The authors wish to acknowledge support from Rioja Salud Foundation (Fondos FEDER) and the Spanish Ministerio de Ciencia, Innovaci&#243;n y Universidades (MCIU) (Grant PGC2018-094276-B-I00). Cristina Romero Aranda is funded by a fellowship from the AECC (Asociaci&#243;n Espa&#241;ola Contra el C&#225;ncer).

1. Grow C. elegans on Petri dishes
<ol>
	<li>Prepare NGM plates and seed them with E. coli OP50 as the food source (Figure 1). Grow and maintain C. elegans as described17. Store seeded plates at 4 &#176;C for up to one month.</li>
	<li>Adjust the plates to the desired temperature before adding the worms.
	<ol>
		<li>To transfer the worms, remove a chunk of agar from an old plate and place it on a fresh plate.</li>
		<li>Alternatively, capture single animals with a sterile worm picker (a 1-inch piece of 32-gauge platinum wire with a flattened tip, mounted onto the tip of a Pasteur pipette) and place them on the new plate.</li>
	</ol>
	</li>
	<li>Grow the worms at the desired temperature.
	<ol>
		<li>Grow C. elegans worms at 20 &#176;C, the standard temperature. However, to analyze the development of thermo-sensitive mutants, perform an overnight incubation, usually at 25 &#176;C. Adjust the duration and temperature of this incubation as needed, depending on the specific mutant.</li>
	</ol>
	</li>
</ol>
2. Prepare the 4D microscopy recording before mounting the embryos (Figure 2)
<ol>
	<li>Set up the microscope and temperature controls before preparing the embryo. C. elegans embryos divide very quickly. Be prepared to begin recording immediately after mounting the embryos.</li>
	<li>Adjust the recording temperature to either 15 &#176;C, 20 &#176;C or 25 &#176;C.
	<ol>
		<li>Routinely record the embryos at 25 &#176;C. Record thermo-sensitive mutants at the restrictive temperature to show their phenotypes. Record a WT control (if done in a different preparation) at the same temperature as the mutants. 
		NOTE: WT embryos develop faster at 25 &#176;C and slower at 15 &#176;C without additional differences in cell lineage. Place microscopes in a temperature-controlled room. Additional control by cooling or heating the slide is highly desirable. This can be achieved by circulating water at a specific temperature through a metal ring around the microscope objective and condenser. The objective and the preparation are in direct contact through the immersion oil and the temperature transfer is efficient. This system allows for precise control of the recording temperature and for performing temperature shifts during embryo development.</li>
	</ol>
	</li>
	<li>Define the record parameters in the microscopy software.
	<ol>
		<li>For a standard C. elegans recording (without fluorescent scans), select: 
		z-stacks of 30 focal planes, at a distance of 1 micron each. 
		30 second intervals between the beginning of each z-stack. 
		1500 z-stacks (12.5 hours of record). 
		NOTE: Both commercial as well as open source microscope control programs can be used to define this workflow for capturing images. Now the microscope is ready to record.</li>
	</ol>
	</li>
</ol>
3. Prepare and mount the embryos
<ol>
	<li>Prepare a thin, homogeneous agar pad as the first step in obtaining a nice image (Figure 3).
	<ol>
		<li>Prepare 50 ml of a 4.5% agar solution in deionized water. Heat to boiling in the microwave and pour 0.5 &#8211; 1 ml into 3 ml glass tubes.</li>
		<li>Carefully seal the tubes with wax film to avoid desiccation. Sealed agar tubes can be stored at room temperature for up to two months.</li>
		<li>On the lab bench, have a heat block at 80 &#176;C with: 
		a test tube of pure petroleum jelly (melted), with a fine paint brush inside. 
		a test tube with distilled water containing a Pasteur pipette. 
		NOTE: This ensures that all the required materials will be hot, and the agar will not solidify in the process of making the pad.</li>
		<li>Remove the wax film from the top of one of the agar tubes, and carefully heat it over an alcohol burner to melt the agar. Exercise caution as hot agar expelled from the glass tube could cause burns.</li>
		<li>Once the agar is melted, place the tube in the heat block to keep the agar in liquid form.</li>
		<li>Alternatively, place the agar tubes into the heat block 1h before the experiment to melt them without using a burner. The melted agar should be discarded after one day.</li>
		<li>Place a microscope slide (Slide A) between two others on a piece of plastic.</li>
		<li>Take another slide (Slide B) and hold it with your fingers in one hand.</li>
		<li>With the other hand, place a small drop of melted agar in the center of Slide A using the warm Pasteur pipette.</li>
		<li>Immediately press slide B onto the agar drop to create a very thin pad between Slides A and B. Keep these slides sandwiched together until step 3.2.3.</li>
	</ol>
	</li>
	<li>Mount the embryos.
	<ol>
		<li>Collect 5-10 gravid hermaphrodites with the picker and place them in a watchmaker glass filled with water.</li>
		<li>Use a scalpel to cut open the hermaphrodite nematodes and extract early eggs (1-4 cells) from the uterus, under the stereomicroscope.</li>
		<li>Take the slide from step 3.1.10 and gently slip Slide B off to expose the agar pad on Slide A.</li>
		<li>Place an early egg in the center of the agar pad by pipetting with a capillary tube.</li>
		<li>Alternatively, pipette a drop containing a set of embryos onto the agar pad and then search for early-stage embryos. Do this step under the stereomicroscope.</li>
		<li>If necessary, move the egg by nudging it with an eyelash glued to the end of a toothpick.</li>
		<li>Remove excess water with the capillary pipette. 
		NOTE: A capillary pipette can easily be prepared by heating a Pasteur pipette over an alcohol burner and pulling it from both ends.</li>
