This research was supported by the National Science Foundation through grants PHY-1205494 and MCB-1413134 (EL) and by the National Research Foundation of Korea grant 2017R1D1A1B03035671 (KSL).

The protocol below uses WormSpa5, a previously-described microfluidic device for longitudinal imaging of worms. Fabrication of WormSpa (starting with CAD files that can be obtained from the authors upon request) is straightforward but requires some expertise. In most cases, fabrication can readily be done by a core facility or by a commercial company that provides such services. When fabricating the device, make sure to specify that the height of the features is 50 &#181;m.
1. Experimental Setup
<ol>
	<li>Mount the microfluidic device on an inverted fluorescence microscope with a motorized stage.</li>
	<li>Place an orbital shaker close to the microscope, but make sure the vibration from the shaker does not directly affect the microscope. For example, put the shaker on a shelf fixed on a wall that makes no contact with the optical table.</li>
	<li>Place a syringe pump on the orbital shaker. Tape the pump to the shaker so that it does not jiggle while shaking.</li>
</ol>
2. Preparation of the Microfluidic Environment
NOTE: During the experiment, four syringes are connected to the microfluidic device via syringe tubes, as outlined in Figure 1. This step describes the preparation of these syringes. Unless mentioned otherwise, keep the syringe tubes as short as possible without making them taut.
<ol>
	<li>Prepare an outlet syringe and a syringe tube. This syringe (S1 in Figure 1) will later be connected to the outlet of the device through the syringe tube, and used for temporary storage of fluid coming out of the device.
	<ol>
		<li>Make an adapter needle by removing plastic parts of a 20 G x 1/2 inch syringe tip, keeping only the needle (the metal part). Do so by holding the plastic and metal parts with pliers and pulling them apart. Remaining plastic residue on the needle can be burnt off with a Bunsen burner.</li>
		<li>Bend the needle according to the geometry of the experimental setup while holding its two ends with pliers. In the design described here, bending the needle in its middle to an angle of ~110&#176; is most convenient.</li>
		<li>Plug the adapter needle half way into the tube. The other half will later be plugged into the device. The syringe tube should be compatible with a 20 G syringe tip without a leak (for example, tubing with inner diameter of 0.86 mm and outer diameter of 1.32 mm). The recommended tube length is ~50 cm.</li>
		<li>Connect a 10 mL luer-lock syringe to the other (open) end of the syringe tube via a 20 G x 1/2 inch syringe tip.</li>
	</ol>
	</li>
	<li>Prepare buffer-inlet syringes and syringe tubes. One of these syringes (S2 in Figure 1) is used as a reservoir for the fluid that goes into the device and to control the injection flow. The other syringe (S3 in Figure 1) is required for buffer exchange, as explained in Step 3.2.
	<ol>
		<li>Connect a 10 mL luer-lock syringe (S2) directly to a 3-way valve and connect another syringe (S3) to the same valve via a syringe tube (Figure 1): connect S3 to the tube via a syringe tip, and connect the tube to the valve via another syringe tip. 
		NOTE: The order in which these materials are connected does not matter.</li>
		<li>Connect a syringe tube to the remaining port of the 3-way valve. Prepare another adapter needle as in Step 2.1.2, and plug the needle half-way into the other (open) end of the syringe tube.</li>
		<li>Set the valve such that the syringe tube in Step 2.2.2 and S2 are connected while S3 is closed (Figure 1).</li>
	</ol>
	</li>
	<li>Prepare a worm-inlet syringe and a syringe tube. This syringe (S4 in Figure 1) is used for loading worms into the device.
	<ol>
		<li>Connect a long syringe tube to a 10 mL luer-lock syringe (S4). Determine the length of this tube from the volume of the fluid that will be used for loading; for example, 100 cm of tubing supports over 2 mL of liquid (also, see step 4.2).</li>
		<li>Prepare another adapter needle as in Step 2.1.2, and plug one half of the needle to the other (open) end of the syringe tube.</li>
	</ol>
	</li>
	<li>Rinse all the syringes and tubing with S-medium12 twice. Fill the syringes and the tubes with S-medium, taking care to remove all air bubbles.</li>
	<li>Connect the outlet syringe tube to the device outlet.
	<ol>
		<li>Plug in the adapter needle at the end of the syringe tube to the outlet port.</li>
		<li>Inject a small volume of S-medium through the outlet to fill up the device, until a little S-medium comes out of both the buffer inlet and the worm inlet.</li>
	</ol>
	</li>
	<li>Connect the buffer-inlet syringe tube to the buffer-inlet port of the device, while plugging the worm-inlet with a pin (stainless steel dowel pin with a 1/32-inch diameter).</li>
	<li>Inject a constant flow of buffer into the device. Load the buffer-inlet syringe S2 onto a syringe pump13, and set the pump such that the medium in S2 is continuously injected into the device at a flow rate of 3 &#181;L/min.</li>
</ol>
3. Loading Bacteria into the Microfluidic Device
<ol>
	<li>Prepare Escherichia coli OP50 suspended in S-medium12.
	<ol>
		<li>Following a standard protocol, inoculate a 250 mL flask with 50 mL of LB and a single colony, and incubate overnight at 37 &#176;C.</li>
		<li>Centrifuge the overnight culture at 4000 rpm for 10 minutes and resuspend the pellet in 30 mL of S-medium.</li>
