We thank the reviewers for helpful comments and suggestions. We thank Aleen Saxton, Brandon Henderson, and Jade Stair for outstanding technical assistance. We thank Brian Kraemer and Rebecca Kow for assistance developing these assays. This material is the result of work supported with resources and the use of facilities at the VA Puget Sound Health Care System. This work was supported by a grant from the United States (U.S.) Department of Veterans Affairs (VA) Biomedical Laboratory Research and Development Service [Merit Review Grant #I01BX004044 to N.F.L.]

<ol>
	<li>Wijesekera, L. C., Leigh, P. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Amyotrophic+lateral+sclerosis.">Amyotrophic lateral sclerosis.</a> Orphanet Journal of Rare Diseases. 4, 3 (2009).</li><li>Arai, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=TDP-43+is+a+component+of+ubiquitin-positive+tau-negative+inclusions+in+frontotemporal+lobar+degeneration+and+amyotrophic+lateral+sclerosis.">TDP-43 is a component of ubiquitin-positive tau-negative inclusions in frontotemporal lobar degeneration and amyotrophic lateral sclerosis.</a> Biochemical and Biophysical Research Communication. 351 (3), 602-611 (2006).</li><li>Neumann, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ubiquitinated+TDP-43+in+frontotemporal+lobar+degeneration+and+amyotrophic+lateral+sclerosis.">Ubiquitinated TDP-43 in frontotemporal lobar degeneration and amyotrophic lateral sclerosis.</a> Science. 314 (5796), 130-133 (2006).</li><li>Mejzini, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ALS+genetics,+mechanisms,+and+therapeutics:+Where+are+we+now.">ALS genetics, mechanisms, and therapeutics: Where are we now.</a> Frontiers in Neuroscience. 13, 1310 (2019).</li><li>Tan, R. H., Ke, Y. D., Ittner, L. M., Halliday, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ALS/FTLD:+Experimental+models+and+reality.">ALS/FTLD: Experimental models and reality.</a> Acta Neuropathologica. 133 (2), 177-196 (2017).</li><li>Kim, W., Underwood, R. S., Greenwald, I., Shaye, D. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=OrthoList+2:+A+new+comparative+genomic+analysis+of+human+and+Caenorhabditis+elegans+genes.">OrthoList 2: A new comparative genomic analysis of human and Caenorhabditis elegans genes.</a> Genetics. 210 (2), 445-461 (2018).</li><li>White, J. G., Southgate, E., Thomson, J. N., Brenner, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+structure+of+the+nervous+system+of+the+nematode+Caenorhabditis+elegans.">The structure of the nervous system of the nematode Caenorhabditis elegans.</a> Philosophical Transaction of the Royal Society of London B: Biological Sciences. 314 (1165), 1 (1986).</li><li>Caldwell, K. A., Willicott, C. W., Caldwell, G. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modeling+neurodegeneration+in+Caenorhabditis+elegans.">Modeling neurodegeneration in Caenorhabditis elegans.</a> Disease Models and Mechanisms. 13 (10), (2020).</li><li>Liachko, N. F., Guthrie, C. R., Kraemer, B. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Phosphorylation+promotes+neurotoxicity+in+a+Caenorhabditis+elegans+model+of+TDP-43+proteinopathy.">Phosphorylation promotes neurotoxicity in a Caenorhabditis elegans model of TDP-43 proteinopathy.</a> Journal of Neuroscience. 30 (48), 16208-16219 (2010).</li><li>Robatzek, M., Thomas, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium/calmodulin-dependent+protein+kinase+II+regulates+Caenorhabditis+elegans+locomotion+in+concert+with+a+G(o)/G(q)+signaling+network.">Calcium/calmodulin-dependent protein kinase II regulates Caenorhabditis elegans locomotion in concert with a G(o)/G(q) signaling network.</a> Genetics. 156 (3), 1069-1082 (2000).</li><li>Porta-de-la-Riva, M., Fontrodona, L., Villanueva, A., Ceron, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Basic+Caenorhabditis+elegans+methods:+synchronization+and+observation.">Basic Caenorhabditis elegans methods: synchronization and observation.</a> Journal of Visualized Experiments. (64), e4019 (2012).</li><li>Husson, S. J., Costa, W. S., Schmitt, C., Gottschalk, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Keeping+track+of+worm+trackers.">Keeping track of worm trackers.</a> WormBook. , 1-17 (2013).</li><li>Ringstad, N., Horvitz, B., Koelle, M., Hart, A. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> WormBook. , 1551 (2006).</li></ol>

The authors declare that they have no disclosures.

