The Max Planck Society and a G&#246;ttingen Graduate School Junior Group stipend to H.B. funded this work.

1. Instruments, Culture Media, and Dishes
<ol>
	<li>Use a microscope that is capable of keeping the sample in focus and that is equipped with an automatic stage. Build a custom-made lid heater or purchase a commercial solution. Set up an LED-EMCCD camera system in which the exposure of the camera triggers LED illumination through a TTL signal using manufacturer&#39;s instructions. 
	Note: See Discussion for details.</li>
	<li>Polydimethylsiloxan (PDMS) stamps:
	<ol>
		<li>Fabricate PDMS stamps in a microfluidics facility or have the PDMS stamps produced by a commercial foundry. To use a commercial foundry, send an autocad file to the company and specify the depth (15 &#181;m, for example) of the device. After delivery from the foundry, cut the PDMS chip into its 16 stamps using a scalpel.</li>
		<li>Bond each individual stamp to a glass slide using air plasma. For bonding, expose both the PDMS stamp and the glass slide to air plasma for about 1 min (using the highest plasma settings at 0.5 mbar). Ensure that the surfaces for bonding face upwards during plasma treatment. Then, place the PDMS stamp onto the glass slide.</li>
	</ol>
	</li>
	<li>Take a 3.5 cm bottom dish and cut out a square area of 18 x 18 mm from the center of the bottom of the dish using a vertical milling machine and sharp rotary cutters. Use low rotation speed of the cutter and slow feeding to prevent the plastic from melting due to heat produced by friction. Prepare several dishes at once. 
	Note: The bottom dish can be reused many times after the experiment if it is cleaned after soaking in pure ethanol O/N.</li>
	<li>Agarose:
	<ol>
		<li>Dissolve 3 g high-melting point agarose in 100 ml S-Basal (5.85 g NaCl, 1 g K2HPO4, 6 g KH2PO4, 1 ml cholesterol (5 mg/ml in ethanol), H2O to 1 L, sterilize by autoclaving) by boiling. Make aliquots of the dissolved agarose in 2 ml Eppendorf tubes and store. Also, prepare a batch of low-melting point agarose exactly the same way as the high-melting point agarose.</li>
		<li>Prior to use, place 3 aliquots of high-melting point agarose onto a heating block at 95 - 98 &#176;C. Prior to use, place one aliquot of low-melting point agarose onto a heating block at 95 - 98 &#176;C until it has melted and then onto a heating block at 30 - 35 &#176;C.</li>
	</ol>
	</li>
</ol>
2. Selection of Animals
<ol>
	<li>Grow worms at a low density on seeded NGM plates to obtain clean animals. Make sure that there is plenty of food and only few animals on the plate.</li>
	<li>For imaging L1 larvae, transfer about 30 eggs containing embryos at the pretzel stage onto a fresh seeded NGM plate. For transferring later larval stages or adult C. elegans, transfer about 30 worms onto a fresh seeded NGM plate.</li>
</ol>
3. Preparation of Agarose Microchambers
<ol>
	<li>Preparation of the dish:
	<ol>
		<li>Take a plastic dish with a square opening of 18 x 18 mm at the bottom and place it upside down with the opening facing upwards. Close the opening by placing a piece of double-sided sticky tape of 20 x 20 mm onto the opening. Turn the dish around so that the sticky tape is on the bottom and place the dish onto a hard surface. Cut the opening free using a scalpel.</li>
		<li>Using a P1000 pipette, fill 2 ml of 3% high-melting point agarose in S-Basal into the dish. Place the agarose onto the area that surrounds the opening and let solidify. Wait until the agarose is solid.</li>
		<li>Turn around the dish and peel off the protective film that covers the double-sided sticky tape so that the sticky side will remain on the dish and will be exposed. 
		Note: As a result, a fine ring of sticky tape will surround the outside of the opening. The agarose serves as a moisture reservoir that will later surround the sample.</li>
	</ol>
	</li>
	<li>Casting of microchambers:
	<ol>
		<li>Expose the PDMS surface for molding with air plasma for 20 - 60 sec. 
		Note: This plasma treatment renders the PDMS surface hydrophilic, which prevents the trapping of air bubbles and produces sharper prints.</li>
		<li>Construct two spacers of equal height by stacking 5 - 9 glass slides. Place, in parallel to their long sides, the first spacer stack, then a single glass slide, then again a spacer stack. Place the glass slide that contains the PDMS stamp orthogonally across the spacers. Adjust the height of the spacers so that there is a space of about 1.5 mm between the molding surface of the PDMS stamp and the single glass slide.</li>
		<li>Place a drop of hot liquid high-melting point agarose onto the single glass slide near the PDMS stamp and quickly slide the PDMS stamp vertically into the liquid agarose. Let the agarose solidify. Check that it gets an opaque appearance, which usually takes about 2 min. Pull off the stamp vertically with one move. 