		<li>Filter the suspension through a 5 &#181;m syringe filter. Measure the bacterial concentration by its optical density at 600 nm (OD600), and adjust the concentration to an OD600 of 3. This high density is required to keep the worms well-fed.</li>
	</ol>
	</li>
	<li>Exchange the buffer in the device with the OP50 suspension.
	<ol>
		<li>Prepare a clean syringe (S5) filled with the OP50 suspension. Hold the syringe vertically and tap it several times to collect all the air bubbles at the top.</li>
		<li>Turn the 3-way valve of the buffer-inlet syringes to close the connection to the device, and connect the two syringes, S2 and S3. Disengage S2 from the syringe pump.</li>
		<li>Disconnect S2 from the valve and connect S5 to the valve. Push out all the air collected at the top of S5 to S3 through the valve until a little bit of the OP50 buffer goes into S3. In doing so, make sure that no air bubble remains in S5 or the valve.</li>
		<li>Turn the 3-way valve back to the original position to close the connection between S3 and S5 and connect S5 to the device. Load S5 onto the syringe pump. Turn on the orbital shaker that holds the pump. Set the shaking speed to approximately 200 rpm.</li>
		<li>Set the flow rate to be 100 &#181;L/min. The time it takes for the new buffer to arrive to the worms in the device depends on the length of the buffer-inlet syringe tube (around 5 minutes with the tubing described above). A higher flow rate can be employed if a quicker buffer exchange is required.</li>
		<li>Once the OP50 buffer spreads throughout the device, set the flow rate back to 3 &#181;L/min.</li>
	</ol>
	</li>
</ol>
4. Loading Worms into the Microfluidic Device
<ol>
	<li>Prepare a small low binding microcentrifuge tube (650 &#181;L) and fill it with 100 &#181;L of OP50 suspension. Transfer around 20 - 30 age-synchronized young adult worms (46 hours post L1 larval arrest at 25 &#176;C) from an NGM (nematode growth medium) agar plate to the tube by picking them one by one with a worm pick. Age-synchronization can be done following a standard protocol12.</li>
	<li>Fill the worm-inlet syringe and syringe tube (S4 in Figure 1) with the OP50 suspension. Inject ~500 &#181;L of the fluid into the microcentrifuge tube with worms. Draw the suspension with worms into the worm-inlet syringe tube. Make sure that the syringe tube is long enough (&#62;1 m) and that drawn worms stay in the syringe tube and do not make all the way to the syringe S4.</li>
	<li>Connect S4 to the worm-inlet. Inject all the worms in the worm-inlet syringe tube into the microfluidic device. Disconnect S4 and the syringe tube. Plug the worm-inlet with a pin.</li>
	<li>Disengage the buffer-inlet syringe S2 from the mechanical pump ( Figure 1) and manually control S1 and S2 to push all the worms into separate channels. Pressing S2 will move worms into the channels, while pressing S1 will move them in the reverse direction. Once a channel is loaded, the worm in each channel will block the entry of other worms.</li>
	<li>Let worms adjust to the new environment for 2 - 3 hours.</li>
</ol>
5. Setting Up an Imaging Protocol
<ol>
	<li>Find all worms that are securely located in the device, and set an imaging position for each of them in the image acquisition software that controls your microscope. Note that in some cases (e.g. when higher magnification is desired, or when using cameras with a smaller CCD sensor) multiple imaging positions are required for each channel. 
	NOTE: While this can be done manually, some acquisition software packages can load a text file that lists the positions where images should be taken. Since these positions are ordered regularly in WormSpa, a user can write a small script (e.g., in Python or commercial software) that automatically creates such a list. The precise format of this script would depend on the file format expected by the acquisition software (see an example in Supplementary Material 1).</li>
	<li>Set the required frequency of time-lapse imaging. For example, imaging of immune gene response to infection is done at a frequency of 10 minutes per frame. Make sure that all necessary channels (e.g. bright-field and the GFP fluorescence) are acquired at every time point.</li>
</ol>
6. Host Response to Pseudomonas Infection
Note: This step is specific for studying host-pathogen interactions. Alternatively, one can prepare a buffer that contains other environmental cues of interest (biotic and abiotic stressors, drugs, signaling molecules, etc.).
<ol>
	<li>Prepare Pseudomonas aeruginosa PA14 suspended in SK-medium14.
	<ol>
		<li>Inoculate a 250 mL flask with 50 mL of LB and a single colony, and incubate overnight at 37 &#176;C.</li>
		<li>Inoculate another 250 mL flask with 50 mL of LB and an aliquot of the culture, and incubate overnight at 37 &#176;C.</li>
		<li>Centrifuge the overnight culture at 4000 rpm for 10 minutes and resuspend the pellet in 10 mL of SK-medium. Measure the bacterial concentration and adjust the concentration to OD600 = 4.</li>
		<li>Transfer the suspension to a clean flask, and incubate at 37 &#176;C for 24 hours and at 25 &#176;C for another 24 hours while shaking. This step mimics the plate-incubation step in the standard killing assay14.</li>
		<li>Filter the suspension through a 5 &#181;m syringe filter. This is to prevent bacterial aggregates from clogging the device.</li>
	</ol>
	</li>
	<li>Replace the OP50 suspension in the device with the PA14 suspension, following the same procedure as in Step 3.2. Keep imaging for 10 hours.</li>
</ol>

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The authors have nothing to disclose.

