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ven-1 encodes a novel secreted protein required for the proper positioning and attachment of the ventral nerve cord and body wall muscles in C. elegans

Go Shioi 1,2 and Shin Takagi 1
1. Division of Biological Science, Nagoya University Graduate School of Science, Nagoya, Japan
2. Present address: Laboratory for Comprehensive Bioimaging, RIKEN Center for Biosystems Dynamics Research, Kobe, Japan
Correspondence to: Go Shioi (go.shioi@riken.jp)

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.

Figures



Figure 1: Cloning of the ven-1 gene
(A) Schematic representation of the genetic and physical maps of ven-1. All the ven-1 alleles were genetically mapped between unc-112 and unc-76. ven-1 was complemented by the deficiency yDf9, but not by yDf8. Two cosmids, R02D5 and D1027 (asterisk), and a 900 bp genomic fragment derived from R02D5 rescued the ven-1 defects. The ven-1 cDNA, composed of three exons, is shown at the bottom. Box: exons; black box: coding region of the ven-1 open reading frame (ORF).
(B) Sequence of the ven-1 transcript and ORF. The ven-1 transcript is 345 nt long and contains an ORF of 258 bp. The putative signal sequence is shown in italic font. nc25 and nc26 are missense mutations, whereas nc27 is a nonsense mutation. nc28 is a point mutation in the splicing acceptor on the second intron.


Figure 2: Expression pattern and rescue experiments of the ven-1 gene
(A) Schematic illustration of the ven-1::HA and ven-1p::GFP constructs
(B) Expression pattern of ven-1. (a, b) The GFP signal in worms carrying ven-1p::GFP was expressed in ventral hypodermal cells from the comma stage. The arrowhead indicates the ventral side. (c, d, e, f) Worms expressing VEN-1::HA were doubly stained with the HA antibody (c, e, f) and phalloidin (d). Scale bar: 100 µm.
(C) Rescue experiments with ven-1 cDNA expressed in ven-1(nc25) under the control of heterologous tissue-specific promoters: epidermis (dpy-7 promoter, 94.7%, n = 227), body wall muscles (unc-54 promoter, 91.1%, n = 158), neurons (jkk-1 promoter, 40.9%, n = 227), and intestine (elt-2 promoter, 16.3%, n = 469). VEN-1::HA (79.6%, n = 141) and the 900 bp genomic fragment (93.0%, n = 127) corresponded to the rescue region in (A).
(D) Rescue experiments with ven-1 cDNA expressed in ven-1(nc25) under the control of a heat shock promoter. Eggs at 0–6 h after being laid were grown at 30°C until hatching: 0 h: 100.0%, n = 94; 1 h: 100.0%, n = 240; 2 h: 99.4%, n = 190; 3 h: 99.2%, n = 274; 4 h: 72.7%, n = 191; 5 h: 1.1%, n = 286; 6 h: 0.1%, n = 152.


Figure 3: Antibody staining of attachment components of body wall muscles in ven-1 mutants.
(Left) Antibody staining. MH4: intermediate filament; MH3: perlecan; MH46: myotactin. Immunofluorescence signals were detected on body wall muscles. (Right) Phalloidin staining. Scale bars: 100 µm.

References

Shioi, G., M. Shoji, M. Nakamura, T. Ishihara, I. Katsura, H. Fujisawa, and S. Takagi (2001). Mutations affecting nerve attachment of Caenorhabditis elegans. Genetics 157, 1611-1622.  PubMed

Moerman, D. G. and Williams, B. D. (2006). Sarcomere assembly in C. elegans muscle, WormBook, ed. The C. elegans Research Community, WormBook  PubMed

Labouesse, M. (2006). Epithelial junction and attachments, WormBook, ed. The C. elegans Research Community, WormBook  PubMed

Plenefisch, J. D., X. Zhu and E. M. Hedgecock (2000). Fragile skeletal muscle attachments in dystrophic mutants of Caenorhabditis elegans: isolation and characterization of the mua genes. Development 127, 1197-1207.  PubMed

  PubMed

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Measuring decreases in gene expression following RNA interference in C. elegans

Megan M. Senchuk1, and Jeremy M. Van Raamsdonk1,2,3,4,5
1. Laboratory of Aging and Neurodegenerative Disease, Center for Neurodegenerative Science, Van Andel Research Institute, Grand Rapids MI, USA
2. Department of Neurology and Neurosurgery, McGill University, Montreal, Quebec, Canada
3. Metabolic Disorders and Complications Program, and Brain Repair and Integrative Neuroscience Program, Research Institute of the McGill University Health Centre, Montreal, Quebec, Canada
4. Division of Experimental Medicine, McGill University, Montreal, Quebec, Canada
5. Department of Genetics, Harvard Medical School, Boston MA, USA
Correspondence to: Jeremy Van Raamsdonk (jeremy.vanraamsdonk@mcgill.ca)
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.

