We thank Yishi Jin and Andrew Chisholm for the initial support and the strain used in the study. The bacterial strain OP50 was commercially availed from Caenorhabditis Genetics Center (CGC) funded by NIH Office of Research Infrastructure Programs (P40 OD010440). We also thank Dharmendra Puri for the standardization of the experimental procedures. The study is funded by the core grant of National Brain Research Centre (supported by Department of Biotechnology, Govt. of India), DBT/Wellcome Trust India Alliance Early Career Grant (Grant # IA/E/18/1/504331) to S.D., Wellcome Trust-DBT India Alliance Intermediate Grant (Grant # IA/I/13/1/500874) to A.G.-R and a grant from Science and Engineering Research Board (SERB: CRG/2019/002194) to A.G.-R.

1. Reporter strain: Culture and maintenance
NOTE: To measure the microtubule dynamics and orientation in the PLM neurons, we used the worm strain expressing EBP-2::GFP under the touch neuron specific promoter mec-4 (juIs338 allele)18,25,26. We use standard worm culture and maintenance methods for this strain27.
<ol>
	<li>Prepare the Nematode Growth Medium (3.0 g/L sodium chloride, 2.5 g/L peptone, 20.0 g/L agar, 10 mg/L cholesterol (diluted from a stock solution of 10 mg/mL)) in water and autoclave the mixture.</li>
	<li>Cool the mixture briefly for 10-15 min before supplementing it with 1 mM calcium chloride, 1 mM magnesium sulfate and 25.0 mM potassium phosphate monobasic pH 6.0.</li>
	<li>Pour around 9 mL of the mixture into 60 mm Petri dishes using a sterile dispenser followed by storage at room temperature and sterile conditions for a day and subsequent storage at 4 &#176;C.</li>
	<li>Prepare 50 mL of B Broth (5.0 g/L sodium chloride, 10.0 g/L Bacto-Tryptone), autoclave and cool it down before inoculating with a single colony of Escherichia coli (OP50 strain).</li>
	<li>Culture the bacteria at 37 &#176;C for 12 hours. Bring the refrigerated NGM plates to room temperature for seeding and using a sterilized glass spreader make a smear of OP50 bacteria onto the NGM plate.</li>
	<li>Keep the inoculated NGM plates at 37 &#176;C for 12 hours, which will allow the formation of a bacterial lawn for the culturing of C. elegans. These plates can be stored in a 20 &#176;C incubator until usage.</li>
	<li>Rear the transgenic strain on the seeded NGM plates.</li>
	<li>Sterilize a platinum wire pick on flame and pick 20-25 gravid adult hermaphrodites for transfer to a freshly seeded plate.</li>
	<li>Once transferred, shift the plate to a 20 &#176;C incubator for 1 h. Then, extract the adults from these plates. These plates will now contain eggs that are age-matched.</li>
	<li>Shift the plates to the 20 &#176;C incubator for rearing until they reach the developmental stage of interest which in this case is the L4 stage at 43.5 hours after egg-laying. For more worms, rear multiple plates to avoid crowding.</li>
	<li>At the desired developmental stage, these worms can be mounted for imaging of the EBP-2::GFP comets.</li>
</ol>
2. Sample preparation: Mounting of worms for imaging of EBP-2 comets
NOTE: To enable live observation of the EBP comets in the PLM neurons, we mounted the worms on agarose pads to minimize their mobility while not compromising the physiology of the neuron. Among the various immobilization methods, we have chosen 0.1 &#181;m Polystyrene bead solution readily available commercially. We have outlined the mounting procedure used specifically for EBP-2::GFP observation.
<ol>
	<li>Prepare a solution of 10% agarose by melting 0.2 g of agarose in 2 mL of 1x M9 buffer (0.02 M KH2PO4, 0.02 M Na2HPO4, 0.008 M NaCl, 0.02 M NH4Cl) in a 5 mL glass test tube over a flame. This test tube can easily fit into a heating block which keeps the agarose in the molten state until usage.</li>
	<li>Using a wide-mouthed dropper, dispense a drop (around 100 &#181;L) of this molten agarose over a clean glass slide of 35 mm x 25 mm dimensions.</li>
	<li>While it is in the molten state, press the drop with another glass slide to create a thin film of the agarose between the glass slides. The agarose solidifies as a film of around 0.5 - 1.0 mm in thickness in around 30 seconds.</li>
	<li>Once the agarose solidifies, hold the bottom slide in the non-dominant hand and the top slide in the dominant hand. Flip up the top slide while holding the bottom slide steady as a result of which the agarose pad sticks to one of the two slides.</li>
	<li>If the formed agarose pad is larger than the coverslip size (18 mm x 18 mm), then trim the edges of the pad to obtain the required dimensions to ensure proper placement of the coverslip. These agarose pads are for the immediate use and not to be stored.</li>
