Search tips
Search criteria 


Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Methods. Author manuscript; available in PMC 2017 April 28.
Published in final edited form as:
PMCID: PMC5408735

Expanding the C. elegans toolbox into a toolshed

Since Sydney Brenner’s proposal to exploit Caenorhabditis elegans to unravel the genetic basis of metazoan development and the nervous system [1], technological developments have turned this model organism into one of the premier resources for studying multiple facets of cell biology. Several excellent compendia outline some of these methodological advancements [28]. In addition, the WormMethods section of WormBook ( provides a rich compilation of C. elegans methodology. The goal of this edition of Methods is to highlight recent innovations that have expanded the repertoire of techniques available to C. elegans investigators, and to update methods not covered extensively by these other references.

The study of gene function by transgenesis is one of the most powerful technologies available to C. elegans investigators. Recently, the application of genome editing has enhanced significantly the scope and precision of these methods. The first 8 articles of this edition of Methods provide added insight into these technologies. Waaijers and Boxem, discuss practical considerations in the use of the CRISPR/Cas9 system to engineer the genome. They describe non-homologous end-joining or homologous recombination methods in combination with modified DNA templates [9]. As an alternative approach, Yi et al. review the use of transcription activator-like effector nucleases (TALENs) for genome editing, especially in somatic tissues [10]. Another, powerful strategy for genome manipulation employs the Mos1 transposon to introduce single copy transgene insertions, gene deletions or gene conversions. These techniques were recently reviewed in Methods [11] and in other journals, and will not be described here [1215]. Kage-Nakadai et al. provide a simplified method for single or low copy transgene integration from extrachromosomal arrays by treatment with trimethylpsoralen and less DNA damaging, long-wave ultraviolet illumination [16]. Antibiotic selection for transgene maintenance or integration, a mainstay of mammalian transgenic systems, has finally made its way into C. elegans. Two groups report on the use of single or dual drug selection on solid or liquid media and after microinjection or microparticle bombardment [17,18]. Of major interest in C. elegans transgenesis is better spatial and temporal control of transgene expression. Hubbard describes intersectional expression strategies using the FLP or Cre recombinases to direct cell specific re-arrangement of transgenes flanked by FRT and loxP recombination sites, respectively [19]. Okazaki et al. detail an optogenetic technique that utilizes transgenes harboring light-driven proton pumps to selectively silence subsets of cells [20]. Adaptation of a microscope to deliver a laser pulse at the appropriate wavelength could be used to activate optogenetic transgenes or ones driven by heat-shock promoters in order to direct expression in single cells [21].

Forward genetic screens in C. elegans continue to provide important unbiased insights into biological pathways. While the screens can be tedious and labor-intensive, the biggest bottleneck has been mapping and identifying the mutants. Whole-genome DNA sequencing has helped streamline and accelerate this process. Hu outlines two different crossing strategies for maximizing the chances of identifying the correct mutation without SNP mapping [22].

C. elegans was one of the first organisms used to study the genetic basis of aging, especially in response to different types of stressors. One article examines methods for performing genotoxic stress assays using different mutagens [23], while two manuscripts enumerate techniques used to induce and assay proteotoxicity: heat shock [24] and transgenes encoding aggregation-prone proteins [25]. The effect of these and others stressors can have dramatic effects on the lifespan or healthspan of animals. Two papers describe the multiple methods for determining these parameters including assays amenable to automation [26,27].

