JoVE Logo
Authors
Contact Us

Sign In

A subscription to JoVE is required to view this content. Sign in or start your free trial.

In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

This study presents protocols for two semi-automated locomotor activity analysis approaches in C. elegans complex I disease gas-1(fc21) worms, namely, ZebraLab (a medium-throughput assay) and WormScan (a high-throughput assay) and provide comparative analysis among a wide array of research methods to quantify nematode behavior and integrated neuromuscular function.

Abstract

Caenorhabditis elegans is widely recognized for its central utility as a translational animal model to efficiently interrogate mechanisms and therapies of diverse human diseases. Worms are particularly well-suited for high-throughput genetic and drug screens to gain deeper insight into therapeutic targets and therapies by exploiting their fast development cycle, large brood size, short lifespan, microscopic transparency, low maintenance costs, robust suite of genomic tools, mutant repositories, and experimental methodologies to interrogate both in vivo and ex vivo physiology. Worm locomotor activity represents a particularly relevant phenotype that is frequently impaired in mitochondrial disease, which is highly heterogeneous in causes and manifestations but collectively shares an impaired capacity to produce cellular energy. While a suite of different methodologies may be used to interrogate worm behavior, these vary greatly in experimental costs, complexity, and utility for genomic or drug high-throughput screens. Here, the relative throughput, advantages, and limitations of 16 different activity analysis methodologies were compared that quantify nematode locomotion, thrashing, pharyngeal pumping, and/or chemotaxis in single worms or worm populations of C. elegans at different stages, ages, and experimental durations. Detailed protocols were demonstrated for two semi-automated methods to quantify nematode locomotor activity that represent novel applications of available software tools, namely, ZebraLab (a medium-throughput approach) and WormScan (a high-throughput approach). Data from applying these methods demonstrated similar degrees of reduced animal activity occurred at the L4 larval stage, and progressed in day 1 adults, in mitochondrial complex I disease (gas-1(fc21)) mutant worms relative to wild-type (N2 Bristol) C. elegans. This data validates the utility for these novel applications of using the ZebraLab or WormScan software tools to quantify worm locomotor activity efficiently and objectively, with variable capacity to support high-throughput drug screening on worm behavior in preclinical animal models of mitochondrial disease.

Introduction

Caenorhabiditis elegans is widely recognized as an outstanding model in neuroscience based on it having 302 neurons that coordinate all worm behaviors, including mating, feeding, egg-laying, defecation, swimming, and locomotion on solid media1. These hermaphroditic nematodes are also widely used to understand a wide array of human disease mechanisms, made possible by its well-characterized genome and high homology of ~80% genes between C. elegans and humans2,3,4. C. elegans have long been used to interrogate human mitochondrial disease5,6,7,8,9,10, which is a highly genetically and phenotypically heterogeneous group of inherited metabolic disorders that share impaired capacity to generate cellular energy and often clinically present with substantially impaired neuromuscular function, exercise intolerance, and fatigue11,12,13,14.To this end, the use of C. elegans models enable preclinical modeling of quantitative aspects of animal activity and neuromuscular function in different genetic subtypes of mitochondrial disease, as well as their response to candidate therapies that may improve their neuromuscular function and overall activity.

Neuromuscular activity in C. elegans is objectively measurable by a range of experimental methodologies, including both manual and semi-automated approaches that allow functional analyses in either solid or liquid media (Table 1)1,15. Accurate quantitation of C. elegans activity has proven important to enable discoveries related to function and development of the muscular and nervous system16,17,18. This study summarizes and compares experimental requirements, advantages, and limitations of 17 different assays that can be performed in research laboratories to evaluate neuromuscular function and activity on four key outcomes in C. elegans diseases models, both at the baseline at a range of developmental stages and ages as well as in response to candidate therapies (Table 1). Indeed, the study provides a detailed overview of the range of available experimental approaches to characterize rates of C. elegans thrashing (body bends per minute), locomotor activity, pharyngeal pumping, and chemotaxis-in each case specifying the experimental and analytic methodology used, the advantages and limitations of each method, the equipment and software needed to perform and analyze each assay, and the throughput capacity of each method to support its use for high-throughput genetic or drug screening purposes. The throughput capacity of each assay is described as low, medium, or high based on the experimental protocol complexity, including worm maintenance, processing time, the use of single or multi-well plates, and/or experimenter time needed to complete the experimental setting and data analyses.