Microfluidic tools provide multiple benefits in studying worms. Imaging in a PDMS device offers higher imaging quality as compared with a standard NGM agar plate. Multiple images can be taken from a single worm, in contrast with traditional methods in which animals are picked from the plate and mounted on a microscope slide for imaging. In addition, the microenvironment in which worms reside can be kept constant or modulated as desired, permitting precise mapping between the composition of the environment and the response of the animals.
Here we demonstrated how to perform a longitudinal measurement of host response to bacterial infection by imaging the response of multiple individual worms to the opportunistic pathogen P. aeruginosa PA14 using a custom microfluidic device (WormSpa). Both bright-field and fluorescence images were taken at multiple times without perturbing the ongoing experiment. In particular, the kinetics of egg-laying and the irg-1 gene expression were probed every 10 minutes, a significant increase in temporal resolution over traditional methods14,15.
In this device, longitudinal measurements of multiple worms can be performed by revisiting the saved location of each worm at any given time. This was demonstrated in Figures 2C and 3B, where we quantify the irg-1::GFP fluorescence and the eggs laid by individual worms. These figures show that worms display broad range of kinetic profiles. As previously shown in single-cell studies19,20,21, this individuality can be used to understand the design of genetic circuits, as well as the correlation between distinct pathways19,22.
For successful experiments, it is critical to load worms into the microfluidic device quickly. While worms crawl on agar surface, they tend to thrash in liquid environment, and this behavior induces a genetic program in which AMP activated protein kinases are involved23,24. To minimize this undesirable perturbation, it is important to place worms in their separate chambers quickly, as the microstructures in the chamber and the air permeability of PDMS provide worms with an environment that mimics solid surface.5 Although applying higher pressure helps loading worms faster, pressurizing the device too much could induce mechanical stress on the worms. Worms that are severely stressed do not feed or lay eggs, resulting in a clearly observable bagging phenotype25. After loading, worms are fed E. coli OP50 for 2-3 hours, allowing the worms to adjust to the microfluidic environment and providing the researcher an opportunity to observe the animals and discard those that were damaged.
The size and spacing of microstructures in each chamber are optimized for the worms grown for 46 hours from L1 arrest at 25 &#176;C. The structures may be scaled up to study older worms or reduced to examine L4 larvae which are smaller in size. However, worms are not able to successfully molt in the chamber, precluding study of post-embryonic development in this device. For such applications, methods such as WorMotel26 may be considered. Since worms are confined but not immobilized in WormSpa, a successful durable experiment depends on whether the animals stay in the chamber. We find that this can be challenging for early-stage larvae, for male adults, and for some hermaphrodite mutants.
Instead of using a single buffer inlet (at the center of the top, Figure 1), the two upside down balloon-shaped inlets can be used simultaneously. Filling the two syringes with different buffers, one can generate temporally structured environments by pressing on different syringes at different time. This, for example, allows study of adaptation to changing environment.27,28
As mentioned above, the microfluidic device described here can be used for a broad range of other applications. These include studying responses to other environmental cues. Such responses may include changes in gene expression and egg-laying behaviors, as exemplified here, but also changes in metabolism and fat accumulation (e.g. through fat staining by Nile Red), neuronal physiology and signaling (e.g. by calcium imaging and optogenetic perturbations), changes in motor behaviors such as pumping, defecation and head movement (via automated quantitative imaging of time-lapse sequences), and more.
Finally, this approach is largely automated and reduces the labor required for long experiments over many hours, even days. Once the locations of individual worms are saved in the data acquisition software, the experiment consists of automatic recording of images while the mechanical pump controller maintains the constant flux of buffer into the device. Importantly, fabrication of a WormSpa device does not require any more than standard soft lithography protocol, and its design and operation are simple enough even to researchers with no previous experience in microfluidics. With the various advantages discussed above and relatively simple preparation steps, this approach can be helpful to those who are considering longitudinal measurement of live worms under precisely controlled conditions.

Changes in environmental conditions may lead to activation of genetic programs accompanied by induction and repression of the expression of specific genes1,2. These kinetic changes may be variable among tissues in the same animals and between different animals. Studies of such genetic programs therefore call for methods that allow longitudinal imaging of individual animals and provide precise dynamical control of environmental conditions.
In recent years, microfabricated fluidic devices have been used to study many aspects of response and behavior in small animals, including worms, flies, water bears and more3,4,5,6,7. Applications include, for example, deep phenotyping, optogenetic recording of neuronal activity in response to chemical stimuli, and tracking of motor behaviors such as locomotion and pumping8,9,10,11.
Microfluidic-based approaches hold many properties that could benefit long-term longitudinal imaging of response to environmental cues, including precise dynamical control of the local microenvironment, flexible design that allows maintenance of individual animals in separate quarters, and favorable attributes for imaging. However, maintaining animals in a microfluidic chamber for a long time with minimal adverse impact on their well-beings is a challenge, which requires particular care in the design of the microfluidic device as well as in the execution of the experiment.
Here we demonstrate the use of WormSpa, a microfluidic device for longitudinal imaging of Caenorhabditis elegans.5 Individual worms are confined in chambers. A constant low flow of liquid and bacterial suspension guarantees that worms are well-fed and sufficiently active to maintain good health and alleviate stress, and the structure of the chambers allows worms to lay eggs. The simplicity of the design and operation of WormSpa should allow researchers with no previous experience in microfluidics to incorporate this device into their own research plans.