Radial locomotion: 
The resolution of this assay is easily controlled by changing the time variable. Increasing the length of time makes it easier to observe differences between animals with severe phenotypes, thereby identifying subtle differences. However, because this assay measures displacement, if the assay time is extended too long, animals with normal motility, such as N2, will travel to the edges of the plate, and foraging behavior will lead to backtracking. This will artificially decrease the measurement of distance traveled. Time periods that are too long may result in the disappearance of differences between strains, particularly between animals with less severe motor phenotypes, as animals become evenly dispersed across the plate. Shortening the time variable will prevent more active worms from finding the edges of the plate. This method does not track the total distance traveled for each worm but compresses the distance traveled for each worm into a linear distance from the center of the plate. As such, it is inherently less robust than a method that records the total track length of individual worms. However, the radial locomotion assay requires very little researcher training, uses relatively affordable reagents that are commonly available in most worm labs, and is sensitive enough to produce significant and reproducible results. For labs that prefer automated video tracking, several methods have been previously established to track and analyze crawling movements12, or the software parameters used for the swimming assay in this paper could be altered to permit crawling detection and analysis.
This experiment is usually done in triplicate independent replicates, with a set of 30-40 worms per replicate. Each replicate is split onto two different 100 mm or 150 mm plates, with 15-20 worms per plate. Using more worms than recommended per plate can make it hard to score efficiently. A total number scored of 90+ is sufficiently powered to establish significance for mild, moderate, or severe motility impairment. Being consistent with timing between strains scored is essential for accuracy and reproducibility. 30 minutes is generally long enough to establish differences between moderate to severe phenotypes such as transgenic strains expressing human mutant TDP-43 compared to wild-type worms (Figure 4). If the time variable will be extended, it is advisable to also increase the size of the plate from 100 mm to 150 mm. Environmental factors such as temperature and humidity can affect this assay, which is typically conducted at ambient room temperature, so it is important to always use a wild-type (N2) control when comparing across replicates. In addition, this assay can measure the motility of some strains that do not exhibit normal swimming behavior in liquid (thrashing), making it a useful complement to the swimming assay.
Swimming assay: 
The use of the imaging and acquisition system to automate tracking and analysis of worm swimming allows for rigorous and unbiased data. However, there are several factors during initial setup of the experiment that needs to be controlled between samples. These include the time to acclimate to liquid before beginning recording, ambient conditions (i.e., temperature, humidity), and consistent light and recording settings. On the recording stage, there are several features that help reduce variability between plates. These include an integrated track-mounted camera and bright-field stage that makes recording video consistent between plates, shielding around the stage that prevents reflections, glare, and air movements while recording, and a robust software package that reliably detects worms and allows for manual correction of tracks in video post-processing. In this protocol, videos of a 35 mm plate with worms are recorded for 1 minute and then processed using the software package. After processing, manual correction of tracks ensures that worm behaviors are recorded accurately without confounding tracking errors. Turn count and track duration data are used to determine turns per minute as a final readout. To ensure reproducibility, data is collected over a minimum of 3 independent replicate experiments, each with 40-50 animals scored, to achieve a combined final number of animals 120-150. This number is sufficient to discriminate small differences in swimming behavior from control worms. Some worms have motor deficits too severe to be captured by swimming assays. For example, if the animals placed in a liquid medium curl instead of performing the expected thrashing response, this assay will not record those movements accurately and another movement assay, such as radial locomotion, may better capture those motility defects. The provided protocol uses a commercially available imaging system (see Table of Materials for more details), but other worm tracking systems may provide a similar utility, with some being open source12. Previously published methods describe manual scoring of worm thrashing13. While the automated analysis produces a number of metrics for each individual worm, detection of body bends which is measured in thrashes per minute, provides consistent results between experiments and tracks well with conventional scoring of worm thrashes by eye.

