Name: High-Resolution C. elegans Imaging Across All Larval Stages (Video) | JoVE Journal
Uploaded: 2025-05-23T14:20:00.000+00:00
Duration: 7 min 49 s
Description: Learn microfluidic-based time-lapse imaging for developmental biology using C. elegans. This method enables high-resolution observation of organ formation and cell divisions while maintaining animal viability across all larval stages.

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5 minutes

We wish to thank the members of the Hajnal laboratory for critical discussion and comments on the manuscript. We are also grateful to the C. elegans Genetics Center CGC, funded by the NIH Office of Research Infrastructure Programs (P40 OD010440). Furthermore, we wish to acknowledge the members of the Galli lab (Hubrecht Institute), the Conradt lab (University College London), and the van den Heuvel lab (Utrecht University) for valuable input on the presented protocol. This work was supported by grants from the Swiss National Science Foundation no. 31003A-166580 to AH, the Swiss Cancer League no. 4377-02-2018 to AH, and funding by ETH Z&#252;rich to AdM.

The details of the reagents and the equipment used in this study are listed in the Table of Materials.
1. Device fabrication
NOTE: All devices were fabricated using the protocol below, however, devices are directly available from the authors. A separate wafer is fabricated for each device type, with each device wafer fabricated from two layers of SU8 of different heights (i.e., food layer and trap layer). The valve wafer is fabricated on a separate wafer, with a single height of SU-8, omitting steps 1.3-1.6. The silanes are toxic and release corrosive vapors. This step, therefore, needs to be performed in a fume hood. For details on the fabrication procedure, refer to Berger et al.18.
<ol>
	<li>Clean wafers using air plasma.</li>
	<li>Spin coat the first SU8 layer, then perform a soft bake at 95 &#176;C.</li>
	<li>Expose the first layer, followed by a post-bake at 95 &#176;C and development.</li>
	<li>Perform plasma cleaning again after wafer development and drying.</li>
	<li>Spin coat the second SU8 layer, followed by a soft bake at 95 &#176;C (if necessary, conduct an initial bake at 65 &#176;C).</li>
	<li>Align the second mask to the existing features using alignment markers on either side of the wafer, then expose the second layer.</li>
	<li>Post-bake and develop the wafer, then perform a hard bake at 200 &#176;C.</li>
	<li>Treat the wafer with chlorotrimethyl silane for at least 2 h.</li>
	<li>Prepare a batch of 20 g PDMS pre-polymer (Part A) and thoroughly mix it with the 1 g of the cross-linker (ratio 20:1, Part B) (see Table of Materials).</li>
	<li>Degas for 10 min or until all bubbles disappear.</li>
	<li>Remove 2 mL of the PDMS mixture and reserve for step 1.13.</li>
	<li>Add another 3 g of cross-linker to the remaining PDMS mixture, and thoroughly mix (final ratio approx. 5:1).</li>
	<li>Using the batch of 20:1 PDMS, spin coat the device wafer at 750 rpm for 30 s.</li>
	<li>Place the valve wafer in an aluminum dish and cast the 5:1 PDMS onto it. Degas the device wafer for approximately 10 min. After degassing, remove any remaining bubbles using a pipette.</li>
	<li>Bake both the valve and device layers at 70 &#176;C for approximately&#160;15/20 min respectively.</li>
	<li>Assess the curing of the device and valve layers by gently probing them with a pipette tip or tweezers. The valve layer should be solid, while the device layer should remain slightly tacky.</li>
	<li>Detach the valve layer and trim it to the required size using a scalpel or razor blade.</li>
	<li>Punch an access hole into the valve channel (gauge 20).</li>
	<li>Position the device wafer under a stereomicroscope equipped with top illumination.</li>
	<li>Carefully place the PDMS piece containing the valve layer onto the device wafer.</li>
	<li>Align the valve and device features precisely, ensuring the removal of all trapped air between the layers.</li>
	<li>Transfer the assembled device to an oven and bake at 70 &#176;C overnight.</li>
	<li>Detach the device from the wafer.</li>
	<li>Punch all remaining access holes (gauge 20).</li>
	<li>Bond the PDMS to a cover glass using air plasma, then place the bonded device in an oven at 70 &#176;C for a few hours.</li>
</ol>
2. Worm preparation
NOTE: This section describes worm preparation by bleaching, which allows the generation of large numbers (hundreds) of animals. Other synchronization methods, such as manual picking, are possible but are typically more time-consuming and labor-intensive. All worms used in the protocol are maintained according to standard protocols at 20 &#176;C and on NGM plates seeded with E. coli OP50.
<ol>
	<li>Starting from plates with many gravid animals.</li>
	<li>Wash animals off the plate using M9 buffer.</li>
	<li>Add 5% NaClO and 5 M of NaOH to the worm suspension (200 &#181;L/100 &#181;L for every 1 mL worm suspension).</li>
	<li>Gently shake the bleaching mix until the animals begin to break apart (~10 min).</li>
	<li>Centrifuge bleaching mix at room temperature, 1300 x g for 1 min.</li>
	<li>Remove the supernatant using a pipette and add an equal amount of fresh M9 buffer.</li>
	<li>Again, centrifuge at room temperature, 1300 x g for 1 min.</li>
	<li>Remove the supernatant and transfer the pelleted worms to a 15 mL tube with 5 mL of fresh M9 buffer.</li>
	<li>Shake overnight at 20 &#176;C.</li>
	<li>Pass the worm suspension through a 10 &#181;m cell strainer.</li>
	<li>Centrifuge the worm suspension at 1300 &#215; g for 1 min at room temperature.</li>
	<li>Discard the supernatant and resuspend the worms in an equal volume of M9 buffer.</li>
	<li>Centrifuge the worm mixture again at 1300 &#215; g for 1 min at room temperature.</li>