		Note: It is convenient to glue the spacer slides together with double-sided sticky tape. The vertical movement of the stamp prevents air bubbles from getting trapped in the agarose.</li>
	</ol>
	</li>
	<li>Transfer the eggs or worms together with OP50 bacteria onto the agarose using a fine platinum wire pick. Distribute one egg or one worm per chamber together with food using an eyelash. Fill around 30 eggs onto one agarose pad.</li>
	<li>Cut the agarose slab containing the filled microchambers into a square of about 15 x 15 mm so that it fits nicely into the opening of the dish. Pick up the square agarose slab with forceps and place it upside down onto a glass coverslip of 20 x 20 mm. Once dropped, do not lift it up again or slide it around because this may cause the bacteria and worms to be pushed out of their chambers.</li>
	<li>Assembly of the dish:
	<ol>
		<li>Place the glass coverslip onto the opening of the plastic dish. Gently press down the glass coverslip onto the ring made of double-sided sticky tape. Take care not to break the glass.</li>
		<li>Turn the dish upside down and use a P1000 pipette to fill the gap between the agar slab containing the microchambers and the agarose reservoir with liquid low-melting point agarose cooled to about 30 &#176;C. Wait until the agarose has solidified.</li>
		<li>Seal the dish with a lid. For inverted microscopes use a heated lid. For upright microscopes use a normal lid and seal the dish with Parafilm.</li>
	</ol>
	</li>
	<li>After finishing the preparation of microchambers, check them under a stereomicroscope. The correct filling is crucial. See the Discussion for details.</li>
</ol>
4. Calcium Imaging
<ol>
	<li>Use transgenic strains expressing genetically encoded calcium sensors such as HBR16 (goeIs5[pnmr-1::SL1-GCaMP3.35-SL2::unc-54-3&#39;UTR, unc-119(+)])27.</li>
	<li>Use a compound microscope equipped for wide-field epifluorescence. Connect the TTL output of the EMCCD camera to the TTL input of the LED, so that each time the camera records a frame the sample will be illuminated. Use an exposure time of about 5 msec. Use EM gain in the range of 50 - 300.</li>
	<li>Specify a burst movie running for 24 hr with each worm being imaged every 15 - 30 min first for 20 sec with DIC, then for 20 sec with a GFP fluorescence to record GCaMP, and then take a final image of the mKate2 signal is taken to control for expression levels. Use a frame rate of 2/sec during each burst.</li>
	<li>For visual data inspection, use a false-color map to enhance the visibility of small changes in fluorescence intensity. Plot fluorescent data as &#916;F/F, with F being the average baseline value of fluorescence. A detailed description of calcium data analysis can be found in the literature20.</li>
</ol>
5. Parallel AMI of Multiple Worms
<ol>
	<li>Place the dish containing the microchambers onto the microscope, focus on the sample and engage the autofocus. Set up a software protocol so that the camera acquires a burst of 40 image frames in 20 sec every half an hour for 24 hr, which will result in 1,920 frames per worm, a reasonable amount of data. Set up the scan so that it visits each worm using the stage. Aim to film about 30 worms in one run.</li>
	<li>Image multiple worms by zooming out, i.e., using a lower magnification. Use a lower magnification to cover several microchambers with the camera chip and film several adjacent microchambers simultaneously. After the end of image acquisition, separate the data for each individual chamber by cropping a region of interest covering one animal.</li>
	<li>To quickly assess mobility data, use frame subtraction28-32.</li>
</ol>
6. Imaging Different Life Stages of C. elegans
<ol>
	<li>Use agarose microchambers for all life stages of C. elegans from L1 to adult and including dauers. Use the appropriate chamber dimensions for different life stages that are shown in Table 1.</li>
</ol>
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Life stage</td>
			<td>Chamber size</td>
			<td>Chamber depth</td>
			<td>Typical recording time</td>
			<td>Magnification</td>
		</tr>
		<tr>
			<td>L1</td>
			<td>190 x 190 &#956;m</td>
			<td>10 - 15 &#956;m</td>
			<td>egg - mid L2</td>
			<td>400X</td>
		</tr>
		<tr>
			<td>L2</td>
			<td>370 x 370 &#956;m</td>
			<td>15 &#956;m</td>
			<td>egg - L3</td>
			<td>200X</td>
		</tr>
		<tr>
			<td>Dauer larva</td>
			<td>370 x 370 &#956;m</td>
			<td>25 &#956;m</td>
			<td>4 days</td>
			<td>200X</td>
		</tr>
		<tr>
			<td>L3</td>
			<td>700 x 700 &#956;m</td>
			<td>45 &#956;m</td>
			<td>L2 - L4</td>
			<td>200X</td>
		</tr>
		<tr>
			<td>L4</td>
			<td>700 x 700 &#956;m</td>
			<td>45 &#956;m</td>
			<td>young L4 - young adult</td>
			<td>100X</td>
		</tr>
		<tr>
			<td>Young adult</td>
			<td>700 x 700 &#956;m</td>
			<td>45 &#956;m</td>
			<td>12 - 24 hr</td>
			<td>100X</td>
		</tr>
	</tbody>
</table>
Table 1. Chamber sizes for different life stages. Shown are chamber dimensions and magnifications that are useful for various stages optimized for a 8 x 8 mm camera chip.