Amyotrophic Lateral Sclerosis (ALS) is a debilitating, aging-related neurodegenerative disease with a particular impact on motor neurons. The disease features loss of motor neurons in the brain and spinal cord and progressive motor impairment. This results in major functional disability and premature death, typically within 3-5 years after diagnosis1. Mutations in at least 38 genes can cause ALS; however, most patients with ALS accumulate ubiquitinated inclusions of the protein TDP-43 as their primary pathology in neurons and glial cells2,3,4. A number of animal models have been developed to study the underlying mechanisms causing or contributing to ALS in vivo (reviewed in5). In C. elegans, these models include genetic loss-of-function mutations in homologs of ALS-causing genes or transgenic expression of human ALS genes. There are numerous advantages to modeling ALS in C. elegans. C. elegans are a tractable simple animal with a differentiated nervous system, well-characterized behavioral paradigms, and considerable genetic homology to humans6,7. Many tools exist for working with C. elegans, including robust genome editing capabilities, in vivo fluorescent reporters of neurodegeneration, RNAi screening paradigms, tractable genetics, and established behavioral and phenotypic assays. C. elegans models of ALS recapitulate aspects of human disease, including accumulation of insoluble protein, neurodegeneration, and early death8,9. Further, motor dysfunction featuring disturbance of both crawling and swimming behaviors is present in many C. elegans ALS models.
This protocol describes two methods to characterize C. elegans motor phenotypes: the radial locomotion assay for evaluating crawling on a solid surface and the assessment of swimming in liquid (thrashing) using the WormLab automated tracking and analysis. These sensitive methods for characterizing motor deficits allow comparisons of severity and offer tools for measuring suppression and enhancements of motor phenotypes. The radial locomotion assay quantifies differences in crawling motility (sinusoidal movement on a solid surface) among populations of worms. This assay takes advantage of C. elegans natural unstimulated exploration behavior by placing worms in a single location on a plate and marking their final location after a given period of time10. Alternatively, swimming in liquid (thrashing) assays count body bends of individual worms over a set period of time. The manual counting of body bends by the human eye is time-intensive and typically exhibits considerable variability between experimenters. The use of computer-assisted automated tracking and analysis can eliminate much of that variability. In addition to the characterization of baseline motor impairment of ALS models, both radial locomotion and swimming assays can detect modulation of distinct locomotor phenotypes from genetic or small molecule interventions. These methods have utility for studying ALS&#160;models and any C. elegans strain that exhibits altered motility.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>C. elegans</td>
			<td>Caenorhabditis Genetics Center (CGC)</td>
			<td>-</td>
			<td>Aquire your strains as desired, N2 is a useful control strain</td>
		</tr>
		<tr>
			<td>Disposable pasteur pipets, borosilicate glass</td>
			<td>VWR</td>
			<td>14673-010</td>
			<td>Glass pipet used to create worm pick - hold glass pipette in one hand and ~1&#34; of platinum wire (held by pliers) in the other over a flame to join.&#160;</td>
		</tr>
		<tr>
			<td>Disposable petri dishes, 35x10mm</td>
			<td>VWR</td>
			<td>10799-192</td>
			<td>Assay plates for WormLab Imaging System</td>
		</tr>
		<tr>
			<td>Disposable petri dishes, 60x15mm</td>
			<td>VWR</td>
			<td>25384-090</td>
			<td>Stock plates for worms</td>
		</tr>
		<tr>
			<td>Disposable petri dishes, 100x15mm</td>
			<td>VWR</td>
			<td>25384-302</td>
			<td>Standard radial locomotion assay plate</td>
		</tr>
		<tr>
