamics during neurodevelopment.

Video: Isotropic Light-Sheet Microscopy and Automated Cell Lineage Analyses to Catalogue Caenorhabditis elegans Embryogenesis with Subcellular Resolution

Comprehensive Assessment of Germline Chemical Toxicity Using the Nematode Caenorhabditis elegans

Identifying the reproductive toxicity of the thousands of chemicals present in our environment has been one of the most tantalizing challenges in the field of environmental health. This is due in part to the paucity of model systems that can (1) accurately recapitulate keys features of reproductive processes and (2) do so in a medium- to high-throughput fashion, without the need for a high number of vertebrate animals.
We describe here an assay in the nematode C. elegans that allows the rapid identification of germline toxicants by monitoring the induction of aneuploid embryos. By making use of a GFP reporter line, errors in chromosome segregation resulting from germline disruption are easily visualized and quantified by automated fluorescence microscopy. Thus the screening of a particular set of compounds for its toxicity can be performed in a 96- to 384-well plate format in a matter of days. Secondary analysis of positive hits can be performed to determine whether the chromosome abnormalities originated from meiotic disruption or from early embryonic chromosome segregation errors. Altogether, this assay represents a fast first-pass strategy for the rapid assessment of germline dysfunction following chemical exposure.

Comprehensive Assessment of Germline Chemical Toxicity Using the Nematode Caenorhabditis elegans

We describe the detailed steps of a high-throughput chemical assay in the nematode Caenorhabditis elegans used to assess germline toxicity. In this assay, disruption of germline function following chemical exposure is monitored using a fluorescent reporter specific to aneuploid embryos.

Identifying the reproductive toxicity of the thousands of chemicals present in our environment has been one of the most tantalizing challenges in the ...

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 ...

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 ...

Cell Lineage Analyses and Gene Function Studies Using Twin-spot MARCM

Mosaic analysis with a repressible cell marker (MARCM) is a positive mosaic labeling system that has been widely applied in Drosophila neurobiological studies to depict intricate morphologies and to manipulate the function of genes in subsets of neurons within otherwise unmarked and unperturbed organisms. Genetic mosaics generated in the MARCM system are mediated through site-specific recombination between homologous chromosomes within dividing precursor cells to produce both marked (MARCM clones) and unmarked daughter cells during mitosis. An extension of the MARCM method, called twin-spot MARCM (tsMARCM), labels both of the twin cells derived from a common progenitor with two distinct colors. This technique was developed to enable the retrieval of useful information from both hemi-lineages. By comprehensively analyzing different pairs of tsMARCM clones, the tsMARCM system permits high-resolution neural lineage mapping to reveal the exact birth-order of the labeled neurons produced from common progenitor cells. Furthermore, the tsMARCM system also extends gene function studies by permitting the phenotypic analysis of identical neurons of different animals. Here, we describe how to apply the tsMARCM system to facilitate studies of neural development in Drosophila.

Here, we present a protocol for a mosaic labeling technique that permits the visualization of neurons derived from a common progenitor cell in two distinct colors. This facilitates neural lineage analysis with the capability of birth-dating individual neurons and studying gene function in the same neurons of different individuals.

Mosaic analysis with a repressible cell marker (MARCM) is a positive mosaic labeling system that has been widely applied in Drosophila neurobiological ...

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. ...

High-resolution Episcopic Microscopy (HREM) - Simple and Robust Protocols for Processing and Visualizing Organic Materials

We provide simple protocols for generating digital volume data with the high-resolution episcopic microscopy (HREM) method. HREM is capable of imaging organic materials with volumes up to 5 x 5 x 7 mm3 in typical numeric resolutions between 1 x 1 x 1 and 5 x 5 x 5 &#181;m3. Specimens are embedded in methacrylate resin and sectioned on a microtome. After each section an image of the block surface is captured with a digital video camera that sits on the phototube connected to the compound microscope head. The optical axis passes through a green fluorescent protein (GFP) filter cube and is aligned with a position, at which the bock holder arm comes to rest after each section. In this way, a series of inherently aligned digital images, displaying subsequent block surfaces are produced. Loading such an image series in three-dimensional (3D) visualization software facilitates the immediate conversion to digital volume data, which permit virtual sectioning in various orthogonal and oblique planes and the creation of volume and surface rendered computer models. We present three simple, tissue specific protocols for processing various groups of organic specimens, including mouse, chick, quail, frog and zebra fish embryos, human biopsy material, uncoated paper and skin replacement material.

High-resolution Episcopic Microscopy HREM - Simple and Robust Protocols for Processing and Visualizing Organic Materials

We provide simple and robust protocols for processing biopsy material of various species, embryos of biomedical model organisms and samples of other organic tissues in order to permit digital volume data generation with the high-resolution episcopic microscopy method.

We provide simple protocols for generating digital volume data with the high-resolution episcopic microscopy (HREM) method. HREM is capable of imaging ...

Correlative Super-resolution and Electron Microscopy to Resolve Protein Localization in Zebrafish Retina

We present a method to investigate the subcellular protein localization in the larval zebrafish retina by combining super-resolution light microscopy and scanning electron microscopy. The sub-diffraction limit resolution capabilities of super-resolution light microscopes allow improving the accuracy of the correlated data. Briefly, 110 nanometer thick cryo-sections are transferred to a silicon wafer and, after immunofluorescence staining, are imaged by super-resolution light microscopy. Subsequently, the sections are preserved in methylcellulose and platinum shadowed prior to imaging in a scanning electron microscope (SEM). The images from these two microscopy modalities are easily merged using tissue landmarks with open source software. Here we describe the adapted method for the larval zebrafish retina. However, this method is also applicable to other types of tissues and organisms. We demonstrate that the complementary information obtained by this correlation is able to resolve the expression of mitochondrial proteins in relation with the membranes and cristae of mitochondria as well as to other compartments of the cell.

