Over the past 15 years, the free-living nematode, Caenorhabditis elegans has become an important model system for exploring eukaryotic innate immunity to bacterial and fungal pathogens. More recently, infection models using either natural or non-natural nematode viruses have also been established in C. elegans. These models offer new opportunities to use the nematode to understand eukaryotic antiviral defense mechanisms. Here we report protocols for the infection of C. elegans with a non-natural viral pathogen, vesicular stomatitis virus (VSV) through microinjection. We also describe how recombinant VSV strains encoding fluorescent or luciferase reporter genes can be used in conjunction with simple fluorescence-, survival-, and luminescence-based assays to identify host genetic backgrounds with differential susceptibilities to virus infection.

Assays for superoxide dismutase (SOD) and catalase (CAT) activities are widely employed to indicate antioxidant responses underlying the toxic effects of test chemicals. Yet, earlier studies mainly described the procedures as performed according to manufacturer’s instructions without modifications that are specific to any organisms. The present protocol describes the steps in analyzing the superoxide dismutase (SOD) and catalase (CAT) activities in C. elegans, which is a model organism that can be used to study effects of pharmaceutical compounds and environmental pollutants. The main steps include: (1) sample preparation; (2) total protein assay; (3) SOD activity assay; (4) CAT activity assay; and (5) medium list and formula, and also data analysis and performance notes.

The nematode Caenorhabditis elegans is a popular model organism for studies of developmental biology, neurology, ageing and other fields of basic research. Because many developmental processes are regulated by glycosaminoglyans (GAGs) on cell surfaces and in the extracellular matrix, methods to isolate and analyze C. elegans GAGs are needed. Such methods have previously been optimized for other species such as mice and zebrafish. After modifying existing purification protocols, we could recently show that the nematodes also produce chondroitin sulfate, in addition to heparan sulfate, thus challenging the view that only non-sulfated chondroitin was synthesized by C. elegans. We here present our protocol adapted for C. elegans. Since the purification strategy involves separation of non-sulfated and sulfated GAGs, it may also be useful for other applications where this approach could be advantageous.

Post-translational protein modifications (PTMs) orchestrate the activity of individual proteins and ensure their proper function. While modifications such as phosphorylation or glycosylation are well understood, more unusual modifications, including nitrosylation or AMPylation remain comparatively poorly characterized. Research on protein AMPylation–which refers to the covalent addition of an AMP moiety to the side chains of serine, threonine or tyrosine–has undergone a renaissance (Yarbrough et al., 2009; Engel et al., 2012; Ham et al., 2014; Woolery et al., 2014; Preissler et al., 2015; Sanyal et al., 2015; Truttmann et al., 2016; Truttmann et al., 2017). The identification and characterization of filamentation (fic) domain-containing AMPylases sparked new interest in this PTM (Kinch et al., 2009; Yarbrough et al., 2009). Based on recent in vivo and in vitro studies, we now know that secreted bacterial AMPylases covalently attach AMP to members of the Rho family of GTPases, while metazoan AMPylases modify HSP70 family proteins in the cytoplasm and the endoplasmic reticulum (ER) (Itzen et al., 2011; Hedberg and Itzen, 2015; Truttmann and Ploegh, 2017). AMPylation is thought to trap HSP70 in a primed yet transiently disabled state that cannot participate in protein refolding reactions (Preissler et al., 2015). In vitro AMPylation experiments are key to assess the activity, kinetics and specificity of protein AMPylation catalyzed by pro- and eukaryotic enzymes. These simple assays require recombinant AMPylases, target proteins (Rho GTPases, HSP70s), as well as ATP as a nucleotide source. Here, we describe strategies to qualitatively and quantitatively study protein AMPylation in vitro.