			<td>Disposable petri dishes, 150x15mm</td>
			<td>VWR</td>
			<td>25384-326</td>
			<td>Longer time frame radial locomotion assay plate</td>
		</tr>
		<tr>
			<td>Dissecting microscope</td>
			<td>Leica</td>
			<td>M80</td>
			<td>Scope for maintaining worms and setting up radial locomotion assays</td>
		</tr>
		<tr>
			<td>Fine-tipped markers</td>
			<td>VWR</td>
			<td>52877-810</td>
			<td>Need at least 2 colors for radial locomotion assays. Fine tips required for accuracy.</td>
		</tr>
		<tr>
			<td>Flatbed Scanner</td>
			<td>Amazon</td>
			<td>Epson Perfection V850</td>
			<td>Optional for radial locomotion assay. Protocol assumes a resolution of 300dpi, most scanners would work fine</td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td>NIH</td>
			<td>-</td>
			<td>Optional free software provided by the NIH - https://imagej.nih.gov/ij/</td>
		</tr>
		<tr>
			<td>M9 buffer</td>
			<td>VWR</td>
			<td>IC113037012</td>
			<td>Medium used for swimming assay. Can be made from scratch, see WormBook: Maintenance of C. elegans</td>
		</tr>
		<tr>
			<td>NGM (Nematode Growth Medium)</td>
			<td>VWR</td>
			<td>76347-412</td>
			<td>Medium used to cultivate C. elegans. Can be made from scratch, see WormBook: Maintenance of C. elegans</td>
		</tr>
		<tr>
			<td>OP50&#160; bacteria</td>
			<td>Caenorhabditis Genetics Center (CGC)</td>
			<td>OP50</td>
			<td>Primary food source for C. elegans</td>
		</tr>
		<tr>
			<td>p1000 pipettor</td>
			<td>VWR</td>
			<td>76207-552</td>
			<td>Pipettor, used in swimming assay</td>
		</tr>
		<tr>
			<td>p1000 tips</td>
			<td>VWR</td>
			<td>83007-384</td>
			<td>Tips for pipettor, used in swimming assay</td>
		</tr>
		<tr>
			<td>Platinum wire, 0.2032mm diameter</td>
			<td>VWR</td>
			<td>BT136585-5M</td>
			<td>Fine gauge platinum wire used to create worm pick - hold glass pipette in one hand and ~1&#34; of platinum wire (held by pliers) in the other over a flame to join.&#160;</td>
		</tr>
		<tr>
			<td>Ruler</td>
			<td>VWR</td>
			<td>56510-001</td>
			<td>Need to score radial locomotion assays</td>
		</tr>
		<tr>
			<td>WormLab Imaging System</td>
			<td>MBF Bioscience</td>
			<td>WormLab</td>
			<td>The Imaging System includes WormLab hardware (bright field stage, camera, and housing) and WormLab software. https://www.mbfbioscience.com/wormlab-imaging-system</td>
		</tr>
	</tbody>
</table>

evaluation of motor impairment in c. elegans models of amyotrophic lateral sclerosis

The neurodegenerative disease amyotrophic lateral sclerosis (ALS) features progressive loss of motor neurons accompanied by muscle weakness and motor impairment that worsens with time. While considerable advances have been made in determining genetic drivers of ALS for a subset of patients, the majority of cases have an unknown etiology. Further, the mechanisms underlying motor neuron dysfunction and degeneration are not well understood; therefore, there is an ongoing need to develop and characterize representative models to study these processes. Caenorhabditis elegans can adapt their movement to the physical constraints of their surroundings, with two primary movement paradigms studied in a laboratory environment- crawling on a solid surface and swimming in liquid. These represent a complex interplay between sensation, motor neurons, and muscles. C. elegans models of ALS can exhibit impairment in one or both of these movement paradigms. This protocol describes two sensitive assays for evaluating motility in C. elegans: an optimized radial locomotion assay measuring crawling on a solid surface and an automated method for tracking and analyzing swimming in liquid (thrashing). In addition to the characterization of baseline motor impairment of ALS models, these assays can detect suppression or enhancement of the phenotypes from genetic or small molecule interventions. Thus, these methods have utility for studying ALS&#160;models and any C. elegans strain that exhibits altered motility.