	<li>Visually inspect the agarose pad for a smooth surface which are moist and suitable for mounting. If the agarose pad is exposed to air and becomes dried, the surface of the agarose pad appears wrinkled. In such a scenario, prepare fresh agarose pad as mentioned in steps 2-6 of this section.</li>
	<li>On the agarose pad, put 2 &#181;L of 0.1 &#181;m polystyrene bead solution as the mounting medium.</li>
	<li>Pick 3-4 worms using a platinum wire pick and shift them onto an unseeded NGM plate. This process ensures the removal of surface bacteria that hinders the immobilization during mounting.</li>
	<li>Once the worms crawl out of their surface bacteria, pick them up for mounting using a platinum wire pick and resuspend them in the drop of polystyrene beads on the agarose pad.</li>
	<li>Carefully, place an 18 mm x 18 mm coverslip on the worms. To prevent evisceration of the mounted worms, do not move the coverslip after placement.</li>
	<li>Mount the prepared slide on the microscope for imaging.</li>
</ol>
3. Imaging setup and acquisition
NOTE: EBP comets travel with an approximate velocity of 0.22 &#181;m/s as observed in the mammalian and PLM neurons of C. elegans16,18. To optimally sample the events in a time lapse acquisition as per Nyquist criteria28, spatial and temporal scales of 0.09 &#181;m and 0.43 s, respectively, are required. For the prevention of phototoxicity or photobleaching, we used Spinning Disk acquisition. We have described our imaging setup and acquisition settings below.
<ol>
	<li>Capture the images using an inverted microscope equipped with a spinning disk unit and camera. Turn on the setup as per the manufacturer&#39;s instruction.</li>
	<li>Turn on the software that controls the setup.</li>
	<li>As this is an inverted microscope, place the slide with the mounted worms on the stage with a low magnification objective (5x or 10x) with the coverslip facing down.</li>
	<li>Set up the light path to eye or visible mode which uses illumination from halogen lamp for transmitted light and mercury arc lamp for reflected light.</li>
	<li>Focus and center a worm in the view field under the brightfield.</li>
	<li>Put the immersion oil and change the magnification to 63x (1.4NA/Oil) and refocus the worm tail in the brightfield for the PLM neurons.</li>
	<li>Turn on the fluorescence of the mercury arc lamp for a brief refocusing of the PLM neurons. Place the correct set of reflector (AF488) in the light path for the visualization of GFP.</li>
	<li>Set the microscope to the camera acquisition by directing the light path to the camera. 
	NOTE: Controlling the abovementioned steps 4-7 through the Thin Film Transistor (TFT) screen of the microscope is quick and prevents unnecessary exposure to the illumination. Alternatively, control the illumination, reflector through the software.</li>
	<li>After the focusing of the PLM neuron in the view field, in the software interface select the illumination of 488 nm laser with 30% power and 300 ms exposure for the camera with Electron Multiplying (EM) gain value 70. Depending upon the reporter expression, vary these parameters as required. Store the settings as a configuration for subsequent imaging sessions.</li>
	<li>The Channels tab stores the tracks each with a specific illumination and camera setting. The highlighted track allows a live visualization whereas for acquiring an image the track needs to be ticked on. Highlight a channel with brightfield (DIC) for live visualization of the tail of the worm in the workspace. The sample is focused and centered in the live imaging window of the workspace.</li>
	<li>Highlight the channel with 488 nm laser excitation for a quick focusing of the PLM neuron in the live imaging window. Once focused, begin the time lapse acquisition by setting the time series parameters as mentioned in the next step.</li>
	<li>Using 488 nm laser illumination, set up an experiment with time series for 2 minutes and no time interval between frames. To maintain the temporal scales of image acquisition, we only acquired GFP fluorescence only.</li>
	<li>Using the Start Experiment tab, begin the acquisition. Using a 63x magnification restricts the spatial resolution to 0.21 &#181;m which is sampled by the camera at the pixel size of 0.09 &#181;m. Acquire a time lapse at 3.3 frames/s for optimally sample the event. The time lapse image had 396 time frames acquired over a duration of 2 min. Following the acquisition, save the image in the default&#160;format, which can be later be accessed by the default software&#160;or ImageJ.</li>
</ol>
4. Observation and analysis
<ol>
	<li>Preview the timelapse acquisition in the default software or open it in Image J through its Bioformats plug-in. 
