Proper positioning through attachment to the body is essential for tissue and organ function. We had previously reported the lethal mutant, ven (for ventral cord abnormal), exhibiting nerve cord displacement or detachment (Shioi et al., 2001). Herein, we report our identification of the ven-1 gene through genetic/physical mapping as well as transformation rescue of the Ven phenotype.

We cloned a genomic fragment that rescued the Ven phenotype (Figure 1A). All the mutated alleles were detected in the genomic region. From cDNA libraries, we isolated a 345 bp cDNA clone corresponding to the C53A.2 gene assigned on WormBase. RNA interference experiments using this cDNA sequence reproduced the Ven phenotype. The ven-1 transcript encoded a polypeptide of 86 amino acid residues, including the signal sequence at the N-terminus, suggesting that VEN-1 is a secreted protein (Figure 1B). VEN-1 appears to be conserved in Nematoda but not in other phyla.

To gain insight into ven-1 expression, we constructed ven-1p::GFP, in which the 472 bp upstream region of the ven-1 open reading frame was fused to green fluorescent protein (GFP) cDNA (Figure 2A). The GFP signal was first detected at the comma stage before ventral enclosure and remained until the 3-fold stage (Figure 2B). In the comma stage, it was expressed in hypodermal cells: Abpl/raapppp (H7), P1/2 L/R, P3/4 L/R, P5/6 L/R, P7/8 L/R, P9/10 L/R, and P11/12.

Next, we constructed ven-1::HA, in which the C-terminus of VEN-1 in the rescue fragment was tagged with an influenza hemagglutinin (HA) peptide. The ven-1::HA sequence rescued the Ven phenotype, and the HA epitope was detected in hypodermal cells (Figure 2B). The signal consisted of short oblique parallel lines arranged in four rows along the lateral body wall, corresponding to the position of muscle cells. The periodic nature of staining suggests that it may correspond to structures such as M lines and/or dense bodies (Moerman and Williams, 2006).

To examine the possibility of VEN-1 being a secreted protein, we tested whether forced expression of the ven-1 transcript under the control of heterologous promoters could rescue the Ven phenotype in tissues. The forced expression of the transcript in body wall muscle tissue rescued the Ven phenotype as efficiently as the genomic fragment did (Figure 2C). Its forced expression in neuronal and intestinal tissues also resulted in partial rescue of the phenotype. These results support the hypothesis that VEN-1 is a secreted protein.

We also examined the temporal requirement of ven-1 activity by expressing the transcript using a heat shock promoter. Its expression after egg laying completely rescued the Ven phenotype (Figure 2D). However, the recovery ratio gradually decreased after the comma stage, and expression of the transcript after the 1.5-fold stage failed to rescue the Ven phenotype. Therefore, ven-1 expression is required during the embryonic stages when nerves and muscles differentiate and attach to the body wall.

The ven-1 mutants also showed muscle attachment defects (Mua) (Shioi et al., 2001). Therefore, we examined the localization of attachment components of body wall muscles in ven-1 mutants using monoclonal antibodies against intermediate filaments (MH4), perlecan (MH3), and myotactin (MH46) (Labouesse, 2006; Moerman and Williams, 2006). Staining with the antibodies was detected in the displaced muscles (Figure 3), suggesting that ven-1 mutants have major defects in the hypodermis.

Previous studies have reported that a mutation in the gene encoding myosin-4 (unc-54) suppresses the Mua phenotype of mua-1 mutants (Plenefisch et al., 2000). However, we found that the ven-1(nc25); unc-54(n190) double mutant had a stronger phenotype than each single mutant, with its penetrance of the Ven phenotype being higher than that of the ven-1 mutant and its growth being much slower than that of the unc-54 mutant. These results indicate that the function of VEN-1 is different from that of MUA-1.