Both radial locomotion and swimming assays offer sensitive detection of motility impairment (Figure 4 and Figure 5). To investigate the mechanisms underlying pathological TDP-43 in ALS, C. elegans models have been developed that express wild-type or ALS-mutant human TDP-43 pan-neuronally. These animals display molecular and cellular characteristics reminiscent of ALS, including motor dysfunction9. Importantly, they exhibit moderate motility impairment with the expression of wild-type human TDP-43 and more severe motility impairment in animals expressing ALS-mutant TDP-43 using both radial locomotion and swimming assays. Some mutant or transgenic animals will have greater impairment in crawling than swimming, or vice versa. By using two different motility assays, a clearer picture of phenotypic differences between strains is obtained.
Radial locomotion 
When placed on a seeded agar plate, C. elegans explore their surroundings, including seeking out the boundaries of their food source. Radial locomotion assays are one way to take advantage of that behavior as a metric for physical fitness. By analyzing the locomotor behavior (crawling on a solid surface) in a controlled and quantifiable way, the radial locomotor assay offers a simple and effective tool for assessing the severity of motor deficits and other motor-related phenotypes. Radial locomotion assays capture differences in motility in moderately or severely impaired ALS model worms and offer a baseline to compare modulation of motility phenotypes or changes in motility with age (Figure 6). This strategy can be applied to quantify the crawling behavior of any strain that has altered movement from wild type (N2) or control worms. However, this method may not be a good choice to assess animals unable to crawl normally, such as roller mutants or animals that are paralyzed. Typically, wild-type worms will present an average displacement between 200-300 &#181;m/min when raised and tested at 20 &#176;C. The example data presented in Figure 4 show expected results comparing N2, two different transgenic strains expressing wild-type human TDP-43 with a mild phenotype [TDP-43(WT-mild), CK402 (bkIs402[Psnb-1::TDP-43; Pmyo-3::GFP])] or stronger phenotype [TDP-43(WT-moderate), CK410 (bkIs402[Psnb-1::TDP-43; Pmyo-2::GFP])], a transgenic strain expressing mutant human TDP-43 with a severe phenotype [TDP-43(M337V), CK423 (bkIs423[Psnb-1::TDP-43; Pmyo-2::GFP]), and a transgenic strain expressing another neurodegenerative-disease associated protein, wild-type human tau [tau(WT), CK144 (bkIs144[Paex-3::tau(4R1N); Pmyo-2::GFP])]. The two strains expressing wild-type TDP-43 have different degrees of impairment as detected by radial locomotion. TDP-43(WT-low) is not significantly different from N2, while TDP-43(WT-high) does exhibit clear differences in motility. TDP-43(M337V) and tau (WT) strains have more severe impairments in motility.
Swimming assay 
C. elegans engage in stereotypical swimming (thrashing) movement when immersed in liquid. Immediately upon submersion, worms begin to characteristically bend head and tail towards each other with a bend angle of approximately 45&#176;, with the angle vertex being the midpoint of the worm. Worms alternate bending in the ventral and dorsal directions. One thrash, as measured by the software, represents the movement of going from straight to a body bend angle of 20&#176; or greater, irrespective of directionality (angle thresholding can be adjusted post-processing within the Analysis and Plotting window [Workflow | Analyze Data | Body Shape | Bending Angle | Mid-Point | Amplitude Threshold: # degrees]. The swimming assay described here uses automated computer-based tracking and analysis to provide unbiased scoring of swimming activity. It is expected that wild-type (N2) worms will average between 150-200 thrashes per min when raised and recorded at 20 &#176;C. The example data presented in Figure 5, shows expected results comparing N2, two different transgenic strains expressing wild-type human TDP-43 with a mild phenotype [TDP-43(WT-mild), CK402 (bkIs402[Psnb-1::TDP-43; Pmyo-3::GFP])] or stronger phenotype [TDP-43(WT-moderate), CK410 (bkIs402[Psnb-1::TDP-43; Pmyo-2::GFP])], and a transgenic strain expressing another neurodegenerative-disease associated protein, wild-type human tau [tau(WT), CK144 (bkIs144[Paex-3::tau(4R1N); Pmyo-2::GFP])]. The transgenic strain expressing ALS-mutant TDP-43 [TDP-43(M337V)] does not thrash in liquid, and so it is denoted on the graph as ND (no data). This assay can discriminate different phenotypes than the radial locomotion assay shown in Figure 4. For example, in the radial locomotion assay (Figure 4), TDP-43(WT-low) was not significantly different from N2. However, in the swimming assay (Figure 5), both TDP-43(WT-low) and TDP-43(WT-high) are significantly different from N2, as well as significantly different from one another. Also, despite both TDP-43(M337V) and tau(WT) having severe crawling impairments by radial locomotion (Figure 4), only tau(WT) is able to thrash enough to be tracked by the software (Figure 5). TDP-43(M337V) animals are unable to thrash, and software-based analysis does not detect or track these worms accurately. Thus, data on these worms were not collected (ND, no data).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62699/62699fig01.jpg" /> 
Figure 1: Radial locomotion assay workflow. Panels A-E show the generalized steps of the radial locomotion assay. The steps are as follows: (A) NGM Plates, with OP50 seeded to the edges, are prepared by labeling and marking central dot, (B) worms are placed in the center of the agar as marked by the central dot, (C) worms are allowed to move freely for a set amount of time, (D) plate is flipped bottom up and the final location of each worm is marked in a different color than the central dot, (E) The distance from the center dot to each final worm location in measured either by hand or digitally. <a href="https://www.jove.com/files/ftp_upload/62699/62699fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62699/62699fig02.jpg" /> 
Figure 2: Digital measurement of final location using ImageJ.&#160;Distance from center measurements can be measured by hand or digitally, to measure digitally using ImageJ (A)&#160;scan backside of the plate with a ruler in the frame. (B) Draw a known length using the line tool. (C) Use the known length drawn in (B) to set scale [Analyze | Set Scale&#8230;]. (D)&#160;Use the line tool to draw a line from the center point to a final location mark, repeat for each mark. (E)&#160;A paintbrush line marking the first mark measured aids in scoring (squiggly black line near the 1 mark). Final location marks are numbered in yellow to illustrate the directionality of scoring. (F)&#160;Measurements are recorded in the Results window. Save results elsewhere for statistical analysis. <a href="https://www.jove.com/files/ftp_upload/62699/62699fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62699/62699fig03.jpg" /> 