Manual analyses of thrashing19, locomotor activity20, pharyngeal pumping17,21, and chemotaxis22,23 are well-established methodologies to evaluate worm activity that require a stereomicroscope24. While measuring thrashing activity of worms requires analysis in liquid media to determine the frequency of body bends per minute, worm locomotor activity may be measured either on solid media or in liquid media. However, manual analyses of individual worm activity are inherently time-consuming and involves unavoidable user-generated bias. Automation of worm activity analyses minimizes user-generated bias and can greatly increase experimental throughput25. Video recordings of worm thrashing activity in liquid media can be analyzed using wrMTrck, an ImageJ plugin26. However, the original experimental settings that were developed for wrMTrck limited its utility, since too many worms in a single liquid drop led to overlapping of worms that made accurate tracking difficult. While this experimental limitation has been resolved27, the wrMTrck method is not able to support high-throughput screening.

A range of methods exist to quantify worm locomotor activity at baseline and in response to candidate therapies in C. elegans mitochondrial disease models. These include ZebraLab (ViewPoint Life Sciences), Tierpsy Tracker28, wide field-of-view nematode tracking platform (WF-NTP)29, WormMotel, WormWatcher30, WormLab31, Infinity Chip32, and WMicrotracker One33 (Table 1). These methods enable concurrent analysis of locomotion in multiple worm strains or conditions, typically on multi-well plates, thereby supporting higher-throughput drug screening applications. Some of these methods have unique considerations that may limit or enhance their general utility, such as the need for expensive equipment versus open-access software, and varying ease of performing experimental protocols. Overall, no single experimental system or protocol is ideally suited to all C. elegans locomotor activity experiments. Rather, it is important to carefully choose which method is best suited to the specific investigator's experimental goals and requirements.

Pharyngeal pumping represents another important outcome to assess neuromuscular activity in C. elegans. The C. elegans pharynx is composed of 20 muscle cells, 20 neurons, and 20 other cells that enable ingestion of Escherichia coli (E. coli) at the anterior end of the worm's alimentary tract34,35,36. Several manual methods have been established to determine pharyngeal pumping rates17,21,37,38. Most methods are based on the use of a stereomicroscope and camera to visualize and record pharyngeal pumping frequency with direct counting by the experimental observer21. Automated pharyngeal pumping rate analysis is possible by performing an extracellular recording termed electropharyngeogram (EPG), which provides additional information on the duration of each pump39. Pharyngeal pumping rate analysis is also possible in a microfluidic system, WormSpa, where individual worms are confined in chambers40,41. A commercial method available to facilitate analysis of the pharyngeal pump rate is the ScreenChip System (InVivo Biosystems), which measures, visualizes, and analyzes the neuromuscular aspects of feeding behavior in a single worm that is immobilized in a custom chip. This pharyngeal pumping quantitation approach can be used to assess both neuronal and physiological responses to drugs, aging, and other factors42,43,44,45.

Chemotaxis describes the movement of C. elegans in response to an odorant placed away from the worms in a defined area of the nematode growth media (NGM) plate. Assessing the chemotaxis response provides an integrated measure of worm neuronal and neuromuscular activity that is quantifiable by observing and measuring the physical distance traveled by worms toward the odorant in a defined time period46. The Multi-Worm Tracker is an automatic method that can be used to improve the experimental efficiency of quantifying the distance traveled by worms toward an attractant or from a repellant47.

Here, the detailed protocol for two novel, semi-automated methods established for quantifying worm activity is described. The first approach utilizes ZebraLab a commercial software that was originally developed to study swimming activity of Danio rerio (zebrafish), for a novel medium-throughput application to quantify overall locomotor activity in liquid media of C. elegans based on pixel changes during movement (Table 1, Figure 1). Data output is quickly obtained from a large number of concurrent conditions and samples analyzed on a glass slide, although this method is not suitable to a multi-well plate format. The second approach is a novel adaptation of the WormScan methodology48,49 (Figure 2), which uses a flatbed scanner to create a differential image of two sequential scans that can variably be used with open-source software to enable semi-automated quantitative analysis of integrated physiologic outcomes such as fecundity and survival. Here, a novel high-throughput adaptation of the WormScan methodology to quantify worm locomotor activity in liquid media in populations of fifteen larval-stage 4 (L4) worms per well of a 96-well, flat-bottom plate was developed. This semi-automated and low-cost WormScan methodology can be readily adapted to high-throughput drug screens, as well as to analyses of various animal stages and ages48,49.