<table><tbody><tr><td>WormSpa</td><td>N/A</td><td>N/A</td><td>The CAD file for WormSpa is available from the Levine lab.</td></tr><tr><td>Compound Microscope</td><td>Zeiss</td><td>AxioObserver Z1</td><td>An inverted fluorescence microscope with a motorized stage</td></tr><tr><td>Syringe Pump</td><td>New Era Pump Systems</td><td>NE-501</td><td></td></tr><tr><td>Tubing</td><td>SCI Scientific Commodities Inc.</td><td>BB31695-PE/5</td><td>0.034&amp;rdquo; (0.86 mm) I.D. x 0.052&amp;rdquo; (1.32 mm) O.D</td></tr><tr><td>Syringe Tip</td><td>CMLsupply</td><td>901-20-050</td><td>20 Gauge x 1/2&amp;rdquo; blunt tip stainless steel canula</td></tr><tr><td>Syringe Filter</td><td>PALL</td><td>4650</td><td>Acrodisc 32 mm Syringe Filter with 5 um Supor Membrane</td></tr><tr><td>Syringe</td><td>Qosina</td><td>C3307</td><td>10 mL Male Luer Lock Syringe</td></tr><tr><td>3 Way Valve</td><td>ColeParmer</td><td>FF-30600-23</td><td>Large-bore 3-way, male-lock, stopcocks, 10/pack, Non-sterile</td></tr><tr><td>Dowel Pin</td><td>McMaster-Carr</td><td>90145A317</td><td>18-8 Stainless Steel Dowel Pins (1/32&quot; Dia. x 1/2&quot; Lg.)</td></tr><tr><td>Low Binding Microcentrifuge Tube</td><td>Corning</td><td>CL S3206</td><td>0.65 mL low binding snap cap microcentrifuge tube</td></tr></tbody></table>

a microfluidic platform for longitudinal imaging in caenorhabditis elegans

In the last decade, microfluidic techniques have been applied to study small animals, including the nematode Caenorhabditis elegans, and have proved useful as a convenient live imaging platform providing capabilities for precise control of experimental conditions in real time. In this article, we demonstrate live imaging of individual worms employing WormSpa, a previously-published custom microfluidic device. In the device, multiple worms are individually confined to separate chambers, allowing multiplexed longitudinal surveillance of various biological processes. To illustrate the capability, we performed proof-of-principle experiments in which worms were infected in the device with pathogenic bacteria, and the dynamics of expression of immune response genes and egg laying were monitored continuously in individual animals. The simple design and operation of this device make it suitable for users with no previous experience with microfluidic-based experiments. We propose that this approach will be useful for many researchers interested in longitudinal observations of biological processes under well-defined conditions.