	NOTE: We have described the process of image analysis in Image J.</li>
	<li>Install the Bioformats plug-in in ImageJ.
	<ol>
		<li>Alternatively, export the default&#160;format file into .tif for file compatibility in ImageJ. Export function in ZEN2 results in individual frames of the image as discrete .tif files. We avoid this method to prevent the formation of a large number of files.</li>
	</ol>
	</li>
	<li>Open the image in &#39;.czi&#39; format directly in Image J through its &#39;Bioformats&#39; plugin. The image opens as a multi-image stack. If using the &#34;export&#39; function of the default software, then reconstitute the images as a multi-image stack using the following workflow in ImageJ: ImageJ Program &#62; Image Menu &#62; Stacks Submenu &#62; Images To Stack.</li>
	<li>Preview the image as a movie using the Play icon at the left bottom corner of the image window. The comets can be seen as bright mobile punctae in the cell body and processes of the PLM neuron. If the comets are visible clearly, proceed to step 8 otherwise follow steps 5-7 of this section.</li>
	<li>Convert the color profile of the image using the Lookup Table (LUT) to a grayscale image: ImageJ program &#62; Image menu &#62; Lookup Tables submenu &#62; Select Grays.</li>
	<li>Invert the LUT profile of the image for easy visualization: ImageJ program &#62; Image menu &#62; Lookup Tables submenu &#62; Invert LUT.</li>
	<li>Adjust the brightness and the contrast of the image for seeing the comets: ImageJ program &#62; Image menu &#62; Adjust submenu &#62; Brightness/Contrast.
	<ol>
		<li>Move the sliders of Minimum, Maximum, Brightness, and Contrast for the desired scaling of the intensities across the image. We quantified the images with comets visible distinctly.</li>
	</ol>
	</li>
	<li>The basic version of ImageJ has startup tools represented as icons. Locate and right-click the icon with a line to open a dropdown menu and select the Segmented line. Draw the segmented line on the regions of interest (ROI) with its origin on or near the cell body of the neuron.
	<ol>
		<li>For later considerations, save the segmented line ROI through the ROI manager using the following workflow: ImageJ &#62; Analyze &#62; Tools &#62; ROI manager &#62; Select the drawn trace in the image and click Add in the ROI manager window &#62; More in the ROI manager window &#62; Save.</li>
	</ol>
	</li>
	<li>Before generating the kymograph, as the spatial and temporal scales are different remove the scaling to obtain the measurements in pixels using the following: 
	ImageJ &#62; Analyze &#62; Set scale &#62; Click to remove scale &#62; Ok.</li>
	<li>Using the Reslice function under the Image &#62; Stacks dropdown menu, generate a kymograph. The kymograph is a representation of the segmented ROI in time. The horizontal axis represents the distance and the vertical axis represents the time. The diagonal traces represent the moving comets. Generally, the kymograph takes the color profile of the image otherwise assign a color profile as described in the preceding steps 5-6 for the visualization of the diagonal traces.</li>
	<li>Using the Straight line function in the drop-down menu of the Line tool (refer to step 9 of this section) draw straight lines over each of the diagonal traces and append them in the ROI manager.</li>
	<li>Under the Analyze tab, go to Set Measurements and select Mean Intensity and Bounding Rectangle parameters for measuring the traces.</li>
	<li>In the ROI manager window, click Measure which will open a Result window which will yield the mean intensity of ROI, the X and Y coordinates of the line traces, Width, Height, Angle, and Length of the line traces. This window can be saved as a .csv format and later imported into a data analysis program.</li>
	<li>In the spreadsheet program, the results window opens with the values distributed in rows and columns. The relevant values are the width, height, and angle of the traces. The width gives the displacement of the trace, height gives the duration and the angle represents the vector. As the width and height values are in pixels, multiply them with the spatial and temporal scales, respectively, for obtaining the standard measurable parameters.</li>
	<li>As the left side of the kymograph is proximal to the cell body, depending on the angle, classify comets into plus-end-out or minus-end-out (i.e., away from the cell body or towards the cell body, respectively). Classify angles between 1&#176; to -89&#176; and -91&#176; to -179&#176; as plus-end-out and minus-end-out, respectively. Collate data from multiple kymographs in a spreadsheet for a study and statistically represent.</li>
</ol>