Here, the protocol and efficacy of analyzing C. elegans locomotor activity using both ZebraLab and WormScan semi-automated methods is demonstrated in a well-established C. elegans model for mitochondrial complex I disease, gas-1(fc21). gas-1 (K09A9.5 gene) is an ortholog of human NDUFS2 (NADH: ubiquinone oxidoreductase core (iron-sulfur protein) subunit 2) (Figure 3). The C. elegans gas-1(fc21) mutant strain carries a homozygous p.R290K missense mutation in the human ortholog of NDUFS250, causing significantly decreased fecundity and lifespan, impaired respiratory chain oxidative phosphorylation (OXPHOS) capacity51, as well as decreased mitochondrial mass and membrane potential with increased oxidative stress5,8. Despite its well-established use over the past two decades to study mitochondrial disease, locomotor activity of gas-1(fc21) mutants was not previously reported. Here, ZebraLab and WormScan methods were applied to independently quantify the locomotor activity of gas-1(fc21) as compared to wild-type (WT, N2 Bristol) worms, both as a way to validate the methods as well as to demonstrate their comparative utility and efficiency of the experimental protocols and informatics analyses. ZebraLab software allowed rapid quantitation of several concurrent conditions of worm locomotor activity in C. elegans mitochondrial disease models, with potential application for targeted drug screening or validation studies. WormScan analysis, in particular, is well-suited to readily enable high-throughput drug screens of compound libraries and prioritize leads that improve the animal neuromuscular function and the locomotor activity in preclinical C. elegans models of primary mitochondrial disease.