	<li>Remove the supernatant and transfer the worms to NGM plates. If L1 larvae are needed, use them immediately.</li>
	<li>Once the worms reach the desired developmental stage, wash them off the plate using fresh S-Basal buffer filtered through a 0.2 &#181;m filter.</li>
	<li>Allow the worms to sediment by gravity or pellet them by centrifugation at 750 &#215; g for 1 min at room temperature.</li>
	<li>Discard the supernatant and add an equal volume of fresh S-Basal buffer.</li>
	<li>Repeat the sedimentation or centrifugation steps (steps 2.16 and 2.17)</li>
	<li>Remove the supernatant and add an equal volume of fresh S-Basal buffer.</li>
	<li>Perform the sedimentation or centrifugation steps one final time.</li>
	<li>Remove most of the supernatant.</li>
	<li>Keep the worms in the tube until the experiment begins.</li>
</ol>
3. Bacteria preparation
NOTE: Different bacteria strains (e.g., OP50, NA22, HT114, etc.) can be used on the chip. Adjust the proportion of density gradient medium added to the food mix to compensate for differences in density. All buffers are passed through a 0.2 &#181;m filter to remove particulates.
<ol>
	<li>Prepare 40 mL of LBroth, either in 2 x 20 mL in centrifuge tubes, sterile Erlenmeyer flasks, etc.</li>
	<li>Inoculate LB with bacteria picked from a stock plate.</li>
	<li>Grow the bacteria while shaking overnight at 37 &#176;C (~16 h) (OD600 = 1.9).</li>
	<li>Centrifuge bacteria at 3000 x g for 10 min.</li>
	<li>Remove the supernatant and add 2 mL of fresh S-Basal buffer to each tube. Combine all tubes.</li>
	<li>Mechanically resuspend the bacteria using a pipette.</li>
	<li>Centrifuge the bacteria at 3000 x g for 5 min.</li>
	<li>Remove the supernatant and add 2 mL of fresh S-Basal buffer again.</li>
	<li>Mechanically resuspend the bacteria using a pipette.</li>
	<li>Centrifuge the bacteria at 3000 x g for 5 min.</li>
	<li>Remove as much of the supernatant as possible and add 1 mL of fresh S-Basal buffer before mechanically resuspending all bacteria.</li>
	<li>Combine 0.60 mL of density gradient medium, 0.38 mL of S-Basal + 1% by weight Pluronic F-127 (a non-ionic triblock copolymer, represented here as PF) with 1 mL of OP50 bacteria suspension, or 0.65 mL of density gradient medium, 0.33 mL of S-Basal + 1% by weight PF with 1 mL of NA22 or HT114 bacteria suspension.</li>
	<li>Thoroughly mix the food preparation using a vortex mixer.</li>
	<li>Filter the food through a 5 &#181;m or 10 &#181;m cell strainer. A smaller strainer is especially advised when using smaller devices, e.g., L1 devices. 
	NOTE: Food can be kept at room temperature for up to 2 days.</li>
</ol>
4. Chip preparation
NOTE: Refer to Figure 2 for a schematic overview of the device, tubing, and connections. All buffers are passed through a 0.2 &#181;m filter to remove dust. The steel pin mentioned in the following steps refers to a short, hollow tube that is used to interface the larger OD tubing (1/16&#34;) to the small diameter holes fabricated in the PDMS device. These steel pins are simply inserted into the inner diameter of the tubing and then pushed into the PDMS material, effectively connecting the two.
<ol>
	<li>Begin by filling a syringe with deionized (DI) water.</li>
	<li>Attach a 23 G needle and a long piece of 1/16&#34; tubing, with a hollow steel pin (bent at a 90&#176; angle) at the end. The tubing should be long enough to extend from the solenoid to the microscope stage.</li>
	<li>Fill the tubing with DI water from the syringe and connect it to the valve inlet by inserting the steel pin into the punched hole (Figure 2A,J).</li>
	<li>Remove the syringe and needle, then attach the tubing to the off-chip solenoid (Figure 2A,J).</li>
	<li>Using the imaging software, turn on the solenoid and pressurize the device for several minutes to expel all air from the valve. Verify completion by visually checking the air-water interface on the chip, which should appear dark and disappear into the PDMS material (Figure 2B).</li>
	<li>Turn off the solenoid (Figure 2C).</li>
	<li>Fill a 1 mL syringe with the filtered bacteria solution (~0.5 mL of food is sufficient).</li>
	<li>Attach a 30 G needle and a long piece of 1/32&#34; tubing to the needle.</li>
	<li>Press the plunger to fill both the needle and attached tubing, ensuring no air remains in the syringe or tubing.</li>
	<li>Insert the 1/32&#34; tubing directly into the food inlet of the microfluidic device, using tweezers to prevent damage to the tubing (SMD tweezers are recommended) (Figure 2D,K).</li>
	<li>Place the syringe on the syringe pump (Figure 2D).</li>
	<li>Press the syringe plunger using the thumbscrew at the back to fill the device with liquid. A drop of liquid should appear at each open connection (Figure 2D, arrow).</li>
	<li>Block both the worm inlet and outlet with a sealed steel pin (a steel pin with a short piece of 1/16&#34; tubing attached and sealed by burning) (Figure 2E,L).</li>
	<li>Apply additional pressure using the thumbscrew to remove any remaining air in the device (Figure 2E, arrow).</li>
	<li>Remove the blocked pin at the outlet and attach the waste container (made from a cryovial or microcentrifuge tube, with two steel pins stuck through the lid and secured with glue). The waste container is connected using a short piece of 1/16&#34; tubing, with one end attached to the container and the other end connected to a hollow steel pin (bent 90&#176;) (Figure 2F,G,M).</li>
	<li>Push the syringe to ensure the waste container is properly connected and that no blockages exist in the system. For this, push a small amount of liquid through the system (turn the thumbscrew on the syringe pump) until liquid flows through the waste tubing (Figure 2G, arrow).</li>
	<li>Remove the second blocked steel pin.</li>
