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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!

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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

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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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Immobilizing nematodes for live imaging using an agarose pad generated with a Vinyl Record

Katherine Andrea Rivera-Gomez and Mara Schvarzstein
CUNY Brooklyn College, NY, USA
Correspondence to: Mara Schvarzstein (maraschvarzstein@gmail.com)
In the absence of microfluidic rigs, most live imaging protocols utilize flat agarose pads along with anesthetics and/or microbeads. It is often difficult to manipulate the worms position in this set up and thus a higher number of worms need to be mounted to ensure a few showing an optimal position for imaging. This is problematic when trying to image nematodes with genotypes that are scarce within a strain, such as m+z- sterile or embryonic/larval lethal, because finding enough animals to mount can be challenging. In order to image worms of genotypes that are hard to find, we slightly modified a previously published protocol (Zhang, M. et al., 2008) to make agarose pads that allow us to maintain L4 or adult hermaphrodites and adult males straight during imaging for at least 2 hours. These pads also appear to reduce rolling of the worms during imaging.

Materials:
A 12-inch long-playing (LP) vinyl record, labeling tape, Pasteur pipette with bulb, agar (at the concentration in which you prefer to use, we use 4% agar), microscope slides, your preferred anesthetic (we use either levamisole or serotonin), a minutien pin (eyelash pick), coverslips, a petroleum jelly (we use Vaseline) filled syringe with a flat end needle.

Procedure:
1. Place strips of tape 1.5-2 inches apart perpendicular to the grooves on the vinyl record’s grooved surface, as shown in Figure 1.A.
2. Pipette a drop of melted agarose onto the vinyl record between the strips of tape, and immediately add a microscope slide on top of the drop to make a pad against the vinyl record.
3. After cooling, remove microscope slide from the vinyl record by lifting from one side as shown in Figure 1.B. If the pad is too large, it can be trimmed down by using another microscope slide’s flat edge to make straight cuts.
4. Add a volume of 2-5l of anesthetic at a time, depending on the size of your pad. Then pick worms onto anesthetic droplet (the grooves are an optimal width for hermaphrodites from L4-adult).
5. Position worms into the grooves in the agarose pad utilizing the minutien pin (Figure 1.C). Worms are easier to position when most of the anesthetic has been absorbed into the pad, but don’t allow it to dry completely, as this will form air bubbles in the adjacent grooves.
6. Make a circle line around the pad with the syringe filled with Vaseline before placing the coverslip as shown in Figure 1D. Slightly press the coverslip over the pad and vaseline to keep the worms in place and make a seal around the pad to prevent evaporation of the anesthetics mix.

This protocol can be used in conjunction with microbeads, but in our hands they do not seem to improve the immobilization of the worms.

The vinyl record grooved pads function, in part, as a poor-person’s microfluidics for live imaging. It allows the placing of worms into single channels organized as in a microfluidic chip, thus reducing the time searching for the worms for imaging on the slide. Addition of drugs or food (for prolonged imaging) could potentially be done with a fine needle inserted through the Vaseline ring. An additional advantage of these pads is that there is no difference in fluorescence brightness loss compared to the conventional agarose pad.

This protocol’s affordability, the ease of use and training has simplified our live imaging of m+z- males expressing low fluorescence fusion proteins in the germline significantly.

We thank the Mikaela Murph and Erlyana Clarke for trying the protocol and providing us with feedback.

Figures



Figure 1: Figure 1: Overview of the process for making agar pads using a vinyl record surface. A. Laboratory label tape placed on vinyl disc gives reference where to place the agar drop and also establishes an even pad thickness. B. A drop of hot agar is covered with the microscope slide. Once agar has cooled down slide is lifted with the attached agar pad. C. Anesthetized C. elegans nematode aligned on a groove made with vinyl record. D. Vaseline perimeter is applied around the agar pad and slide cover is pressed slightly to seal in the humidity to prevent the nematodes from drying over long periods of imaging.
 

References

Zhang, M., Chung, S. H., Fang-Yen, C., Craig, C., Kerr, R. A., Suzuki, H., … Schafer, W. R. (2008). A self-regulating feed-forward circuit controlling C. elegans egg-laying behavior. Current Biology : CB, 18(19), 1445–1455.   PubMed

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An Improved Method for Freezing Nematodes Using a Trehalose-DMSO Cryoprotectant

Kevin F. O’Connell
Laboratory of Biochemistry and Genetics, National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health-USA, Bethesda MD
Correspondence to: Kevin F. O’Connell (kevino@mail.nih.gov)

One of the many advantage of working with C. elegans is the ability to make frozen stocks from which viable animals can be recovered as needed. While we often brag about this to our fly colleagues, we all know that the truth is that worms really don’t freeze all that well. When thawing an aliquot of frozen stock, we are happy if we can recover more than just a few viable larvae amongst the mass of dead worms.  Hoping to develop a better method for freezing worms, I turned to the literature on freezing plant cells which are difficult to freeze as cryoprotectants are often unable to penetrate the thick cell walls.   Since the cuticle of nematodes might also block the penetration of some cryoprotectants, I decided to try a cryoprotectant that works on plant cells.The following is a protocol that I have modified for use in worms and have employed for the past 5 years.  I have found that it is consistently far superior to the traditional glycerol-based method and allows the recovery of a sizable recovery of the frozen worms including all larval stages and adults.

 

Freezing Worms with Trehalose-DMSO

  1. Grow worms on 1-3 100 mm plates until freshly starved. Generally the more worms the better.
  1. Wash worms off plates in M9 buffer and collect by centrifugation (5000 rpms for 3 minutes is sufficient.
  1. Wash once in 4 ml room temperature Trehalose-DMSO freezing buffer.
  1. Re-suspend worms in 4 ml room temperature Trehalose-DMSO freezing buffer. Dispense 1ml per tube in 4 cryogenic vials.
  1. Incubate at room temperature for 30 minutes.  This sometimes increases the rate of survival.
  1. Put vials into Nalgene Cryo 1 C Freezing Container (Cat. No. 5100-0001). Put container in -80°C freezer.
  1. 1 week later, test thaw one of the vials. Many animals should survive.

 

Trehalose-DMSO freezing buffer (Good for six months when stored at room temperature):

Add 15.1 g Trehalose (Sigma T0167) and 17.7 ml DMSO (14.13M) to enough M9 buffer so total volume equals 500ml.  Filter-sterilize and store at room temperature.

Final concentration:  0.08M Trehalose, 0.5M DMSO.

 

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