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The authors declare that they have no competing financial interests.

Hardware
The focus of a microscope will typically drift during long-term image acquisition. For compound microscopes, focus-keeping systems can be purchased from the major microscope manufacturers. In case a compound microscope with focus control is too pricy, a simple alternative would be to use a stereomicroscope. Compound microscopes allow the use of objectives with high numerical aperture (NA) and can be easily automated. 40X oil objectives are well suited. Water immersion lenses are not ideal because of evaporation during long-term imaging. Differential interference contrast (DIC) generates a nice contrast that helps to follow morphological and behavioral changes, but simple bright field imaging can also be used. For DIC or bright field imaging, use red light by placing a red light filter into the dia-illumination path of the microscope. For scanning of several microchambers an automated stage is necessary. Best performance is achieved when a stage is used that has nonlinear acceleration and deceleration and can be set to low scan speed to prevent disturbing the animals during scanning. We have not observed behavioral responses or calcium increase in mechanosensitive neurons (ALM and PLM) during scanning, suggesting that slow scanning indeed does not activate the mechanosensitive system of the worm (data not shown). Commercial LED systems can be used. Several companies offer ready-to-use solutions that include the LEDs at different wavelengths. The LED should have the option of externally triggering the LED with a TTL signal. A highly sensitive camera is needed for calcium imaging of moving animals. EMCCD cameras are the most sensitive cameras on the market. The camera needs to have a TTL output during exposure (also called &#8220;fire&#8221; output). A lid heater is required to prevent condensation on the lid if using an inverted microscope. On an upright microscope, the dish will be placed so that the lid will be at the bottom and thus condensation is prevented and no lid heating is required. The lid should tightly close the dish to prevent evaporation of water during long-term imaging.
PDMS stamps
The surface of the PDMS stamp that contains the structure for molding the agarose is faced away from the glass slide, which supports the PDMS stamp. A list with companies that offer custom microfluidic chips can be found on Wikipedia (<a href="http://en.wikipedia.org/wiki/List_of_microfluidics_related_companies" target="_blank">http://en.wikipedia.org/wiki/List_of_microfluidics_related_companies</a>).
Filling the nematodes into their chambers
This is the most critical step in the protocol and the following points should be considered: A) The moistness of the agarose is crucial for transferring the worms and for imaging. If the agarose is too moist, i.e., there is a liquid covering an area of several microchambers, it will not be possible to distribute bacterial food and worms in a controlled manner because the liquid will make the bacteria flow away. If the agarose is too dry then the bacteria and worms may not get off the pick easily which means that increased force needs to be used to drop the worms and bacteria which easily causes damage to the agarose. When filling in a lot of worms the agarose may dry up. In the worst case the agarose will be so dry in the end that the chambers will collapse. If the agarose is too dry it can be rehydrated by placing a small drop (about 2 &#181;l) of S-Basal onto the side of the chip where there are no worms. Then dip the platinum wire pick into the liquid and even pull some of the liquid into the area where the chambers are filled with worms. B) The amount of food is critical for successful long-term imaging. If there is not enough food, the worms may run out of it. If there is excessive food the cavity of the chamber will not behave like a liquid but rather like a solid and will allow worms to escape from the chamber by pushing off the bacteria. Aim to obtain a bacterial suspension that fills the entire chamber. The worms are in constant physical contact with the agar and glass surface along most of their length during the experiment and the crawling behavior appears to be similar to movement on a plate and dissimilar to thrashing in liquid. C) Mechanical damaging of the agarose can ruin the experiment. A fine pick is essential. It should not have sharp corners. A soft eyelash attached to a pipet tip can be used to move bacteria or worms into the chambers instead of using a platinum pick. In most cases, however, overdried agarose is the reason for damage. The pick or eyelash should, ideally, only barely touch the agarose itself, and the water film on the agarose surface should pull off worms and bacteria. When sealing the chambers, again, the moisture of the agarose is essential. There should not be any free liquid on the agarose surface because this may wash away bacteria and worms during sealing. Large bubbles may be removed by gently lifting a corner of the agar slab. Small bubbles that are smaller than the chamber often get trapped inside the chambers. These bubbles are not a problem and will disappear by absorption. After assembling the dish, check again that the agarose is not too dry or too wet. The chambers should be nicely sealed and there should not be any flow of liquid between chambers. If the agarose is too wet, the chambers will not seal properly. Worms may escape or their food will be washed away. If the sample is too moist, simple open the lid and let the agarose dry for a minute or two. If the agarose is too dry, the chambers may collapse and the worms will escape.
Scaling up by zooming out
For fluorescence imaging, zooming out is limited by the low amount of light obtained at lower magnifications. Also, EMCCD cameras are optimized for sensitivity and often have a relatively low resolution. However, scaling up imaging four-fold is well possible. For instance, four chambers of 190 &#181;m x 190 &#181;m can fit onto one frame when using 140X magnification (achieved by using a 20X Objective and a 0.7X camera mount) and a 512 x 512 pixel, 8 x 8 mm camera. A high-resolution camera with a large chip (such as sCMOS cameras, 16.6 mm x 14 mm chip, 2560 x 2560 pixels = 5.5 Megapixel) optimizes scaling up of DIC and bright field imaging. For instance, up to 30 L1 worms in 190 &#181;m x 190 &#181;m chambers can fit onto each frame of this camera when using a 100x magnification (see Figure 3). In principle, zooming out and scanning can also be combined to obtain even greater numbers of animals. Most neurons stay quite well in focus, so that only one focal plane needs to be imaged. If neurons are found to move out of the focus, a z scan using a piezo drive can be taken at each time point.
Adaptation to different behaviors
This protocol gives a good idea of the behavior across long time scales. Obviously, the timing of the bursts needs to be adapted to different behaviors and life stages.
Limitations of the technique
Several factors limit the duration of imaging. The most important is the amount of food. Once the food is consumed, larvae stop developing. Thus, in small chambers (190 x 190 &#181;m) worms develop until the L3 stage and then arrest. If longer imaging time is required, larger chambers have to be used. The maximum duration of long-term imaging is in the range of 2.5 - 3 days. If longer imaging is required, the worms need to be recovered and placed into new chambers. When imaging adult worms, another limitation is caused by the offspring of these worms. Adult worms lay eggs from which larvae hatch. These larvae will also stay inside the chamber, consume food, and may disturb the image analysis. If offspring is a problem, a solution is to either use sterile adults or to repeatedly place the worms into fresh chambers. To recover worms, the agarose slab containing the chamber is cut free with a scalpel, is pulled off the coverslip, and is placed onto an NGM plate from which the worms can be recovered. Another limit is the restriction of the worms to relatively small areas. This may be a problem if long-range movement needs to be analyzed. While animals in the chambers can be stimulated mechanically and optogenetically21,27,34,35, the sealed nature of the chamber will make it difficult to apply soluble or volatile stimulants. Biologically important gases such as oxygen or carbon dioxide can diffuse freely in the agar. The large air reservoir in the dish should keep the gas concentrations in the chambers constant over the time needed for experiments. However, it should be kept in mind that the local oxygen concentration in the chamber may resemble more the conditions found in liquid culture than culture on the plate.
Significance with respect to existing methods
Microfluidic devices have greatly advanced behavioral and developmental studied in C. elegans. Often, microfluidic structures are made of PDMS12. Here we describe a protocol for generating microfluidic culture chambers made from agarose. The strength of this technique is the combination of high imaging quality, correlation of behavior with physiological measurements, long-term imaging, and a reasonably high throughput. High image quality is achieved by imaging through the glass coverslip using high NA objectives. As a result, fluorescence imaging such as calcium imaging and confocal imaging of subcellular structures can be performed. Because the animals are not immobilized like in other systems, it allows a correlation of behavior with physiological measurements. Because the animals have ample food, they continue developing allowing long-term imaging. This system can image many worms in one run because animals are restricted to their defined chambers. Thus, this method can be easily scaled up9,27,34-36.
Future applications
So far, this system has been used mainly to study sleep behavior in C. elegans L1 larvae. However, the adaptation to all stages will make it possible to study a wide range of behaviors also in dauers and adults. A wide array of behaviors can be studied with this technique ranging from mating to egg laying.

