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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

“N2 male” is a long-lived fln-2 mutant

Yuan Zhao1, Hongyuan Wang1, Richard J. Poole2, David Gems1*
1 Institute of Healthy Ageing, and 2 Department of Cell and Developmental Biology, University College London, London, UK
Correspondence to: David Gems (david.gems@ucl.ac.uk)

To distinguish wild-type and mutant genes, one needs to define a wild-type strain. To this end, we worm breeders use the convention that N2 is the wild type. Now, when people order N2 from the Caenorhabditis Genetics Center, they usually request a hermaphrodite stock, but there is also a male stock available (strain: “N2 male”); for convenience, we will refer to these two lines as N2H and N2M. A problem here is that these lines are not genetically identical: N2M hermaphrodites are longer lived (+11% median lifespan) (Gems and Riddle, 2000). So which is wild type, N2H or N2M? This is a problem, and we have been looking into it.

Under standard culture conditions N2 hermaphrodites exhibit two forms of death, with either a swollen or an atrophied pharynx (P and p death, respectively). P death occurs earlier and results from bacterial infection, facilitated by mechanical senescence of the pharynx due to the high, wild-type pumping rate (Zhao et al., 2017). We have found that the greater lifespan of N2M hermaphrodites is due to lower P death frequency (and therefore reduced early mortality).

This turns out to be due to a single recessive mutation in the X-linked gene fln-2 (filamin) in N2M, as revealed initially by Variant Discovery Mapping. fln-2(ot611) is a nonsense allele resulting from a C to A transversion, creating a stop codon at Y800 in the FLN-2A isoform. Thus, N2H is wild type and N2M is mutant. This conclusion is confirmed by examination of the fln-2 sequence in multiple C. elegans wild isolates (Cook et al., 2017). It would therefore be advisable to discontinue the use of N2M (“N2 male”).

Although people mainly use N2H as their wild type, N2M is sometimes used for strain construction and backcrossing. To get an idea of the prevalence of fln-2(ot611) we checked a sample of strains in our collection and found fln-2(ot611) in 23/50, particularly in strains generated by the C. elegans Gene Knockout Project and the C. elegans Expression Project.

Variation at the fln-2 locus is a potential confounding variable, especially in studies of lifespan genetics. We have so far found it to confound the effects on lifespan of alteration of eat-2, sir-2.1 and daf-12. For this reason we advise the use of routine checks of fln-2 genotype in studies of the genetics of lifespan. Here is information about how to do this.

The relevant fln-2 sequence information is:
Wild-type fln-2: GGCGCTGGTCAATA[C]AAAATCCACGTTCTT
fln-2(ot611): GGCGCTGGTCAATA[A]AAAATCCACGTTCTT with a C to A change in the bracket.

To genotype fln-2, we used the following primers to PCR amplify the region containing the mutation: forward 5’-GGTGTTCGATTCTGGTCTGG; reverse 5’-ACATCGACGAGAAGACAACAC. The PCR product can be sequenced using the primer 5’-TGTACCCAGAAATTGACAAGATAC.

Allele-specific PCR can also be used to discriminate between the alleles, using the following primers: wild-type-specific forward 5′-taccattccgagcttattgattgttacctGGACGGCGCTGGTCCATAC-3′; ot611-specific forward 5′-GGACGGCGCTGGTCTATAA-3′; reverse 5′-ATCGCATGAACCATAAATGATG-3′. Each forward primer contains an additional mismatch to make it different from both the template and the other forward primer to ensure allele specificity (Neagu and Maier, 2011). The wild-type PCR product is 30bp longer than the mutant product, and can be readily distinguished on a 2% agarose gel.

References

Cook D, Zdraljevic S, Roberts J, and Andersen E. (2017). CeNDR, the Caenorhabditis elegans natural diversity resource. Nucleic Acids Research. 45, D650-D657.  PubMed

Gems D and Riddle DL. (2000). Defining wild-type life span in Caenorhabditis elegans. J. Gerontol. A Biol. Sci. Med. Sci. 55, B215-B219.  PubMed

Neagu A and Maier W. (2011). snPCR for reliable one-step genotyping of single nucleotide differences. The Worm Breeder’s Gazette 19, 1.