Translational medicine attempts to solve clinical problems by bridging the gap between basic bench research and the bedside. Traditionally, translational medicine has relied on mammalian models of human disease. The ability to study normal and human disease-related physiological processes in C. elegans has transformed this organism into a powerful translational tool [28]. For example, elevations of blood pCO2 occur in a number of respiratory illnesses and this increase can have profound effects on cellular metabolism. Noam et al. outline several techniques to study the effects of hypercapnea in C. elegans and show that many of these responses are conserved in higher eukaryotes [29]. Obesity is a major public health concern. Wahlby et al. expand upon their earlier studies and describe Oil Red O staining and automated imaging methods to study fat metabolism in the nematode [30]. C. elegans has been used a model to study muscle physiology and movement extensively. In contrast, C. elegans can also be used to study quiescence, and Nagy et al. outline mounting and imaging technologies that permit quantitative analysis of this important behavioral phenotype [31]. Derangements of intracellular pH, redox status and levels of ATP, NADPH and calcium are important indicators of cellular injury and stress. Wang et al. show how these types of parameters can be studied in C. elegans in real-time by automated imaging using fluorescent ratiometric reporters [32]. Similarly, endocytosis plays a crucial role in diverse cellular processes including nutrient intake and cell signaling. Wang and Audhya describe transgenic approaches using fluorescently-tagged proteins and different imaging techniques to follow several steps in endomembrane trafficking [33].

High-throughput and high-content drug screening, once the domain of protein biochemists and cell biologists, are now being adapted for use in C. elegans for modeling different aspects of human disease or mammalian physiology [34]. Benson et al. details assay development for a semi-automated high-content drug screen [35]. While these phenotype-based screens are extremely powerful in identifying hits for lead development, they do not yield the actual drug targets. Miedel et al. provide a combined biochemical purification and mass spectroscopy approach to this problem by constructing a triple affinity tag that can be adapted for detection of drug-target or protein–protein interactions [36]. Live animal imaging also has become an important part of the drug discovery process in C. elegans, especially during the validation phase. Luke et al. describe a simple non-microfluidic means of immobilizing animals for relatively long periods of time that allow for widefield or confocal imaging following drug treatments [37].

Collectively, the authors of these manuscripts have shed more light on the vast array of experimental techniques now available to conduct studies in C. elegans. Combined with forward and reverse genetic approaches, C. elegans will continue to be one of the premier tools for understanding basic cellular biology under physiological and pathological conditions.

Contributor Information

Arjumand Ghazi, Department of Pediatrics, University of Pittsburgh School of Medicine,Pittsburgh, PA, United States. Department of Cell Biology, University of Pittsburgh School of Medicine, Pittsburgh, PA, United States. Department of Developmental Biology, University of Pittsburgh School of Medicine, Pittsburgh, PA, United States.

Judith Yanowitz, Department of Cell Biology, University of Pittsburgh School of Medicine, Pittsburgh, PA, United States. Department of Developmental Biology, University of Pittsburgh School of Medicine, Pittsburgh, PA, United States. Department of Obstetrics and Gynecology, University of Pittsburgh School of Medicine, Pittsburgh, PA, United States.

Gary A. Silverman, Department of Pediatrics, University of Pittsburgh School of Medicine, Pittsburgh, PA, United States. Department of Cell Biology, University of Pittsburgh School of Medicine, Pittsburgh, PA, United States.