This protocol describes the necessary steps to obtain subcellular protein localization results on zebrafish retina by correlating super-resolution light microscopy and scanning electron microscopy images.

We present a method to investigate the subcellular protein localization in the larval zebrafish retina by combining super-resolution light microscopy and ...

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

Axonal transport and intraflagellar transport (IFT) are essential for axon and cilia morphogenesis and function. Kinesin superfamily proteins and dynein are molecular motors that regulate anterograde and retrograde transport, respectively. These motors use microtubule networks as rails. Caenorhabditis elegans (C. elegans) is a powerful model organism to study axonal transport and IFT in vivo. Here, I describe a protocol to observe axonal transport and IFT in living C. elegans. Transported cargo can be visualized by tagging cargo proteins using fluorescent proteins such as green fluorescent protein (GFP). C. elegans is transparent and GFP-tagged cargo proteins can be expressed in specific cells under cell-specific promoters. Living worms can be fixed by microbeads on 10% agarose gel without killing or anesthetizing the worms. Under these conditions, cargo movement can be directly observed in the axons and cilia of living C. elegans without dissection. This method can be applied to the observation of any cargo molecule in any cells by modifying the target proteins and/or the cells they are expressed in. Most basic proteins such as molecular motors and adaptor proteins that are involved in axonal transport and IFT are conserved in C. elegans. Compared to other model organisms, mutants can be obtained and maintained more easily in C. elegans. Combining this method with various C. elegans mutants can clarify the molecular mechanisms of axonal transport and IFT.

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

Caenorhabditis elegans (C. elegans) is a good model to study axonal and intracellular transport. Here, I describe a protocol for in vivo recording and analysis of axonal and intraflagellar transport in C. elegans.

Axonal transport and intraflagellar transport (IFT) are essential for axon and cilia morphogenesis and function. Kinesin superfamily proteins and dynein ...

Analysis of Cardiac Chamber Development During Mouse Embryogenesis Using Whole Mount Epifluorescence

The goal of this protocol is to describe a method for the dissection of mouse embryos and visualization of embryonic mouse ventricular chambers during heart development using ventricular specific fluorescent reporter knock-in mice (MLC-2v-tdTomato mice). Heart development involves a linear heart tube formation, the heart tube looping, and four chamber septation. These complex processes are highly conserved in all vertebrates. The mouse embryonic heart has been widely used for heart developmental studies. However, due to their extremely small size, dissecting mouse embryonic hearts is technically challenging. In addition, visualization of cardiac chamber formation often needs in situ hybridization, beta-galactosidase staining using LacZ reporter mice, or immunostaining of sectioned embryonic hearts. Here, we describe how to dissect mouse embryonic hearts and directly visualize ventricular chamber formation of MLC-2v-tdTomato mice using whole mount epifluorescent microscopy. With this method, it is possible to directly examine heart tube formation and looping, and four chamber formation without further experimental manipulation of mouse embryos. Although the MLC-2v-tdTomato reporter knock-in mouse line is used in this protocol as an example, this protocol can be applied to other heart-specific fluorescent reporter transgenic mouse lines.

We present the protocols to examine mouse heart development using whole mount epifluorescent microscopy on mouse embryos dissected from ventricular specific MLC-2v-tdTomato reporter knock-in mice. This method allows us to directly visualize each stage of the ventricular formation during mouse heart development without labor-intensive histochemical methods.

The goal of this protocol is to describe a method for the dissection of mouse embryos and visualization of embryonic mouse ventricular chambers during ...

Measuring Sperm Guidance and Motility within the Caenorhabditis elegans Hermaphrodite Reproductive Tract

Successful fertilization is fundamental to sexual reproduction, yet little is known about the mechanisms that guide sperm to oocytes within the female reproductive tract. While in vitro studies suggest that sperm of internally fertilizing animals can respond to various cues from their surroundings, the inability to visualize their behavior inside the female reproductive tract creates a challenge for understanding sperm migration and mobility in its native environment. Here, we describe a method using C. elegans that overcomes this limitation and takes advantage of their transparent epidermis. C. elegans males stained with a mitochondrial&#160; dye are mated with adult hermaphrodites, which act as modified females, and deposit fluorescently labeled sperm into the hermaphrodite uterus. The migration and motility of the labeled sperm can then be directly tracked using an epi-fluorescence microscope in a live hermaphrodite. In wild-type animals, approximately 90% of the labeled sperm crawl through the uterus and reach the fertilization site, or spermatheca. Images of the uterus can be taken 1 h after mating to assess the distribution of the sperm within the uterus and the percentage of sperm that have reached the spermatheca. Alternatively, time-lapse images can be taken immediately after mating to assess sperm speed, directional velocity and reversal frequency. This method can be combined with other genetic and molecular tools available for the C.elegans to identify novel genetic and molecular mechanisms that are important in regulating sperm guidance and motility within the female reproductive tract.

Measuring Sperm Guidance and Motility within the Caenorhabditis elegans Hermaphrodite Reproductive Tract

Sperm must successfully navigate through the oviduct to fertilize an oocyte. Here, we describe an assay for measuring sperm migration within the C. elegans hermaphrodite uterus. This assay can provide quantitative data on sperm distribution within the uterus after mating, as well as on speed, directional velocity, and reversal frequency.

Successful fertilization is fundamental to sexual reproduction, yet little is known about the mechanisms that guide sperm to oocytes within the female ...