Zhao Y, Gilliat AF, Ziehm M, Turmaine M, Wang H, Ezcurra M, Yang C, Phillips G, McBay D, Zhang WB, Partridge L, Pincus Z, and Gems D. (2017). Two forms of death in ageing Caenorhabditis elegans. Nature Comm. 8, 15458.  PubMed

Review of 3rd Parasitic Nematodes: Bridging the Divide workshop

Mostafa Zamanian 1, William Sullivan 2, Jordan D. Ward 2*
1 Department of Pathobiological Sciences School of Veterinary Medicine,University of Wisconsin, Madison, USA
2 Department of Molecular, Cell, and Developmental Biology, University of California, Santa Cruz, USA
Correspondence to: Jordan D. Ward (jward2@ucsc.edu)
On Wednesday June 21st, 2017 before the start of the International C. elegans Meeting, we held a workshop designed to bring parasitologists and C. elegans researchers together. The workshop was made possible by a grant from the National Science Foundation Division of Integrative Organismal Systems. The goal was to cover the broad theme of C. elegans as a model for parasitic nematode biology. We had six invited speakers covering a range of nematodes and approaches and also featured four lightning talks from submitted abstracts.

Soil-dwelling helminths such as hookworms, whipworms, and Ascaris infect hundreds of millions of people globally, and also infect livestock and crops. Dick Davis (University of Colorado School of Medicine) gave an overview of Ascaris, highlighting resources, tools, and key questions in this system. Two fascinating features of the system he discussed were somatic DNA elimination, in which 15% of the genome is removed in somatic cells during embryogenesis, and early zygotic genome transcription commencing remarkably early (immediately following fertilization). Elissa Hallem (UCLA) discussed parasitic nematode sensory behaviors with respect to host seeking. Using both Strongyloides nematodes and C. elegans, her group is exploring the neurobiology of how parasites find their hosts. Tiffany Baiocchi, a PhD student with Adler Dillman (University of California, Riverside) presented her work on novel odorants produced from nematode­-parasitized insect cadavers and the behavioral responses this odorant triggers in both free-­living and parasitic nematodes. Jonathan Ewbank (Centre d’Immunologie de Marseille- Luminy) gave a lightning talk on assembling the Nippostrongyloides brasieliensis genome and dealing with complex repeats; this nematode is a parasite of rodents and a useful model for human hookworm infection.

Two experts discussed filarial parasites, a group of nematodes that cause lymphatic filariasis and onchocerciasis. Several of these species carry obligate endosymbiotic bacteria (Wolbachia), which is an attractive therapeutic target. Malina Bakowski (California Institute for Biomedical Research) discussed her high-throughput screens to identify potent anti-Wolbachia molecules. Sara Lustigman (New York Blood Center) presented on the role of “omics” and molecular tools in Brugia species efforts to eliminate nematodes that cause the human disease filariasis. Conor Caffrey (University of California, San Diego) also presented a high-content screening system for flatworms which cause schistosomiasis.

Adrian Wolstenholme (University of Georgia) discussed the problem of resistance to anthelminthics and the mechanisms of resistance. His group uses a combination of heterologous studies in C. elegans and work in parasites such as Brugia malayi and Haemonchus contortus to explore potential mechanisms of resistance to drugs such as ivermectin and diethylcarbamizine. Nidhi Sharma, a Ph.D student with John Gilleard (University of Calgary) presented a lightning talk on the role of UDP-glucuronosyltransferase (UGT) enzymes in benzimidazole drug biotransformation by nematodes. Keeping with the theme of glycosylation,  Patricia Berninsone (University of Nevada, Reno) gave a lightning talk on her identification of nematode phosphorylcholine-modified N-glycoproteins using C. elegans.

This was the third time that the workshop was held and it brought together 77 participants to explore a range of approaches and systems to explore parasitic nematode biology and to define areas where C. elegans researchers can make significant impact. Going forward, we aim to integrate the workshop into regular program of the worm meeting as having it beforehand limits the ability of some researchers attend. Look for us at the 2019 Worm Meeting!

Troubleshooting the positive butanone associative memory assay

Charline Borghgraef 1, Rachel Arey 2, Liliane Schoofs 1, Liesbet Temmerman 1* and Coleen Murphy 2*
1 Animal Physiology and Neurobiology, University of Leuven (KU Leuven), Leuven, Belgium
2 LSI Genomics & Dept. of Molecular Biology, Princeton University, Princeton NJ, USA
Correspondence to: Coleen Murphy (ctmurphy@princeton.edu) – Liesbet Temmerman (liesbet.temmerman@kuleuven.be)

Learning assays for C. elegans are known to be challenging due to the fragility of the phenotype, hence, the ease by which it can be disturbed. For several months, our teams at KU Leuven struggled to implement the Murphy lab’s short-term associative memory assay, failing to obtain proper control data despite having implemented other learning assays without noteworthy incidents. We are aware that others in the field have struggled with this assay as well.

