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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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 &#174; 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 &#174; Core &#8482; 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

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2.9K Views.  DBT-National Brain Research Centre. 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.

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

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

In vivo Laser Axotomy in C. elegans

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron is damaged by injury or disease, it may regenerate. Cell-intrinsic and extrinsic factors influence the ability of a neuron to regenerate and restore function. Recently, the nematode C. elegans has emerged as an excellent model organism to identify genes and signaling pathways that influence the regeneration of neurons1-6. The main way to initiate neuronal regeneration in C. elegans is laser-mediated cutting, or axotomy. During axotomy, a fluorescently-labeled neuronal process is severed using high-energy pulses. Initially, neuronal regeneration in C. elegans was examined using an amplified femtosecond laser5. However, subsequent regeneration studies have shown that a conventional pulsed laser can be used to accurately sever neurons in vivo and elicit a similar regenerative response1,3,7.
We present a protocol for performing in vivo laser axotomy in the worm using a MicroPoint pulsed laser, a turnkey system that is readily available and that has been widely used for targeted cell ablation. We describe aligning the laser, mounting the worms, cutting specific neurons, and assessing subsequent regeneration. The system provides the ability to cut large numbers of neurons in multiple worms during one experiment. Thus, laser axotomy as described herein is an efficient system for initiating and analyzing the process of regeneration.

In vivo Laser Axotomy in C. elegans

A protocol to cut neurons in C. elegans with a MicroPoint pulsed laser is presented. We describe setting up the system, immobilizing worms, and severing labeled neurons. Advantages include a relatively low-cost system and the ability to sever neuronal processes or ablate cells in vivo.

Neurons communicate with other cells via axons and dendrites, slender membrane extensions that contain pre- or post-synaptic specializations. If a neuron ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

In vivo Neuronal Calcium Imaging in C. elegans

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

In vivo Neuronal Calcium Imaging in C. elegans

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

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

A Caenorhabditis elegans Model System for Amylopathy Study

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

A Caenorhabditis elegans Model System for Amylopathy Study

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

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

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

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 ...

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic tectum, is a popular model to study how neural circuits self-assemble. The ability to carry out whole cell patch clamp recordings from tectal neurons and to record RGC-evoked responses, either in vivo or using a whole brain preparation, has generated a large body of high-resolution data about the mechanisms underlying normal, and abnormal, circuit formation and function. Here we describe how to perform the in vivo preparation, the original whole brain preparation, and a more recently developed horizontal brain slice preparation for obtaining whole cell patch clamp recordings from tectal neurons. Each preparation has unique experimental advantages. The in vivo preparation enables the recording of the direct response of tectal neurons to visual stimuli projected onto the eye. The whole brain preparation allows for the RGC axons to be activated in a highly controlled manner, and the horizontal brain slice preparation allows recording from across all layers of the tectum.

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

In this paper, we discuss three brain preparations used for whole cell patch clamp recording to study the retinotectal circuit of Xenopus laevis tadpoles. Each preparation, with its own specific advantages, contributes to the experimental tractability of the Xenopus tadpole as a model to study neural circuit function.

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic ...

Ballistic Labeling of Pyramidal Neurons in Brain Slices and in Primary Cell Culture

It has been reported that the size and shape of dendritic spines is related to their structural plasticity. To identify the morphological structure of pyramidal neurons and dendritic spines, a ballistic labeling technique can be utilized. In the present protocol, pyramidal neurons are labeled with DilC18(3) dye and analyzed using neuronal reconstruction software to assess neuronal morphology and dendritic spines. To investigate neuronal structure, dendritic branching analysis and Sholl analysis are performed, allowing researchers to draw inferences about dendritic branching complexity and neuronal arbor complexity, respectively. The evaluation of dendritic spines is conducted using an automatic assisted classification algorithm integral to the reconstruction software, which classifies spines into four categories (i.e., thin, mushroom, stubby, filopodia). Furthermore, an additional three parameters (i.e., length, head diameter, and volume) are also chosen to assess alterations in dendritic spine morphology. To validate the potential of wide application of the ballistic labeling technique, pyramidal neurons from in vitro cell culture were successfully labeled. Overall, the ballistic labeling method is unique and useful for visualizing neurons in different brain regions in rats, which in combination with sophisticated reconstruction software, allows researchers to elucidate the possible mechanisms underlying neurocognitive dysfunction.

We present a protocol to label and analyze pyramidal neurons, which is critical for evaluating potential morphological alterations in neurons and dendritic spines that may underlie neurochemical and behavioral abnormalities.

It has been reported that the size and shape of dendritic spines is related to their structural plasticity. To identify the morphological structure of ...