The author deeply thanks Dr. Asako Sugimoto (Tohoku University) for her helpful discussion. wyIs251 was a generous gift from Dr. Kang Shen (Stanford University). mnIs17 was provided by the CGC, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440). This work was supported by JSPS KAKENHI grant #17H05010 and #16H06536 and Daiichi Sankyo foundation, Brain Science Foundation and Naito Foundation.

1. Sample Preparation
<ol>
	<li>Generate a transgenic C. elegans strain of interest using transgenic methodologies18. Alternatively, obtain appropriate strains from the Caenorhabditis Genetics Center (CGC). To visualize axonal transport of synaptic vesicle precursors, use the transgenic line wyIs251 [Pmig-13::gfp::rab-3]7. To observe IFT, obtain the transgenic line mnIs17 [osm-6::gfp]19. These strains are used in this protocol. 
	NOTE: Either extrachromosomal array, genomic integration, or even GFP knock-in works. To reduce background signals, use cell-specific promoters, rather than pan-neuronal promoters. 
	NOTE: Maintenance of worms is described in other protocols20. Strains were maintained on lawns of Escherischia coli OP50 feeder on nematode growth medium (NGM) plates (1.7% (w/v) agarose, 50mM NaCl, 0.25% (w/v) peptone, 1 mM CaCl2, 5 mg/mL cholesterol, 25 mM KH2PO4, 1 mM MgSO4 on 60 mm diameter plastic dishes) at 20 &#176;C.</li>
	<li>Grow worms at 20 &#176;C on NGM plates to any life cycle stage of the worms. 
	NOTE: One can observe both axonal transport and IFT at any stage. Late L4 to young adult is a good positive control because multiple publications have used these stages and visualized trafficking events14,21,22,23.</li>
</ol>
2. Preparation of 10% Agarose
NOTE: Many vendors provide similar products such as agar, agar powders, and agarose. Use electrophoresis grade agarose (gel strength &#62;1200 g/cm2). Cheap agar powders do not work because the resulting gel is not strong enough to immobilize worms.
<ol>
	<li>Mix 0.4 g agarose in 4 mL distilled water (DW) in a glass tube (1.5 cm x 10.5 cm). Put a lid on the glass tube to avoid evaporation.</li>
	<li>Put the glass tube on a 95 &#176;C heat block for 90 min. In addition, prepare another glass tube (1.5 cm x 10.5 cm) filled with DW and keep it at 95 &#176;C. Put a Pasteur pipet in the 95 &#176;C DW (Figure 1A). A lot of small bubbles will be seen in the agarose tube and the solution should be turbid. Leave it until the solution becomes clear. Then, mix by pipetting up and down until the agarose solution becomes homogeneous. 
	NOTE: Microwaves cannot be used to prepare 10% agarose solution when made from DW and agarose. Small bubbles caused by microwaves prevent agarose melting. However, solidified 10% agarose gel can easily be melted again using a microwave. Glass tubes containing 10% agarose gel can be prepared ahead of time and stored at 4 &#176;C for at least 6 months. If doing so, melt the stock using a microwave when needed and start from step 3 in this protocol. 
	NOTE: Agarose gradually loses the ability to solidify if incubated at 95 &#176;C for a long time or after repeated solidification and melting. If this happens, prepare new 10% agarose.</li>
</ol>
3. Preparation of Agarose Pad
<ol>
	<li>Using the warmed Pasteur pipet prepared in step 2.2, place 2 drops of 10% agarose on a 76 x 22 mm slide.</li>
	<li>Quickly cover with a second 76 x 22 mm2 slide before the agarose drop solidifies as shown in Figure 1B, and push down on the second slide to flatten the agarose solution. The thickness should be 0.5-1 mm.</li>
	<li>Quickly flush the Pasteur pipette by pipetting 95 &#176;C DW up and down until the agarose solution is washed out. Otherwise, the pipet will become clogged by agarose gel. Leave the pipette standing in the 95 &#176;C DW.</li>
	<li>Wait for 1 min. The agarose pad forms and becomes cloudy. Then, slide the slides apart by hand (Figure 1C).</li>
</ol>
4. Mounting of Worms
NOTE: Even small amounts of worm movement prevent good observation. Levamisole has traditionally been used to prevent worm movement on the agarose pad24,25. However, Levamisole inhibits neuronal receptors in C. elegans, and therefore may affect trafficking events in neurons15. Using polystyrene microbeads described hereis a good alternative17.
<ol>
	<li>Under a stereo microscope equipped with a slidable mirror transillumination, transfer ~30 transgenic worms expressing the appropriate markers to a new NGM plate without bacteria using a platinum wire pick. 
	NOTE: A platinum wire pick is made of platinum wire and Pasteur pipette (5 inches), and the tip of the platinum wire was flattened. The preparation of the platinum wire picks and handling of worms with these are described in the Wormbook (http://www.wormbook.org).</li>
	<li>Leave the plate for 1 min. Bacteria are automatically removed from the surfaces of worms as the worms move on NGM agarose gel without bacteria.</li>