Caenorhabditis elegans has been established as a model system to study behavior1. Due to its amenability to genetics, the molecular and cellular mechanisms underlying behavior can be studied. Many behaviors occur on long time scales. Two principal approaches can be used to observe motile animals over long time scales. The first approach is following the animal during movement using an automated stage or camera2-8 and the second is to restrict the movement to a range that is at least as small as the field of view of the camera9-14. Both methods have their advantages. Tracking allows following an animal over long spatial ranges but limits the number of animals that can be discerned in one experiment. Restricting the movement of the animals allows scaling up the observation to many individuals at the same time by using arrays of restricted compartments. 
Because the nematode is transparent, live fluorescent imaging can be performed non-invasively15. Calcium imaging provides a functional readout for the activity of excitable cells and is established for C. elegans16-20. Calcium enters the cell via channels in the plasma membrane that open upon depolarization. Thus, calcium acts as a proxy for neural activity. Calcium sensors can be grouped in two major classes, ratiometric and non-ratiometric sensors. Both classes employ conformational changes of calcium-binding proteins induced upon binding of calcium. Ratiometric sensors contain two fluorescent proteins. When the lower-wavelength fluorescent protein is excited, a part of the light energy is transmitted to the higher wavelength fluorescent protein as a function of their distance in a process called Fluorescence Resonance Energy Transfer or F&#246;rster Resonance Energy Transfer (FRET)21. Non-ratiometric sensors are based on circularly permuted GFP and employ de-quenching of the fluorophore caused by calcium binding22. Each class has its advantages. While ratiometric sensors are less sensitive to movement or expression artifacts, non-ratiometric sensors typically have a higher dynamic range. Both ratiometric and non-ratiometric sensors have been useful to study the activity of excitable cells in C. elegans16-20,23,24. 
When doing long-term fluorescence imaging, the experimenter will have to deal with several potential challenges: 1. Disturbance of behavior through excitation light: Worms are sensitive to short-wavelength fluorescence excitation light and avoid light from the violet to blue range that is used for calcium imaging25,26. Worms respond with either a backward or a forward escape response25,26. Thus, the amount of light needs to be controlled. 2. Bleaching of the fluorescent sensor protein: Often, bleaching of the fluorescent protein hampers long-term imaging. Typically, however, light intensities that are needed to observe bleaching are higher than the light intensities that cause disturbing effects on the behavior of the animals. Thus, bleaching is only a theoretical problem in this type of calcium imaging. 3. Worms inside the microchambers are not fixed and move constantly during wake behavior and images may appear blurred if the worm is not immobilized. All these challenges can be solved using extremely short exposure times with low light intensity. This can be realized by using a highly sensitive EMCCD camera with short exposure times and external triggering of a light emitting diode (LED). To expose the worms to the illumination light as briefly as possible (only during the exposure time of the camera) the LED is externally triggered using a transistor-transistor logic (TTL) signal that the camera emits during exposure resulting in an illumination of the worm for precisely the time the camera chip is exposed. This also means that the EMCCD chip will be dark during data readout, which is optimal for readout performance of this chip. 
Previously, agarose microchambers have been developed for long-term fluorescence imaging of C. elegans larvae9. Here, it is described how agarose micro-chambers can be used for long-term calcium imaging for any life stage of C. elegans, how calcium imaging can be performed, and how this method can be upscaled to assay many individual worms in parallel.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>high-melting point agarose</td>
			<td>Fisher Bioreagents</td>
			<td>BP(E)164-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Top Vision Low Melting Point Agarose</td>
			<td>Thermo Scientific</td>
			<td>R0801</td>
			<td></td>
		</tr>
		<tr>
			<td>LED system</td>
			<td>CoolLED</td>
			<td>pE-2</td>
			<td></td>
		</tr>
		<tr>
			<td>compound microscope</td>
			<td>Nikon</td>
			<td>TiE</td>
			<td></td>
		</tr>
		<tr>
			<td>stereomicroscope</td>
			<td>Leica</td>
			<td>M5</td>
			<td></td>
		</tr>
		<tr>
			<td>EMCCD camera</td>
			<td>Andor</td>
			<td>iXon 897</td>
			<td>now sold as iXon Ultra 897</td>
		</tr>
		<tr>
			<td>sCMOS camera</td>
			<td>Andor</td>
			<td>Neo</td>
			<td></td>
		</tr>
		<tr>
			<td>plasma cleaner</td>
			<td>Harrick</td>
			<td>PDC-23G2</td>
			<td></td>
		</tr>
		<tr>
			<td>lid heater</td>
			<td>MPI workshop</td>
			<td>custom made</td>
			<td></td>
		</tr>
		<tr>
			<td>plastic dish</td>
			<td>Falcon</td>
			<td>08-757-100A</td>
			<td></td>
		</tr>
		<tr>
			<td>z-stage for the microscope</td>
			<td>Prior</td>
			<td>Nano Scan Z</td>
			<td></td>
		</tr>
		<tr>
			<td>x-y-stage for the microscope</td>
			<td>Prior</td>
			<td>Proscan III</td>
			<td></td>
		</tr>
		<tr>
			<td>PDMS stamp</td>
			<td>Abbott Laboratories&#160;</td>
			<td>custom made</td>
			<td></td>
		</tr>
		<tr>
			<td>red light filter</td>
			<td>Chroma</td>
			<td>ET660/50m</td>
			<td></td>
		</tr>
		<tr>
			<td>35mm petri dish</td>
			<td>Falcon</td>
			<td colspan="2">Falcon Disposable Petri Dishes, Sterile, Corning</td>
		</tr>
		<tr>
			<td>double sided sticky tape</td>
			<td>Sellotape/Henkel</td>
			<td>double sided 12mm x 33 m</td>
			<td>alternatively Advance double sided sticky tape from Rubance Adhesives can be used.</td>
		</tr>
	</tbody>
</table>