Figure 3: Materials and hardware adjustments for using the software. (A)&#160;Materials used for the software-assisted swimming assay. (B)&#160;General set up of the software and needed materials, note that the housing is in the raised position. (C) The lens mounting bracket height can be adjusted. This image shows the adjustable track and a tape indicator marking the preferred height for performing swimming assays. It is necessary to adjust the brightfield light and camera focus before recording. (D) A 35 mm assay plate is placed on the stage, animals in M9 are added to the plate, and a 1 min timer is started. (E)&#160;It is sometimes advantageous to gently swirl the plate to move worms closer to the center - observe location on the live capture screen; alternatively, a few drops of M9 from a pipettor can be used to separate worms. (F) Adjust the focus ring on the camera lens, observe worms on the screen to determine optimal focus. <a href="https://www.jove.com/files/ftp_upload/62699/62699fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62699/62699fig04.jpg" /> 
Figure 4: Radial locomotion assays detect differences in crawling speed. Unstimulated dispersion of developmentally staged L4 larvae was measured using the radial locomotion assay described above and graphed as &#181;m/min traveled (A-B).&#160;The same data plotted in (A)&#160;and&#160;(B) demonstrate two different possible graphical presentations.&#160;In (A),&#160;data are displayed as a bar graph, making relative differences between strains clearer. In (B),&#160;the final displacement of each worm isplotted within the graph, allowing the variation within the population to be better visualized. To evaluate significance among strains tested, one-way analysis of variance (ANOVA) with Tukey&#39;s multiple comparison test was used. **p=0.0022, ****p&#60;0.0001, ns=not significant. TDP-43(M337V) and tau(WT) are also significantly different from N2, p&#60;0.0001. Error bars in (A)&#160;are standard error of the mean (SEM) and in (B)&#160;are standard deviation. <a href="https://www.jove.com/files/ftp_upload/62699/62699fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/62699/62699fig05.jpg" /> 
Figure 5: Swimming assays detect differences of thrashing in liquid. Rates of swimming (thrashing or undulation frequency in liquid) were measured using unbiased computer-assisted scoring and analysis described above and graphed as thrashes/min (A-B). Same data plotted in (A)&#160;and (B). In (A),&#160;data are graphed as a bar graph, which makes relative differences between strains easier to see. In (B),&#160;data from each individual worm scored are plotted within the graph, allowing the variation within the population to be better visualized. To evaluate significance among strains tested, a one-way analysis of variance (ANOVA) with Tukey&#39;s multiple comparison test was used. ****p&#60;0.0001, ns=not significant. TDP-43(WT-moderate) and tau(WT) are also significantly different from N2, p&#60;0.0001.Error bars in (A)&#160;are standard error of the mean (SEM) and in (B)&#160;are standard deviation. <a href="https://www.jove.com/files/ftp_upload/62699/62699fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/62699/62699fig06.jpg" /> 
Figure 6:&#160;Mutant TDP-43 worms travel less than wild-type worms. Plates showing a representative difference in the final location of staged L4 N2 (wild-type) and TDP-43(M337V) (CK423 (bkIs423[Psnb-1::TDP-43; Pmyo-2::GFP]), a strain expressing mutant human TDP-43 with severely impaired motor function after 1 hour of unstimulated crawling at room temperature. Plates are marked with a red dot for the center point and blue dots for the final location of the animals. <a href="https://www.jove.com/files/ftp_upload/62699/62699fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1: Recording swimming behavior using the imaging and tracking software. (A)&#160;The tracking workflow and the location of the video capture icon. (B) The Video Capture window is shown in Live View, and the most important aspects are highlighted. The green &#34;Live view&#34; words will change to a red &#34;Recording&#34; when the record button is activated. <a href="https://www.jove.com/files/ftp_upload/62699/Currey2021_JoVEManuscript_SuppFigure1.pdf" target="_blank">Please click here to download this File.</a>
Supplementary Figure 2: Preparing and tracking swimming behavior. This figure shows a series of screenshots to aid in setting up a video image sequence for tracking. The general protocol for preparing a video sequence is to open each workflow menu in this order: Import Image Sequence | Set Sequence Info | Adjust Image | Detect and Track | Save Project. The Analyze Data window can only be used after a project file is tracked. (A) shows an opened .avi (video) file after importing the sequence into the software from within the Track workflow. (B)&#160;The Set Sequence Info window provides a place for notes, set/change the scale, and to check the metadata for a video sequence. Instructions for changing the scale are found in this window and should be followed when the camera is lowered or raised (B). Image (C) shows the Adjust Image window settings. (D)&#160;Screenshot showing the detection tab of the Detect and Track window. Detecting worms is used to train the software and must be set for each video sequence. Additional detection parameters can be set here if desired. (E) Screenshot of Tracking tab of the Detect and Track window with the recommended settings for tracking swimming behavior. This is the last step in setting up a video sequence. Save the sequence as a project using the Save Project window. Repeat these steps for each video sequence. Settings can be saved as a configuration to reduce workload and ensure that all sequences are treated the same (not shown). To track project files for analysis, navigate to the Batch Workflow. (F)&#160;shows the icon used to navigate to the Batch workflow, and (G)&#160;shows the batch processing window, highlights the add and start buttons, and shows the expected appearance of a project file that has been tracked. <a href="https://www.jove.com/files/ftp_upload/62699/Currey2021_JoVEManuscript_SupFigure2.pdf" target="_blank">Please click here to download this File.</a>
Supplementary Figure 3: Swimming behavior analysis. After a project file has been tracked, it can be opened using [File | Open Project]. Worms will appear in green as shown in (A). Scrubbing through the video offers a quick check that the video processed as expect, green highlighting will disappear while scrubbing. The Data Analysis and Plotting window is opened from the Analyze Data workflow item. (B) shows the Data Analysis and Plotting window with Position view opened (default), all tracks are highlighted. Below the datapoints is a plot showing the recorded track for each highlighted worm. The track summary analysis is used to calculated turns per minute. (C-D) show the track summary report and export details. (E-F) show the track summary in a spreadsheet and how to calculate turns per minute, the output measured in this assay. <a href="https://www.jove.com/files/ftp_upload/62699/Currey2021_JoVEManuscript_SuppFigure3.pdf" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Evaluation of Motor Impairment in C. elegans Models of Amyotrophic Lateral Sclerosis at JoVE.com