	<li>Put a 0.5-1.0 &#181;L drop of 2.6% 100 nm diameter polystyrene microbeads in water onto the agar pad (Figure 1D). Transfer 10-20 worms from the bacteria-free NGM plate to the microbeads. Drop and spread the worms using the platinum wire pick to avoid the overlapping of worm bodies (Figure 1E).</li>
	<li>Apply a 22 x 40 mm2 coverslip (Figure 1F). Push gently to remove air bubbles, but avoid crushing worms. The sample is now ready for imaging. 
	NOTE: Because no anesthesia is administered, worms are fixed only by the frictional force generated by the agar pad, polystyrene microbeads, and coverslip17. The presence of feeder bacteria and/or too much solution will make the frictional force weaker. 
	NOTE: Different sizes of coverslips (e.g., 22 x 22 mm2, 18 x 18 mm2) work. However, larger coverslips can prevent immersion water or oil from the objective lens leaking into samples during the observation.</li>
</ol>
5. Observation
NOTE: Appropriate imaging parameters (laser power, gain, binning, etc.) will differ for each microscope system and camera. Here, a widefield microscope equipped with a spinning disk confocal scanner and a digital CCD camera is used.
<ol>
	<li>Set the temperature control of the stage to 20 &#176;C, or adjust the room temperature to 20 &#176;C.</li>
	<li>Put the sample slide on the microscope stage. Use 100x (numerical aperture = 1.3 or better) objective lens.</li>
	<li>Look for the areas indicated in Figure 2A (for wyIs251 and axonal transport) or Figure 2B (for mnIs17 and IFT) as positive controls. Other areas can be analyzed as well22.</li>
	<li>Set parameters (CCD Gain = 200, Binning = 1 or 2, laser power at 488 nm = 1.0-1.3 mW). Perform time lapse recording at 4 frames/s for up to 1 min. Save files as mutli-TIFF format. 
	NOTE: If GFP signals disappear and it is difficult to continue to observe GFP::RAB-3 or OSM-6::GFP for 30 s, the laser power is too strong. In that case, reduce the laser power. Conversely, if no vesicular movement is recorded, the laser power is too weak. In that case, increase the laser power.</li>
	<li>Observe a different worm and repeat 5.3 and 5.4. The observations can last for up to 20 min, but each worm should be recorded only once to avoid the effects of photo damage. 
	NOTE: Worms have been observed for at least 20 min without significant defects7,27,28. Longer observation may be possible but haven&#39;t been tested yet. A clear sign of neuronal death is the change of neuronal morphology such as neurite swelling. As weak GFP signals can be seen in axons and dendrites in both wyIs251 and mnIs17, neurite morphology can be easily checked by just looking at axons or dendrites. Neuronal morphology is shown in Figure 2.</li>
	<li>Generate movie files.
	<ol>
		<li>Open the multi-TIFF file using Fiji software. Go to File&#62;Open then select the multi-TIFF file saved in step 5.4. 
		NOTE: Fiji can be downloaded at jttps://fiji.sc. Image J and plugins can also be used to generate kymographs. If doing so, follow the relevant instructions for the chosen plugin.&#160;</li>
		<li>Click the &#34;Rectangular selection&#34; button (Figure 3A, arrow) and select the area of interest by drawing a rectangle with a computer mouse (yellow rectangle in Figure 3A). Go to Image&#62;Stacks&#62;Tools&#62;Make Substacks and select the slice numbers that are included in the movie. A substack is generated.</li>
		<li>Activate the substack window by clicking and go to Image&#62;Adjust&#62;Brightness/Contrast. A window to adjust the brightness and contrast shows up. In the window, slide the top bar (Minimum) to the right so that the background signal disappears and the second top bar (Maximum) to the left so that the moving vesicles and particles can be seen quite well over the background.</li>
		<li>Go to Image&#62;Stacks&#62;Time Stamper. As the movie is recorded at 4 frames/s in step 5.4, the time interval is 0.25 s. Thus, type &#34;0.25&#34; in Interval.</li>
		<li>Generate a movie file by selecting Save as&#62;AVI from the File menu (Movie 1 and 2). 
		NOTE: By using appropriate plugins, you can save the file as other formats such as &#34;mpeg4&#34; or &#34;mov&#34; files.</li>
	</ol>
	</li>
	<li>Generate kymographs.
	<ol>
		<li>Open the original movie files using FIji software again and repeat steps 5.6.2 and 5.6.3 to adjust the brightness and contrast.</li>
		<li>Click &#34;segmented line&#34; (Figure 3B, arrow). Using a mouse, draw a segmented line along the cilia or axon (yellow line, in Figure 3B).</li>
		<li>Go to Analyze&#62;Multi Kymograph&#62;Multi Kymograph (Figure 3C). Line width window appears (Figure 3D). Type &#34;3&#34; in the Linewidth window and Click OK (Figure 3D).</li>
		<li>Kymograph window is created. Save the image in TIFF or JPEG format.</li>
	</ol>
	</li>
</ol>