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 can be applied to any life stage of C. elegans. As can be seen in Figure 1A, larval development and sleep behavior during L1 lethargus can be observed. Shown are chamber sizes that are 190 x 190 &#181;m, 10 &#181;m deep. If longer time scales are required, larger chambers can be used. As can be seen in Figure 1B, using a chamber with larger dimensions (370 x 370 &#181;m, 25 &#181;m deep) allows development of C. elegans from egg until adult. Figure 1C shows an analysis of long-term changes in behavior using frame subtraction of a worm grown in a chamber from egg until adult. Shown are mean intensities after frame subtraction for selected bursts during wake and lethargus. The lower the mean intensity after frame subtraction values is, the lower the mobility of the worm. Shown are selected burst traces from one wake condition and one lethargus condition per larval stage. The observed increase in mean intensity during development is mostly due to increased contrast and size of the animal. Figure 1D shows a dauer larva, an alternative life stage that is engaged during adverse environmental conditions33. Chamber size is 370 x 370 &#181;m, 15 &#181;m deep. Note that the dauer larva was placed into the chamber without bacterial food, which would cause exit from the dauer larval stage. Figure 1E shows an adult worm that has already laid many eggs into the chamber. Chamber dimensions used for the adult were 700 x 700 &#181;m, 45 &#181;m deep. Finally, Figure 1F shows an adult hermaphrodite and a male mating inside an adult chamber.
Calcium imaging in motile worms is possible with GCaMP calcium sensors. Figures 2A,B show calcium imaging of the command interneuron AVA for an L1 larva27. Figures 2C,D show calcium activity for the same type of neuron in an adult animal.
Scaling up long-term imaging is possible by scanning and by zooming out. Figure 3A shows the image quality of simultaneous imaging of 30 worms on one camera chip with a 5 megapixel camera. Figure 3B shows the image quality of 4 worms calcium imaged simultaneously.
<img alt="Figure 1" src="/files/ftp_upload/52742/52742fig1highres.jpg" width="700" /> 
Figure 1. Adaptation of AMI to all life stages of C. elegans. (A) Larva imaged from early L1 until early L2 stage in 190 x 190 &#181;m chambers. (B) Development from egg until adult in a 370 x 370 &#181;m chamber. (C) Mobility of a worm assessed using frame subtraction. Shown is the mean intensity of all pictures of the image after frame subtraction for one selected burst movie for each larval stage for wake and lethargus behavior. (D) A dauer larva in the absence of food in a 370 x 370 &#181;m chamber. (E) Adult hermaphrodite with many eggs laid into a 700 x 700 &#181;m chamber. (F) Mating of a male and a hermaphrodite inside a 700 x 700 &#181;m chamber. YA means young adult. <a href="https://www.jove.com/files/ftp_upload/52742/52742fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/52742/52742fig2highres.jpg" width="700" /> 
Figure 2. Calcium imaging with AMI. (A) Calcium imaging with GCaMP3 in the command interneuron AVA of L1 larvae using the nmr-1 promoter. Shown are two false-color images. In the left image, the activity of AVA is low and the worm is not making a backwards movement. In the right image the activity of AVA is high, and the worm is moving backwards. (B) Calcium transients over time for an L1 worm. (C,D) Calcium transients in an adult animal. <a href="https://www.jove.com/files/ftp_upload/52742/52742fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/52742/52742fig3highres.jpg" width="700" /> 
Figure 3. Imaging multiple worms in parallel by zooming out. (A) Simultaneous imaging of 30 L1 worms per frame using 190 x 190 &#181;m chambers and a 100X magnification (using a 10X objective) and a 16.6 x 14 mm camera chip. (B) Simultaneous imaging of four L3 worms per frame using 370 x 370 &#181;m chambers and 100X magnification and a region of interest of a 16.6 x 14 mm camera chip. <a href="https://www.jove.com/files/ftp_upload/52742/52742fig3highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans at JoVE.com

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.

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 ...

agarose-microchambers-for-long-term-calcium-imaging-caenorhabditis

Research

JoVE Journal

Neuroscience

9.4K Views.  Max Planck Institute for Biophysical Chemistry. 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. 