Protocol

1. Worm locomotor activity analysis in liquid media on glass slides using ZebraLab software

  1. Nematode growth and handling
    1. Grow C. elegans on Petri plates containing nematode growth media (NGM) and spread with Escherichia coli OP50 as food source. Maintain worm culture at 20 °C, as previously described8.
    2. Synchronize worms performing a timed egg lay52 and study worms at the desired stage. In this protocol, L4 stage worms were analyzed.
    3. Grow control and mutant worm strains on NGM plates with and without drug treatments to be tested or buffer control. To evaluate the drug treatment effects, prepare the desired drug stock concentration in S. basal solution; spread the calculated specific volume onto the NGM plates and allow it to dry. Transfer the worms at a specific larval or adult stage and maintain on the drug treatment plate for the desired duration before analysis.
  2. Experimental set-up of worms for locomotor activity video recording and ZebraLab analysis
    1. Pick 5 synchronized L4 worms per strain and condition using a worm pick. Pipette a single 20 µL drop of S. basal solution onto a glass slide located under a stereomicroscope connected to a camera and transfer 5 worms into it (Figure 1A,B). Transfer the 5 worms from the Petri dish containing NGM and E. coli OP50 to the liquid drop only at the moment preceding the recording.
      NOTE: Continue to maintain the other worms on the Petri dish until the prior video is recorded. This will avoid the damage caused on the other worms due to the drying up of the 20 µl drops during the procedure (dry time ~15-20 min).
    2. Pipette multiple drops on one slide to obtain multiple technical replicates (Figure 1A). Select worms from different NGM plates (biological replicate). Do not use cover slip.
    3. Adjust the microscope's working distance to visualize the complete area of a single drop. Set and maintain a low video resolution (<1024 x 768) to upload the files in the software.
    4. Allow worms to acclimate on the slide at room temperature for 1 min before recording.
    5. Record the worm swim activity in 1 drop for 1 min at 15 frames per second (fps). Repeat the imaging for each additional drop on the plate.
  3. C. elegans locomotor activity recording analysis in ZebraLab software
    1. Use the option ZEBRALAB AVI to upload videos to the software. Click on the option Quantization with AVI Files (Figure 1C).
    2. To create a new protocol, select File > Generate Protocol, and then add the number of areas selected for the analysis. Choose Location Count: 1.
    3. Open Protocol Parameters and select 1 min in the Experiment Duration window. Select a different experimental duration for different experimental durations. Select or deselect the window No Time Bin and choose Integration Period, depending on the data output desired. In this study No Time Bin was selected (Figure 1D).
      ​NOTE: Time bin is the time over which the activity will be averaged.
    4. If a protocol was already created, select Open Protocol and select the saved protocol (in .vte format).
    5. Select File and Open a Movie to upload each individual video file that was previously recorded.
    6. Select the Area icon indicated in Figure 1E (black arrow) to build a single area of detection and create an area around the whole liquid drop where worms are located. Click on Select, then the green circle icon (gray arrow) under Areas > Build > Clear marks.
      NOTE: The activity of all worms in the defined drop will be detected in the selected area (Figure 1E,F).
    7. Go to Calibration > Draw scale (Figure 1E) and draw a horizontal line from the left to the right of the video area. Indicate the real distance to calibrate. Then select Apply to Group.
    8. Unselect the icon selected to build the area (arrow in Figure 1E) and select or deselect Transparent.
      NOTE: In this study, Transparent was selected and gave better results.
    9. Adjust Detection Sensitivity and Activity Threshold to allow the detection of all the different C. elegans worm strains analyzed.
      NOTE: In this experiment, the Detection Sensitivity was set at 8 with Burst and Freezing values of 15 and 2, respectively (Figure 1F).
    10. Set Display Scale at 70 to visualize the track made by the animal while activity analysis is underway. Then select Apply to Group (Figure 1F).
    11. Click on Experiment > Execute > Save as, and then on Start. A window opens. Choose Do you want to process the video media at maximum computer speed? to analyze the video quickly (e.g., a 1 min video recording is analyzed by the in ZebraLab software in 5 s).
    12. Another window opens: Running Experiment; click on Start to proceed with the experiment.
    13. After the video recording is complete, the analysis stops. Click on Experiment > Stop. This saves the activity analyzed from a single drop in a spreadsheet.
    14. Repeat the analysis for each video of individual drop. Each drop is one technical replicate.
  4. Output and analysis of ZebraLab data
    NOTE: After the experiment, data from each video is individually saved as separate spreadsheets in the chosen folder. In the data output file, the integrated activity level of all worms moving in an individual drop is recorded as pixel changes under actinteg.
    1. Open each spreadsheet obtained from the analysis of each video. Compile them manually into a single file.
      ​Normalize the mutant and wild-type data to a percent of control. Here, statistical analyses were performed to compare mean activity levels between groups.

2. Worm locomotor activity analysis in liquid media in 96-well plate format by WormScan software analysis

  1. Nematode growth and handling
    1. Grow C. elegans as described in section 1.1.1.
    2. Synchronize worms as described in section 1.1.2.
    3. Grow worms on specific media as described in section 1.1.3 until L4 stage or day 1 adult.
  2. Experimental set-up of worms in 96-well plate for WormScan activity analysis
    1. Add 50 µL of 2% weight per volume of E. coli OP50 in liquid suspension in S. basal medium to each well of a 96-well, clear, flat-bottom, microplate, as previously described49,53.
    2. Under a stereomicroscope, manually pick 15 synchronized worms at L4 stage or day 1 adult from their NGM plates into liquid media within each experimental well of the 96-well microplate. Allow the worms to acclimate to the liquid media for 20 min before scanning.
      NOTE: Other animal stages and ages can be readily substituted for study.
  3. WormScan activity analysis in 96-well plate and data export to spreadsheet.
    1. Scan each 96-well, clear, flat-bottom, microplate twice sequentially using a standard flatbed scanner, with less than 10 s between scans.
      NOTE: Here, the photo scanner with resolution of 1,200 dots/in and 16-bit grayscale was used to produce jpeg images. Time required to scan four 96-well plates using the photo scanner is less than 10 min.
    2. Align the two sequential scans (Figure 2A) using open source software49.
      NOTE: The software generates a difference image to evaluate pixel changes between the two sequential images for a region of interest (Figure 2B) and a relative WormScan Score. This WormScan Score is equivalent to changes in locomotor response based on the light intensity produced by the scanner when set to a pixel threshold of 5 (Figure 2C).
    3. Export the data from WormScan as a spreadsheet. Save the spreadsheet containing the data to the local computer. Normalize the data as percentage of control (POC) and compare across biological replicate experiments for diverse mutant or treatment conditions. Perform statistical analysis to compare mutant and control means using student t-test.