		<li>Carefully cover the preparation with a coverslip. To avoid air bubbles, place one edge of the coverslip on the slide and gently slide a scalpel along the adjacent edge to slowly and obliquely drape the coverslip onto the preparation.</li>
		<li>Use a pipette to fill 3/4 of the space surrounding the agar pad with water. Leave 1/4 of the space with air.</li>
		<li>Seal the coverslip with petroleum jelly to avoid desiccation during long recording periods.</li>
		<li>Use the fine brush to extend a thin layer of melted petroleum jelly around the edge of the coverslip. 
		NOTE: Now the preparation is ready for recording.</li>
	</ol>
	</li>
</ol>
4. Adjust the DIC and start the 4D microscopy recording
<ol>
	<li>Place the slide on the microscope stage. Focus the embryo using the low magnification objective (5x or 10x).</li>
	<li>Change to the 100x immersion objective.</li>
	<li>Adjust the optical components of the microscope to get a Nomarski image.
	<ol>
		<li>Focus the condenser.</li>
		<li>Completely open the aperture of the condenser and close the field diaphragm (this will provide a higher numerical aperture and therefore, greater resolution).</li>
		<li>Check that both polarizers on the microscope are oriented to cause the maximum light extinction.</li>
		<li>Turn the Wollaston prism to get a nice three-dimensional image of the embryo, illuminated on one side. Turn the prism in the other direction to get the effect of having the embryo illuminated on the other side. 
		NOTE: These steps can be performed on a test sample before the recording so that only fine tuning is required on the specimen being analyzed.</li>
	</ol>
	</li>
	<li>Launch the image capture in the microscope.</li>
</ol>
5. Analyze the 4D-movie (Figure 4).
NOTE: Once the recording is complete, use cell lineage tracing software to reconstruct and analyze cell lineage.
Cell lineage tracing software is a powerful tool for performing detailed analyses of embryonic development or dynamics in cell cultures or tissue fragments. The program extracts and quantifies several data sets on the sample&#8217;s cellular dynamics that include generation of the complete cell lineage of each and every recorded cell, including cell divisions, cell cycle length, migration or apoptosis as well as its kinetics. In addition, cell differentiation can be scored by the cell&#8217;s morphological changes or by expression of specific markers. Basically, the software screen displays two windows: on the left window, the 4D movie can be played forward and backward or up and down to either the top or bottom levels so that each cell can be followed in time and space throughout the recording. On the right widow, the cell lineage is generated. Clicking on a cell nucleus in the 4D movie generates a point in the lineage window that stores the information of the cell name, fate and spatial coordinates. The cell lineage of a specific cell is generated by playing the 4D movie forward and clicking periodically on the nucleus to mark the mitosis of that specific cell over time. Repetition of this process for each of the recorded cells generates the complete cell lineage of the embryo or sample. The stored information for spatial coordinates of each cell is later used to reconstruct 3D embryo models and cell migration paths.
<ol>
	<li>Open the lineage tracing software and create a new project by going to the upper bar menu and selecting: 
	File | New project.</li>
	<li>Select the cell lineage template depending on the recording temperature: DB08 for recording at 25 &#176;C, DB10 for recording at 20 &#176;C and DB12 for recording at 15 &#176;C.</li>
	<li>Set the recording parameters in the emerging window: scan count (usually 1500), time between scans (30 seconds), level count (30) and distance between levels (1 micron).</li>
	<li>Select the image file and format.
	<ol>
		<li>Select the image directory where the images were saved.</li>
		<li>Choose whether images should be saved as single images (one image per level and time) or as multi-image z-stacks.</li>
		<li>Determine the file naming and image format. Routinely, single images are saved under the following names: 
		X0000L00C1 (for scan 0, level 0, channel 1) 
		X0000L01C1 (for scan 0, level 1, channel 1) 
		X0000L02C1 (for scan 0, level 2, channel 1) 
		... 
		X0001L00C1 (for scan 1, level 0, channel 1) 
		X0001L01C1 (for scan 1, level 1, channel 1) 
		... 
		X0300L04C1 (for scan 300, level 4, channel 1) 
		... 
		NOTE: Saving the images in a compressed format saves space on your hard disk.</li>
	</ol>
	</li>
	<li>Define the light channels: 1 for DIC optics, 2 for GFP, 3 for RFP, etc. Add those that were used in the 4D-recording. Click on &#8220;channel processing enabled&#8221; to detect them.</li>
	<li>Start tracing the cell lineage of the embryo. The screen now contains two major windows: the video window and the cell lineage window.
	<ol>
		<li>On the lineage window, select a lineage branch and use the mouse to click the cell nucleus corresponding to this cell on the video window.</li>
		<li>Follow the cell spatially and over time by playing the 4D-movie forward, backward, or up or down a level, using the cursor keys.</li>
		<li>Periodically click on the cell nucleus. This generates a point in the lineage branch and registers the spatial coordinates of the cell at this time. As a result, cell lineage progresses, and 3D reconstructions of the embryo are possible.</li>
		<li>Mark mitosis by clicking the return key. Then select one of the daughter cells and follow it as before.</li>
	</ol>
	</li>
	<li>Repeat the process (steps 5.6.1 to 5.6.4) for the rest of the embryonic cells to trace the complete cell lineage, or to follow specific cells of interest such as those undergoing apoptosis.</li>
	<li>Compare the mutant lineage with the stereotyped WT C. elegans cell lineage.</li>
</ol>