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The author has nothing to disclose.

Limitation with respect to existing methods 
The method described here is optimized to observe fast events such as axonal transport and IFT. Thus, immobilization is more prioritized than longer incubation. While we have been able to observe trafficking events for at least 20 min without significant perturbation, this method may not be always suitable to observe slow events requiring longer observations, such as axon elongation and cell migration. For longer observations, one needs to optimize the conditions by reducing the percentage of the agarose (i.e., increasing the water to avoid drying up and reducing the pressure that potentially causes damage). Alternatively, the microfluidic device described above15 or immobilization of worms using agarose pads and anesthesia may be more suitable16 while side effects should be carefully assessed.
Critical steps within the protocol 
For adequate observation, markers are critical. Markers need to be incorporated into vesicles or particles and be of sufficient brightness. Either extrachromosomal arrays or integrated lines can be used. For visualization of the axonal transport of synaptic vesicle precursors in DA9 neurons, the wyIs251 strain is useful7,23. jsIs821 has been used to observe axonal transport in mechanosensory neurons in some studies26,27. When other neurons are analyzed, synaptic vesicle proteins should be expressed in interested neurons using cell specific promoters. These promoters should be strong enough. RAB-3 and SNB-1 are widely used to label synaptic vesicle precursors22,23,26. To observe IFT, the components of the IFT complex should be labeled. As a positive control, the mnIs17 strain is a good choice29. Cilia emanating from 8 cells (ASE, ASG, ASI, ASL, ASH, ASK, ADF, and ADL) are bundled together in the head (amphid) while cilia from 2 cells (PHA and PHB) are bundled in the tail (phasmid). Because OSM-6::GFP is expressed in all ciliated neurons in mnIs17, it may be difficult to analyze the IFT in detail in head cilia. However, in the tail region, only PHA and PHB neurons are labeled and IFT in each cell can easily be resolved. Other IFT proteins labeled by fluorescent proteins can be used to observe IFT21,28. If one wishes to analyze head neurons, cell-specific promoters would be useful. While head cilia have varying morphologies, transport events can be recorded29. By crossing mutants with these markers, the functions of any genes in axonal transport and IFT can be analyzed in vivo. The parameters that can be analyzed have been described in previous work22,23,26,27. GFP is the best and primary choice. mCherry and tdTomato can also be used22, however, mCherry is much dimmer than GFP. tdTomato is a tandem dimer and larger than monomeric fluorescent proteins, which sometimes affects the dynamics of fusion proteins30. Therefore, the results should be analyzed and interpreted cautiously when tdTomato is used. mScarlet, a bright monomeric red fluorescent protein which has recently been developed, is an alternative choice31.
While spinning disk confocal microscopy is widely used to observe axonal transport and IFT7,21, the equipment is expensive and may be difficult to access in some research environments. However, these phenomena can also be recorded and analyzed in C. elegans using standard fluorescent microscopes, if equipped with high NA lenses, because C. elegans is a small and transparent animal and easy to observe.
Future application 
By the similar immobilization method described here, IFT was observed at the single molecule resolution in vivo32. Moreover, several types of super-resolution microscopy have been developed. One can directly apply the methods described here to super-resolution microscopic analysis. In my experience, the fast mode of Airyscan33 works very well to analyze IFT. Theoretically, a spinning disk super-resolution microscope (SDSRM) would be a good choice as well34. In conclusion, by combining mutant libraries and these new techniques, the method described here should be useful for clarifying the molecular mechanisms of axonal transport and IFT in vivo.