Evaluation of Motor Impairment in C. elegans Models of Amyotrophic Lateral Sclerosis

This protocol describes two sensitive assays for discriminating among mild, moderate, and severe motor impairment in C. elegans models of amyotrophic lateral sclerosis, with general utility for C. elegans strains, with altered motility.

Evaluation of Motor Impairment in C. elegans Models of Amyotrophic Lateral Sclerosis

The neurodegenerative disease amyotrophic lateral sclerosis (ALS) features progressive loss of motor neurons accompanied by muscle weakness and motor ...

evaluation-motor-impairment-c-elegans-models-amyotrophic-lateral

Research

JoVE Journal

Neuroscience

3.9K Views.  Veterans Affairs Puget Sound Health Care System. This protocol describes two sensitive assays for discriminating among mild, moderate, and severe motor impairment in C. elegans models of amyotrophic lateral sclerosis, with general utility for C. elegans strains, with altered motility.

Video: Evaluation of Motor Impairment in C. elegans Models of Amyotrophic Lateral Sclerosis

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 ...

Automated Quantification of Synaptic Fluorescence in C. elegans

Synapse strength refers to the amplitude of postsynaptic responses to presynaptic neurotransmitter release events, and has a major impact on overall neural circuit function. Synapse strength critically depends on the abundance of neurotransmitter receptors clustered at synaptic sites on the postsynaptic membrane. Receptor levels are established developmentally, and can be altered by receptor trafficking between surface-localized, subsynaptic, and intracellular pools, representing important mechanisms of synaptic plasticity and neuromodulation. Rigorous methods to quantify synaptically-localized neurotransmitter receptor abundance are essential to study synaptic development and plasticity. Fluorescence microscopy is an optimal approach because it preserves spatial information, distinguishing synaptic from non-synaptic pools, and discriminating among receptor populations localized to different types of synapses. The genetic model organism Caenorhabditis elegans is particularly well suited for these studies due to the small size and relative simplicity of its nervous system, its transparency, and the availability of powerful genetic techniques, allowing examination of native synapses in intact animals.
Here we present a method for quantifying fluorescently-labeled synaptic neurotransmitter receptors in C. elegans. Its key feature is the automated identification and analysis of individual synapses in three dimensions in multi-plane confocal microscope output files, tabulating position, volume, fluorescence intensity, and total fluorescence for each synapse. This approach has two principal advantages over manual analysis of z-plane projections of confocal data. First, because every plane of the confocal data set is included, no data are lost through z-plane projection, typically based on pixel intensity averages or maxima. Second, identification of synapses is automated, but can be inspected by the experimenter as the data analysis proceeds, allowing fast and accurate extraction of data from large numbers of synapses. Hundreds to thousands of synapses per sample can easily be obtained, producing large data sets to maximize statistical power. Considerations for preparing C. elegans for analysis, and performing confocal imaging to minimize variability between animals within treatment groups are also discussed. Although developed to analyze C. elegans postsynaptic receptors, this method is generally useful for any type of synaptically-localized protein, or indeed, any fluorescence signal that is localized to discrete clusters, puncta, or organelles.
The procedure is performed in three steps: 1) preparation of samples, 2) confocal imaging, and 3) image analysis. Steps 1 and 2 are specific to C. elegans, while step 3 is generally applicable to any punctate fluorescence signal in confocal micrographs.

Automated Quantification of Synaptic Fluorescence in C. elegans

The abundance of neurotransmitter receptors clustered at synapses strongly influences synaptic strength. This method quantifies fluorescently-labeled neurotransmitter receptors in three dimensions with single-synapse resolution in C. elegans, allowing hundreds of synapses to be rapidly characterized within a single sample without distortions introduced by z-plane projection.

Synapse strength refers to the amplitude of postsynaptic responses to presynaptic neurotransmitter release events, and has a major impact on overall ...

C. elegans Tracking and Behavioral Measurement

We have developed instrumentation, image processing, and data analysis techniques to quantify the locomotory behavior of C. elegans as it crawls on the surface of an agar plate. For the study of the genetic, biochemical, and neuronal basis of behavior, C. elegans is an ideal organism because it is genetically tractable, amenable to microscopy, and shows a number of complex behaviors, including taxis, learning, and social interaction1,2. Behavioral analysis based on tracking the movements of worms as they crawl on agar plates have been particularly useful in the study of sensory behavior3, locomotion4, and general mutational phenotyping5. Our system works by moving the camera and illumination system as the worms crawls on a stationary agar plate, which ensures no mechanical stimulus is transmitted to the worm. Our tracking system is easy to use and includes a semi-automatic calibration feature. A challenge of all video tracking systems is that it generates an enormous amount of data that is intrinsically high dimensional. Our image processing and data analysis programs deal with this challenge by reducing the worms shape into a set of independent components, which comprehensively reconstruct the worms behavior as a function of only 3-4 dimensions6,7. As an example of the process we show that the worm enters and exits its reversal state in a phase specific manner.

C. elegans Tracking and Behavioral Measurement

We have developed a video-rate tracking microscope system that can record and quantify C. elegans behavior at high resolution and high speeds. We have also developed computational methods to reduce the dimensionality of the worm images to a fundamental set of measurements that completely describe the shape of the worm.

We have developed instrumentation, image processing, and data analysis techniques to quantify the locomotory behavior of C. elegans as it crawls on the ...

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

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

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

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

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

In vivo Neuronal Calcium Imaging in C. elegans

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

In vivo Neuronal Calcium Imaging in C. elegans

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

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

Visualizing Neuroblast Cytokinesis During C. elegans Embryogenesis

This protocol describes the use of fluorescence microscopy to image dividing cells within developing Caenorhabditis elegans embryos. In particular, this protocol focuses on how to image dividing neuroblasts, which are found underneath the epidermal cells and may be important for epidermal morphogenesis. Tissue formation is crucial for metazoan development&#160;and relies on external cues from neighboring tissues. C. elegans is an excellent model organism to study tissue morphogenesis in vivo due to its transparency and simple organization, making its tissues easy to study via microscopy. Ventral enclosure is the process where the ventral surface of the embryo is covered by a single layer of epithelial cells. This event is thought to be facilitated by the underlying neuroblasts, which provide chemical guidance cues to mediate migration of the overlying epithelial cells. However, the neuroblasts are highly proliferative and also may act as a mechanical substrate for the ventral epidermal cells. Studies using this experimental protocol could uncover the importance of intercellular communication during tissue formation, and could be used to reveal the roles of genes involved in cell division within developing tissues.

Visualizing Neuroblast Cytokinesis During C. elegans Embryogenesis

This protocol describes how to image dividing cells within a tissue in Caenorhabditis elegans embryos. While several protocols describe how to image cell division in the early embryo, this protocol describes how to image cell division within a developing tissue during mid late embryogenesis.

This protocol describes the use of fluorescence microscopy to image dividing cells within developing Caenorhabditis elegans embryos. In particular, this ...

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 ...