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J. C., Chan, R. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+C.+elegans+Embryos+using+an+Epifluorescent+Microscope+and+Open+Source+Software.">Imaging C. elegans Embryos using an Epifluorescent Microscope and Open Source Software.</a> Journal of Visualized Experiments. (49), e2625 (2011).</li><li>Boyd, L., Hajjar, C., O'Connell, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Time-lapse+Microscopy+of+Early+Embryogenesis+in+Caenorhabditis+elegans.">Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans.</a> Journal of Visualized Experiments. (54), e2852 (2011).</li><li>Urbach, R., Schnabel, R., Technau, G. 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G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Buccal+capsule+development+as+a+consideration+for+phylogenetic+analysis+of+Rhabditida+(Nemata).">Buccal capsule development as a consideration for phylogenetic analysis of Rhabditida (Nemata).</a> Development Genes and Evolution. 208 (9), 495-503 (1998).</li><li>Houthoofd, W., Jacobsen, K., Mertens, C., Vangestel, S., Coomans, A., Borgonie, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Embryonic+cell+lineage+of+the+marine+nematode+Pellioditis+marina.">Embryonic cell lineage of the marine nematode Pellioditis marina.</a> Developmental Biology. 258 (1), 57-69 (2003).</li><li>Hejnol, A., Schnabel, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+eutardigrade+Thulinia+stephaniae+has+an+indeterminate+development+and+the+potential+to+regulate+early+blastomere+ablations.">The eutardigrade Thulinia stephaniae has an indeterminate development and the potential to regulate early blastomere ablations.</a> Development. 132 (6), 1349-1361 (2005).</li><li>Hejnol, A., Schnabel, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=What+a+couple+of+dimensions+can+do+for+you:+Comparative+developmental+studies+using+4D+microscopy-examples+from+tardigrade+development.">What a couple of dimensions can do for you: Comparative developmental studies using 4D microscopy-examples from tardigrade development.</a> Integrative and Comparative Biology. 46 (2), 151-161 (2006).</li><li>Bischoff, M., Parfitt, D. E., Zernicka-Goetz, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Formation+of+the+embryonic-abembryonic+axis+of+the+mouse+blastocyst:+relationships+between+orientation+of+early+cleavage+divisions+and+pattern+of+symmetric/asymmetric+divisions.">Formation of the embryonic-abembryonic axis of the mouse blastocyst: relationships between orientation of early cleavage divisions and pattern of symmetric/asymmetric divisions.</a> Development. 135 (5), 953-962 (2008).</li><li>Mora-Lorca, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glutathione+reductase+gsr-1+is+an+essential+gene+required+for+Caenorhabditis+elegans+early+embryonic+development.">Glutathione reductase gsr-1 is an essential gene required for Caenorhabditis elegans early embryonic development.</a> Free Radical Biology and Medicine. 96, 446-461 (2016).</li></ol>

The authors have nothing to disclose.