Results

Analysis of C. elegans locomotor activity in the liquid media could easily capture an integrated phenotype of mitochondrial disease worm models that may not be easily quantifiable on solid media. ZebraLab was used to quantify locomotor activity of the well-established mitochondrial complex I disease gas-1(fc21) strain relative to WTworms in liquid media at the L4 larval stage. The activity of 5 worms in a single liquid drop was recorded over 1 min, with a total of 19 videos (technical replicate...

Discussion

Here, the study summarized detailed information and rationales for studying C. elegans neuromuscular activity at the level of diverse outcomes, including worm thrashing, locomotion, pharyngeal pumping, and chemotaxis. The comparison of 16 different activity analysis methodologies was performed in terms of the relative throughput, advantages, and limitations of quantify nematode activities in a single worm or worm populations at different ages and experimental durations. Among these, two novel adaptations and...

Disclosures

M.L., N.D.M., N.S., and E.N.-O. have no relevant financial disclosures. M.J.F. is a co-founder of MitoCUREia, Inc., scientific advisory board member with equity interest in RiboNova, Inc., and scientific board member as paid consultant with Khondrion, and Larimar Therapeutics. M.J.F. has previously been or is currently engaged with several companies involved in mitochondrial disease therapeutic preclinical and/or clinical stage development as a paid consultant (Astellas [formerly Mitobridge] Pharma Inc., Cyclerion Therapeutics, Epirium Bio, Imel Therapeutics, Minovia Therapeutics, NeuroVive, Reneo Therapeutics, Stealth BioTherapeutics, Zogenix, Inc.) and/or a sponsored research collaborator (AADI Therapeutics, Cardero Therapeutics, Cyclerion Therapeutics, Imel Therapeutics, Minovia Therapeutics Inc., Mission Therapeutics, NeuroVive, Raptor Therapeutics, REATA Inc., RiboNova Inc., Standigm Therapeutics, and Stealth BioTherapeutics).

Acknowledgements

We are grateful to Anthony Rosner, PhD., with his organizational support for the early preparation of this project, and to Erin Haus for contributing to protocol analysis. This work was funded by the Juliet's Cure FBXL4 Mitochondrial Disease Research Fund, the Jaxson Flynt C12ORF65 Research Fund, and the National Institutes of Health (R01-GM120762, R01-GM120762-08S1, R35-GM134863, and T32-NS007413). The content is solely the responsibility of the authors and does not necessarily represent the official views of the funders or the National Institutes of Health.

Materials

NameCompanyCatalog NumberComments
C. elegans wild isolate Caenorhabditis Genetics Center (CGC)N2 Bristol
CameraOlympusDP73
gas-1(fc-21)CGCCW152
Microscope slidesThermoFisher4951PLUS
Nematode Growth Medium (NGM)Research Products International Corp.N81800-1000.0
OP50 Escherichia coliCGCUracil auxotroph E. coli strain
Petri dishes (60 mm) VWR international25373-085
S. BasalVWR 5.85 g NaCl, 1 g K2 HPO4, 6 g KH2PO4, and 5 mg cholesterol, in 1 l H2OVWR 101175-162, 103467-156, EM1.09828.1000, 97061-660
ScannerEPSONV800
StereomicroscopeOlympusMVX10 microscope
96-well flat bottom VWR international29442-056
WormScan softwareMathew et al. 45S1 Standalone Java platformSoftware for automation of difference image of scanned plates
ZebraLab softwareViewPointSoftware for automated quantization and tracking of zebrafish behavior, designed by ViewPoint (http://www.viewpoint.fr/en/p/software/zebralab-zebrafish-behavior-screening) and here applied to C. elegans. This system is applicable for high-throughput behavioral analysis