	<li>Push the syringe until a small drop of liquid appears at the worm inlet (Figure 2H,N).</li>
	<li>Attach a longer piece of 1/16&#34; tubing (15-20 cm) to a 1 mL syringe filled with S-Basal buffer, using a 23 G needle (Figure 2H,N).</li>
	<li>Attach a straight 23 G steel pin to the other end of the tubing.</li>
	<li>Fill the needle and tubing with buffer from the syringe, ensuring no air remains in the syringe or tubing.</li>
	<li>Insert the steel pin at the end of the tubing into the tube containing the worms.</li>
	<li>Push a small amount of liquid through the tubing, ensuring no air is left.</li>
	<li>Pull the worms into the tubing, but do not pull them into the syringe.</li>
	<li>Push the syringe connected to the worms until a small drop of liquid appears on the steel pin (Figure 2H).</li>
	<li>Insert the steel pin into the worm inlet (Figure 2I). 
	NOTE: Device preparation can be performed prior to the final worm preparation and left on the microscope until the worms are ready.</li>
</ol>
5. Worm loading and imaging
<ol>
	<li>Place the device onto a microscope at low magnification (5x or 10x) or on a dissection microscope.</li>
	<li>Identify the device unit currently in use and find the worm inlet using brightfield illumination.</li>
	<li>Position the device such that the inlet is visible on one side of the field of view and the back of the trap channel inlet is visible on the other side.</li>
	<li>Gently push on the worm syringe&#39;s plunger. The liquid will noticeably flow from the inlet through the channel array towards the outlet. The presence of bacteria and the difference in refractive index due to the addition of the density gradient medium make the liquid flow apparent.</li>
	<li>Ensure that the worms appear from the inlet and flow toward the back of the channel array.</li>
	<li>Gently push the animals toward the channel array. Once an animal faces the channel, push it into the channel, and repeat for additional animals. 
	NOTE: Animals need to be oriented with their heads toward the outlet to have easy access to food. They can be oriented by pushing and pulling on the plunger and by their swimming.</li>
	<li>Once sufficient animals have been trapped, place the syringe, still attached to the worm inlet, on the microscope stage, where it will remain throughout the experiment. Briefly detaching the worm tubing from the syringe may be necessary to release any pressure buildup, then carefully reattach it to the blunt needle.</li>
	<li>If loading was performed on a dissection microscope, transfer the device to the imaging microscope. The tubing can remain attached to the chip, and the syringe is connected to the pump.</li>
	<li>Switch on the syringe pump and run it at the preset rate of 1 &#181;L/h for 0.5 &#181;L, increase the rate by 100 &#181;L/h for 0.5 &#181;L, and reduce it back to 1 &#181;L/h. This cycle is automatically repeated for the entire experiment. For information on how to program the pump, see the user manual of the equipment.</li>
	<li>Place the device onto the microscope stage and ensure it is firmly held (suggestions for mounting mechanisms can be found in Figure 3A).</li>
	<li>If loading was not done on the imaging microscope, identify the device unit of interest at low magnification.</li>
	<li>Switch to the desired imaging magnification. 
	NOTE: The devices are compatible with all objective types.</li>
	<li>Identify the animals and regions of interest within the trap channel array and set up the desired imaging conditions.</li>
	<li>Image at the desired imaging conditions, with the on-chip valve, actuated through the solenoid 10 s before image acquisition, so the animals are held in place. 
	NOTE: Actuating the on-chip valve will elongate and flatten the animals, possibly requiring adjustments to the imaging region of interest (ROI).</li>
</ol>
6. Setup preparation
NOTE: This section refers to the initial preparation of the device control mechanism and does not need to be repeated before each experiment. Refer to Figure 3B,C. Because many solenoids require a higher voltage, the solenoid will most likely not be connected directly to the microcontroller but via a relay or MOSFET (see Figure 3C).
<ol>
	<li>Identify a pressurized air source (e.g., wall supply installation or gas cylinder).</li>
	<li>Connect the pressure source to a suitable pressure regulator (0-2 bar pressure range) using 6 mm tubing (wall source) or directly attach a suitable regulator to the gas cylinder.</li>
	<li>Connect the pressure regulator to the solenoid valve input (6 mm tubing, see Figure 3B).</li>
	<li>Place the solenoid as close to the microscope stage as possible (~0.5-1 m from the stage is suggested, though longer distances are possible if needed).</li>
	<li>Connect the solenoid valve electronically to the selected controller. Different options are available, such as controlling it using a separate microcontroller board (e.g., Arduino) or an existing DAq card.</li>
</ol>
7. Cleanup and storage
<ol>
	<li>Once the experiment is completed, remove the device from the microscope.</li>
	<li>Remove the immersion liquid. Immersion oil can be removed with organic solvents (e.g., acetone, methanol, etc.).</li>
	<li>Place the device on a flat surface and disconnect all tubing except the valve tubing. To safely disconnect each tubing, press down on the PDMS close to it and gently pull. Do not lift and bend the device, as this may cause the cover glass to break.</li>
	<li>Clean the PDMS surface with 70% ethanol.</li>
	<li>Store the device at room temperature with the valve tubing attached. 
	NOTE: The device can be reused until all units have been used, likely without needing to refill the valve tubing. However, the water in the valve tubing will evaporate over an extended period and may eventually need to be refilled.</li>
</ol>