One of the major challenges in modern biology is understanding the development of multicellular organisms. C. elegans has emerged as one of the best suited models for studying the fine coordination between cell proliferation and cell differentiation in the developing embryo. From an optical point of view, its transparent body and its small size make this nematode an ideal specimen for DIC microscopy. Other organisms with similar characteristics have also been subjected to 4D microscopy analysis11,12,13,14,15,16.
For those developmental studies, gene inactivation by either forward or reverse genetics provides a clue to its involvement in embryogenesis. Once a gene has been proven to play a role in development, the next step is to define its exact role in the establishment of the correct body plan. Immunostaining is the selected approach for most models. This technique elucidates problems in cell differentiation or expression of specific markers. However, a major limitation of this approach is that it only provides a static view of the expression of a single or more markers at a fixed point in development. A dynamic view of these markers throughout development can only be obtained by staining different embryos at different time points. In addition, cell lineage reconstruction is not possible in such fixed samples.
4D microscopy is a complementary approach for studying embryonic development. This technique reveals development dynamics at a cell level resolution. Any defect in the embryo such as problems in spindle orientation, cell migration, apoptosis, cell fate specification, etc. will show up in a 4D movie that can be visualized forward and backward, quantified and scored by the researcher. Using this technique, virtually each and every cell in the embryo can be followed up to the moment that the embryo begins to move. Embryos subjected to 4D microscopy with only visible light and Nomarski optics do not incur photodamage. Fluorescent scans can also be intercalated within the recording to detect when and where a gene is expressed. Embryos that suffer significant photodamage are identified by the cell cycle extension that causes strong UV irradiation compared to a standard WT lineage embryo. In that case, photodamage can be reduced by lowering the UV lamp intensity and increasing camera sensitivity or exposure time. Morphological characteristics and molecular markers can help clarify the embryonic development of any mutant.
Setting up a 4D microscopy system is easy to implement in the lab and, after some practice, enables an unmatched analysis of cell dynamics and lineage tracing of cell cultures and living transparent specimens at a resolution level of each and every cell in the microscope field. Cell lineage tracing on DIC images is still processed by hand. It is time consuming and, although the software detects lineage errors such as different lineage branches marking the same cell, mistakes are possible. While automatic detection of GFP-labeled cells is well developed2, complementary lineage tracing software based on unmarked cells and visible light images is still in the early stage and not really useful for a full embryo analysis. Without any doubt, application of image recognition systems to the field of visible light microscopy will bring about a great advance in this field.

4D microscopy is a multifocal time-lapse recording system that allows researchers to register and quantify the cell dynamics of a biological sample both spatially and over time. Cell cultures, yeasts or living tissues can be subjected to 4D analysis but this technique is especially suited for analyzing the development of living embryos. The resolution of this analysis reaches the level of every single cell of the embryo. Each cell division can be detected, and cell movements can be traced over time. Cell fates are assessed according to the position and shape that cells acquire. The use of Nomarski optics enhances the contrast of unstained transparent samples using orthogonally polarized light beams that interfere at the focal plane. The resulting images appear three-dimensional, illuminated on one side.
Other methods based on the use of confocal microscopy and GFP transgenic animals for automatic detection of nuclei and generation of cell lineages have been developed1,2. The advantage of those systems is obvious: the software greatly overrides the need for manually marking each nucleus over a period of time (although some manual supervision is required during the late stages). However, cellular processes involving changes in cell shape or membrane dynamics, such as those occurring during cell differentiation, migration, apoptosis or corpse engulfment, remain hidden as a black background in the fluorescent-labeled nuclei images.
In contrast, 4D Nomarski microscopy (also called DIC microscopy, Differential Interference Contrast microscopy) shows both nuclei and cell shape changes that occur during the development of wild type or mutant animals. This allows cell lineage tracing using standard microscopes, employing only transmitted light. There is no general need to use transgenic animals except to show specific expression patterns, in which case fluorescent scans can be intercalated. Therefore, this could be the optimal approach for many labs working on dynamic cell processes such as embryogenesis or apoptosis that can be highlighted under DIC microscopy3,4,5,6,7.
Several flexible and user-friendly programs are available for capturing microscopic images and reconstructing cell lineages, 3D models, cell migration paths, etc. in the recorded sample. In a standard experiment, images are acquired in a series of focal planes, at a constant distance, the number of which depends on the sample thickness. Temporal resolution of the analysis can be optimized by increasing scan frequency. There is virtually no limit for the duration of the recording other than computer storage capacity. For example, for a C. elegans embryo development analysis, we routinely acquire images on 30 focal planes (1 micron-step each), every 30 seconds for 12 hours.
These systems have been applied to the analysis of several animal embryos such as Caenorhabditis elegans8,9,10, Drosophila melanogaster11, other nematode embryos12,13, tardigrades14,15 and even early mouse embryos16. The only requirement is having a transparent embryo able to develop on the slide preparation under the microscope.
In summary, DIC based 4D microscopy is especially useful for 1) analyzing embryonic development of small, transparent animals: tracing cell lineage, cell migration paths, generating 3D models, etc; 2) defining gene expression patterns; 3) studying cell culture dynamics, from yeast to human cells; 4) analyzing tissue dynamics or embryo fragments; 5) quantifying cell death kinetics and corpse engulfment; and 6) performing comparative phylogeny analysis based on embryonic developmental characteristics. If there is interest in any of these topics (or similar ones), 4D microscopy can be used.