To address this issue, the Murphy lab supported us in troubleshooting their protocol both in Belgium as well as during a stay at their lab in the USA. We discovered several small and seemingly trivial differences which showed to be of major importance for the success of this assay. These observations allowed us to successfully perform the assay in Belgium, where it had not worked previously.

In order to enable others to smoothly implement this assay in their labs, the updated protocol is described on the Murphy and Schoofs labs’ webpages.

Weblink Murphy lab: http://www.molbio1.princeton.edu/labs/murphy/protocols.html

Weblink Schoofs lab: https://bio.kuleuven.be/df/ls/research/positive-butanone-associative-memory-assay

Preparation of Nuclei Using the Balch Homogenizer

Oxana Radetskaya1 and Shane L. Rea1

1 The Barshop Institute for Longevity and Aging Studies, University of Texas Health Science Center at San Antonio (UTHSCSA), San Antonio, USA

Correspondence to: Shane L. Rea (reas3@uthscsa.edu)

We have previously described use of a Balch homogenizer for the rapid, cost-effective and freeze-thaw independent homogenization of Caenorhabditis elegans (Bhaskaran, S., et al., 2011). This tool operates through shear forces generated by placing a ball bearing of known diameter within a confined chamber and then using two disposable syringes to manually advance worm samples past the micron sized gap that is formed between the chamber wall and the bearing. Using this tool we showed that worms of any developmental stage could be equally well homogenized after selecting an appropriately sized bearing to precisely control the wall-to-bearing clearance. Here we describe the use of the Balch Homogenizer for isolation of nuclei from C. elegans.

Since the primary goal of this study was to identify conditions that permitted rapid, freeze-thaw independent isolation of structurally-sound nuclei from any stage of worm development, our choice of buffers as well as centrifuge spin speeds to pellet nuclei were the same as those described in the embryonic and oocyte nuclei purification protocol of Mains and McGhee (1999). In their method, several approaches to disrupt worm samples were described, including use of a vintage motorized Stansted Cell Disruptor, a French Press, sonication and a mortar & pestle. Each method had its associated limitations, ranging from cost to sample heating. We established the following protocol for nuclei extraction which is rapid, does not require prior sample freezing, and can be applied equally to all stages of nematode development. Samples of nuclei collected using this protocol have been utilized in a number of downstream applications, including preparation of nuclear extracts for use in gel shift assays.

 

Buffers

– 2x Nuclear Preparation Buffer (2xNPB): 20 mM HEPES (pH 7.6), 20 mM KCl, 3 mM MgCl2, 2 mM EGTA, 0.5 M sucrose, 1 M dithiothreitol, 1 mM PMSF, 20 μM E-64, (pre-chill to 4°C)

– Nuclear Extraction Buffer (NEB): 20 mM HEPES (pH7.6), 350 mM NaCl, 2 mM EDTA, 25% glycerol, 0.5 mM dithiothreitol, 0.5 mM PMSF, 10 μM E-64, (pre-chill to 4°C)

– Phosphate Buffered Saline (PBS), pH 7.4 (keep at room temperature)

 

Equipment

– Balch Homogenizer (Isobiotec, Heidelberg, Germany), including 12 & 18 μm ball bearings. Refer to (Bhaskaran, S., et al., 2011) for full specification details of the homogenizer. (Pre-chill to 4°C)

– Bench top centrifuges

 

Worm Strain

– SD1084 [gaIs148 [Pges-1::FLAG::PAB-1 + sur-5::GFP] (Pauli, F., et al., 2006)

 

For the following protocol we utilized a mixed-stage population of SD1084 worms containing a sur-5::GFP reporter gene. Use of this strain allowed us to monitor the degree of disruption of the nuclear membrane at each step of the protocol, as SUR-5::GFP is soluble. We have previously published ball bearing clearances required to either fracture or homogenize synchronized populations of worms at any stage of larval development [refer to Table 1 in (Bhaskaran, S., et al., 2011)]. The following protocol can therefore be easily modified to accommodate worms of any size. Nuclei in C. elegans range from ~1.8 – 7μm (Chen, L. et al., 2013).