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<li>Agarwal, P., Berger, S., Shemesh, T., Zaidel-Bar, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Active+nuclear+positioning+and+actomyosin+contractility+maintain+leader+cell+integrity+during+gonadogenesis%5BTitle%5D)+AND+%22Curr+Biol%22%5BJournal%5D)">Active nuclear positioning and actomyosin contractility maintain leader cell integrity during gonadogenesis.</a> Curr Biol. 34 (11), 2373-2386 (2024).</li>
<li>Segos, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Non-random+segregation+of+mitochondria+during+asymmetric+cell+division+contributes+to+cell+fate+divergence+in+daughter+cells%5BTitle%5D)+AND+%22bioRxiv%22%5BJournal%5D)">Non-random segregation of mitochondria during asymmetric cell division contributes to cell fate divergence in daughter cells.</a> bioRxiv. , 2024-2109 (2024).</li>
<li>Brenner, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+genetics+of+Caenorhabditis+elegans%5BTitle%5D)+AND+%22Genetics%22%5BJournal%5D)">The genetics of Caenorhabditis elegans.</a> Genetics. 77 (1), 71-94 (1974).</li>
<li>Stiernagle, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Maintenance+of+C.+elegans%5BTitle%5D)+AND+%22WormBook%3A+The+Online+Review+of+C.+elegans+Biology%22%5BJournal%5D)">Maintenance of C. elegans.</a> WormBook: The Online Review of C. elegans Biology. , (2006).</li>
<li>Pepin, B. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aPepin%2C+B+%22GPU+Deconvolution+Library+with+C+and+Matlab+API+for+Nvidia+GPUs%22">GPU Deconvolution Library with C and Matlab API for Nvidia GPUs</a>. , https://github.com/bobpepin/YacuDecu (2025).</li>
<li>Podbilewicz, B., White, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cell+fusions+in+the+developing+epithelia+of+C.+elegans%5BTitle%5D)+AND+%22Dev+Biol%22%5BJournal%5D)">Cell fusions in the developing epithelia of C. elegans.</a> Dev Biol. 161 (2), 408-424 (1994).</li>
<li>Nusser-Stein, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cell-cycle+regulation+of+NOTCH+signaling+during+C.+elegans+vulval+development%5BTitle%5D)+AND+%22Mol+Syst+Biol%22%5BJournal%5D)">Cell-cycle regulation of NOTCH signaling during C. elegans vulval development.</a> Mol Syst Biol. 8 (1), 618(2012).</li>
<li>Euling, S., Ambros, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Heterochronic+genes+control+cell+cycle+progress+and+developmental+competence+of+C.+elegans+vulva+precursor+cells%5BTitle%5D)+AND+%22Cell%22%5BJournal%5D)">Heterochronic genes control cell cycle progress and developmental competence of C. elegans vulva precursor cells.</a> Cell. 84 (5), 667-676 (1996).</li>
<li>Mok, D. Z., Sternberg, P. W., Inoue, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Morphologically+defined+sub-stages+of+C.+elegans+vulval+development+in+the+fourth+larval+stage%5BTitle%5D)+AND+%22BMC+Dev+Biol%22%5BJournal%5D)">Morphologically defined sub-stages of C. elegans vulval development in the fourth larval stage.</a> BMC Dev Biol. 15, 1-8 (2015).</li>
<li>Laranjeira, A. C., Berger, S., Kohlbrenner, T., Greter, N. R., Hajnal, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Nutritional+vitamin+B12+regulates+RAS%2FMAPK-mediated+cell+fate+decisions+through+one-carbon+metabolism%5BTitle%5D)+AND+%22Nat+Commun%22%5BJournal%5D)">Nutritional vitamin B12 regulates RAS/MAPK-mediated cell fate decisions through one-carbon metabolism.</a> Nat Commun. 15 (1), 8178(2024).</li>
</ol>