<table>
	<tbody>
		<tr>
			<td>Caenorhabditis elegans (N2)</td>
			<td>GCG (Caenorhabditis Genetics Center)</td>
			<td>N2</td>
			<td>WT C. elegans strain. Can be requested at GCG (Caenorhabditis Genetics Center): https://cgc.umn.edu/</td>
		</tr>
		<tr>
			<td>Caenorhabditis elegans (VZ454)</td>
			<td>GCG (Caenorhabditis Genetics Center)</td>
			<td>VZ454</td>
			<td>gsr-1(tm3574) C. elegans mutant strain. Can be requested at GCG (Caenorhabditis Genetics Center): https://cgc.umn.edu/</td>
		</tr>
		<tr>
			<td>Cell Lineage Tracing software</td>
			<td>SIMI</td>
			<td>Simi BioCell</td>
			<td>This is the software to reconstruct the embryo cell lineage. For a detailed explanation check at: http://www.simi.com/en/products/cell-research/simi-biocell.html</td>
		</tr>
		<tr>
			<td>Microscope camera</td>
			<td>Hamamatsu</td>
			<td>Orca-R2</td>
			<td>Miscroscope camera for both transmitted and UV light</td>
		</tr>
		<tr>
			<td>Microscope control software</td>
			<td>Caenotec</td>
			<td>Time to Live</td>
			<td>This software controls the microscope to perform the 4D image capture. Can be requested at: Caenotec Prof. Ralf Schnabel 
			Kleine Dorfstr. 9 38312 B&#246;r&#223;um, Germany, Ph: ++49 151 11653356 r.schnabel(at)tu-bs.de</td>
		</tr>
		<tr>
			<td>Microscope control software</td>
			<td>Micro-manager</td>
			<td>Micro-manager</td>
			<td>This software controls the microscope to perform the 4D image capture. Can be downloaded at: https://micro-manager.org/</td>
		</tr>
		<tr>
			<td>Motorized microscope</td>
			<td>Leica</td>
			<td>Leica DM6000</td>
			<td>Motorized upright microscope to perform 4D microscopy</td>
		</tr>
		<tr>
			<td>Standard equipment in a Molecular Biology lab.</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>Leica</td>
			<td>MZ16FA</td>
			<td>Steromicroscope to manipulate nematodes and prepare embryos.</td>
		</tr>
	</tbody>
</table>

4d microscopy: unraveling caenorhabditis elegans embryonic development using nomarski microscopy

4D microscopy is an invaluable tool for unraveling the embryonic developmental process in different animals. Over the last decades, Caenorhabditis elegans has emerged as one of the best models for studying development. From an optical point of view, its size and transparent body make this nematode an ideal specimen for DIC (Differential Interference Contrast or Nomarski) microscopy. This article illustrates a protocol for growing C. elegans nematodes, preparing and mounting their embryos, performing 4D microscopy and cell lineage tracing. The method is based on multifocal time-lapse records of Nomarski images and analysis with specific software. This technique reveals embryonic developmental dynamics at the cellular level. Any embryonic defect in mutants, such as problems in spindle orientation, cell migration, apoptosis or cell fate specification, can be efficiently detected and scored. Virtually every single cell of the embryo can be followed up to the moment the embryo begins to move. Tracing the complete cell lineage of a C. elegans embryo by 4D DIC microscopy is laborious, but the use of specific software greatly facilitates this task. In addition, this technique is easy to implement in the lab. 4D microscopy is a versatile tool and opens the possibility of performing an unparalleled analysis of embryonic development.