 

Protocol

  1. Wash mixed-stage, SD1084 worms off 5 x 10cm NGM plates using 5 ml PBS/plate. Pool, then pellet worms (800 x g, 3 minutes, RT), wash and re-pellet thrice more (3 x 15 mls PBS, RT). Transfer worms to ice (4°C). (Note: At this point we have also snap frozen worms in liquid nitrogen for later nuclei extraction. Freezing risks fracturing the nuclear membrane.)
  2. Add an equal volume of ice-cold 2xNPB to the worm pellet.
  3. Homogenize worms using a pre-chilled Balch Homogenizer (30 strokes /18 μm ball bearing, followed by 25 strokes / 12 μm ball bearing.)
  4. Transfer homogenate into a 15 ml falcon tube. Rinse the Balch Homogenizer with 2 ml of 1xNPB and combine with the first homogenate. Pellet nuclei (4000 x g, 5 minutes, 4°C). (Note: The supernatant corresponds to a crude cytoplasmic extract. To clarify, spin for 30 minutes at 35,000 x g at 4° Take the resulting supernatant, avoid the white flocculant, add glycerol to 25%, then snap freeze.)
  5. Resuspend nuclei in 5 ml 1xNPB containing 0.25% NP-40 + 0.1% Triton-X-100. To remove any large debris, centrifuge sample at 100 x g, 5 minutes, 4° Transfer supernatant containing nuclei to a fresh tube. (Note: NP-40 and Triton-X-100 are harsh detergents that will ultimately dissolve the nuclear membrane. If used at higher concentrations, or for longer periods, they will also disrupt protein structure. Use Tween-20, Saponin or Digitonin (0.2% – 0.5%, up to 30 min.) for milder membrane disruption.)
  6. To recover nuclei trapped within large debris, wash the low speed pellet twice with 5 ml 1xNPB containing 0.25% NP-40 + 0.1% Triton-X-100.
  7. Combine all supernatants and centrifuge at 4000 x g, 5 minutes, 4° Resuspend the pellet in 5 ml 1xNPB sans detergents. (Note: This sample represents the nuclei-enriched fraction.)
  8. To prepare a nuclear extract, re-pellet the nuclei (4000 x g, 5 minutes, 4°C).
  9. Resuspend the pellet in 2-4 volumes of NEB. Transfer to a microfuge tube, extract by gentle rotation 45 minutes at 4°
  10. Centrifuge at 10,000 x g for 14 minutes at 4°C to pellet debris.
  11. Carefully remove the supernatant and store in aliquots at -80°C (Note: This sample represents a crude nuclear extract positive for GFP (Fig. 1A). Surprisingly it also contains buoyant structures that stain positive with propidium iodide (Fig. 1B). These structures are chromatin skeletons (Fig. 1C) and they presumably avoid precipitation, in part because of the high glycerol content of the extraction buffer.)

Figures

figure_1

Figure 1: Characterization of extraction fractions using immunofluorescence and western analysis. (A) α-GFP western blot of various nuclei extraction fractions obtained using SD1084 worms. (Monoclonal GFP antibody obtained from Antibodies Inc., cat# 73-131). The precise identity of the expressed SUR-5::NLS::GFP fusion protein is unknown, but its full length predicted size is 104 kDa. (Yochem, J., et al., 1998). (B) Fluorescent images of mounted extract from Step 11 showing propidium iodide (PI) positive structures and SUR-5::GFP staining. (C) Higher magnification image of PI-positive structures shown in (B). Very few structures are both PI-positive and sur-5::GFP positive (compare left and right arrows), presumably due to detergent extraction of sur-5::GFP, but their presence nonetheless underscores their nuclear origin.

References

Bhaskaran S, Butler JA, Becerra S, Fassio V, Girotti M, Rea SL (2011). Breaking Caenorhabditis elegans the easy way using the Balch homogenizer: an old tool for a new application. Analytical Biochemistry. 413, 123-132  PubMed

Chen L, Chan LL, Zhao Z, Yan H (2013). A novel cell nuclei segmentation method for 3D C. elegans embryonic time-lapse images. BMC Bioinformatics. 14, 328.  PubMed

Mains PE, McGhee JD (1999). Biochemistry of C. elegans. In C. elegans : a practical approach. (IA Hope, ed). Oxford: Oxford University Press.  PubMed

Pauli F, Liu Y, Kim YA, Chen PJ, Kim SK (2006). Chromosomal clustering and GATA transcriptional regulation of intestine-expressed genes in C. elegans. Development. 133, 287-295  PubMed

Yochem J, Gu T, Han M (1998). A new marker for mosaic analysis in Caenorhabditis elegans indicates a fusion between hyp6 and hyp7, two major components of the hypodermis. Genetics. 149, 1323-1334.  PubMed

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