The authors declare no competing or financial interests.

This article has described the operation of a unique microfluidic long-term imaging platform, which is suitable for high-resolution imaging and tracking of various developmental processes in C. elegans during all four larval stages into early adulthood18. The ability to follow developmental processes over time in vivo, using this imaging method and others, has become indispensable in many fields of C. elegans research and has allowed previously inaccessible questions to be answered23,24,29.
In contrast to traditional imaging methods using agar pads and previously published microfluidic-based long-term imaging strategies, this approach preserves worm identity and orientation throughout the experiment, enabling the tracking of complex developmental processes over time. Regardless of developmental stage, animals are confined to one of 41 parallel trap channels, each connected to an on-chip food supply and covered by a large hydraulic valve. Immobilization of all trapped worms occurs through the inflation of the hydraulic valve, pressing the worms onto the cover glass only during image acquisition. High-throughput imaging is possible due to the narrowly spaced trap channel array, with multiple animals visible within a single field of view. The long-term imaging devices are compatible with most microscope setups, requiring minimal additional hardware (less than $1000) and no modifications to existing hardware. The operation of the devices is intentionally simple, so all users should quickly be able to learn it by following the above protocol and outlined results. 
Briefly, animals can be imaged on-chip across up to 2.5 larval stages (Figure 1C). Animals are initially synchronized and either directly loaded on-chip (L1 device) or seeded onto the NGM plate until the desired developmental stage (L1-L4 and L2-A devices). Animals on-chip are fed using a highly concentrated bacteria suspension and immobilized using an on-chip hydraulic valve actuated by the imaging software (Figure 1B). Animals are maneuvered into the trap channels using a syringe, carefully orienting animals with the head toward the food supply and finally trapping animals at the end of the trap channel (Figure 1A and Figure 2). Care must be taken so that no air is trapped in the syringes, blunt needles, and tubing, as air bubbles will readily disturb the animal's position. Devices are compatible with most imaging modalities18,19,24, as the PDMS material is highly transparent, non-fluorescent, and sealed against a thin cover glass (Figure 4, Supplementary Movie 1, Supplementary Movie 2, and Supplementary Movie 3). However, care must be taken when selecting imaging conditions, as long exposure times and high excitation intensities will readily affect animal viability and result in photobleaching.
In addition to the presented version of this imaging method, several modifications to the protocol are possible and already available. The most straightforward modification is replacing the standard bacteria strains, e.g., with RNAi-expressing bacteria18 or bacteria with different nutritional values32. Bacterial strains can readily be exchanged by adjusting the Optiprep content to account for differences in density. Similarly, any number of compounds can be added to the bacterial food, e.g., auxin, nutrients, or small molecule drug compounds, and their effect is observed directly in vivo, without any need for any changes to the protocol. In particular, a media exchange device was developed for auxin-induced protein degradation or any application that needs timed or reversible exposure by adding a food inlet to the device layout without changing the geometry or operation. The additional inlet allows simple switching between two different conditions, e.g., auxin and no auxin, attached to the device in the same manner as shown here for a single food source, and the resulting changes are observed directly. Device variants were also developed where the hydraulic valve is not shared across all units on a device but separated for each unit, such that multiple conditions can be imaged within the same device and experiment individually, reducing the potentially harmful effects of long valve actuation and acquisition times. Finally, beyond the original device variants described in Berger et al.18, two new device variants were developed, L3-A and L4-A (introduced here), allowing imaging from the mid-L3 or mid-L4 stage until adulthood. The purpose of these modifications is to delay the start time of an experiment to a later larval stage, such that larger animals can be loaded in applications where the earlier developmental stages are not of interest.

C. elegans imaging can be carried out in several ways depending on the application, desired throughput, and resolution. When a high resolution is needed, the standard method utilizes agar pads1,2,3, a simple construction of glass slide, agarose, and cover glass. Animals on these slides are immobilized by the pressure exerted by a cover glass. Immobilization can further be improved by the addition of anesthetizing agents, e.g., levamisole or sodium azide1,2, such that animals on agar pads are perfectly still and can be imaged at high resolution, including various super-resolution methods2. Unfortunately, agar pads are known to affect developmental progression, with the combination of high applied pressures and anesthetizing agents retarding development or causing complete developmental arrest4,5.
Besides these standard methods, several microfluidic methodologies have emerged in recent years, each with its advantages and disadvantages or specialized use cases6,7,8,9,10,11,12,13,14,15,16,17,18,19. For instance, Gritti et al.20 introduced a method in which worms are confined within large chambers. Embryos placed in these chambers develop and move freely while remaining within the designated region of interest on a microscope. While generally effective, this approach is limited to the use of bright fluorescent markers. More recently, Keil et al.21 proposed a modified version of this method, incorporating an on-chip hydraulic valve to restrain animals during image acquisition, thereby addressing some limitations of the original approach. However, since both methods utilize larger chambers than those used for trapped worms, the animals continue to move and rotate throughout the experiment. As a result, tracking developmental processes becomes challenging and requires extensive image post-processing to correct for animal motion.
To address these limitations, the approach introduced by Berger et al.18 (Figure 1), where animals are not held in a large chamber but housed in a trap channel designed to fit animals within a shorter developmental window (up to 2.5 larval stages) is adapted. Unlike existing microfluidic methods, this approach preserves animal orientation and identity throughout an experiment and allows the acquisition of high-resolution images of multiple worms in parallel and across multiple larval stages within a single device.
The animals are trapped using a specialized channel geometry where the channel height is close to the thickness of an animal at the beginning of an experiment, thus preventing rotation. Channel widths and lengths are selected so that animals can move and grow over time, facilitating molting and the transition from one larval stage to the next. Parallelization is then achieved simply by placing multiple trap channels next to each other, such that multiple animals are visible in a single field of view (FOV, Figure 1A&#39;), and up to 41 animals can be imaged in a single device unit (Figure 1A). Animals on the chip are continually supplied a highly concentrated bacterial food suspension via a low-height channel, which at the same time helps confine animals at the end of the trap channel array (height step, Figure 1A, red). Similar to the approach taken by Keil et al.21, a large hydraulic, on-chip valve is implemented, which can be inflated during image acquisition, pressing the animals against the coverglass, thus improving animal immobilization (Figure 1B). Limiting hydraulic valve actuation to only the image acquisition period minimizes any adverse effect on animal development, ensuring reliable development across all larval stages. Unlike the methods described by Gritti et al.20 and Keil et al.21, animals cannot be imaged across the entire post-embryonic development within the same device. Instead, five distinct devices are introduced, termed L1, L1-4, L2-A, L3-A, and L4-A, covering the whole L1 stage and early L2 stage, the mid-late L1 stage up to the mid-L4 stage, the mid-late L2 up to young adulthood, the mid-late L3 up to young adulthood, and the late L4 up to young adulthood, respectively (Figure 1C), which allow uninterrupted imaging of large parts of the post-embryonic development. This approach is taken due to the large increase in animal size from L1 to adulthood, such that a channel suitable for L1 animals would be much too small for adults, resulting in developmental arrest, and a channel fitting adults would be much too large for L1s, resulting in excessive movement, rotation and animals escaping the trap channels.
Using this approach, reliable developmental progression for a variety of tissues is achieved18, e.g., the post-embryonic development of the hypodermis (L1 to L4 stage), induction of the vulval cell fates (L2 and L3 stage), vulval morphogenesis (L3 to adulthood19) and anchor cell invasion. Since their introduction, these long-term imaging devices have been used to study a variety of processes22,23, with the platform even being combined with techniques such as AiryScan super-resolution microscopy24, enabling numerous discoveries previously impossible.
In the following, the operation aspects of these long-term imaging devices are introduced in a step-by-step protocol, highlighting the expected results and some potential challenges when setting up the system.