To characterize embryonic development of a C. elegans mutant for the gene gsr-1, that encodes the enzyme glutathione reductase, required to regenerate reduced glutathione (GSH) and involved in maintaining redox homeostasis in the nematode, we performed 4D microscopy of a gsr-1 (tm3574) deletion mutant that is a loss of function allele causing an early embryonic arrest phenotype18. Both WT and balanced gsr-1 (tm3574) mutant C. elegans nematodes were grown on NGM plates seeded with E. coli OP50 as the food source17. gsr-1 (tm3574) worms were grown as heterozygous at 20 &#176;C for two generations and then segregating homozygous worms (which are able to grow up to adulthood thanks to the maternal load) were shifted to 25 &#176;C for an overnight incubation prior to embryo analysis. Worm plates were incubated within cardboard boxes to avoid condensation (Figure 1). Gravid nematodes were cut open to extract young embryos.
To compare embryonic development of the mutant versus the stereotyped WT under identical conditions, a WT (as control) and a gsr-1 (tm3574) embryo were placed on the same preparation next to each other. 4D microscopy workflow was run on a standard motorized upright microscope outfitted with DIC optics. The selected recording parameters on the microscope control program were: z-stacks of 30 focal planes at 1 micron distance each, 30 second intervals between the beginning of each z-stack and 1500 z-stacks (12.5 hours of recording). The recording temperature was adjusted to 25 &#176;C (both in the room and on the microscope stage) (Figure 2).
Once the recording was completed, the images file was opened, and cell lineage was reconstructed using lineage tracing software by clicking on cell nuclei shown in the video window (Figure 4). The traced gsr-1 (tm3574) mutant embryonic cell lineage was compared with the C. elegans WT lineage depicted in the background. A major result was the detection of a progressive delay of the cell cycle during embryonic development. As a consequence, mutant embryos arrested at intermediate stages whereas WT embryos progressed and finally hatched as larvae.
Preparation and direct observation of embryos under the microscope or immunostaining with antibodies against late embryonic markers could reveal the presence of a high percentage of young embryos in the mutant compared to the WT. Embryo arrest could then be inferred as the most plausible explanation. However, direct proof and exact quantification of the cell cycle delay can only be elegantly and easily shown and quantified through a 4D microscopy experiment. Other important features of embryonic development such as cell differentiation or apoptosis (Figure 5) can also be visualized in a dynamic way using 4D microscopy which offers a detailed analysis of multiple aspects of development in a single experiment.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61736/61736fig01.jpg" /> 
Figure 1: C. elegans nematodes growing under laboratory conditions.&#160;Nematodes are grown on E. coli-seeded NGM plates, stored in cardboard boxes and incubated either at 15 &#176;C, 20 &#176;C or 25 &#176;C. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61736/61736fig01large.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/61736/61736fig02.jpg" /> 
Figure 2: Screenshot of 4D microscopy recording software.&#160;Example of two different microscope control software programs (A and B). These programs create workflows to control the microscope and image capturing during 4D microscopy recording. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61736/61736fig02large.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/61736/61736fig03.jpg" /> 
Figure 3: Serial photographs of agar pad preparation and mounting of the C. elegans embryo showing.&#160;A. Prepared agar tubes. B-C. Preparation of the agar pad. D. Slide partially filled with water. E. Sealing the slide with petroleum jelly. F. Final preparation. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61736/61736fig03large.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/61736/61736fig04.jpg" /> 
Figure 4: Serial screen shots of cell lineage tracer software.&#160;The program allows reconstruction of the embryonic cell lineage of a cell cycle delay mutant (left) and a WT (right) C. elegans embryo. A. An early step of the development. B-C. Development of both embryos progresses over time. D. WT embryo develops properly and starts elongation whereas the mutant arrests. In all cases the program displays the video window and the lineage window. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61736/61736fig04large.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/61736/61736fig05v2.jpg" /> 
Figure 5: Lentil refractile shape of apoptotic cells in a C. elegans WT embryo.&#160;Cell fate, defined by morphological characteristics, can be assessed by 4D microscopy. The image shows a C. elegans embryo in the bean stage. Living cells show smooth-shaped nuclei surrounded by a granular cytoplasm. In contrast, apoptotic cells (yellow arrows) condense and adopt a lentil-like, refractile shape. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61736/61736fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about 4D Microscopy: Unraveling Caenorhabditis elegans Embryonic Development Using Nomarski Microscopy at JoVE.com

4D Microscopy: Unraveling Caenorhabditis elegans Embryonic Development Using Nomarski Microscopy

Here, we present a protocol for preparing and mounting Caenorhabditis elegans embryos, recording development under a 4D microscope and tracing cell lineage.

4D Microscopy: Unraveling Caenorhabditis elegans Embryonic Development Using Nomarski Microscopy

4D microscopy is an invaluable tool for unraveling the embryonic developmental process in different animals. Over the last decades, Caenorhabditis elegans ...

4d-microscopy-unraveling-caenorhabditis-elegans-embryonic-development

Research

JoVE Journal

Developmental Biology

2.5K Views.  CIBIR (Center for Biomedical Research of La Rioja), La Rioja, Spain. Here, we present a protocol for preparing and mounting Caenorhabditis elegans embryos, recording development under a 4D microscope and tracing cell lineage.

Video: 4D Microscopy: Unraveling Caenorhabditis elegans Embryonic Development Using Nomarski Microscopy

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 ...

Analysis of Cardiomyocyte Development using Immunofluorescence in Embryonic Mouse Heart

During heart development, the generation of myocardial-specific structural and functional units including sarcomeres, contractile myofibrils, intercalated discs, and costameres requires the coordinated assembly of multiple components in time and space. Disruption in assembly of these components leads to developmental heart defects. Immunofluorescent staining techniques are used commonly in cultured cardiomyocytes to probe myofibril maturation, but this ex vivo approach is limited by the extent to which myocytes will fully differentiate in culture, lack of normal in vivo mechanical inputs, and absence of endocardial cues. Application of immunofluorescence techniques to the study of developing mouse heart is desirable but more technically challenging, and methods often lack sufficient sensitivity and resolution to visualize sarcomeres in the early stages of heart development. Here, we describe a robust and reproducible method to co-immunostain multiple proteins or to co-visualize a fluorescent protein with immunofluorescent staining in the embryonic mouse heart and use this method to analyze developing myofibrils, intercalated discs, and costameres. This method can be further applied to assess cardiomyocyte structural changes caused by mutations that lead to developmental heart defects.

Mutations that lead to congenital heart defects benefit from in vivo investigation of cardiac structure during development, but high-resolution structural studies in the mouse embryonic heart are technically challenging. Here we present a robust immunofluorescence and image analysis method to assess cardiomyocyte-specific structures in the developing mouse heart.