Cells and organisms face constant exposure to reactive oxygen species (ROS), either from the environment or as a by-product from internal metabolic processes. To prevent cellular damage from ROS, cells have evolved detoxification mechanisms. The activation of these detoxification mechanisms and their downstream responses represent an overlapping defense response that can be tailored to different sources of ROS to adequately adapt and protect cells. In this protocol, we describe how to measure the sensitivity to oxidative stress from two different sources, arsenite and tBHP, using the nematode C. elegans.

Single-molecule RNA fluorescence in situ hybridization (smFISH) is a technique to visualize individual RNA molecules using multiple fluorescently-labeled oligonucleotide probes specific to the target RNA (Raj et al., 2008; Lee et al., 2016a). We adapted this technique to visualize RNAs in the C. elegans whole adult worm or its germline, which enabled simultaneous recording of nascent transcripts at active transcription sites and mature mRNAs in the cytoplasm (Lee et al., 2013 and 2016b). Here we describe each step of the smFISH procedure, reagents, and microscope settings optimized for C. elegans extruded gonads.

Optogenetics is a powerful tool for manipulating neuronal activity with high temporal and spatial precision. In the nematode C. elegans optogentics is especially useful and easy to apply. This is because C. elegans is translucent, so its neurons are highly accessible to optic stimulation. In addition, many of its neurons can be exclusively targeted using cell-specific promoters. We have recently taken advantage of optogenetics to deliver artificial patterns of prolonged activation to a class of mechanosensory neurons, called touch receptor neurons (TRNs) in worms that lack touch sensation due to a genetic mutation. Our aim was to examine whether we can counteract the effects of sensory loss by artificially activating the sensory neurons. Here we describe in detail the various components of the protocol that we used. This consists of exposing worms expressing the light-sensitive ion channel Channelrohdopsin 2 (ChR2) in TRNs to long-term random flashes of light.

This protocol describes two biological assays to evaluate pathogenicity of Burkholderia cepacia complex (Bcc) strains against the nematode Caenorhabditis elegans. Specifically, these two assays allow one to identify if the under-investigated Bcc strains are able to kill the nematodes by intestinal colonization (slow killing assay, SKA) or by toxins production (fast killing assay, FKA). The principal differences between the two assays rely on the different killing kinetics for worms.

In this method, the intestinal lumen of Caenorhabditis elegans (C. elegans) is labeled with a fluorescent fluid-phase marker, Texas Red-dextran. Since dextran conjugates are membrane impermeable, animals fed with it show a red fluorescent signal in the lumen of the intestine. Texas Red-dextran in the lumen is not efficiently endocytosed by intestinal cells and is not effectively transported to the body cavity paracellularly. It is useful to determine whether round-shaped membrane structures are invaginations from the apical membrane or cytoplasmic vesicles. If the barrier function of the intestinal epithelium is impaired, Texas Red-dextran can leak from the intestinal lumen to the body cavity. Therefore, this method can be used to visualize apical membrane morphology in intestinal cells and to investigate the barrier properties of the intestinal epithelium.

The Caenorhabditis elegans (C. elegans) vulva is a well-established system to study organ development as the molecular mechanisms that govern its formation are conserved in animals. Of special interest is the EGFR/RAS/MAPK signaling pathway that is required for fate acquisition and morphogenesis of the vulva. let-23 encodes the sole homologue of the epidermal growth factor receptor (EGFR), is expressed at the plasma membrane of the vulval precursor cells (VPCs) and is activated by LIN-3 EGF at the end of the L3 larval stage to initiate vulva development. LET-23 activity can be modulated through altering its subcellular and plasma membrane localization. To study the trafficking of EGF receptor in a living organism, we created a functional LET-23::GFP translational reporter worm line (Haag et al., 2014) and quantified the mobile fraction of LET-23::GFP at the basolateral membrane of the VPCs by fluorescence recovery after photobleaching (FRAP). Here we describe the protocol for LET-23::GFP FRAP at the basolateral membrane of the VPCs and the data analysis using FIJI (ImageJ).