References

  1. Husson, S. J., Costa, W. S., Schmitt, C., Gottschalk, A. Keeping track of worm trackers. WormBook. , 1-17 (2013).
  2. Shaye, D. D., Greenwald, I. OrthoList: a compendium of C. elegans genes with human orthologs. PLoS One. 6 (5), 20085 (2011).
  3. van Ham, T. J., et al. C. elegans model identifies genetic modifiers of alpha-synuclein inclusion formation during aging. PLoS Genetics. 4 (3), 1000027 (2008).
  4. Kim, W., Underwood, R. S., Greenwald, I., Shaye, D. D. OrthoList 2: A new comparative genomic analysis of human and Caenorhabditis elegans genes. Genetics. 210 (2), 445-461 (2018).
  5. Dingley, S., et al. Mitochondrial respiratory chain dysfunction variably increases oxidant stress in Caenorhabditis elegans. Mitochondrion. 10 (2), 125-136 (2010).
  6. Polyak, E., Zhang, Z., Falk, M. J. Molecular profiling of mitochondrial dysfunction in Caenorhabditis elegans. Methods in Molecular Biology. 837, 241-255 (2012).
  7. McCormick, E., Place, E., Falk, M. J. Molecular genetic testing for mitochondrial disease: from one generation to the next. Neurotherapeutics. 10 (2), 251-261 (2013).
  8. McCormack, S., et al. Pharmacologic targeting of sirtuin and PPAR signaling improves longevity and mitochondrial physiology in respiratory chain complex I mutant Caenorhabditis elegans. Mitochondrion. 22, 45-59 (2015).
  9. Polyak, E., et al. N-acetylcysteine and vitamin E rescue animal longevity and cellular oxidative stress in pre-clinical models of mitochondrial complex I disease. Molecular Genetics and Metabolism. 123 (4), 449-462 (2018).
  10. Guha, S., et al. Pre-clinical evaluation of cysteamine bitartrate as a therapeutic agent for mitochondrial respiratory chain disease. Human Molecular Genetics. 28 (11), 1837-1852 (2019).
  11. Gorman, G. S., et al. Prevalence of nuclear and mitochondrial DNA mutations related to adult mitochondrial disease. Annals of Neurology. 77 (5), 753-759 (2015).
  12. Mancuso, M., Orsucci, D., Filosto, M., Simoncini, C., Siciliano, G. Drugs and mitochondrial diseases: 40 queries and answers. Expert Opinion on Pharmacotherapy. 13 (4), 527-543 (2012).
  13. Gai, X., et al. Mutations in FBXL4, encoding a mitochondrial protein, cause early-onset mitochondrial encephalomyopathy. American Journal of Human Genetics. 93 (3), 482-495 (2013).
  14. Dillin, A., et al. Rates of behavior and aging specified by mitochondrial function during development. Science. 298 (5602), 2398-2401 (2002).
  15. Yemini, E., Jucikas, T., Grundy, L. J., Brown, A. E., Schafer, W. R. A database of Caenorhabditis elegans behavioral phenotypes. Nature Methods. 10 (9), 877-879 (2013).
  16. Bargmann, C. I., Avery, L. Laser killing of cells in Caenorhabditis elegans. Methods in Cell Biology. 48, 225-250 (1995).
  17. Avery, L., Horvitz, H. R. Effects of starvation and neuroactive drugs on feeding in Caenorhabditis elegans. Journal of Experimental Zoology. 253 (3), 263-270 (1990).
  18. Chalfie, M., et al. The neural circuit for touch sensitivity in Caenorhabditis elegans. Journal of Neuroscience. 5 (4), 956-964 (1985).
  19. Ghosh, R., Emmons, S. W. Episodic swimming behavior in the nematode C. elegans. Journal of Experimental Biology. 211 (23), 3703-3711 (2008).
  20. Rankin, C. H., Beck, C. D., Chiba, C. M. Caenorhabditis elegans: a new model system for the study of learning and memory. Behavioural Brain Research. 37 (1), 89-92 (1990).
  21. Avery, L. Motor neuron M3 controls pharyngeal muscle relaxation timing in Caenorhabditis elegans. Journal of Experimental Zoology. 175, 283-297 (1993).