Live cell imaging is an essential tool for analyzing intracellular transport. In neuronal cell biology, analyses of axonal transport with live cell imaging are essential for understanding neuronal function and morphogenesis1. Defects in axonal transport underlie several neurodegenerative disorders2. Kinesin superfamily proteins and dynein carry out axonal transport anterogradely and retrogradely, respectively1,2.
Cilia are another cellular compartment in which the microtubule network and trafficking machinery are highly developed3. Protein synthesis machinery is not localized in cilia, which means that ciliary proteins must be transported from the cytoplasm to the cilia tips. Cilia-specific kinesin and dynein, called kinesin-2 and cytoplasmic dynein-2, respectively, transport the components of cilia4, in a phenomenon called intraflagellar transport (IFT)5. Impairment of IFT causes a spectrum of diseases called ciliopathies6. Thus, analysis of the IFT mechanism by live cell imaging is required to understand basic mechanisms of ciliary formation and pathogenesis.
Caenorhabditis elegans (C. elegans) is a good model to study axonal transport and IFT7,8,9. To observe IFT, Chlamydomonas has been widely used as a model organism5,6. As Chlamydomonas is a unicellular organism, the relationship of IFT with aging, neuronal function, and behavior would be difficult to analyze. In addition, essential genetic techniques such as CRISPR/Cas9 have not been applied to Chlamydomonas. In higher model organisms, such as mice and Drosophila, disruption of axonal transport and IFT often causes lethal phenotypes because axonal transport and IFT are essential for the morphogenesis and homeostasis of the animals 10,11. In the case of mice, cell culture and transfection is generally needed to observe axonal transport and IFT, which requires many skills and extensive time12,13. In addition, a lot of important physiological context may be lost in cultured cells and cell lines. Because the nervous system is not essential for the survival of the worms, C. elegans mutants in which axonal transport or IFT are disrupted are often not lethal7,9,14. Axonal transport and IFT can be directly observed in vivo without dissection because C. elegans is transparent and it is therefore easy to observe GFP-tagged markers.
There are several protocols to immobilize C. elegans, such as using a microfluidic device15, agarose pads with anesthesia16, or microbeads17. The inclusion of anesthesia may inhibit the trafficking events in neurons15. A clear drawback of the microfluidic-device method is that preparing a microfluidic device is not always easy. Instead, immobilization by agarose pads and microbeads is a convenient and easy way to perform time lapse imaging in C. elegans. Here, I describe this basic protocol to immobilize C. elegans and visualize axonal transport and IFT in vivo in C. elegans. Compared to the other methods, the method described here does not require special equipment and is much cheaper and easier to perform.

<table><tbody><tr><td>Slideglass (76&amp;nbsp; x 26 mm)</td><td>Matsunami</td><td>S1111</td><td></td></tr><tr><td>Coverglass (22&amp;nbsp; x 40 mm)</td><td>Matsunami</td><td>C024401</td><td></td></tr><tr><td>Agarose</td><td>Wako</td><td>318-01195</td><td></td></tr><tr><td>Polystylene microbeads 0.1 micron</td><td>Polysciences</td><td>#00876</td><td></td></tr><tr><td>Heat block</td><td>TAITEC</td><td>0063288-000</td><td>CTU-mini</td></tr><tr><td>Microscope</td><td>Olympus</td><td>IX-71</td><td>widefield microscope</td></tr><tr><td>Spinning disk Scanner</td><td>Yokogawa</td><td>CSU-X1</td><td>spinning disc confocal scanner&amp;nbsp;</td></tr><tr><td>Digital CCD camera</td><td>Hamamatsu Photonics</td><td>C10600-10B</td><td>ORCA-R2 degital CCD camera</td></tr><tr><td>Objective lens (x100, NA1.4)</td><td>Olympus</td><td>UPLSAPO 100XO</td><td></td></tr><tr><td>Pasteur pipette (5 inch)</td><td>IWAKI</td><td>IK-PAS-5P</td><td></td></tr><tr><td>Glass tube (1.5 cm diameter x 10.5 cm)</td><td>IWAKI</td><td>9820TST15-105NP</td><td></td></tr><tr><td>TV9211: &lt;em&gt;wyIs251&lt;/em&gt;</td><td>Laboratory of Kang Shen</td><td>N/A</td><td></td></tr><tr><td>OTL11: &lt;em&gt;mnIs17&lt;/em&gt;</td><td>Ref. 27, 29</td><td>N/A</td><td>SP2101 was backcrossed with wild type for 6 times</td></tr><tr><td>Stereo microscope</td><td>Carl Zeiss</td><td>435064-9000-000</td><td>STEMI 508&amp;nbsp;</td></tr><tr><td>Mirror transillumination unit</td><td>Carl Zeiss</td><td>435425-9010-000</td><td></td></tr><tr><td>platinum wire (0.2 mm)</td><td>Nilaco Corporation</td><td>m78483501</td><td></td></tr><tr><td>60 mm plastic dish</td><td>Falcon</td><td>#351007</td><td></td></tr><tr><td>Fiji</td><td>N/A</td><td>N/A</td><td>https://fiji.sc/</td></tr><tr><td>nematode growth medium (NGM)&amp;nbsp;&amp;nbsp;</td><td></td><td></td><td>1.7% (w/v) agarose, 50mM NaCl, 0.25% (w/v) Peptone, 1 mM CaCl2, 5 mg/mL Cholesterol, 25 mM KH2PO4, 1 mM MgSO4&amp;nbsp;</td></tr></tbody></table>