Automated Analysis of C. elegans Swim Behavior Using CeleST Software

Dissecting the neuronal and neuromuscular circuits that regulate behavior remains a major challenge in biology. The nematode Caenorhabditis elegans has proven to be an invaluable model organism in helping to tackle this challenge, from inspiring technological approaches,&#160;building the human brain connectome, to actually shedding light on the specific molecular drivers of basic functional patterns. The bulk of the behavioral studies in C. elegans have been performed on solid substrates. In liquid, animals exhibit behavioral patterns that include movement at a range of speeds in 3D, as well as partial body movements, such as a posterior curl without anterior shape change, which introduce new challenges for quantitation. The steps of a simple procedure, and use of a software that enables high-resolution analysis of C. elegans swim behavior, are presented here. The software, named CeleST, uses a specialized computer program that tracks multiple animals simultaneously and provides novel measures of C. elegans locomotion in liquid (swimming). The measures are mostly grounded in animal posture and based on mathematics used in computer vision and pattern recognition, without computational requirements for threshold cut-offs. The software tool can be used to both assess overall swimming prowess in hundreds of animals from combined small batch trials and to reveal novel phenotypes even in well-characterized genetic mutants. The preparation of specimens for analysis with CeleST is simple and low-tech, enabling wide adaptation by the scientific community. Use of the computational approach described here should therefore contribute to the greater understanding of behavior and behavioral circuits in the C. elegans model.

Automated Analysis of C. elegans Swim Behavior Using CeleST Software

An efficient and simple methodology for computer-based analysis of nematode swimming behavior in liquid is described. The method requires little to no investment for C. elegans laboratories. The hardware used is standard, and the computer software for the behavioral analysis (CeleST) is an open source one.

Dissecting the neuronal and neuromuscular circuits that regulate behavior remains a major challenge in biology. The nematode Caenorhabditis elegans has ...

Comprehensive Autopsy Program for Individuals with Multiple Sclerosis

We describe a rapid tissue donation program for individuals with multiple sclerosis (MS) that requires scientists and technicians to be on-call 24/7, 365 days a year. Participants consent to donate their brain and spinal cord. Most patients were followed by neurologists at the Cleveland Clinic Mellen Center for MS Treatment and Research. Their clinical courses and neurological disabilities are well-characterized. Soon after death, the body is transported to the MS Imaging Center, where the brain is scanned in situ by 3 T magnetic resonance imaging (MRI). The body is then transferred to the autopsy room, where the brain and spinal cord are removed. The brain is divided into two hemispheres. One hemisphere is immediately placed in a slicing box and alternate 1 cm-thick slices are either fixed in 4% paraformaldehyde for two days or rapidly frozen in dry ice and 2-methylbutane. The short-fixed brain slices are stored in a cryopreservation solution and used for histological analyses and immunocytochemical detection of sensitive antigens. Frozen slices are stored at -80 &#176;C and used for molecular, immunocytochemical, and in situ hybridization/RNA scope studies. The other hemisphere is placed in 4% paraformaldehyde for several months, placed in the slicing box, re-scanned in the 3 T magnetic resonance (MR) scanner and sliced into centimeter-thick slices. Postmortem in situ MR images (MRIs) are co-registered with 1 cm-thick brain slices to facilitate MRI-pathology correlations. All brain slices are photographed and brain white-matter lesions are identified. The spinal cord is cut into 2 cm segments. Alternate segments are fixed in 4% paraformaldehyde or rapidly frozen. The rapid procurement of postmortem MS tissues allows pathological and molecular analyses of MS brains and spinal cords and pathological correlations of brain MRI abnormalities. The quality of these rapidly-processed postmortem tissues (usually within 6 h of death) is of great value to MS research and has resulted in many high-impact discoveries.

Multiple sclerosis is an inflammatory demyelinating disease with no cure. Analysis of brain tissue provides important clues to understanding the pathogenesis of the disease. Here we discuss the methodology and downstream analysis of MS brain tissue collected through a unique rapid autopsy program in operation at the Cleveland Clinic.

We describe a rapid tissue donation program for individuals with multiple sclerosis (MS) that requires scientists and technicians to be on-call 24/7, 365 ...

3D Kinematic Analysis for the Functional Evaluation in the Rat Model of Sciatic Nerve Crush Injury

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of sciatic nerve injury rodent models. In this protocol, we describe a novel kinematic analysis method that uses a three-dimensional (3D) motion capture apparatus for functional evaluations using a rat sciatic nerve crush injury model. First, the rat is familiarized with treadmill walking. Markers are then attached to the designated bone landmarks and the rat is made to walk on the treadmill at the desired speed. Meanwhile, the posterior limb movements of the rat are recorded using four cameras. Depending on the software used, marker tracings are created using both automatic and manual modes and the desired data are produced after subtle adjustments. This method of kinematic analysis, which uses a 3D motion capture apparatus, offers numerous advantages, including superior precision and accuracy. Many more parameters can be investigated during the comprehensive functional evaluations. This method has several shortcomings that require consideration: The system is expensive, can be complicated to operate, and may produce data deviations due to skin shifting. Nevertheless, kinematic analysis using a 3D motion capture apparatus is useful for performing functional anterior and posterior limb evaluations. In the future, this method may become increasingly useful for generating accurate assessments of various traumas and diseases.

We introduce a kinematic analysis method that uses a three-dimensional motion capture apparatus containing four cameras and data processing software for performing functional evaluations during fundamental research involving rodent models.

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of ...

Evaluation of Motor Impairment in C. elegans Models of Amyotrophic Lateral Sclerosis

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