  22. Ward, S. Chemotaxis by the nematode Caenorhabditis elegans: identification of attractants and analysis of the response by use of mutants. Proceedings of the National Academy of Sciences of the United States of America. 70 (3), 817-821 (1973).
  23. Bargmann, C. I., Thomas, J. H., Horvitz, H. R. Chemosensory cell function in the behavior and development of Caenorhabditis elegans. Cold Spring Harbor Symposia on Quantitative Biology. 55, 529-538 (1990).
  24. Anne, C. H. Behavior. WormBook: The Online Review of C. elegans Biology. 2005-2018, (2006).
  25. Biston, M. C., et al. An objective method to measure cell survival by computer-assisted image processing of numeric images of Petri dishes. Physics in Medicine & Biology. 48 (11), 1551-1563 (2003).
  26. Nussbaum-Krammer, C. I., Neto, M. F., Brielmann, R. M., Pedersen, J. S., Morimoto, R. I. Investigating the spreading and toxicity of prion-like proteins using the metazoan model organism C. elegans. Journal of Visualized Experiments: JoVE. (95), e52321 (2015).
  27. Shi, W., Qin, J., Ye, N., Lin, B. Droplet-based microfluidic system for individual Caenorhabditis elegans assay. Lab on a Chip. 8 (9), 1432-1435 (2008).
  28. Javer, A., et al. An open-source platform for analyzing and sharing worm-behavior data. Nature Methods. 15 (9), 645-646 (2018).
  29. Koopman, M., et al. Assessing motor-related phenotypes of Caenorhabditis elegans with the wide field-of-view nematode tracking platform. Nature Protocols. 15 (6), 2071-2106 (2020).
  30. Churgin, M. A., et al. Longitudinal imaging of Caenorhabditis elegans in a microfabricated device reveals variation in behavioral decline during aging. eLife. 6, 26652 (2017).
  31. Angstman, N. B., Kiessling, M. C., Frank, H. G., Schmitz, C. High interindividual variability in dose-dependent reduction in speed of movement after exposing C. elegans to shock waves. Frontiers in Behavioral Neuroscience. 9, 12 (2015).
  32. Rahman, M., et al. NemaLife chip: a micropillar-based microfluidic culture device optimized for aging studies in crawling C. elegans. Scientific Reports. 10 (1), 16190 (2020).
  33. Bianchi, J. I., Stockert, J. C., Buzzi, L. I., Blazquez-Castro, A., Simonetta, S. H. Reliable screening of dye phototoxicity by using a Caenorhabditis elegans fast bioassay. PLoS One. 10 (6), 0128898 (2015).
  34. Albertson, D. G., Thomson, J. N. The pharynx of Caenorhabditis elegans. Philososophical Transactions of the Royal Society of London. Series B, Biological Sciences. 275 (938), 299-325 (1976).
  35. Raizen, D. M., Avery, L. Electrical activity and behavior in the pharynx of Caenorhabditis elegans. Neuron. 12 (3), 483-495 (1994).
  36. Avery, L., You, Y. J. C. elegans feeding. WormBook. , 1-23 (2012).
  37. Morck, C., Rauthan, M., Wagberg, F., Pilon, M. pha-2 encodes the C. elegans ortholog of the homeodomain protein HEX and is required for the formation of the pharyngeal isthmus. Developmental Biology. 272 (2), 403-418 (2004).
  38. Song, B. M., Avery, L. Serotonin activates overall feeding by activating two separate neural pathways in Caenorhabditis elegans. TheJournal of Neuroscience. 32 (6), 1920-1931 (2012).
  39. Avery, L., Raizen, D., Lockery, S. Electrophysiological methods. Methods in Cell Biology. 48, 251-269 (1995).
  40. Kopito, R. B., Levine, E. Durable spatiotemporal surveillance of Caenorhabditis elegans response to environmental cues. Lab in a Chip. 14 (4), 764-770 (2014).
  41. Lee, K. S., et al. Serotonin-dependent kinetics of feeding bursts underlie a graded response to food availability in C. elegans. Nature Communications. 8, 14221 (2017).