immobilization of caenorhabditis elegans to analyze intracellular transport in neurons

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

Axonal transport in DA9 neuron 
Using the wyIs251 line, both the anterograde and retrograde axonal transport of GFP::RAB-3 can be simultaneously recorded in the DA9 motor neuron. The average speed of anterograde and retrograde transport in the proximal dorsal axon of the DA9 neuron is about 1.8 and 2.6 &#956;m/s, respectively22. The number of moving vesicles is about 0.03 and 0.018 per &#956;m of axon per s. Thus, for a 30 s observation of 10 &#956;m of DA9 axon using a 100x lens, one can find about 9 and 5 moving vesicles in the axon (Figure 4 and Movie 1). If no vesicular movement can be observed, it is possible that the laser power is too weak. In addition to long range runs, short range movement of vesicle pools can be recorded (Movie 1). Some vesicles stop at these vesicle pools, while others dissociate from there7,23. These parameters can be monitored and used to analyze mutant phenotypes.
IFT in tail neurons 
Cilia are localized to the tip of dendrites in C. elegans (Figure 2B). The mnIs17 strain expresses GFP-tagged OSM-6, one of the IFT subunits29. OSM-6 is expressed in both head and tail neurons19. While many head cilia are labeled by mnIs17, only two cilia are clearly observed in the tail neuron. Thus, observing these cilia is easier than head cilia (see also Discussion). OSM-6::GFP is diffuse in the neuronal cytoplasm such as that found in the axon, cell body, and dendrite. However, in cilia, OSM-6::GFP is concentrated to cilia and incorporated in moving particles. The length of PHA and PHB cilia are about 6.5 &#956;m. Anterograde IFT is biphasic 8,9. The speed of anterograde IFT is about 0.7 and 1.3 &#956;m/s in the middle and distal segments of the cilia, respectively (Figure 5 and Movie 2).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56690/56690fig1.jpg" /> 
Figure 1: Demonstration of the sample preparation. (A) Hot DW, Pasteur pipet and 10% agarose on heat block. (B and C) Preparation of agarose pads: An agarose pad is formed between slides (B). The upper slide is removed after an agarose pad forms (C). (D) Schematic drawing showing how polystyrene bead solution is put on the agarose pad. (E) Schematic drawing showing how worms are put in the polystyrene bead solution using a platinum wire pick. (F) A coverslip (22 x 44 mm2) is put on the agarose pad.&#160;<a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56690/56690fig1large.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/56690/56690fig2.jpg" /> 
Figure 2: Imaging area. (A) Schematic drawing of DA9 and VA12 neurons and a representative image of wyIs251. In DA9 neurons, the ventral axon, commissure, and proximal axon does not have mature synapses. But synaptic vesicle precursors are transported from the cell body to synapses via these axonal regions. The region that should be observed is shown by boxes. (B) Schematic drawing of PHA and PHB neurons and their cilia. The region that should be observed is shown by boxes. Scale bar = 10 &#956;m.&#160;<a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56690/56690fig2large.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/56690/56690fig3.jpg" /> 
Figure 3: Generation of movies and kymographs using Fiji software. (A) Click the &#34;Rectangular selection&#34; button (arrow) and select the area one would like to focus on by drawing a rectangle (Yellow box) to generate substacks. (B) Click the segmented line button (arrow) and draw a segmented line along the cilia using a mouse (Yellow line). (C) Select &#34;Analyze&#34;, &#34;Multi Kymograph&#34;, and &#34;Multi Kymograph&#34; to start a plugin to generate kymographs. (S) Input linewidth, click &#34;OK&#34; and generate kymographs (Figure 5 and Figure 7).&#160;<a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56690/56690fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56690/56690fig4.jpg" /> 
Figure 4: Representative kymograph of axonal transport of synaptic vesicle precursors. GFP::RAB-3 movement is observed at the proximal asynaptic region of DA9 neuron and the kymograph was generated with Multi Kymograph of Fiji. Horizontal and vertical bars represent length and time, respectively.&#160;<a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56690/56690fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56690/56690fig5.jpg" /> 
Figure 5: Representative kymograph of IFT. OSM-6::GFP is observed in PHA and PHB cilia and this kymograph is generated from the trafficking event in either one of them. The kymograph was generated with Multi Kymograph of Fiji. Horizontal and vertical bars represent length and time, respectively.&#160;<a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56690/56690fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Movie 1" src="/files/ftp_upload/56690/56690movie1.jpg" /> 
Movie 1: Representative movie of axonal transport of synaptic vesicle precursors. GFP::RAB-3 movement is observed at the proximal asynaptic region of the DA9 neuron. GFP::RAB-3 is incorporated into synaptic vesicle precursors and transported. Both anterograde and retrograde axonal transport are recorded. Some vesicles stop but restart again. The movie was recorded at 4 frames/s and plays at 15 frames/s. Scale bar = 5 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56690/movie1.mp4" target="_blank">Please click here to view this video. (Right-click to download.)</a>
<img alt="Movie 2" src="/files/ftp_upload/56690/56690Movie2.jpg" /> 
Movie 2: Representative movie of IFT. OSM-6::GFP is observed in PHA and PHB cilia. OSM-6::GFP is incorporated into the IFT complex and moves along the cilia. The movie was recorded at 4 frames/s and plays at 15 frames/s. Scale bar = 5 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/56690/Movie2.mov" target="_blank">Please click here to view this video. (Right-click to download.)</a>