In C. elegans, RNAi is a useful tool for decreasing the expression of a gene of interest. In order to properly interpret the results of an RNAi experiment it is important to quantify the level of gene knockdown typically using quantitative RT-PCR (qPCR). In some cases, qPCR primers will detect the RNA produced by the RNAi bacteria thereby leading to a false measurement of the mRNA levels of the gene of interest. To circumvent this problem, it is essential to design primer pairs in which at least one of the primers binds outside of the region targeted by the RNAi clone. As an example, we treated worms with an RNAi clone targeting the mitochondrial superoxide dismutase gene sod-2. When we compared gene expression to worms grown on EV RNAi using qPCR primers that bind within the sequence targeted by the RNAi clone, we detected no decrease in sod-2 mRNA level (Figure 1). However, when we examined sod-2 mRNA levels using primers that bind outside of the region targeted by the RNAi clone, we found that the sod-2 RNAi treatment did effectively reduce sod-2 mRNA levels (Figure 1). This illustrates the importance of designing primers such that they only amplify mRNA and not the RNA generated by the RNAi bacteria in order to properly measure the levels of knockdown.

Our standard approach to primer design for qPCR:
   1. Go to www.wormbase.org
   2. Search for gene of interest (e.g. sod-2)
   3. Click on “Location”. This will show you if there are multiple transcripts. You may want to target a specific transcript or try to design primers that target all of them.
   4. Click on “Sequences”. Under “Coding Sequence” all of the transcripts for the gene of interest will be displayed.
   5. Copy the spliced RNA + UTR sequence to microsoft word (in order to keep a record of where your primers are targeting).
   6. If you are designing qPCR primers to measure RNAi knockdown, identify the region of the transcript where the RNAi clone is targeting. At least one primer should be outside of this region. We find that often primers targeting the untranslated regions (UTR) bind outside of the RNAi targeted region.
   7. We use Primer 3 to pick primers (bioinfo.ut.ee/primer3-0.4.0/). If possible, it is best to pick at least one primer that overlaps spans two different exons. By designing a primer that contains parts of two exons, this primer will not be able to bind to DNA, which has introns present. We normally aim for the amplified region to be 100-200 bp. In the word file, we normally indicate the primer locations and note the expected size of the amplified region.
   8. As a final check, we run the primer pair through an in silico PCR website: http://genome.ucsc.edu/cgi-bin/hgPcr. Be sure to select C. elegans. Because the primers are specific for spliced RNA, they should not amplify any DNA sequences.

To allow everybody a detailed analysis of nematode embryogenesis, we designed a program to analyze our 4-D recordings on standard Windows- or Mac-based computers.

Below access is given to download (i) a detailed manual explaining how to use this program including several figures and many questions that can be addressed plus some background information, (ii) The Viewer Program (executable JAR file), (iii) videos of 4 different nematodes, and (iv) an extensive review of nematode development (Schierenberg and Sommer, 2013).

For storage and handling of the 4 videos 8 GB RAM and about 6-8 GB of free space on your hard disc is required. Like with the expensive commercial 4-D software you can move forward and reverse through the videos quickly or image by image plus up and down within the developing embryo. The program allows studying cell division by division and events within blastomeres as far as visible with Nomarski optics. In addition, running time is shown and time intervals between selected events (e.g., cell cycle lengths) can be easily measured.

The material provided should allow teaching respective lab courses even by instructors not particularly familiar with nematode development. In contrast to live material developmental steps can be analyzed again and again in slow motion. It is suited for individual work in the classroom or at home, for instance, by addressing questions asked in the manual.

From the films of 3 other nematodes (their phylogenetic relationships are indicated in the manual) it becomes obvious that various deviations from the C. elegans pattern exist already during early embryogenesis–some of them rather prominent–demonstrating considerable evolutionary modifications within the phylum Nematoda. A more detailed discussion of this issue can be found in the added review.

How to circumvent potential problems with uploading films into the viewer program once stored on your computer (particularly Plectus) is described in the manual. Therefore, it is advised to study at least the introductory part of it first. Computer freaks: Any suggestions, how to fix this bug? On demand I will be happy to share the source code of the viewer program.

As not everybody has access to the WBG you are welcome to spread the message to potentially interested colleagues. Please note that the whole package is for non-commercial use, only.