Figures



Figure 1: Using primers targeting outside of the RNAi-targeted sequence demonstrates that sod-2 RNAi effectively decreases sod-2 mRNA levels. Worms were grown on either empty vector (EV) or sod-2 RNAi bacteria beginning at the parental L4 stage. Gravid adults were transferred to a new plate, allowed to lay eggs for 24 hours and then removed. The resulting progeny were collected when they reached adulthood. RNA was isolated and converted to cDNA for quantification. Measuring sod-2 mRNA levels using primers that target the sequence targeted by the RNAi clone falsely shows little or no effect of sod-2 RNAi on sod-2 levels. In contrast, using primers that target outside of the RNAi-targeted sequence shows that sod-2 RNAi decrease sod-2 mRNA levels.
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An easy-to-use tool for studying and teaching C. elegans embryogenesis and evolution of nematode development

Einhard Schierenberg1
1 Department of Zoology, University of Cologne, Germany
Correspondence to: Einhard Schierenberg (e.schierenberg@uni-koeln.de)

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 (philipp.schiffer@gmail.com). Any other feedback should go to me (see top).

 

References

Schierenberg, E., and Sommer, R.J. (2013); Handbook of Zoology, vol. Nematoda, A. Schmidt-Rhaesa, ed., (Berlin: de Gruyter), pp. 61-108.

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Worm Cool Kit: An Online CRISPR Planner of Point Mutations to Facilitate Modeling of Human Genetic Variations in C. elegans Orthologs

Liran Avda1, Anat Nitzan1, Ronen Zaidel-Bar1
1 Department of Cell and Developmental Biology, Faculty of Medicine, Tel Aviv University, Tel Aviv, Israel
Correspondence to: Ronen Zaidel-Bar (zaidelbar@tauex.tau.ac.il)
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.

Figures



Figure 1:

References

A Transparent Window into Biology: A Primer on Caenorhabditis elegans. Corsi AK, Wightman B, Chalfie M. Genetics. 2015 Jun;200(2):387-407. PubMed

OrthoList: a compendium of C. elegans genes with human orthologs. Shaye DD, Greenwald I. PLoS One. 2011;6(5):e20085. PubMed

OrthoList 2: A New Comparative Genomic Analysis of Human and Caenorhabditis elegans Genes. Kim W, Underwood RS, Greenwald I, Shaye DD. Genetics. 2018 Oct;210(2):445-461. PubMed

Cas9-assisted recombineering in C. elegans: genome editing using in vivo assembly of linear DNAs. Paix A, Schmidt H, Seydoux G. Nucleic Acids Res. 2016 Sep 6;44(15):e128. PubMed

Precision genome editing using CRISPR-Cas9 and linear repair templates in C. elegans. Paix A, Folkmann A, Seydoux G. Methods. 2017 May 15;121-122:86-93. PubMed

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Forward and unbiased RNA-interference screen

Wadim J. Kapulkin1, 2, 3
1 Department of Virology and Immunopathology, National Institute of Public Health-PZH, Chocimska 24, 00-791 Warsaw, Poland
2 Department of Infectious Diseases, Microbiology and Parasitology, Faculty of Veterinary Medicine, Grochowska 272, 03-849 Warsaw, Poland
3 || Present address: Veterinary Consultancy, Conrada 01-922 Warsaw, Poland
Correspondence to: Wadim J. Kapulkin (wadim_kapulkin@yahoo.co.uk)

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.

References

Fire A, Xu S, Montgomery MK, Kostas SA, Driver SE, Mello CC. Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature. 1998 Feb 19;391(6669):806-11. PubMed

Kamath RS, Fraser AG, Dong Y, Poulin G, Durbin R, Gotta M, Kanapin A, Le Bot N, Moreno S, Sohrmann M, Welchman DP, Zipperlen P, Ahringer J. Systematic functional analysis of the Caenorhabditis elegans genome using RNAi. Nature. 2003 Jan 16;421(6920):231-7. PubMed

Kapulkin WJ, Hiester BG, Link CD. Compensatory regulation among ER chaperones in C. elegans. FEBS Lett. 2005 Jun 6;579(14):3063-8. Erratum in: FEBS Lett. 2007 Dec 22;581(30):5952. Kapulkin, Vadim [corrected to Kapulkin, Wadim J]. PubMed

Rual JF, Ceron J, Koreth J, Hao T, Nicot AS, Hirozane-Kishikawa T, Vandenhaute J, Orkin SH, Hill DE, van den Heuvel S, Vidal M. Toward improving Caenorhabditis elegans phenome mapping with an ORFeome-based RNAi library. Genome Res. 2004 Oct;14(10B):2162-8. PubMed

Timmons L, Court DL, Fire A. Ingestion of bacterially expressed dsRNAs can produce specific and potent genetic interference in Caenorhabditis elegans. Gene. 2001 Jan 24;263(1-2):103-12. PubMed

Timmons L, Fire A. Specific interference by ingested dsRNA. Nature. 1998 Oct 29;395(6705):854. PubMed

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