Watch this Scientific Journal Video about Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons at JoVE.com

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

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

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

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

immobilization-caenorhabditis-elegans-to-analyze-intracellular

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

8.0K Views.  Tohoku University. Caenorhabditis elegans (C. elegans) is a good model to study axonal and intracellular transport. Here, I describe a protocol for in vivo recording and analysis of axonal and intraflagellar transport in C. elegans.

Video: Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

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 ...

Bioengineering

In Vivo Calcium Imaging in C. elegans Body Wall Muscles

The model organism C. elegans provides an excellent system to perform in vivo calcium imaging. Its transparent body and genetic manipulability allow for the targeted expression of genetically encoded calcium sensors. This protocol outlines the use of these sensors for the in vivo imaging of calcium dynamics in targeted cells, specifically the body wall muscles of the worms. By utilizing the co-expression of presynaptic channelrhodopsin, stimulation of acetylcholine release from excitatory motor neurons can be induced using blue light pulses, resulting in muscle depolarization and reproducible changes in cytoplasmic calcium levels. Two worm immobilization techniques are discussed with varying levels of difficulty. Comparison of these techniques demonstrates that both approaches preserve the physiology of the neuromuscular junction and allow for the reproducible quantification of calcium transients. By pairing optogenetics and functional calcium imaging, changes in postsynaptic calcium handling and homeostasis can be evaluated in a variety of mutant backgrounds. Data presented validates both immobilization techniques and specifically examines the roles of the C. elegans sarco(endo)plasmic reticular calcium ATPase and the calcium-activated BK potassium channel in the body wall muscle calcium regulation.

In Vivo Calcium Imaging in C. elegans Body Wall Muscles

This method provides a way to couple optogenetics and genetically encoded calcium sensors to image baseline cytosolic calcium levels and changes in evoked calcium transients in the body wall muscles of the model organism C. elegans.

The model organism C. elegans provides an excellent system to perform in vivo calcium imaging. Its transparent body and genetic manipulability allow for ...

Biochemistry

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, ...

Neuroscience

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 ...