<table><tbody><tr><td>3/2 solenoid valve&amp;nbsp;</td><td>Distrelec</td><td>154-22-898 and 154-22-899</td><td>Alternative to above. Solenoid valve connecting the pressure regulator to the on-chip valve.</td></tr><tr><td>Arduino</td><td>Distrelec</td><td>301-01-956</td><td>Optional microcontroller depending on the selected controller and interface.</td></tr><tr><td>Bacto-Tryptone</td><td>Thermo Fisher Scientific</td><td>211705</td><td>For L Broth production.</td></tr><tr><td>Bacto-Yeast</td><td>Thermo Fisher Scientific</td><td>212750</td><td>For L Broth production.</td></tr><tr><td>Blunt needle 23G</td><td>Gonano Dosiertechnik</td><td>GGA723050&amp;nbsp;</td><td>Blunt needle connecting the 1/16&quot; tubing to a syringe.</td></tr><tr><td>Blunt needle 30G</td><td>Gonano Dosiertechnik</td><td>IG-TE730050</td><td>Blunt needle connecting the 1/32&quot; tubing to a syringe.</td></tr><tr><td>&lt;em&gt;C. elegans&lt;/em&gt; Construct 1</td><td>NA</td><td>NA</td><td>AH5786 LGI: hmr-1(cp21[hmr-1::gfp + LoxP]) Marston, D.J. et al. MRCK-1 drives apical constriction in C. elegans by linking developmental patterning to force generation. Curr. Biol. 26 (16), 2079-2089. 2016.</td></tr><tr><td>&lt;em&gt;C. elegans&lt;/em&gt; Construct 2</td><td>NA</td><td>NA</td><td>AH1187 LGII: arIs92[egl-17::cfp] Yoo, A.S., Bais, C., Greenwald, I. Crosstalk between the EGFR and LIN-12/Notch pathways in C. elegans vulval development. Science. 303 (5658), 663-666. 2004.</td></tr><tr><td>&lt;em&gt;C. elegans&lt;/em&gt; Construct 3</td><td>NA</td><td>NA</td><td>AH5786 LGV: qyIs50[Pcdh-3&gt;mCherry::moeABD, unc-119(+)] Ziel, J.W., Hagedorn, E.J., Audhya, A., Sherwood, D.R. UNC-6 (netrin) orients the invasive membrane of the anchor cell in C. elegans. Nat. Cell Biol., 11 (2), 183-189. 2009.</td></tr><tr><td>&lt;em&gt;C. elegans&lt;/em&gt; Construct 4</td><td>NA</td><td>NA</td><td>ML2615 LGX: dlg-1(mc103[dlg-1::gfp]) Vuong-Brender, T.T.K., Suman, S.K., Labouesse, M. The apical ECM preserves embryonic integrity and distributes mechanical stress during morphogenesis. Development. 144 (23), 4336-4349. 2017.</td></tr><tr><td>Cable</td><td>Distrelec</td><td>143-46-644</td><td>Needed if selecting the solenoid from distrelec. Cable connecting the solenoid to the MOSFET.</td></tr><tr><td>Chlorotrimethyl silane&amp;nbsp;</td><td>Sigma Aldrich</td><td>386529-25ML</td><td>Silane for wafer passivation.</td></tr><tr><td>Cholesterol</td><td>Sigma Aldrich</td><td>C8667</td><td>For&amp;nbsp; S-Basal production.</td></tr><tr><td>Flyback Diode</td><td>Distrelec</td><td>110-52-628</td><td>Flyback diode protecting the microcontroller.</td></tr><tr><td>Hollow steel pin</td><td>Gonano Dosiertechnik</td><td>GGA7R23050&amp;nbsp;</td><td>23G hollow steel pin connecting the 1/16&quot; tubing to the device.</td></tr><tr><td>L Broth</td><td>NA</td><td>NA</td><td>Prepared in house following standard recipe. 10 g Bacto-tryptone, 5 g Bacto-yeast, 5 g NaCl, H2O to 1 litre, pH to 7.0 using 1 M NaOH</td></tr><tr><td>Long-term Imaging Device L1</td><td>NA</td><td>NA</td><td>Long-term imaging device for L1 to early/mid L2 larvae. Made inhouse, available on request. Also available as media exchange variant.</td></tr><tr><td>Long-term Imaging Device L1-4</td><td>NA</td><td>NA</td><td>Long-term imaging device for mid L1 to mid L4 larvae. Made inhouse, available on request. Also available as media exchange variant.</td></tr><tr><td>Long-term Imaging Device L2-A</td><td>NA</td><td>NA</td><td>Long-term imaging device for mid L2 to adulthood (eversion). Made inhouse, available on request. Also available as media exchange variant.</td></tr><tr><td>Long-term Imaging Device L3-A</td><td>NA</td><td>NA</td><td>Long-term imaging device for mid L3 to adulthood (eversion). Made inhouse, available on request. Also available as media exchange variant.</td></tr><tr><td>Long-term Imaging Device L4-A</td><td>NA</td><td>NA</td><td>Long-term imaging device for mid L4 to adulthood (eversion). Made inhouse, available on request. Also available as media exchange variant.</td></tr><tr><td>M9 Buffer</td><td>NA</td><td>NA</td><td>Prepared in house following standard recipe.</td></tr><tr><td>MOSFET</td><td>Distrelec</td><td>303-41-391</td><td>Metal-oxide-semiconductor field-effect transistor, used to switch the 24VDC required for the solenoid.</td></tr><tr><td>NGM plates</td><td>NA</td><td>NA</td><td>Prepared in house following standard recipe.</td></tr><tr><td>Nylon Luer-Fitting</td><td>Fisher Scientific</td><td>11776048</td><td>Adapter from OD 6mm tubing to 23G blunt needle.