1. Brenner S. Genetics. 1974;77:71–94. [PubMed]
2. Emmons SW. The Nematode Caenorhabditis elegans: The Genome. 1. Cold Spring Harbor Laboratory Press; Plainview: 1988.
3. Epstein HF, Shakes DC. Caenorhabditis elegans: Modern Biological Analysis of an Organism. Academic Press; San Diego, CA: 1995.
4. Hope IA. C. elegans: A Practical Approach. Oxford University Press; New York, NY: 1999.
5. Riddle DL, Blumenthal T, Meyer BJ, Priess JR. C. elegans II. Cold Spring Harbor Laboratory Press; Plainview, New York: 1977.
6. Rothman JH, Singson A. Caenorhabditis elegans: Molecular Genetics and Developement. 2. Academic Press; Waltham, MA: 2011.
7. Rothman JH, Singson A. Caenorhabditis elegans: Cell Biology and Physiology. 2. Academic Press; Waltham, MA: 2012.
8. Strange K. C. elegans: Mehods and Applications. Humana Press; Totowa, New Jersey: 2006.
9. Waaijers S, Boxem M. Methods. 2014;68:381–388. [PubMed]
10. Yi P, Li W, Ou G. Methods. 2014;68:389–396. [PubMed]
11. Robert VJ, Katic I, Bessereau JL. Methods. 2009;49:263–269. [PubMed]
12. Frokjaer-Jensen C, Davis MW, Ailion M, Jorgensen EM. Nat Methods. 2012;9:117–118. [PMC free article] [PubMed]
13. Frokjaer-Jensen C, Davis MW, Hopkins CE, Newman BJ, Thummel JM, Olesen SP, Grunnet M, Jorgensen EM. Nat Genet. 2008;40:1375–1383. [PMC free article] [PubMed]
14. Frokjaer-Jensen C, Davis MW, Sarov M, Taylor J, Flibotte S, LaBella M, Pozniakovsky A, Moerman DG, Jorgensen EM. Nat Methods. 2014;11:529–534. [PMC free article] [PubMed]
15. Robert VJ, Davis MW, Jorgensen EM, Bessereau JL. Genetics. 2008;180:673–679. [PubMed]
16. Kage-Nakadai E, Imae R, Yoshina S, Mitani S. Methods. 2014;68:397–402. [PubMed]
17. Cornes E, Quere CA, Giordano-Santini R, Dupuy D. Methods. 2014;68:403–408. [PubMed]
18. Semple JI, Lehner B. Methods. 2014;68:409–416. [PubMed]
19. Hubbard EJ. Methods. 2014;68:417–424. [PMC free article] [PubMed]
20. Okazaki A, Takahashi M, Toyoda N, Takagi S. Methods. 2014;68:425–430. [PubMed]
21. Churgin MA, He L, Murray JI, Fang-Yen C. Methods. 2014;68:431–436. [PMC free article] [PubMed]
22. Hu PJ. Methods. 2014;68:437–440. [PubMed]
23. Kessler Z, Yanowitz J. Methods. 2014;68:441–449. [PubMed]
24. Zevian SC, Yanowitz JL. Methods. 2014;68:450–457. [PMC free article] [PubMed]
25. Volovik Y, Marques FC, Cohen E. Methods. 2014;68:458–464. [PubMed]
26. Amrit FR, Ratnappan R, Keith SA, Ghazi A. Methods. 2014;68:465–475. [PubMed]
27. Keith SA, Amrit FR, Ratnappan R, Ghazi A. Methods. 2014;68:476–486. [PubMed]
28. Silverman GA, Luke CJ, Bhatia SR, Long OS, Vetica AC, Perlmutter DH, Pak SC. Pediatr Res. 2009;65:10–18. [PMC free article] [PubMed]
29. Zuela N, Friedman N, Zaslaver A, Gruenbaum Y. Methods. 2014;68:487–491. [PubMed]
30. Wahlby C, Lee Conery A, Bray MA, Kamentsky L, Larkins-Ford J, Sokolnicki KL, Veneskey M, Michaels K, Carpenter AE, O’Rourke EJ. Methods. 2014;68:492–499. [PMC free article] [PubMed]
31. Nagy S, Raizen DM, Biron D. Methods. 2014;68:500–507. [PMC free article] [PubMed]
32. Wang H, Karadge U, Humphries WH, Iv, Fisher AL. Methods. 2014;68:508–517. [PMC free article] [PubMed]
33. Wang L, Audhya A. Methods. 2014;68:518–528. [PMC free article] [PubMed]
34. Gosai SJ, Kwak JH, Luke CJ, Long OS, King DE, Kovatch KJ, Johnston PA, Shun TY, Lazo JS, Perlmutter DH, Silverman GA, Pak SC. PLoS ONE. 2010;5:e15460. [PMC free article] [PubMed]
35. Benson JA, Cummings EE, O’Reilly LP, Lee MH. Methods. 2014;68:529–535. [PubMed]
36. Miedel MT, Zeng X, Yates NA, Silverman GA, Luke CJ. Methods. 2014;68:536–541. [PMC free article] [PubMed]
37. Luke CJ, Niehaus JZ, O’Reilly LP, Watkins SC. Methods. 2014;68:542–547. [PMC free article] [PubMed]