Video: Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans

C. elegans Positive Butanone Learning, Short-term, and Long-term Associative Memory Assays

The memory of experiences and learned information is critical for organisms to make choices that aid their survival. C. elegans navigates its environment through neuron-specific detection of food and chemical odors1, 2, and can associate nutritive states with chemical odors3, temperature4, and the pathogenicity of a food source5.
Here, we describe assays of C. elegans associative learning and short- and long-term associative memory. We modified an aversive olfactory learning paradigm6 to instead produce a positive response; the assay involves starving ~400 worms, then feeding the worms in the presence of the AWC neuron-sensed volatile chemoattractant butanone at a concentration that elicits a low chemotactic index (similar to Toroyama et al.7). A standard population chemotaxis assay1 tests the worms' attraction to the odorant immediately or minutes to hours after conditioning.
After conditioning, wild-type animals' chemotaxis to butanone increases ~0.6 Chemotaxis Index units, its "Learning Index". Associative learning is dependent on the presence of both food and butanone during training. Pairing food and butanone for a single conditioning period ("massed training") produces short-term associative memory that lasts ~2 hours. Multiple conditioning periods with rest periods between ("spaced training") yields long-term associative memory (&lt;40 hours), and is dependent on the cAMP Response Element Binding protein (CREB),6 a transcription factor required for long-term memory across species.8
Our protocol also includes image analysis methods for quick and accurate determination of chemotaxis indices. High-contrast images of animals on chemotaxis assay plates are captured and analyzed by worm counting software in MatLab. The software corrects for uneven background using a morphological tophat transformation.9 Otsu's method is then used to determine a threshold to separate worms from the background.10 Very small particles are removed automatically and larger non-worm regions (plate edges or agar punches) are removed by manual selection. The software then estimates the size of single worm by ignoring regions that are above a specified maximum size and taking the median size of the remaining regions. The number of worms is then estimated by dividing the total area identified as occupied by worms by the estimated size of a single worm.
We have found that learning and short- and long-term memory can be distinguished, and that these processes share similar key molecules with higher organisms.6,8 Our assays can quickly test novel candidate genes or molecules that affect learning and short- or long-term memory in C. elegans that are relevant across species.

C. elegans Positive Butanone Learning, Short-term, and Long-term Associative Memory Assays

Here we describe methods to test C. elegans associative learning and short- and long-term associative memory. These population assays employ the worms abilities to chemotax toward volatile odorants, and form positive associations upon pairing food with the chemoattractant butanone. Increasing the number of conditioning periods induces long-term memory.

The memory of experiences and learned information is critical for organisms to make choices that aid their survival. C. elegans navigates its environment ...

A Polished and Reinforced Thinned-skull Window for Long-term Imaging of the Mouse Brain

In vivo imaging of cortical function requires optical access to the brain without disruption of the intracranial environment. We present a method to form a polished and reinforced thinned skull (PoRTS) window in the mouse skull that spans several millimeters in diameter and is stable for months. The skull is thinned to 10 to 15 &mu;m in thickness with a hand held drill to achieve optical clarity, and is then overlaid with cyanoacrylate glue and a cover glass to: 1) provide rigidity, 2) inhibit bone regrowth and 3) reduce light scattering from irregularities on the bone surface. Since the skull is not breached, any inflammation that could affect the process being studied is greatly reduced. Imaging depths of up to 250 &mu;m below the cortical surface can be achieved using two-photon laser scanning microscopy. This window is well suited to study cerebral blood flow and cellular function in both anesthetized and awake preparations. It further offers the opportunity to manipulate cell activity using optogenetics or to disrupt blood flow in targeted vessels by irradiation of circulating photosensitizers.

We present a method to form an imaging window in the mouse skull that spans millimeters and is stable for months without inflammation of the brain. This method is well suited for longitudinal studies of blood flow, cellular dynamics, and cell/vascular structure using two-photon microscopy.

In vivo imaging of cortical function requires optical access to the brain without disruption of the intracranial environment. We present a method to form ...

A Molecular Readout of Long-term Olfactory Adaptation in C. elegans

During sustained stimulation most sensory neurons will adapt their response by decreasing their sensitivity to the signal. The adaptation response helps shape attention and also protects cells from over-stimulation. Adaptation within the olfactory circuit of C. elegans was first described by Colbert and Bargmann1,2. Here, the authors defined parameters of the olfactory adaptation paradigm, which they used to design a genetic screen to isolate mutants defective in their ability to adapt to volatile odors sensed by the Amphid Wing cells type C (AWC) sensory neurons. When wildtype C. elegans animals are exposed to an attractive AWC-sensed odor3 for 30 min they will adapt their responsiveness to the odor and will then ignore the adapting odor in a chemotaxis behavioral assay for ~1 hr. When wildtype C. elegans animals are exposed to an attractive AWC-sensed odor for ~1 hr they will then ignore the adapting odor in a chemotaxis behavioral assay for ~3 hr. These two phases of olfactory adaptation in C. elegans were described as short-term olfactory adaptation (induced after 30 min odor exposure), and long-term olfactory adaptation (induced after 60 min odor exposure). Later work from L'Etoile et al.,4 uncovered a Protein Kinase G (PKG) called EGL-4 that is required for both the short-term and long-term olfactory adaptation in AWC neurons. The EGL-4 protein contains a nuclear localization sequence that is necessary for long-term olfactory adaptation responses but dispensable for short-term olfactory adaptation responses in the AWC4. By tagging EGL-4 with a green fluorescent protein, it was possible to visualize the localization of EGL-4 in the AWC during prolonged odor exposure. Using this fully functional GFP-tagged EGL-4 (GFP::EGL-4) molecule we have been able to develop a molecular readout of long-term olfactory adaptation in the AWC5. Using this molecular readout of olfactory adaptation we have been able to perform both forward and reverse genetic screens to identify mutant animals that exhibit defective subcellular localization patterns of GFP::EGL-4 in the AWC6,7. Here we describe: 1) the construction of GFP::EGL-4 expressing animals; 2) the protocol for cultivation of animals for long-term odor-induced nuclear translocation assays; and 3) the scoring of the long-term odor-induced nuclear translocation event and recovery (re-sensitization) from the nuclear GFP::EGL-4 state.

A Molecular Readout of Long-term Olfactory Adaptation in C. elegans

Here we describe a molecular readout of long-term olfactory adaptation in Caenorhabditis elegans. The Protein Kinase G, EGL-4, is necessary for stable adaptation responses in the primary sensory neuron pair called AWC. During prolonged odor exposure EGL-4 translocates from the cytosol to nucleus of the AWC.

During sustained stimulation most sensory neurons will adapt their response by decreasing their sensitivity to the signal. The adaptation response helps ...