The authors declare no conflicts of interest.

Understanding the microtubule dynamics has been a key focus in the field of cytoskeletal research over the years. Microtubules undergo nucleation and catastrophe along with a continuous process of dynamic instability44,45,46,47. Much of this information has been obtained through in vitro assays like light scattering readouts of free vs polymerized tubulin, microtubu...

Neurons have an elaborate architecture with specialized compartments like dendrites, cell bodies, axons, and synapses. The neuronal cytoskeleton is constituted of the microtubules, microfilaments, and neurofilaments and their distinct organization supports the neuronal compartments structurally and functionally1,2,3,4,5,6,7,8,9,10. Over the years, microtubule organization has been identified as a key determinant of neuronal polarity and function. As neurons undergo structural remodeling during development or regeneration, the microtubule dynamics and orientation determine the identity, polarized transport, growth, and development of various neuronal compartments7. It is, therefore, imperative to assess the microtubule dynamics and orientation in vivo to correlate with the neuronal remodeling process.
Microtubules are composed of protofilaments of &#945; and &#946; Tubulin heterodimers with dynamic plus ends and relatively stable minus ends11,12. The discovery of the plus tip complex and associated end binding proteins have enabled a platform to assess the microtubule organization13. End binding proteins (EBP) transiently associate with the growing plus ends of the microtubule and their association dynamics are correlated to the growth of the microtubule protofilaments14,15. Due to frequent association and dissociation of plus tip complex with the microtubule, the point spread function of GFP-tagged EBP appears as a &#34;comet&#34; in a timelapse movie15,16. Since the pioneering observation in mammalian neurons16, end binding proteins tagged with fluorescent proteins have been used to determine microtubule dynamics across different model systems and neuron types17,18,19,20,21,22,23.
Due to its simple nervous system and transparent body, C. elegans has proven to be an excellent model system to study neuronal remodeling during development and regeneration in vivo. 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 EBP-2::GFP, we imaged the microtubules in the PLM neuron, which allowed us to determine the polarity of the microtubules in two different neurites of this neuron24. This method allows observation and quantification of the EBP comets as a measure of microtubule dynamics in different cellular contexts, for example, the local changes in microtubule behavior that initiates axon regeneration following axotomy can be assessed using our protocol. This assay can be adapted to investigate the regulation of microtubule dynamics in various cellular processes in diverse cell types and genetic backgrounds.