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

It has become increasingly clear that neural circuit activity in behaving animals differs substantially from that seen in anesthetized or immobilized animals. Highly sensitive, genetically encoded fluorescent reporters of Ca2+ have revolutionized the recording of cell and synaptic activity using non-invasive optical approaches in behaving animals. When combined with genetic and optogenetic techniques, the molecular mechanisms that modulate cell and circuit activity during different behavior states can be identified.
Here we describe methods for ratiometric Ca2+ imaging of single neurons in freely behaving Caenorhabditis elegans worms. We demonstrate a simple mounting technique that gently overlays worms growing on a standard Nematode Growth Media (NGM) agar block with a glass coverslip, permitting animals to be recorded at high-resolution during unrestricted movement and behavior. With this technique, we use the sensitive Ca2+ reporter GCaMP5 to record changes in intracellular Ca2+ in the serotonergic Hermaphrodite Specific Neurons (HSNs) as they drive egg-laying behavior. By co-expressing mCherry, a Ca2+-insensitive fluorescent protein, we can track the position of the HSN within ~ 1 &#181;m and correct for fluctuations in fluorescence caused by changes in focus or movement. Simultaneous, infrared brightfield imaging allows for behavior recording and animal tracking using a motorized stage. By integrating these microscopic techniques and data streams, we can record Ca2+ activity in the C. elegans egg-laying circuit as it progresses between inactive and active behavior states over tens of minutes.

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

This protocol describes the use of genetically encoded Ca2+ reporters to record changes in neural activity in behaving Caenorhabditis elegans worms. &#8195;

It has become increasingly clear that neural circuit activity in behaving animals differs substantially from that seen in anesthetized or immobilized ...

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 ...

Live Animal Imaging and Cell Sorting Methods for Investigating Neurodegeneration in a C. elegans Excitotoxic Necrosis Model

Excitotoxic necrosis is a leading form of neurodegeneration. This process of regulated necrosis is triggered by the synaptic accumulation of the neurotransmitter glutamate, and the excessive stimulation of its postsynaptic receptors. However, information on the subsequent molecular events that culminate in the distinct neuronal swelling morphology of this type of neurodegeneration is lacking. Other aspects, such as changes in specific subcellular compartments, or the basis for the differential cellular vulnerability of distinct neuronal subtypes, remain under-explored. Furthermore, a range of factors that come into play in studies that use in vitro or ex vivo preparations might modify and distort the natural progression of this form of neurodegeneration. It is therefore important to study excitotoxic necrosis in live animals by monitoring the effects of interventions that regulate the extent of neuronal necrosis in the genetically amenable and transparent&#160;model system of the nematode Caenorhabditis elegans. This protocol describes methods of studying excitotoxic necrosis in C. elegans neurons, combining optical, genetic, and molecular analysis. To induce excitotoxic conditions in C. elegans, a&#160;knockout of a glutamate transporter gene (glt-3) is combined with a neuronal sensitizing genetic background (nuls5 [Pglr-1::G&#945;S(Q227L)]) to produce glutamate receptor hyperstimulation and neurodegeneration. Nomarski differential interference contrast (DIC), fluorescent, and confocal microscopy in live animals are methods used to quantify neurodegeneration, follow subcellular localization of fluorescently labeled proteins, and quantify mitochondrial morphology in the degenerating neurons. Neuronal Fluorescence Activated Cell Sorting (FACS) is used to distinctly sort at-risk neurons for cell-type specific transcriptomic analysis of neurodegeneration. A combination of live imaging and FACS methods as well as the benefits of the C. elegans model organism allow researchers to leverage this system to obtain reproducible data with a large sample size. Insights from these assays could translate to novel targets for therapeutic intervention in neurodegenerative diseases.

Live Animal Imaging and Cell Sorting Methods for Investigating Neurodegeneration in a C. elegans Excitotoxic Necrosis Model

In a C. elegans excitotoxicity model, this protocol employs in vivo imaging to analyze the regulation of necrotic neurodegeneration, the effect of genes encoding candidate mediators, and involvement of mitochondria. Cell dissociation and sorting is used to specifically obtain at-risk neurons for cell-specific transcriptomic analysis of neurodegeneration and neuroprotection mechanisms.

Excitotoxic necrosis is a leading form of neurodegeneration. This process of regulated necrosis is triggered by the synaptic accumulation of the ...