</td></tr><tr><td>Optiprep</td><td>Sigma Aldrich</td><td>D1556</td><td>Density matching liquid.</td></tr><tr><td>PDMS</td><td>Ameba</td><td>Elastosil RT601A/B</td><td>PDMS elastomer kit.</td></tr><tr><td>Pluronic F-127</td><td>Sigma Aldrich</td><td>P2443</td><td>Non-ionic surfactant.</td></tr><tr><td>Potassium dihydrogen phosphate</td><td>Sigma Aldrich</td><td>1.37039</td><td>For&amp;nbsp; S-Basal production.</td></tr><tr><td>Potassium hydrogen phosphate</td><td>Sigma Aldrich</td><td>1.05099</td><td>For&amp;nbsp; S-Basal production.</td></tr><tr><td>Power Supply 24VDC</td><td>Distrelec</td><td>301-63-304</td><td>24VDC power supply. Select a power supply suitable for the solenoid selected.</td></tr><tr><td>Precision pressure regulator</td><td>Distrelec</td><td>154-23-578</td><td>Alternativ to above. Pressure regulator.</td></tr><tr><td>Pressure Gauge G1/8</td><td>Distrelec</td><td>154-23-643</td><td>Pressure gauge attaches to the pressure regulator.</td></tr><tr><td>PU Tubing OD 6mm</td><td>Distrelec</td><td>154-23-866</td><td>Tubing connecting the pressure source and regulator, as well as regulator and solenoid valve.</td></tr><tr><td>Push-in fitting G1/8 to 6mm</td><td>Distrelec</td><td>301-69-330</td><td>Alternativ to above. 2x Push-in fitting attaches to the pressure regulator and connects to the 6mm tubing.</td></tr><tr><td>Push-in fitting M5 to 6 mm</td><td>Distrelec</td><td>301-69-459</td><td>2x Push-in fitting attaches to the solenoid and the 6mm tubing.</td></tr><tr><td>S-Basal Buffer</td><td>NA</td><td>NA</td><td>Prepared in house following standard recipe. 5.85 g NaCl, 1 g K2HPO4, 6 g KH2PO4, 1 ml cholesterol (5 mg/ml in ethanol), H2O to 1 litre. Sterilize by autoclaving.</td></tr><tr><td>Sodium Chloride</td><td>Sigma Aldrich</td><td>S9625</td><td>For L Broth and S-Basal production.</td></tr><tr><td>Sodium hydroxite (5M)</td><td>Sigma Aldrich</td><td>567530</td><td>Sodium hydroxide used when bleaching C. elegans (prepare solution at 5M concentration)</td></tr><tr><td>Sodium hypochlorite (5%)</td><td>Sigma Aldrich</td><td>1.05614</td><td>Sodium hypochlorite used when bleaching C. elegans (prepare solution at 5% concentration)</td></tr><tr><td>Strainer 10 &amp;micro;m</td><td>pluriSelect</td><td>43-10010-40</td><td>Alternative for bacteria food preparation, especially if smaller devices are used.</td></tr><tr><td>Strainer 5 &amp;micro;m</td><td>pluriSelect</td><td>43-10005-40</td><td>Strainer used to filter L1 larvae after overnight starvation and bacteria after completed food preparation.</td></tr><tr><td>SU-8</td><td>Gersteltec</td><td>GM1050</td><td>SU-8 for height from 1-10 &amp;micro;m.</td></tr><tr><td>SU-8</td><td>Gersteltec</td><td>GM1060</td><td>SU-8 for height from 5-30 &amp;micro;m.</td></tr><tr><td>SU-9</td><td>Gersteltec</td><td>GM1070</td><td>SU-8 for height from 15-200 &amp;micro;m.</td></tr><tr><td>Syringe 1 mL</td><td>Fisher Scientific</td><td>11338763</td><td>1mL syringe. If using a different type ensure that the syringe body and plunger are stiff, such that they won&#039;t be deformed by the syringe pump.</td></tr><tr><td>Syringe Pump</td><td>WPI</td><td>Al1000-220</td><td>Syringe pump (https://www.wpiinc.com/var-2300-aladdin-single-syringe-pump.html).</td></tr><tr><td>Tube 15 mL</td><td>Fisher Scientific</td><td>50-809-220</td><td>Tube used for worm bleaching and bacteria preparation.</td></tr><tr><td>Tube 2 mL</td><td>Fisher Scientific</td><td>NC1186931</td><td>Tube used for worm preparation and bacteria preparation.</td></tr><tr><td>Tube 50 mL</td><td>Fisher Scientific</td><td>50-465-232</td><td>Tube used for bacteria preparation and growth.</td></tr><tr><td>Tweezer SMD</td><td>Br&amp;uuml;tsch R&amp;uuml;egg Werkzeuge</td><td>448980</td><td>Tweezers used to plug the 1/32&quot; tubing into the device.</td></tr><tr><td>Tygon tubing 1/16&quot;</td><td>Fisher Scientific</td><td>642002</td><td>Flexible 1/16&quot; OD tubing connecting most components to the device.</td></tr><tr><td>Tygon tubing 1/32&quot;</td><td>Fisher Scientific</td><td>641900</td><td>Flexible 1/32&quot; OD tubing connecting the bacteria supply to the device.</td></tr><tr><td>UV KUB-3</td><td>Kloe</td><td>NA</td><td>Mask aligner and UV exposure system.</td></tr><tr><td>Wafer</td><td>Siegert Wafer</td><td>4P0/&gt;1/525&amp;plusmn;25/SSP/TTV&amp;lt;10</td><td>Silicon Wafer.</td></tr><tr><td>Waste Container</td><td>NA</td><td>NA</td><td>Prepared in house using a 2mL tube or a cryo vial and hollow steel pins.</td></tr></tbody></table>