<table><tbody><tr><td>CZ18975 worm strain</td><td>Yishi Jin lab</td><td>CZ18975</td><td>Generated by Anindya Ghosh-Roy</td></tr><tr><td>Agarose</td><td>Sigma</td><td>A9539</td><td>Mounting worms</td></tr><tr><td>Coverslip (18 mm x 18 mm)</td><td>Zeiss</td><td>474030-9010-000</td><td>Mounting worms</td></tr><tr><td>Dry bath with heating block</td><td>Neolab</td><td></td><td>Mounting worms</td></tr><tr><td>Glass slides (35 mm x 25 mm)</td><td>Blue Star</td><td></td><td>Mounting worms</td></tr><tr><td>Polystyrene bead solution (4.55 x 10^13 particles/ml in aqueous medium with minimal surfactant)</td><td>Polysciences Inc.</td><td>00876</td><td>Mounting worms</td></tr><tr><td>Test tubes</td><td></td><td></td><td>Mounting worms</td></tr><tr><td>OP50 bacterial strain</td><td>Caenorhabditis Genetics Center (CGC)</td><td>OP50</td><td>Worm handling</td></tr><tr><td>60mm petri plates</td><td>Praveen Scientific</td><td>20440</td><td>Worm handling</td></tr><tr><td>Aspirator/Capillary</td><td>VWR</td><td>53432-921</td><td>Worm handling</td></tr><tr><td>Incubator</td><td>Panasonic</td><td>MIR554E</td><td>Worm handling</td></tr><tr><td>Platinum wire</td><td></td><td></td><td>Worm handling</td></tr><tr><td>Stereomicroscope with fluorescence attachment</td><td>Leica</td><td>M165FC</td><td>Worm handling</td></tr><tr><td>0.3% Sodium Chloride</td><td>Sigma</td><td>71376</td><td>Nematode Growth Medium</td></tr><tr><td>0.25% Peptone</td><td>T M Media</td><td>1506</td><td>Nematode Growth Medium</td></tr><tr><td>10mg/mL Cholesterol</td><td>Sigma</td><td>C8667</td><td>Nematode Growth Medium</td></tr><tr><td>1mM Calcium chloride dihydrate</td><td>Sigma</td><td>223506</td><td>Nematode Growth Medium</td></tr><tr><td>1mM Magnesium sulphate heptahydrate</td><td>Sigma</td><td>M2773</td><td>Nematode Growth Medium</td></tr><tr><td>2% Agar</td><td>T M Media</td><td>1202</td><td>Nematode Growth Medium</td></tr><tr><td>25mM Monobasic Potassium dihydrogen phosphate</td><td>Sigma</td><td>P9791</td><td>Nematode Growth Medium</td></tr><tr><td>0.1M Monobasic Potassium dihydrogen phosphate</td><td>Sigma</td><td>P9791</td><td>1X M9 buffer</td></tr><tr><td>0.04M Sodium chloride</td><td>Sigma</td><td>71376</td><td>1X M9 buffer</td></tr><tr><td>0.1M Ammonium chloride</td><td>Fisher Scientific</td><td>21405</td><td>1X M9 buffer</td></tr><tr><td>0.2M Dibasic Disodium hydrogen phosphate heptahydrate</td><td>Sigma</td><td>S9390</td><td>1X M9 buffer</td></tr><tr><td>Glass bottles</td><td>Borosil</td><td></td><td>Buffer storage</td></tr><tr><td>488 nm laser</td><td>Zeiss</td><td></td><td>Imaging</td></tr><tr><td>5X objective</td><td>Zeiss</td><td></td><td>Imaging</td></tr><tr><td>63X objective</td><td>Zeiss</td><td></td><td>Imaging</td></tr><tr><td>Camera</td><td>Photometrics</td><td>Evolve 512 Delta</td><td>Imaging</td></tr><tr><td>Computer system for Spinning Disk unit</td><td>HP</td><td>Intel &amp;reg; Xeon CPU E5-2623 3.00GHz</td><td>Imaging</td></tr><tr><td>Epifluorescence microscope</td><td>Zeiss</td><td>Observer.Z1</td><td>Imaging</td></tr><tr><td>Halogen lamp</td><td>Zeiss</td><td></td><td>Imaging</td></tr><tr><td>Mercury Arc Lamp</td><td>Zeiss</td><td></td><td>Imaging</td></tr><tr><td>Spinning Disk Unit</td><td>Yokogawa</td><td>CSU-X1</td><td>Imaging</td></tr><tr><td>ZEN2 software</td><td>Zeiss</td><td></td><td>Imaging</td></tr><tr><td>Image J (Fiji Version)</td><td></td><td></td><td>Image analysis and processing</td></tr><tr><td>Adobe Creative Cloud</td><td>Adobe</td><td></td><td>Image analysis and processing</td></tr><tr><td>Computer system for Image Analysis</td><td>Dell</td><td>Intel &amp;reg; Core &amp;trade; i7-9700 CPU 3.00GHz</td><td>Image processing/Representation</td></tr></tbody></table>

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.

