The WBG

An online publication service of WormBook

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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Encapsulation of nematodes into hollow fibers for thin-section transmission electron microscopy

Willisa Liou1 and David Belnap1, 2
1 Electron Microscopy Core Laboratory
2 Department of Biochemistry and School of Biological Sciences University of Utah, Salt Lake City, UT, USA
Correspondence to: Willisa Liou (willisaliou@gmail.com)

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.



Figure 1: (A) Trimmed hollow fibers from a hemodialyzer (KF-201, Asahi Kasei Medical). (B) For easier manipulation of the fiber, an insect pin is inserted into the lumen of the hollow fiber, as shown in the lower one. (C) A worm specimen together with the encapsulating hollow fiber (red arrow) in a resin block. (D) A trimmed block-face showing a worm within a hollow fiber (blue arrow).


Figure 2: Cross section profiles of C. elegans within a hollow fiber. (A) A light microscopic view of a 500 nm section stained with toluidine blue. (B) A TEM view of a 50 nm section stained with uranyl acetate and lead citrate.
Download Movie 1

Movie 1: Encapsulation of nematodes into hollow fibers for thin-section transmission electron microscopy

References

Hall DH, Hartwieg E, and Nguyen KC (2012). Modern electron microscopy methods for C. elegans. Methods Cell Biol. 107, 93-149.
  PubMed

Mulcahy B, Witvliet D,Holmyard D,Mitchell J,Chisholm AD,Meirovitch Y, Samuel ADT, and Zhen M (2018). A pipeline for volume electron microscopy of the Caenorhabditis elegans nervous system.  PubMed

Müller-Reichert T, Hohenberg H, O'Toole E T and McDonald K. (2003). Cryoimmobilization and three-dimensional visualization of C. elegans ultrastructure. J. Microsc. 212(Pt 1), 71-80.  PubMed

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Single worm transfer with a pin hook

Willisa Liou1 and David Belnap1
1 Electron Microscopy Core, University of Utah, UT, USA
Correspondence to: Willisa Liou (willisaliou@gmail.com)

Breeding worms in the laboratory typically involves transferring individual animals from one plate to another. This is usually done with a platinum wire, or sometimes a hair (e.g. eyelash or eyebrow). The wire and hair are both tiny rod-like segments and are generally mounted to a pipette or a stick for easy handling. Hairs are useful only for small larvae transfer. With larger animals, the hair is either too weak to lift them up, or too stiff that the creatures suddenly straighten out and dart away upon touching. A tip-flattened platinum wire a.k.a. “the worm pick” can hold heavier animals. However, they often linger around the pick and refuse to land on the plate. Here we introduce an angle bent pointed hook made of insect pin for single worm transfer that is applicable to all developmental stages of worm from L1 to adult gravid.

The smallest-sized insect pin1 is used. These pins are sold in a package of 500 and are more affordable than platinum wires. 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. Although small, its pointed end is not sharp as to poke holes in the worms and kill them. The pin, being made of stainless steel, can be bent to a fixed angle. To make a hook, we clamp the pin at about 0.3 mm from the pointed end with tweezers, and then pull the shaft to 90° angle. This angle-bent pointed hook, like platinum wire and hair, is then annealed to a pipette tip. An easy way is to pass a p10 polypropylene pipette tip over an alcohol flame and quickly insert the blunt end of the insect pin into it. When in use, the pick is sterilized by dipping in 70% alcohol and let air-dry. Coating the pick with sticky bacteria is not necessary. Because its tip is L-shaped, the pick can be lowered vertically onto plate without poking a hole in agar. The tapered end, now runs parallel to the agar surface, is glided under the worm to shovel it up2. A sudden prodding will elicit an abrupt response; however, the worm remains on the plate locally. For younger animals a couple swipes may be needed to pick them up. For large animals, if the maneuver is slow enough, the worm will tangle with the pick (as if mistaken it for another worm) and be elevated. Once lifted, the worm dangles precariously from the hook. The worm will crawl away immediately when the new plate surface is reached.

1 Minutiens, Austerilitz Insect Pins®

2 Orient the pointed tip at 12 o’clock direction so that the stroke, from lower left to upper right (for right-handed person), is an upward moment. Doing so would avoid poking holes in agar.

Figures



Figure 1: A pin hook is attached to a pipette tip and held in a chopstick.


Figure 2: An L1 worm is being lifted up by a pin hook.
Download Movie 1

Movie 1: Pick up small worm
Download Movie 2

Movie 2: Adult gravid transfer with pin hook
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