During heart development, the generation of myocardial-specific structural and functional units including sarcomeres, contractile myofibrils, intercalated ...

Transcriptome Profiling of In-Vivo Produced Bovine Pre-implantation Embryos Using Two-color Microarray Platform

Early embryonic loss is a large contributor to infertility in cattle. Moreover, bovine becomes an interesting model to study human preimplantation embryo development due to their similar developmental process. Although genetic factors are known to affect early embryonic development, the discovery of such factors has been a serious challenge. Microarray technology allows quantitative measurement and gene expression profiling of transcript levels on a genome-wide basis. One of the main decisions that have to be made when planning a microarray experiment is whether to use a one- or two-color approach. Two-color design increases technical replication, minimizes variability, improves sensitivity and accuracy as well as allows having loop designs, defining the common reference samples. Although microarray is a powerful biological tool, there are potential pitfalls that can attenuate its power. Hence, in this technical paper we demonstrate an optimized protocol for RNA extraction, amplification, labeling, hybridization of the labeled amplified RNA to the array, array scanning and data analysis using the two-color analysis strategy.

Transcriptome Profiling of In-Vivo Produced Bovine Pre-implantation Embryos Using Two-color Microarray Platform

Microarray technology allows quantitative measurement and gene expression profiling of transcripts on a genome-wide basis. Therefore, this protocol provides an optimized technical procedure in a two-color custom made bovine array using Day 7 bovine embryos to demonstrate the feasibility of using low amount of total RNA.

Early embryonic loss is a large contributor to infertility in cattle. Moreover, bovine becomes an interesting model to study human preimplantation embryo ...

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method described here allows time-lapse analysis of normal and impaired kidney development in zebrafish embryos by using a fluorescence dissecting microscope equipped for structured illumination and z-stack acquisition. To visualize nephrogenesis, transgenic zebrafish (Tg(wt1b:GFP)) with fluorescently labeled kidney structures were used. Renal defects were triggered by injection of an antisense morpholino oligonucleotide against the Wilms tumor gene wt1a, a factor known to be crucial for kidney development.
The advantage of the experimental setup is the combination of a zoom microscope with simple strategies for re-adjusting movements in x, y or z direction without additional equipment. To circumvent focal drift that is induced by temperature variations and mechanical vibrations, an autofocus strategy was applied instead of utilizing a usually required environmental chamber. In order to re-adjust the positional changes due to a xy-drift, imaging chambers with imprinted relocation grids were employed.
In comparison to more complex setups for time-lapse recording with optical sectioning such as confocal laser scanning&#160;or light sheet microscopes, a zoom microscope is easy to handle. Besides, it offers dissecting microscope-specific benefits such as high depth of field and an extended working distance.
The method to study organogenesis presented here can also be used with fluorescence stereo microscopes not capable of optical sectioning. Although limited for high-throughput, this technique offers an alternative to more complex equipment that is normally used for time-lapse recording of developing tissues and organ dynamics.

The method described here allows time-lapse analysis of organ development in zebrafish embryos by using a fluorescence dissecting microscope capable of performing optical sectioning and simple strategies of readjustment to correct focal and planar drift.

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method ...

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 ...

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 ...

Multimodal Hierarchical Imaging of Serial Sections for Finding Specific Cellular Targets within Large Volumes

Targeting specific cells at ultrastructural resolution within a mixed cell population or a tissue can be achieved by hierarchical imaging using a combination of light and electron microscopy. Samples embedded in resin are sectioned into arrays consisting of ribbons of hundreds of ultrathin sections and deposited on pieces of silicon wafer or conductively coated coverslips. Arrays are imaged at low resolution using a digital consumer like smartphone camera or light microscope (LM) for a rapid large area overview, or a wide field fluorescence microscope (fluorescence light microscopy (FLM)) after labeling with fluorophores. After post-staining with heavy metals, arrays are imaged in a scanning electron microscope (SEM). Selection of targets is possible from 3D reconstructions generated by FLM or from 3D reconstructions made from the SEM image stacks at intermediate resolution if no fluorescent markers are available. For ultrastructural analysis, selected targets are finally recorded in the SEM at high-resolution (a few nanometer image pixels). A ribbon-handling tool that can be retrofitted to any ultramicrotome is demonstrated. It helps with array production and substrate removal from the sectioning knife boat. A software platform that allows automated imaging of arrays in the SEM is discussed. Compared to other methods generating large volume EM data, such as serial block-face SEM (SBF-SEM) or focused ion beam SEM (FIB-SEM), this approach has two major advantages: (1) The resin-embedded sample is conserved, albeit in a sliced-up version. It can be stained in different ways and imaged with different resolutions. (2) As the sections can be post-stained, it is not necessary to use samples strongly block-stained with heavy metals to introduce contrast for SEM imaging or render the tissue blocks conductive. This makes the method applicable to a wide variety of materials and biological questions. Particularly prefixed materials e.g., from biopsy banks and pathology labs, can directly be embedded and reconstructed in 3D.