As a representative example, we have described in vivo observation of the EBP comets in the steady-state and regenerating axons of the PLM neurons. PLM neurons are located in the tail region of the worm with a long anterior process that forms a synapse and a short posterior process. PLM neurons grow in the anterior-posterior direction close to the epidermis and are responsible for the gentle touch sensation in the worms. Due to their simplified structure, and amenability to imaging and microsurgery, PLM neurons have been...

Watch this Scientific Journal Video about In vivo Assessment of Microtubule Dynamics and Orientation in Caenorhabditis elegans Neurons at JoVE.com

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 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 ...

in-vivo-assessment-microtubule-dynamics-orientation-caenorhabditis

Research

JoVE Journal

Neuroscience

3.0K Views.  DBT-National Brain Research Centre. Het doel van deze procedure is om de organisatie en dynamiek van de microtubuli in vivo te beoordelen. Neuronen zijn gepolariseerde cellen met verschillende axonale, dendritische en synaptische compartimenten, onderhouden door hun onderliggende cytoskelet. Microtubuli vormen de meerderheid van het neuronale cytoskelet en de dynamiek en oriëntatie bepalen belangrijke gebeurtenissen tijdens neuronale ontwikkeling en rijping.Microtubuli bestaan uit alfa- en bèta-tubuline-heterodimeren e...

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

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

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

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

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

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

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

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

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

Immobilization of Caenorhabditis elegans to Analyze Intracellular Transport in Neurons

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

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

Developmental Biology

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, ...

Calcium Imaging in Freely Behaving Caenorhabditis elegans with Well-Controlled, Nonlocalized Vibration

Nonlocalized mechanical forces, such as vibrations and acoustic waves, influence a wide variety of biological processes from development to homeostasis. Animals cope with these stimuli by modifying their behavior. Understanding the mechanisms underlying such behavioral modification requires quantification of neural activity during the behavior of interest. Here, we report a method for calcium imaging in freely behaving Caenorhabditis elegans with nonlocalized vibration of specific frequency, displacement, and duration. This method allows the production of well-controlled, nonlocalized vibration using an acoustic transducer and quantification of evoked calcium responses at single-cell resolution. As a proof of principle, the calcium response of a single interneuron, AVA, during the escape response of C. elegans to vibration is demonstrated. This system will facilitate understanding of neural mechanisms underlying behavioral responses to mechanical stimuli.

Calcium Imaging in Freely Behaving Caenorhabditis elegans with Well-Controlled, Nonlocalized Vibration

Reported here is a system for calcium imaging in freely behaving Caenorhabditis elegans with well-controlled, nonlocalized vibration. This system allows researchers to evoke nonlocalized vibrations with well-controlled properties at nano-scale displacement and to quantify calcium currents during responses of C. elegans to the vibrations.

Nonlocalized mechanical forces, such as vibrations and acoustic waves, influence a wide variety of biological processes from development to homeostasis. ...

Behavior

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

The mechanisms controlling stress-induced phenotypic plasticity in animals are frequently complex and difficult to study in vivo. A classic example of stress-induced plasticity is the dauer stage of C. elegans. Dauers are an alternative developmental larval stage formed under conditions of low concentrations of bacterial food and high concentrations of a dauer pheromone. Dauers display extensive developmental and behavioral plasticity. For example, a set of four inner-labial quadrant (IL2Q) neurons undergo extensive reversible remodeling during dauer formation. Utilizing the well-known environmental pathways regulating dauer entry, a previously established method for the production of crude dauer pheromone from large-scale liquid nematode cultures is demonstrated. With this method, a concentration of 50,000 - 75,000 nematodes/ml of liquid culture is sufficient to produce a highly potent crude dauer pheromone. The crude pheromone potency is determined by a dose-response bioassay. Finally, the methods used for in vivo time-lapse imaging of the IL2Qs during dauer formation are described.

In Vivo Imaging of Dauer-specific Neuronal Remodeling in C. elegans

Following exposure to specific environmental stressors, the nematode Caenorhabditis elegans undergoes extensive phenotypic plasticity to enter into a stress-resistant &#8216;dauer&#8217; juvenile stage. We present methods for the controlled induction and imaging of neuroplasticity during dauer.