Age-synchronized young adult worms (46 hours post L1 larval arrest at 25 &#176;C)12 were loaded in the device, as described in the protocol. The worms were individually located in separate channels, enabling longitudinal measurement of animals&#39; response to the pathogen. When the experiment is successful, most worms remain in their channels for the duration of the experiment. In this case, images of individual worms are taken simply by placing their channel in the imaging path, avoiding the need to actively locate the animal. Figure 2A shows the bright-field images of a single representative worm over the course of 10 hours in the device. While worms are confined within their channels, they can move up- and downstream, and can freely turn their heads. This is critical to guarantee that the device itself does not cause stress or harm the animals.
We monitored response of worms to bacterial infection by P. aeruginosa, one of the best-studied bacterial pathogens of C. elegans. It is also an opportunistic human pathogen which causes infection in cystic fibrosis and immunocompromised patients. Feeding worms with particular strains of P. aeruginosa, such as the clinical isolate PA14, leads to lethal intestinal infection, and the virulence factors involved in this process are also implicated in the infection of mammalian hosts.14,15
To examine the pattern of changes in gene expression during bacterial infection, fluorescent reporters can be employed. Among many genes that display dynamic response to infection, we monitored the transcriptional response of irg-1 (infection response gene 1), by imaging GFP fluorescence from transgenic worms expressing irg-1::GFP (AU0133 [agIs17(pirg-1::GFP; pmyo-2::mCherry)], obtained from the Ausubel lab). irg-1 is known to be strongly induced in the intestine of infected animals at the early stage of infection by P. aeruginosa PA14.16,17 Figure 2B shows time lapse images of a representative worm (the same animal as in Figure 2A). Each fluorescence image was taken immediately after the corresponding bright-field image. In the first 5 hours post infection, GFP fluorescence increases only mildly in the mid body and the tail. Then, a substantial increase in fluorescence is observed, starting from the mid body. The total fluorescence in each worm is plotted in Figure 2C as a function of time post infection. These data illustrate not only the previously-described induction of irg-1 upon PA14 infection, but also the worm-to-worm variability in the timing and extent of induction.
The design of the WormSpa microfluidic device includes filters which collect the eggs laid by each individual worm5. Once eggs hatch, the L1 larvae are washed through the filter and into the outlet tube. This allowed us to manually monitor the egg-laying kinetics of individual worms during bacterial infection. The time lapse images of the eggs laid by the same worm as in Figure 2 are shown in Figure 3A. These images were taken right after the corresponding bright-field images in Figure 2A. The cumulative number of eggs laid by each worm is shown as a function of time post infection in Figure 3B, and the average and standard deviation over the entire population are shown in Figure 3C. These data capture the animal-to-animal variability as well as the previously-described decline in egg-laying as infection progresses. In contrast, in uninfected animals egg-laying rate does not decline until it peaks at ~54 hours post L1 at 25 &#176;C5,18.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57348/57348fig1.jpg" /> 
Figure 1: Experimental scheme of live imaging of worms in a microfluidic device. Worms are loaded into the device via the worm inlet, and are subsequently pushed into separate channels. Bacterial suspension is injected into the device from the buffer inlet by a mechanical pump mounted on an orbital shaker. The buffer exits through the outlet port of the device. Four syringes (S1, S2/S5, S3, and S4) are used to operate the device. The red dashed lines are syringe tubes connecting syringes and the device. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/57348/57348fig1large.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/57348/57348fig2.jpg" /> 
Figure 2:&#160;Representative gene expression kinetics. Time lapse images of a representative worm taken by (A) Bright-field and (B) GFP fluorescence microscopy. The leftmost images were taken just before the pathogen was introduced, whereas the rightmost images were taken 10 hours post infection by PA14. The time interval between images is 1 hour. (C) The time course of total irg-1::GFP fluorescence in individual worms. Each row corresponds to a single animal. Rows were sorted in descending order of mean fluorescence. Fluorescence intensity is color-coded and the color scale is shown on the right. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/57348/57348fig2large.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/57348/57348fig3.jpg" /> 
Figure 3:&#160;Representative egg-laying kinetics. (A) Time lapse images of the number of eggs laid by the same worm shown in Figure 2A and B. The leftmost images were taken just before the pathogen was introduced, whereas the rightmost images were taken 10 hours post infection by PA14. The time interval between images is 1 hour. (B) The time course of cumulative number of eggs in individual worms. Each row corresponds to a single animal. Rows were sorted in descending order of the total number of eggs laid per worm. The number of eggs is color-coded and the color scale is shown on the right. (C) The average number of eggs laid per worm as a function of time. The gray area indicates &#177; one standard deviation. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/57348/57348fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Supplementary Material 1" src="/files/ftp_upload/57348/57348supfig.jpg" /> 
Supplementary Material 1: A sample script that creates a text file containing the imaging positions for all channels in WormSpa device. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/57348/Supp1.EditZeissTiles.m" target="_blank">Please click here to download this file.</a>

Watch this Scientific Journal Video about A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans at JoVE.com

A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

In this article, we demonstrate live imaging of individual worms employing a custom microfluidic device. In the device, multiple worms are individually confined to separate chambers, allowing multiplexed longitudinal surveillance of various biological processes.

A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

In the last decade, microfluidic techniques have been applied to study small animals, including the nematode Caenorhabditis elegans, and have proved ...

a-microfluidic-platform-for-longitudinal-imaging-caenorhabditis

Universidad San Sebastian

Research

JoVE Journal

Bioengineering

6.4K Views.  Kongju National University. In this article, we demonstrate live imaging of individual worms employing a custom microfluidic device. In the device, multiple worms are individually confined to separate chambers, allowing multiplexed longitudinal surveillance of various biological processes.

Video: A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

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

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are available for microfluidic devices using soft lithography techniques 1-3. Microfluidic devices have been used for sub-cellular imaging 4,5, in vivo laser microsurgery 2,6 and cellular imaging 4,7. In vivo imaging requires immobilization of organisms. This has been achieved using suction 5,8, tapered channels 6,7,9, deformable membranes 2-4,10, suction with additional cooling 5, anesthetic gas 11, temperature sensitive gels 12, cyanoacrylate glue 13 and anesthetics such as levamisole 14,15. Commonly used anesthetics influence synaptic transmission 16,17 and are known to have detrimental effects on sub-cellular neuronal transport 4. In this study we demonstrate a membrane based poly-dimethyl-siloxane (PDMS) device that allows anesthetic free immobilization of intact genetic model organisms such as Caenorhabditis elegans (C. elegans), Drosophila larvae and zebrafish larvae. These model organisms are suitable for in vivo studies in microfluidic devices because of their small diameters and optically transparent or translucent bodies. Body diameters range from ~10 &mu;m to ~800 &mu;m for early larval stages of C. elegans and zebrafish larvae and require microfluidic devices of different sizes to achieve complete immobilization for high resolution time-lapse imaging. These organisms are immobilized using pressure applied by compressed nitrogen gas through a liquid column and imaged using an inverted microscope. Animals released from the trap return to normal locomotion within 10 min.
We demonstrate four applications of time-lapse imaging in C. elegans namely, imaging mitochondrial transport in neurons, pre-synaptic vesicle transport in a transport-defective mutant, glutamate receptor transport and Q neuroblast cell division. Data obtained from such movies show that microfluidic immobilization is a useful and accurate means of acquiring in vivo data of cellular and sub-cellular events when compared to anesthetized animals (Figure 1J and 3C-F 4).
Device dimensions were altered to allow time-lapse imaging of different stages of C. elegans, first instar Drosophila larvae and zebrafish larvae. Transport of vesicles marked with synaptotagmin tagged with GFP (syt.eGFP) in sensory neurons shows directed motion of synaptic vesicle markers expressed in cholinergic sensory neurons in intact first instar Drosophila larvae. A similar device has been used to carry out time-lapse imaging of heartbeat in ~30 hr post fertilization (hpf) zebrafish larvae. These data show that the simple devices we have developed can be applied to a variety of model systems to study several cell biological and developmental phenomena in vivo.