  42. Brinkmann, V., Ale-Agha, N., Haendeler, J., Ventura, N. The Aryl Hydrocarbon Receptor (AhR) in the aging process: Another puzzling role for this highly conserved transcription factor. Frontiers in Physiology. 10, 1561 (2019).
  43. Huang, C., et al. Intrinsically aggregation-prone proteins form amyloid-like aggregates and contribute to tissue aging in Caenorhabditis elegans. eLife. 8, 43059 (2019).
  44. Zhu, B., et al. Functional analysis of epilepsy-associated variants in STXBP1/Munc18-1 using humanized Caenorhabditis elegans. Epilepsia. 61 (4), 810-821 (2020).
  45. Weeks, J. C., Robinson, K. J., Lockery, S. R., Roberts, W. M. Anthelmintic drug actions in resistant and susceptible C. elegans revealed by electrophysiological recordings in a multichannel microfluidic device. International Journal of Parasitology. Drugs and Drug Resistance. 8 (3), 607-628 (2018).
  46. Haroon, S., et al. Multiple molecular mechanisms rescue mtDNA disease in C. elegans. Cell Reports. 22 (12), 3115-3125 (2018).
  47. Swierczek, N. A., Giles, A. C., Rankin, C. H., Kerr, R. A. High-throughput behavioral analysis in C. elegans. Nature Methods. 8 (7), 592-598 (2011).
  48. Mathew, M. D., Mathew, N. D., Ebert, P. R. WormScan: a technique for high-throughput phenotypic analysis of Caenorhabditis elegans. PLoS One. 7 (3), 33483 (2012).
  49. Mathew, M. D., et al. Using C. elegans forward and reverse genetics to identify new compounds with anthelmintic activity. PLoS Neglected Tropical Diseases. 10 (10), 0005058 (2016).
  50. Kayser, E. B., Morgan, P. G., Hoppel, C. L., Sedensky, M. M. Mitochondrial expression and function of GAS-1 in Caenorhabditis elegans. Journal Biological Chemistry. 276 (23), 20551-20558 (2001).
  51. Falk, M. J., Kayser, E. B., Morgan, P. G., Sedensky, M. M. Mitochondrial complex I function modulates volatile anesthetic sensitivity in C. elegans. Current Biology. 16 (16), 1641-1645 (2006).
  52. Kwon, Y. J., Guha, S., Tuluc, F., Falk, M. J. High-throughput BioSorter quantification of relative mitochondrial content and membrane potential in living Caenorhabditis elegans. Mitochondrion. 40, 42-50 (2018).
  53. Hirsh, D., Oppenheim, D., Klass, M. Development of the reproductive system of Caenorhabditis elegans. Developmental Biology. 49 (1), 200-219 (1976).
  54. Steele, W. B., Mole, R. A., Brooks, B. W. Experimental protocol for examining behavioral response profiles in larval fish: Application to the Neuro-stimulant caffeine. Journal of Visualized Experiments: JoVE. (137), e57938 (2018).
  55. Carlsson, G., Blomberg, M., Pohl, J., Orn, S. Swimming activity in zebrafish larvae exposed to veterinary antiparasitic pharmaceuticals. Environmental Toxicology and Pharmacology. 63, 74-77 (2018).
  56. Yang, X., et al. High-throughput screening in larval zebrafish identifies novel potent sedative-hypnotics. Anesthesiology. 129 (3), 459-476 (2018).

Reprints and Permissions

Request permission to reuse the text or figures of this JoVE article

Request Permission

Explore More Articles

C ElegansMitochondrial DiseaseLocomotor ActivitySemi automated AssaysDrug ScreeningMotility AssaysNeuromuscular SystemExperimental ConditionsGlass Slide AnalysisZebraLab SoftwareVideo QuantificationActivity ThresholdDetection SensitivityWorm BehaviorSynchronization Techniques

This article has been published

Video Coming Soon

JoVE Logo

Privacy

Terms of Use

Policies

Contact Us

Recommend to library

JoVE NEWSLETTERS

Research

Education

Authors

Librarians

ABOUT JoVE

Copyright © 2025 MyJoVE Corporation. All rights reserved