I am grateful to Philipp Schiffer, University of Cologne, for depositing the 4 files for download on the local SCIEBO server:

https://uni-koeln.sciebo.de/s/u3KmlF37XKeyUNR

On the long run the data may possibly be shifted to another server. In this case go to Philipp’s homepage (https://worm-lab.eu/4d). From there you will be redirected to the new location. If you run into problems with uploading please contact Philipp ([email protected]). Any other feedback should go to me (see top).

 

An exponentially growing number of human patients have their genomes or exomes sequenced. An outstanding challenge is to identify which of the myriad genetic changes identified contribute directly to pathological conditions and how. Basic research in C. elegans has a history of uncovering the molecular basis of essential developmental and physiological processes (Reviewed in 1).

Comprehensive sequence alignments between the human and worm genome identified 7,663 C. elegans coding-genes with human orthologs (2,3), including components of all major signaling pathways and essential cellular machineries. Precise genome editing by CRISPR-Cas9 works efficiently in the C. elegans germline when Cas9 protein is injected along with single guide RNA (sgRNA) and a short linear DNA repair template (4).

To facilitate the use of C. elegans as a model for human genetic disease we launched the WormCoolKit website (http://www.wormcoolkit.com/), which offers bioinformatic tools that assist in finding worm orthologs to human genes and vice versa, selecting CRISPR-Cas9 RNA guides and providing DNA template designs for introducing point mutations in the C. elegans genome.

The WormCoolKit orthologs finder furnishes the user with the most likely C. elegans gene ortholog to their human gene of interest. The tool extracts candidates from the OrthoList2 database and carries several additional filtration steps, aimed to increase the likelihood that these genes are true orthologs to the query.

The amino acid conservation tool is designed to inform whether a specific amino acid in a human protein is conserved in its C. elegans ortholog sequence. If conservation is confirmed, the tool will provide the user with the corresponding site in the worm protein amino acid sequence. If not, the tool will notify whether the amino acid in the worm sequence is similar or not conserved at all. To identify conservation status, the algorithm uses different methods of sequence alignments between the human gene and its ortholog. Therefore, each conserved amino acid is delivered with the number of alignments (out of ~2000) supporting the conservation. The tool also delivers an alignment score for the region surrounding the relevant amino acid, to assess whether the variation lies within a conserved region of the protein.

The WormCoolKit automated CRISPR planner is based on the CRISPR method by Paix A. et al (4,5). The planner is designed to first supply the users with RNA guides suitable for their query (based on IDT’s CRISPR-Cas9 guide RNA design checker) and then, once a guide is chosen, the algorithm defines the relevant parameters such as PAM site, CAS9 double-strand break site and mutation zone. It then goes through all possible options to mutate the strand as needed, including mutations to change the designated amino acid, mutations that prevent re-attachment of the CAS9 complex and insertion or removal of a restriction enzyme site to enable post editing identification of the gene by PCR. Lastly, the tool provides the user with a complete DNA repair template sequence containing the mutations flanked by two homology arms (35nts in length).

The WormCoolKit website was devised to streamline the process of modeling human genetic diseases in C. elegans (figure 1), starting by identifying the most likely worm ortholog of the human gene of interest, checking whether a given amino acid variation in the human gene is in a residue conserved in worms and finally designing the CRISPR strategy to insert the variation into the worm genome. That said, the free tools are generally useful for gene and residue conservation analysis between worms and humans and for providing RNA guides and DNA template designs for CRISPRing any point mutation in a worm gene.

RNA-interference (Fire et al. 1998) is a popular ‘reverse-genetics’ screening strategy. In particular ingested variant of RNAi (Timmons et al. 1999) gained popularity for genome-wide screens, where bacterially expressed dsRNA is administrated per os utilizing modified plasmids and E. coli as a feeding vector (Timmons et al. 2001). Genome-wide RNAi screens are presently carried using RNAi feeding libraries.

Two types of genome-wide RNAi ‘feeding libraries’ entailing PCR-amplified target regions are presently available: library of predicted gene-overlapping segments (Kamath et al. 2003) and amplified cDNA library (Rual et al. 2004). However, available genome-wide PCR-based recombinant RNAi libraries – resources consisting of dsRNA producing plasmids – depend heavily on gene predictions, hence the bias toward certain exon rich gene regions or certain cDNAs. Here, I report on a complementary resource facilitating an approach to RNAi screen relying on unbiased ‘forward-genetics’ strategy.