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

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.

Micro fabricated fluidic devices provide an accessible micro-environment for in vivo studies on small organisms. Simple fabrication processes are ...

Microfluidic-based Electrotaxis for On-demand Quantitative Analysis of Caenorhabditis elegans' Locomotion

The nematode Caenorhabditis elegans is a versatile model organism for biomedical research because of its conservation of disease-related genes and pathways as well as its ease of cultivation. Several C. elegans disease models have been reported, including neurodegenerative disorders such as Parkinson's disease (PD), which involves the degeneration of dopaminergic (DA) neurons 1. Both transgenes and neurotoxic chemicals have been used to induce DA neurodegeneration and consequent movement defects in worms, allowing for investigations into the basis of neurodegeneration and screens for neuroprotective genes and compounds 2,3.
Screens in lower eukaryotes like C. elegans provide an efficient and economical means to identify compounds and genes affecting neuronal signaling. Conventional screens are typically performed manually and scored by visual inspection; consequently, they are time-consuming and prone to human errors. Additionally, most focus on cellular level analysis while ignoring locomotion, which is an especially important parameter for movement disorders.
We have developed a novel microfluidic screening system (Figure 1) that controls and quantifies C. elegans' locomotion using electric field stimuli inside microchannels. We have shown that a Direct Current (DC) field can robustly induce on-demand locomotion towards the cathode ("electrotaxis") 4. Reversing the field's polarity causes the worm to quickly reverse its direction as well. We have also shown that defects in dopaminergic and other sensory neurons alter the swimming response 5. Therefore, abnormalities in neuronal signaling can be determined using locomotion as a read-out. The movement response can be accurately quantified using a range of parameters such as swimming speed, body bending frequency and reversal time.
Our work has revealed that the electrotactic response varies with age. Specifically, young adults respond to a lower range of electric fields and move faster compared to larvae 4. These findings led us to design a new microfluidic device to passively sort worms by age and phenotype 6.
We have also tested the response of worms to pulsed DC and Alternating Current (AC) electric fields. Pulsed DC fields of various duty cycles effectively generated electrotaxis in both C. elegans and its cousin C. briggsae 7. In another experiment, symmetrical AC fields with frequencies ranging from 1 Hz to 3 KHz immobilized worms inside the channel 8.
Implementation of the electric field in a microfluidic environment enables rapid and automated execution of the electrotaxis assay. This approach promises to facilitate high-throughput genetic and chemical screens for factors affecting neuronal function and viability.

Microfluidic-based Electrotaxis for On-demand Quantitative Analysis of Caenorhabditis elegans' Locomotion

A semi-automated micro-electro-fluidic method to induce on-demand locomotion in Caenorhabditis elegans is described. This method is based on the neurophysiologic phenomenon of worms responding to mild electric fields (&#x201C;electrotaxis&#x201D;) inside microfluidic channels. Microfluidic electrotaxis serves as a rapid, sensitive, low-cost, and scalable technique to screen for factors affecting neuronal health.

The nematode Caenorhabditis elegans is a versatile model organism for biomedical research because of its conservation of disease-related genes 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 ...

Developmental Biology

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

C. elegans is the premier system for the systematic analysis of cell fate specification and morphogenetic events during embryonic development. One challenge is that embryogenesis dynamically unfolds over a period of about 13 h; this half day-long timescale has constrained the scope of experiments by limiting the number of embryos that can be imaged. Here, we describe a semi-high-throughput protocol that allows for the simultaneous 3D time-lapse imaging of development in 80&#8211;100 embryos at moderate time resolution, from up to 14 different conditions, in a single overnight run. The protocol is straightforward and can be implemented by any laboratory with access to a microscope with point visiting capacity. The utility of this protocol is demonstrated by using it to image two custom-built strains expressing fluorescent markers optimized to visualize key aspects of germ-layer specification and morphogenesis. To analyze the data, a custom program that crops individual embryos out of a broader field of view in all channels, z-steps, and timepoints and saves the sequences for each embryo into a separate tiff stack was built. The program, which includes a user-friendly graphical user interface (GUI), streamlines data processing by isolating, pre-processing, and uniformly orienting individual embryos in preparation for visualization or automated analysis. Also supplied is an ImageJ macro that compiles individual embryo data into a multi-panel file that displays maximum intensity fluorescence projection and brightfield images for each embryo at each time point. The protocols and tools described herein were validated by using them to characterize embryonic development following knock-down of 40 previously described developmental genes; this analysis visualized previously annotated developmental phenotypes and revealed new ones. In summary, this work details a semi-high-throughput imaging method coupled with a cropping program and ImageJ visualization tool that, when combined with strains expressing informative fluorescent markers, greatly accelerates experiments to analyze embryonic development.