In vivo Neuronal Calcium Imaging in C. elegans

The nematode worm C. elegans is an ideal model organism for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, well-characterized nervous system allows identification and fluorescence imaging of any neuron within the intact animal. Simple immobilization techniques with minimal impact on the animal's physiology allow extended time-lapse imaging. The development of genetically-encoded calcium sensitive fluorophores such as cameleon 1 and GCaMP 2 allow in vivo imaging of neuronal calcium relating both cell physiology and neuronal activity. Numerous transgenic strains expressing these fluorophores in specific neurons are readily available or can be constructed using well-established techniques. Here, we describe detailed procedures for measuring calcium dynamics within a single neuron in vivo using both GCaMP and cameleon. We discuss advantages and disadvantages of both as well as various methods of sample preparation (animal immobilization) and image analysis. Finally, we present results from two experiments: 1) Using GCaMP to measure the sensory response of a specific neuron to an external electrical field and 2) Using cameleon to measure the physiological calcium response of a neuron to traumatic laser damage. Calcium imaging techniques such as these are used extensively in C. elegans and have been extended to measurements in freely moving animals, multiple neurons simultaneously and comparison across genetic backgrounds. C. elegans presents a robust and flexible system for in vivo neuronal imaging with advantages over other model systems in technical simplicity and cost.

In vivo Neuronal Calcium Imaging in C. elegans

With its small transparent body, well-documented neuroanatomy and a host of amenable genetic techniques and reagents, C. elegans makes an ideal model organism for in vivo neuronal imaging using relatively simple, low-cost techniques. Here we describe single neuron imaging within intact adult animals using genetically encoded fluorescent calcium indicators.

The nematode worm C. elegans is an ideal model organism  for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, ...

A Caenorhabditis elegans Model System for Amylopathy Study

Amylopathy is a term that describes abnormal synthesis and accumulation of amyloid beta (A&beta;) in tissues with time. A&beta; is a hallmark of Alzheimer's disease (AD) and is found in Lewy body dementia, inclusion body myositis and cerebral amyloid angiopathy 1-4. Amylopathies progressively develop with time. For this reason simple organisms with short lifespans may help to elucidate molecular aspects of these conditions. Here, we describe experimental protocols to study A&beta;-mediated neurodegeneration using the worm Caenorhabditis elegans. Thus, we construct transgenic worms by injecting DNA encoding human A&beta;42 into the syncytial gonads of adult hermaphrodites. Transformant lines are stabilized by a mutagenesis-induced integration. Nematodes are age synchronized by collecting and seeding their eggs. The function of neurons expressing A&beta;42 is tested in opportune behavioral assays (chemotaxis assays). Primary neuronal cultures obtained from embryos are used to complement behavioral data and to test the neuroprotective effects of anti-apoptotic compounds.

A Caenorhabditis elegans Model System for Amylopathy Study

We describe methods to study aspects of amylopathies in the worm C. elegans. We show how to construct worms expressing human A&beta;42 in neurons and how to test their function in behavioral assays. We further show how to obtain primary neuronal cultures that can be used for pharmacological testing.

Amylopathy is a term that describes abnormal synthesis and accumulation of amyloid beta (A&beta;) in tissues with time. A&beta; is a hallmark of ...

Methods for Studying the Mechanisms of Action of Antipsychotic Drugs in Caenorhabditis elegans

Caenorhabditis elegans is a simple genetic organism amenable to large-scale forward and reverse genetic screens and chemical genetic screens. The C. elegans genome includes potential antipsychotic drug (APD) targets conserved in humans, including genes encoding proteins required for neurotransmitter synthesis and for synaptic structure and function. APD exposure produces developmental delay and/or lethality in nematodes in a concentration-dependent manner. These phenotypes are caused, in part, by APD-induced inhibition of pharyngeal pumping1,2. Thus, the developmental phenotype has a neuromuscular basis, making it useful for pharmacogenetic studies of neuroleptics. Here we demonstrate detailed procedures for testing APD effects on nematode development and pharyngeal pumping. For the developmental assay, synchronized embryos are placed on nematode growth medium (NGM) plates containing APDs, and the stages of developing animals are then scored daily. For the pharyngeal pumping rate assay, staged young adult animals are tested on NGM plates containing APDs. The number of pharyngeal pumps per unit time is recorded, and the pumping rate is calculated. These assays can be used for studying many other types of small molecules or even large molecules.

Methods for Studying the Mechanisms of Action of Antipsychotic Drugs in Caenorhabditis elegans

Approaches for testing the effects of antipsychotic drugs (APDs) in Caenorhabditis elegans are demonstrated. Assays are described for testing drug effects on development and viability and on pharyngeal pumping rate. These methods are also applicable for pharmacogenetic experiments with drug classes other than APDs.

Caenorhabditis elegans is a simple genetic organism amenable to  large-scale forward and reverse genetic screens and chemical genetic screens. The C.  ...

Long-term Behavioral Tracking of Freely Swimming Weakly Electric Fish

Long-term behavioral tracking can capture and quantify natural animal behaviors, including those occurring infrequently. Behaviors such as exploration and social interactions can be best studied by observing unrestrained, freely behaving animals. Weakly electric fish (WEF) display readily observable exploratory and social behaviors by emitting electric organ discharge (EOD). Here, we describe three effective techniques to synchronously measure the EOD, body position, and posture of a free-swimming WEF for an extended period of time. First, we describe the construction of an experimental tank inside of an isolation chamber designed to block external sources of sensory stimuli such as light, sound, and vibration. The aquarium was partitioned to accommodate four test specimens, and automated gates remotely control the animals&#39; access to the central arena. Second, we describe a precise and reliable real-time EOD timing measurement method from freely swimming WEF. Signal distortions caused by the animal&#39;s body movements are corrected by spatial averaging and temporal processing stages. Third, we describe an underwater near-infrared imaging setup to observe unperturbed nocturnal animal behaviors. Infrared light pulses were used to synchronize the timing between the video and the physiological signal over a long recording duration. Our automated tracking software measures the animal&#39;s body position and posture reliably in an aquatic scene. In combination, these techniques enable long term observation of spontaneous behavior of freely swimming weakly electric fish in a reliable and precise manner. We believe our method can be similarly applied to the study of other aquatic animals by relating their physiological signals with exploratory or social behaviors.