In vivo Assessment of Microtubule Dynamics and Orientation in Caenorhabditis elegans Neurons

In neurons, microtubule orientation has been a key assessor to identify axons that have plus-end out microtubules and dendrites that generally have mixed orientation. Here we describe methods to label, image, and analyze the microtubule dynamics and growth during the development and regeneration of touch neurons in C. elegans. Using genetically encoded fluorescent reporters of microtubule tips, we imaged the axonal microtubules. The local changes in microtubule behavior that initiates axon regeneration following axotomy can be quantified using this protocol. This assay is adaptable to other neurons and genetic backgrounds to investigate the regulation of microtubule dynamics in various cellular processes.

In vivo Assessment of Microtubule Dynamics and Orientation in Caenorhabditis elegans Neurons

A protocol for imaging the dynamic microtubules in vivo using fluorescently labeled End binding protein has been presented. We described the methods to label, image, and analyze the dynamic microtubules in the Posterior lateral microtubule (PLM) neuron of C. elegans.

In neurons, microtubule orientation has been a key assessor to identify axons that have plus-end out microtubules and dendrites that generally have mixed ...

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Axonal transport maintains neuronal homeostasis by enabling the bidirectional trafficking of diverse organelles and cargoes. Disruptions in axonal transport have devastating consequences for individual neurons and their networks, and contribute to a plethora of neurological disorders. As many of these conditions involve both cell autonomous and non-autonomous mechanisms, and often display a spectrum of pathology across neuronal subtypes, methods to accurately identify and analyze neuronal subsets are imperative.
This paper details protocols to assess in vivo axonal transport of signaling endosomes and mitochondria in sciatic nerves of anesthetized mice. Stepwise instructions are provided to 1) distinguish motor from sensory neurons in vivo, in situ, and ex vivo by using mice that selectively express fluorescent proteins within cholinergic motor neurons; and 2) separately or concurrently assess in vivo axonal transport of signaling endosomes and mitochondria. These complementary intravital approaches facilitate the simultaneous imaging of different cargoes in distinct peripheral nerve axons to quantitatively monitor axonal transport in health and disease.

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Using transgenic fluorescent mice, detailed protocols are described to assess in vivo axonal transport of signaling endosomes and mitochondria within motor and sensory axons of the intact sciatic nerve in live animals.

Axonal transport maintains neuronal homeostasis by enabling the bidirectional trafficking of diverse organelles and cargoes. Disruptions in axonal ...

Subcellular Imaging of Neuronal Calcium Handling In Vivo

Calcium (Ca2+) imaging has been largely used to examine neuronal activity, but it is becoming increasingly clear that subcellular Ca2+ handling is a crucial component of intracellular signaling. The visualization of subcellular Ca2+ dynamics in vivo,&#160;where neurons can be studied in their native, intact circuitry, has proven technically challenging in complex nervous systems. The transparency and relatively simple nervous system of the nematode Caenorhabditis elegans enable the cell-specific expression and in vivo visualization of fluorescent tags and indicators. Among these are fluorescent indicators that have been modified for use in the cytoplasm as well as various subcellular compartments, such as the mitochondria. This protocol enables non-ratiometric Ca2+ imaging in vivo with a subcellular resolution that permits the analysis of Ca2+ dynamics down to the level of individual dendritic spines and mitochondria. Here, two available genetically encoded indicators with different Ca2+ affinities are used to demonstrate the use of this protocol for measuring relative Ca2+ levels within the cytoplasm or mitochondrial matrix in a single pair of excitatory interneurons (AVA). Together with the genetic manipulations and longitudinal observations possible in C. elegans, this imaging protocol may be useful for answering questions regarding how Ca2+ handling regulates neuronal function and plasticity.

Subcellular Imaging of Neuronal Calcium Handling In Vivo

The current methods describe a non-ratiometric approach for high-resolution, sub-compartmental calcium imaging in vivo in Caenorhabditis elegans using readily available genetically encoded calcium indicators.

Calcium (Ca2+) imaging has been largely used to examine neuronal activity, but it is becoming increasingly clear that subcellular Ca2+ handling is a ...

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

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Chapters in this video

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

In Vivo Calcium Imaging in C. elegans Body Wall Muscles

In vivo Neuronal Calcium Imaging in C. elegans

Agarose Microchambers for Long-term Calcium Imaging of Caenorhabditis elegans

Ratiometric Calcium Imaging of Individual Neurons in Behaving Caenorhabditis Elegans

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

Live Animal Imaging and Cell Sorting Methods for Investigating Neurodegeneration in a C. elegans Excitotoxic Necrosis Model

In vivo Assessment of Microtubule Dynamics and Orientation in Caenorhabditis elegans Neurons

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Subcellular Imaging of Neuronal Calcium Handling In Vivo