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

This work describes a semi-high-throughput protocol that allows simultaneous 3D time-lapse imaging of embryogenesis in 80&#8211;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.

C. elegans is the premier system for the systematic analysis of cell fate specification and morphogenetic events during embryonic development. One ...

Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans

Behavior is controlled by the nervous system. Calcium imaging is a straightforward method in the transparent nematode Caenorhabditis elegans to measure the activity of neurons during various behaviors. To correlate neural activity with behavior, the animal should not be immobilized but should be able to move. Many behavioral changes occur during long time scales and require recording over many hours of behavior. This also makes it necessary to culture the worms in the presence of food. How can worms be cultured and their neural activity imaged over long time scales? Agarose Microchamber Imaging (AMI) was previously developed to culture and observe small larvae and has now been adapted to study all life stages from early L1 until the adult stage of C. elegans. AMI can be performed on various life stages of C. elegans. Long-term calcium imaging is achieved without immobilizing the animals by using short externally triggered exposures combined with an electron multiplying charge-coupled device (EMCCD) camera recording. Zooming out or scanning can scale up this method to image up to 40 worms in parallel. Thus, a method is described to image behavior and neural activity over long time scales in all life stages of C. elegans.

Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans

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.

Behavior is controlled by the nervous system. Calcium imaging is a straightforward method in the transparent nematode Caenorhabditis elegans to measure ...

Neuroscience

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

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

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

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

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

Biology

Using an Adapted Microfluidic Olfactory Chip for the Imaging of Neuronal Activity in Response to Pheromones in Male C. Elegans Head Neurons

The use of calcium indicators has greatly enhanced our understanding of neural dynamics and regulation. The nematode Caenorhabditis elegans, with its completely mapped nervous system and transparent anatomy, presents an ideal model for understanding real-time neural dynamics using calcium indicators. In combination with microfluidic technologies and experimental designs, calcium-imaging studies using these indicators are performed in both free-moving and trapped animals. However, most previous studies utilizing trapping devices, such as the olfactory chip described in Chronis et al., have devices designed for use in the more common hermaphrodite, as the less common male is both morphologically and structurally dissimilar. An adapted olfactory chip was designed and fabricated for increased efficiency in male neuronal imaging with using young adult animals. A turn was incorporated into the worm loading port to rotate the animals and to allow for the separation of the individual neurons within a bilateral pair in 2D imaging. Worms are exposed to a controlled flow of odorant within the microfluidic device, as described in previous hermaphrodite studies. Calcium transients are then analyzed using the open-source software ImageJ. The procedure described herein should allow for an increased amount of male-based C. elegans calcium imaging studies, deepening our understanding of the mechanisms of sex-specific neuronal signaling.

Using an Adapted Microfluidic Olfactory Chip for the Imaging of Neuronal Activity in Response to Pheromones in Male C. Elegans Head Neurons

The use of an adapted &#34;olfactory chip&#34; for the efficient calcium imaging of C. elegans males is described here. Studies of male exposure to glycerol and a pheromone are also shown.

The use of calcium indicators has greatly enhanced our understanding of neural dynamics and regulation. The nematode Caenorhabditis elegans, with its ...

Using a Microfluidics Device for Mechanical Stimulation and High Resolution Imaging of C. elegans

One central goal of mechanobiology is to understand the reciprocal effect of mechanical stress on proteins and cells. Despite its importance, the influence of mechanical stress on cellular function is still poorly understood. In part, this knowledge gap exists because few tools enable simultaneous deformation of tissue and cells, imaging of cellular activity in live animals, and efficient restriction of motility in otherwise highly mobile model organisms, such as the nematode Caenorhabditis elegans. The small size of C. elegans makes them an excellent match to microfluidics-based research devices, and solutions for immobilization have been presented using microfluidic devices. Although these devices allow for high-resolution imaging, the animal is fully encased in polydimethylsiloxane (PDMS) and glass, limiting physical access for delivery of mechanical force or electrophysiological recordings. Recently, we created a device that integrates pneumatic actuators with a trapping design that is compatible with high-resolution fluorescence microscopy. The actuation channel is separated from the worm-trapping channel by a thin PDMS diaphragm. This diaphragm is deflected into the side of a worm by applying pressure from an external source. The device can target individual mechanosensitive neurons. The activation of these neurons is imaged at high-resolution with genetically-encoded calcium indicators. This article presents the general method using C. elegans strains expressing calcium-sensitive activity indicator (GCaMP6s) in their touch receptor neurons (TRNs). The method, however, is not limited to TRNs nor to calcium sensors as a probe, but can be expanded to other mechanically-sensitive cells or sensors.