We describe a set of techniques for studying spontaneous behavior of freely swimming weakly electric fish over an extended period of time, by synchronously measuring the animal&#39;s electric organ discharge timing, body position and posture both accurately and reliably in a specially designed aquarium tank inside a sensory isolation chamber.

Long-term behavioral tracking can capture and quantify natural animal behaviors, including those occurring infrequently. Behaviors such as exploration and ...

Long-term Time Lapse Imaging of Mouse Cochlear Explants

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building the highly ordered, complex structure of the mammalian cochlea proceeds for around ten days. In order to study changes in gene expression over this period and their response to pharmaceutical or genetic manipulation, long-term imaging is necessary. Previously, live imaging has typically been limited by the viability of explanted tissue in a humidified chamber atop a standard microscope. Difficulty in maintaining optimal conditions for culture growth with regard to humidity and temperature has placed limits on the length of imaging experiments. A microscope integrated into a modified tissue culture incubator provides an excellent environment for long term-live imaging. In this method we demonstrate how to establish embryonic mouse cochlear explants and how to use an incubator microscope to conduct time lapse imaging using both bright field and fluorescent microscopy to examine the behavior of a typical embryonic day (E) 13 cochlear explant and Sox2, a marker of the prosensory cells of the cochlea, over 5 days.

Live imaging of the embryonic mammalian cochlea is challenging because the developmental processes at hand operate on a temporal gradient over ten days. Here we present a method for culturing and then imaging embryonic cochlear explant tissue taken from a fluorescent reporter mouse over five days.

Here we present a method for long-term time-lapse imaging of live embryonic mouse cochlear explants. The developmental program responsible for building ...

A Caenorhabditis elegans Nutritional-status Based Copper Aversion Assay

To ensure survival, organisms must be capable of avoiding unfavorable habitats while ensuring a consistent food source. Caenorhabditis elegans alter their locomotory patterns upon detection of diverse environmental stimuli and can modulate their suite of behavioral responses in response to starvation conditions. Nematodes typically exhibit a decreased aversive response when removed from a food source for over 30 min. Observation of behavioral changes in response to a changing nutritional status can provide insight into the mechanisms that regulate the transition from a well-fed to starved state.
We have developed an assay that measures a nematode&#39;s ability to cross an aversive barrier (i.e. copper) then reach a food source over a prolonged period of time. This protocol builds upon previous work by integrating multiple variables in a manner that allows for continued data collection as the organisms shift towards an increasingly starved condition. Moreover, this assay permits an increased sample size so that larger populations of nematodes can be simultaneously evaluated.
Organisms defective for the ability to detect or respond to copper immediately cross the chemical barrier, while wild type nematodes are initially repelled. As wild type worms are increasingly starved, they begin to cross the barrier and reach the food source. We designed this assay to evaluate a mutant that is incapable of responding to diverse environmental cues, including food sensation or detection of aversive chemicals. When evaluated via this protocol, the defective organisms immediately crossed the barrier, but were also incapable of detecting a food source. Hence, these mutants repeatedly cross the chemical barrier despite temporarily reaching a food source. This assay can straightforwardly test populations of worms to evaluate potential pathway defects related to aversion and starvation.

A Caenorhabditis elegans Nutritional-status Based Copper Aversion Assay

Here, we present a Caenorhabditis elegans-specific assay designed to evaluate changes in copper aversion behavior and the ability to locate a common food source, as the organism progresses from a well-fed to starved nutritional state.

To ensure survival, organisms must be capable of avoiding unfavorable habitats while ensuring a consistent food source. Caenorhabditis elegans alter their ...

Generation and Long-term Maintenance of Nerve-free Hydra

The interstitial cell lineage of Hydra includes multipotent stem cells, and their derivatives: gland cells, nematocytes, germ cells, and nerve cells. The interstitial cells can be eliminated through two consecutive treatments with colchicine, a plant-derived toxin that kills dividing cells, thus erasing the potential for renewal of the differentiated cells that are derived from the interstitial stem cells. This allows for the generation of Hydra that lack nerve cells. A nerve-free polyp cannot open its mouth to feed, egest, or regulate osmotic pressure. Such animals, however, can survive and be cultured indefinitely in the laboratory if regularly force-fed and burped. The lack of nerve cells allows for studies of the role of the nervous system in regulating animal behavior and regeneration. Previously published protocols for nerve-free Hydra maintenance involve outdated techniques such as mouth-pipetting with hand-pulled micropipette tips to feed and clean the Hydra. Here, an improved protocol for maintenance of nerve-free Hydra is introduced. Fine-tipped forceps are used to force open the mouth and insert freshly killed Artemia. Following force-feeding, the body cavity of the animal is flushed with fresh medium using a syringe and hypodermic needle to remove undigested material, referred to here as "burping". This new method of force-feeding and burping nerve-free Hydra through the use of forceps and syringes eliminates the need for mouth-pipetting using hand-pulled micropipette tips. It thus makes the process safer and significantly more time efficient. To ensure that the nerve cells in the hypostome have been eliminated, immunohistochemistry using anti-tyrosine-tubulin is conducted.

Generation and Long-term Maintenance of Nerve-free Hydra

Through a double treatment with colchicine, a plant-derived toxin that kills dividing cells, nerve-free Hydra vulgaris can be generated. These Hydra cannot feed or egest on their own. This paper describes an improved method for long-term maintenance of nerve free Hydra vulgaris in the laboratory.

The interstitial cell lineage of Hydra includes multipotent stem cells, and their derivatives: gland cells, nematocytes, germ cells, and nerve cells. The ...