high-resolution c. elegans imaging across all larval stages

High-Resolution C. elegans Imaging Across All Larval Stages

Caenorhabditis elegans has become one of the most widely studied and best-understood animal models in biology. Three features are key to C. elegans' success as a model organism: its invariant cell lineage, transparency, and genetic tractability. These render it ideal for a diverse range of microscopy-based studies directly in vivo. Live C. elegans larvae and adults often need to be immobilized during image acquisition. Traditional immobilization methods adversely affect animal development, especially in time-lapse imaging applications. Here, a detailed setup and operation protocol for a novel microfluidic imaging method is introduced, which addresses the limitations associated with traditional agar-pad-based immobilization and other microfluidic strategies. This approach enables simultaneous live imaging across various larval stages while preserving worm orientation and identity over time. To achieve this, a microfluidic trap channel array is employed, with its geometry precisely designed to maintain a stable worm orientation while accommodating growth and molting. Immobilization is facilitated by an active hydraulic valve that applies pressure to secure worms against the cover glass solely during image acquisition. This design allows high-resolution imaging with minimal effects on worm viability or developmental timing.

This protocol describes microfluidic-based C. elegans time-lapse imaging across the entire post-embryonic development.

Caenorhabditis elegans has become one of the most widely studied and best-understood animal models in biology. Three features are key to C. elegans' ...

high-resolution-c-elegans-imaging-across-all-larval-stages

Research

JoVE Journal

Developmental Biology

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

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

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

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

This article describes a technique for the visualization of the early events of embryogenesis in the nematode Caenorhabditis elegans.

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

A simple microfluidic device has been developed to perform anesthetic free in vivo imaging of C. elegans, intact Drosophila larvae and zebrafish larvae. The device utilizes a deformable PDMS membrane to immobilize these model organisms in order to perform time lapse imaging of numerous processes such as heart beat, cell division and sub-cellular neuronal transport.  We demonstrate the use of this device and show examples of different types of data collected from different model systems.

Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish

This protocol describes how to image dividing cells within a tissue in Caenorhabditis elegans embryos. While several protocols describe how to image cell division in the early embryo, this protocol describes how to image cell division within a developing tissue during mid late embryogenesis.

Visualizing Neuroblast Cytokinesis During C. elegans Embryogenesis

Imaging behavior and neural activity over long time scales without immobilization of the animal is a prerequisite to understand behavior. Agarose microfluidic chambers imaging (AMI) can be used to image neural activity and behavior for all life stages of Caenorhabditis elegans.

Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans

The transparent C. elegans intestine can serve as an "in vivo tissue chamber" for studying apicobasal membrane and lumen biogenesis at the single-cell and subcellular level during multicellular tubulogenesis. This protocol describes how to combine standard labeling, loss-of-function genetic/RNAi and microscopic approaches to dissect these processes on a molecular level.

The C. elegans Intestine As a Model for Intercellular Lumen Morphogenesis and In Vivo Polarized Membrane Biogenesis at the Single-cell Level: Labeling by Antibody Staining, RNAi Loss-of-function Analysis and Imaging

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.

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

This work describes a semi-high-throughput protocol that allows simultaneous 3D time-lapse imaging of embryogenesis in 80–100 C. elegans embryos in a single overnight run. Additionally, image processing and visualization tools are included to streamline data analysis. The combination of these methods with custom reporter strains enables detailed monitoring of embryogenesis.

A Semi-high-throughput Imaging Method and Data Visualization Toolkit to Analyze C. elegans Embryonic Development

The protocol describes a simple microfluidic chip design and microfabrication methodology used to grow C. elegans in presence of a continuous food supply for up to 36 h. The growth and imaging device also enables intermittent long-term high-resolution imaging of cellular and sub-cellular processes during development for several days.

A Simple Microfluidic Chip for Long-Term Growth and Imaging of Caenorhabditis elegans

Super-resolution microscopy can provide a detailed insight into the organization of components within the synaptonemal complex in meiosis. Here, we demonstrate a protocol to resolve individual proteins of the Caenorhabditis elegans synaptonemal complex.

Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline

Learn microfluidic-based time-lapse imaging for developmental biology using C. elegans. This method enables high-resolution observation of organ formation and cell divisions while maintaining animal viability across all larval stages.

High-Resolution C. elegans Imaging Across All Larval Stages

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