Using a Microfluidics Device for Mechanical Stimulation and High Resolution Imaging of C. elegans

New tools for mechanobiology research are needed to understand how mechanical stress activates biochemical pathways and elicits biological responses. Here, we showcase a new method for selective mechanical stimulation of immobilized animals with a microfluidic trap allowing high-resolution imaging of cellular responses.

One central goal of mechanobiology is to understand the reciprocal effect of mechanical stress on proteins and cells. Despite its importance, the ...

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

Caenorhabditis elegans (C. elegans) have proved to be a valuable model system for studying developmental and cell biological processes. Understanding these biological processes often requires long-term and repeated imaging of the same animal. Long recovery times associated with conventional immobilization methods done on agar pads have detrimental effects on animal health making it inappropriate to repeatedly image the same animal over long periods of time. This paper describes a microfluidic chip design, fabrication method, on-chip C. elegans culturing protocol, and three examples of long-term imaging to study developmental processes in individual animals. The chip, fabricated with polydimethylsiloxane and bonded on a cover glass, immobilizes animals on a glass substrate using an elastomeric membrane that is deflected using nitrogen gas. Complete immobilization of C. elegans enables robust time-lapse imaging of cellular and sub-cellular events in an anesthetic-free manner. A channel geometry with a large cross-section allows the animal to move freely within two partially sealed isolation membranes permitting growth in the channel with a continuous food supply. Using this simple chip, imaging of developmental phenomena such as neuronal process growth, vulval development, and dendritic arborization in the PVD sensory neurons, as the animal grows inside the channel, can be performed. The long-term growth and imaging chip operates with a single pressure line, no external valves, inexpensive fluidic consumables, and utilizes standard worm handling protocols that can easily be adapted by other laboratories using C. elegans.

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

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.

Caenorhabditis elegans (C. elegans) have proved to be a valuable model system for studying developmental and cell biological processes. Understanding ...

Long-Term Culture of Individual Caenorhabditis elegans on Solid Media for Longitudinal Fluorescence Monitoring and Aversive Interventions

Caenorhabditis elegans are widely used to study aging biology. The standard practice in C. elegans aging studies is to culture groups of worms on solid nematode growth media (NGM), allowing the efficient collection of population-level data for survival and other physiological phenotypes, and periodic sampling of subpopulations for fluorescent biomarker quantification. Limitations to this approach are the inability to (1) follow individual worms over time to develop age trajectories for phenotypes of interest and (2) monitor fluorescent biomarkers directly in the context of the culture environment. Alternative culture approaches use liquid culture or microfluidics to monitor individual animals over time, in some cases including fluorescence quantification, with the tradeoff that the culture environment is contextually distinct from solid NGM. The WorMotel is a previously described microfabricated multi-well device for culturing isolated worms on solid NGM. Each worm is maintained in a well containing solid NGM surrounded by a moat filled with copper sulfate, a contact repellent for C. elegans, allowing longitudinal monitoring of individual animals. We find copper sulfate insufficient to prevent worms from fleeing when subjected to aversive interventions common in aging research, including dietary restriction, pathogenic bacteria, and chemical agents that induce cellular stress. The multi-well devices are also molded from polydimethylsiloxane, which produces high background artifacts in fluorescence imaging. This protocol describes a new approach for culturing isolated roundworms on solid NGM using commercially available polystyrene microtrays, originally designed for human leukocyte antigen (HLA) typing, allowing the measurement of survival, physiological phenotypes, and fluorescence across the lifespan. A palmitic acid barrier prevents worms from fleeing, even in the presence of aversive conditions. Each plate can culture up to 96 animals and easily adapts to a variety of conditions, including dietary restriction, RNAi, and chemical additives, and is compatible with automated systems for collecting lifespan and activity data.

Long-Term Culture of Individual Caenorhabditis elegans on Solid Media for Longitudinal Fluorescence Monitoring and Aversive Interventions

Here, we present a protocol to culture isolated individual nematodes on solid media for lifelong physiological parameter tracking and fluorescence quantification. This culture system includes a palmitic acid barrier around single-worm wells to prevent animals from fleeing, allowing the use of aversive interventions, including pathogenic bacteria and chemical stressors.

Caenorhabditis elegans are widely used to study aging biology. The standard practice in C. elegans aging studies is to culture groups of worms on solid ...

A Microfluidic Platform for Longitudinal Imaging in Caenorhabditis elegans

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Chapters in this video

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

Microfluidic-based Electrotaxis for On-demand Quantitative Analysis of Caenorhabditis elegans' Locomotion

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

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

Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans

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

Using an Adapted Microfluidic Olfactory Chip for the Imaging of Neuronal Activity in Response to Pheromones in Male C. Elegans Head Neurons

Using a Microfluidics Device for Mechanical Stimulation and High Resolution Imaging of C. elegans

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

Long-Term Culture of Individual Caenorhabditis elegans on Solid Media for Longitudinal Fluorescence Monitoring and Aversive Interventions