This protocol targets specific cells in tissue for imaging at nanoscale resolution using a scanning electron microscope (SEM). Large numbers of serial sections from resin-embedded biological material are first imaged in a light microscope to identify the target and then in a hierarchical manner in the SEM.

Targeting specific cells at ultrastructural resolution within a mixed cell population or a tissue can be achieved by hierarchical imaging using a ...

Assessing Lysosomal Alkalinization in the Intestine of Live Caenorhabditis elegans

The nematode Caenorhabditis elegans (C. elegans) is a model system that is widely used to study longevity and developmental pathways. Such studies are facilitated by the transparency of the animal, the ability to do forward and reverse genetic assays, the relative ease of generating fluorescently labeled proteins, and the use of fluorescent dyes that can either be microinjected into the early embryo or incorporated into its food (E. coli strain OP50) to label cellular organelles (e.g. 9-diethylamino-5H-benzo(a)phenoxazine-5-one and (3-{2-[(1H,1&#39;H-2,2&#39;-bipyrrol-5-yl-kappaN(1))methylidene]-2H-pyrrol-5-yl-kappaN}-N-[2-(dimethylamino)ethyl]propanamidato)(difluoro)boron). Here, we present the use of a fluorescent pH-sensitive dye that stains intestinal lysosomes, providing a visual readout of dynamic, physiological changes in lysosomal acidity in live worms. This protocol does not measure lysosomal pH, but rather aims to establish a reliable method of assessing physiological relevant variations in lysosomal acidity. cDCFDA is a cell-permeant compound that is converted to the fluorescent fluorophore 5-(and-6)-carboxy-2&#39;,7&#39;-dichlorofluorescein (cDCF) upon hydrolysis by intracellular esterases. Protonation inside lysosomes traps cDCF in these organelles, where it accumulates. Due to its low pKa of 4.8, this dye has been used as a pH sensor in yeast. Here we describe the use of cDCFDA as a food supplement to assess the acidity of intestinal lysosomes in C. elegans. This technique allows for the detection of alkalinizing lysosomes in live animals, and has a broad range of experimental applications including studies on aging, autophagy, and lysosomal biogenesis.

Assessing Lysosomal Alkalinization in the Intestine of Live Caenorhabditis elegans

A step-by-step guide to probe loss of lysosomal acidity in the intestine of C. elegans using the pH-sensitive vital dye 5(6)-carboxy-2&#39;,7&#39;-dichlorofluorescein diacetate (cDCFDA)

The nematode Caenorhabditis elegans (C. elegans) is a model system that is widely used to study longevity and developmental pathways. Such studies are ...

Computational Analysis of the Caenorhabditis elegans Germline to Study the Distribution of Nuclei, Proteins, and the Cytoskeleton

The Caenorhabditis elegans (C. elegans) germline is used to study several biologically important processes including stem cell development, apoptosis, and chromosome dynamics. While the germline is an excellent model, the analysis is often two dimensional due to the time and labor required for three-dimensional analysis. Major readouts in such studies are the number/position of nuclei and protein distribution within the germline. Here, we present a method to perform automated analysis of the germline using confocal microscopy and computational approaches to determine the number and position of nuclei in each region of the germline. Our method also analyzes germline protein distribution that enables the three-dimensional examination of protein expression in different genetic backgrounds. Further, our study shows variations in cytoskeletal architecture in distinct regions of the germline that may accommodate specific spatial developmental requirements. Finally, our method enables automated counting of the sperm in the spermatheca of each germline. Taken together, our method enables rapid and reproducible phenotypic analysis of the C. elegans germline.

Computational Analysis of the Caenorhabditis elegans Germline to Study the Distribution of Nuclei, Proteins, and the Cytoskeleton

We present an automated method for three-dimensional reconstruction of the Caenorhabditis elegans germline. Our method determines the number and position of each nucleus within the germline and analyses germline protein distribution and cytoskeletal structure.

The Caenorhabditis elegans (C. elegans) germline is used to study several biologically important processes including stem cell development, apoptosis, and ...

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.

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 ...

4D Microscopy: Unraveling Caenorhabditis elegans Embryonic Development Using Nomarski Microscopy

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Chapters in this video

Comprehensive Assessment of Germline Chemical Toxicity Using the Nematode Caenorhabditis elegans

Analysis of Cardiomyocyte Development using Immunofluorescence in Embryonic Mouse Heart

Transcriptome Profiling of In-Vivo Produced Bovine Pre-implantation Embryos Using Two-color Microarray Platform

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

Using Light Sheet Fluorescence Microscopy to Image Zebrafish Eye Development

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

Multimodal Hierarchical Imaging of Serial Sections for Finding Specific Cellular Targets within Large Volumes

Assessing Lysosomal Alkalinization in the Intestine of Live Caenorhabditis elegans

Computational Analysis of the Caenorhabditis elegans Germline to Study the Distribution of Nuclei, Proteins, and the Cytoskeleton

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