The experiments based on this approach started with the construction of the library of genomic segments incorporated into convergent T7 polymerase binding sites plasmid (PBS) vector (L4440 plasmid ), I ligated with standard, small scale liquid worm DNA prep digested with EcoRI and HindIII* and transformed into HT115(DE3) E. coli (Timmons et al. 2001). Resulting convergent T7 PBS library was estimated as ~95% recombinant. Convergent T7 library clones were fed individually with a standard protocol (as described previously, Kapulkin et al. 2005) into N2-derived worms, to confirm the apparent lethal/sterile phenotype occurs at a frequency of 1-2 per 24 clones tested.

Based on the above experiment, I think the forward RNA interference screening is useful and feasible, with strong expectation the presented screening mode will complement and extend on the existing, currently available, genome-wide RNAi resources.

*Other variants of the experiment explored elsewhere, involve the fragmented DNA ligated with other enzymes and/or linkers.

Transmission electron microscopy (TEM) is used extensively to study the nematode C. elegans. A typical specimen preparation for TEM involves chemical fixing followed by consecutive changes of reagents and embedding in plastic before thin sectioning. To handle this extremely small animal (~ 1 mm in length and 50 μm in diameter for adults), the most popular method is to immerse chemically fixed worms in warm agarose (Hall et al., 2012). Upon cooling, the solidified block is then trimmed to a maneuverable size. The goal and merit of this method is to avoid losing precious specimens during solution change and specimen transfer, as well as to facilitate desired body orientation prior to sectioning. Here we report the use of hollow fibers in lieu of agarose to achieve the same goal.

We used transparent hollow fibers from a hemodialyzer (Figure 1A) to encapsulate the worm. Each fiber has an inner diameter of 175 μm and wall thickness of 25 μm. For easy manipulation of fibers, an insect pin¹ of 100 μm in diameter is inserted into the lumen of the fiber (Figure 1B, Video 1). To encapsulate, a piece of appropriately fixed worm specimen is transferred to a ~ 3μl drop of buffer rinse and then one end of the hollow fiber is submerged into the drop. With capillary action, the specimen is drawn up into the fiber (Video 1). This loaded fiber is handled with tweezers and processed normally as for tissue samples². At the end of resin infiltration, excess length of the fiber is trimmed off and the worm can be oriented as desired in a horizontal mold (Figure 1C). During curing, the specimen will sink into the bottom of the mold due to gravity. The fiber gives sufficient leeway (at least 25 μm) at the bottom of the block for trimming (Figure 1D). Therefore, pre-filling the mold or re-embedding the specimen is not necessary (Mulcahy et al., 2018, Muller-Reichert et al., 2003). This greatly expedites the thin-sectioning process. Figure 2A shows that the hollow fiber, made of ethylene vinyl alcohol copolymer, remains intact after exposure to osmium, uranyl acetate, alcohol, acetone and resin Embed 812. Fine TEM images of the worm (Figure 2B) indicate that the hollow fiber, with its molecular weight cutoff at ~30 kDa, allows for penetration of these chemicals. We have also successfully encapsulated the worm specimens by simple capillary action while they were soaked in 1:1 mixture of resin and acetone. Therefore, encapsulation is applicable to samples that are prepared by other means, for example by high pressure freezing and freeze-substitution with organic solvents.

¹ A Minutiens insect pin is used (size 0.10, Austerilitz Insect Pins^®). Individual pins are about 12 mm in length, 0.1 mm in diameter at the shaft, and 0.0125 mm in diameter at the tip (Figure 1). For easy manipulation of the pin, one end is heat annealed to a p10 pipette tip (Video 1).

² We use ~2.5 cm length of the hollow fiber for encapsulation and place the loaded fiber into 2 ml microfuge tube for solution changes. If the worm is situated in the middle of the fiber, there is no worry of losing the specimen during processing. However, if the hollow fiber were to be trimmed shorter, both ends need to be sealed either by squeezing with forceps (Video 1) or by heating with hot platinum wire to prevent escape of the specimen.