The mechanisms controlling stress-induced phenotypic plasticity in animals are frequently complex and difficult to study in vivo. A classic example of ...

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 ...

3D Particle Tracking for Noninvasive In Vivo Analysis of Synaptic Microtubule Dynamics in Dendrites and Neuromuscular Junctions of Drosophila

Microtubules (MTs) play critical roles in neuronal development, but many questions remain about the molecular mechanisms of their regulation and function. Furthermore, despite progress in understanding postsynaptic MTs, much less is known about the contributions of presynaptic MTs to neuronal morphogenesis. In particular, studies of in vivo MT dynamics in Drosophila sensory dendrites yielded significant insights into polymer-level behavior. However, the technical and analytical challenges associated with live imaging of the fly neuromuscular junction (NMJ) have limited comparable studies of presynaptic MT dynamics. Moreover, while there are many highly effective software strategies for automated analysis of MT dynamics in vitro and ex vivo, in vivo data often necessitate significant operator input or entirely manual analysis due to inherently inferior signal-to-noise ratio in images and complex cellular morphology. &#160;To address this, this study optimized a new software platform for automated and unbiased in vivo particle detection. Multiparametric analysis of live time-lapse confocal images of EB1-GFP labeled MTs was performed in both dendrites and the NMJ of Drosophila larvae and found striking differences in MT behaviors. MT dynamics were furthermore analyzed following knockdown of the MT-associated protein (MAP) dTACC, a key regulator of Drosophila synapse development, and identified statistically significant changes in MT dynamics compared to wild type. These results demonstrate that this novel strategy for the automated multiparametric analysis of both pre- and postsynaptic MT dynamics at the polymer-level significantly reduces human-in-the-loop criteria. The study furthermore shows the utility of this method in detecting distinct MT behaviors upon dTACC-knockdown, indicating a possible future application for functional screens of factors that regulate MT dynamics in vivo. Future applications of this method may also focus on elucidating cell type and/or compartment-specific MT behaviors, and multicolor correlative imaging of EB1-GFP with other cellular and subcellular markers of interest.&#160;

This study presents a noninvasive intravital neuronal imaging strategy combined with a new software strategy to achieve automated, unbiased tracking and analysis of in vivo microtubule (MT) plus-end dynamics in the sensory dendrites and the neuromuscular junctions of Drosophila.

Microtubules (MTs) play critical roles in neuronal development, but many questions remain about the molecular mechanisms of their regulation and function. ...

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 ...

Quantitative Approaches for Studying Cellular Structures and Organelle Morphology in Caenorhabditis elegans

Defining the cellular mechanisms underlying disease is essential for the development of novel therapeutics. A strategy frequently used to unravel these mechanisms is to introduce mutations in candidate genes and qualitatively describe changes in the morphology of tissues and cellular organelles. However, qualitative descriptions may not capture subtle phenotypic differences, might misrepresent phenotypic variations across individuals in a population, and are frequently assessed subjectively. Here, quantitative approaches are described to study the morphology of tissues and organelles in the nematode Caenorhabditis elegans using laser scanning confocal microscopy combined with commercially available bio-image processing software. A quantitative analysis of phenotypes affecting synapse integrity (size and integrated fluorescence levels), muscle development (muscle cell size and myosin filament length), and mitochondrial morphology (circularity and size) was performed to understand the effects of genetic mutations on these cellular structures. These quantitative approaches are not limited to the applications described here, as they could readily be used to quantitatively assess the morphology of other tissues and organelles in the nematode, as well as in other model organisms.

Quantitative Approaches for Studying Cellular Structures and Organelle Morphology in Caenorhabditis elegans

This study outlines quantitative measurements of synaptic size and localization, muscle morphology, and mitochondrial shape in C. elegans using freely available image processing tools. This approach allows future studies in&#160;C. elegans&#160;to quantitatively compare the extent of tissue and organelle structural changes as a result of genetic mutations.

Defining the cellular mechanisms underlying disease is essential for the development of novel therapeutics. A strategy frequently used to unravel these ...

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

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