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Parasitic Nematodes: Bridging the Divide workshop: Call for Lightning Talk Abstracts

Mostafa Zamanian1, William Sullivan2, Jordan D. Ward2
1 Department of Pathobiological Sciences School of Veterinary Medicine,University of Wisconsin, Madison, USA
1 Department of Molecular, Cell, and Developmental Biology, University of California, Santa Cruz, USA
Correspondence to: Jordan D. Ward (jward2@ucsc.edu)
Workshop Date: Wednesday, June 21, 2017
Workshop Time: 2:30pm- 5:30pm
Workshop Location: Sunset Village Grand Horizon Ballroom
Lightning Talk Abstract Submission Deadline: May 12, 2017 (5pm PST)
Notification of Acceptance: May 19, 2017
We are pleased to invite student and postdoc abstract submissions for 5 minute lightning talks to be presented at the NSF-sponsored, 3rd Parasitic Nematodes: Bridging the Divide workshop. We encourage submissions that address the broad theme of C. elegans as a model for parasitic nematode biology, including work relating to human, animal, and plant parasitic nematodes and from both a basic biology standpoint or a translational perspective.
The platform provided by this workshop represents a unique opportunity for early-career scientists to interact with other researchers at the intersection of C. elegans biology and parasitology. These five minute talks will be presented on the first day of the 21st International C. elegans Meeting. The presenting authors must be available during any of the limited number of possible time slots available.
Abstract submissions are due on May 12th at 5pm PST and presenting authors will be notified of decisions by May 19th. Instructions for logistics and formatting of lightning talk presentations will follow acceptance. Abstract Format: Abstracts should be limited to 2,500 characters in total, including the title, authors, author affiliations, main body, and spaces. Abstracts should be sent to the workshop e-mail address (parasiteworkshop@gmail.com) as a single PDF file with subject heading “Lightning Talk Abstract”. A notification of receipt will follow submission. Accepted abstracts will appear online in a Worm Breeder’s Gazette article and titles will be listed on the International Worm Meeting website.

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References

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Experiences with GRASP-based transsynaptic labeling

Meital Oren-Suissa1,2, Emily A. Bayer1, Michael Hart1, Maryam Mayeed1, Abhishek Bhattacharya1, Oliver Hobert1
1 Department of Biological Sciences, HHMI, Columbia University, New York, USA
2 present address Department of Neurobiology, Weizmann Institute of Science, Rehovot, Israel
Correspondence to: Oliver Hobert (or38@columbia.edu)
Reliable and sensitive tools for labeling specific connections between individual neurons are crucial for our understanding of the functionality of the nervous system. The gold standard for visualizing synaptic connections is electron microscopy (EM), wherein the degree of connectivity (number of sections over which en passant synapses are observed) and ultrastructure can be resolved. However, given the substantial time and resource investment, EM reconstruction is impractical for routine analysis of synaptic connections under distinct conditions (e.g. different time points, developmental stages, genetic backgrounds) and even more so for dynamic studies of neuronal connections. The genetically controlled GRASP system (“GFP Reconstitution Across Synaptic Partners”) has been used to visualize synaptic connections in live, transgenic animals (Feinberg et al., 2008). GRASP is based on two non-fluorescent split-GFP fragments, spGFP1-10 and spGFP11, tethered to synaptic membranes in each of two specific neurons or cell-types. When two neurons, each expressing one of the fragments, are tightly apposed across a synaptic cleft, fluorescent GFP is reconstituted. Actual synapses are labeled by targeting the Split GFP molecules to synaptic membranes using the neuroligin protein. C. elegans Neuroligin localizes both pre- and postsynaptically in neurons.

We have recently used GRASP to label a substantial number of synapses in C. elegans (Oren-Suissa et al., 2016)(E.A.B., M.H., M.M., A.B. and O.H., unpubl. data) and want to share a few lessons that we learned:

1) To best visualize the area of contact between the two cells of choice, the neuronal processes should be labeled with a fluorescent cytoplasmic marker. Using both cellular markers in the same color is an option as long as the area of contact is clearly visible. Optimally, promoters should differ from those driving the split GFP fragments. The cell-specificity is less of a problem and promoters for cytoplasmic labeling could be expressed in additional cells.

2) We perform germ line transformation by microinjection. In cases where the GRASP signal is expected in ventral or dorsal areas of the body (for example, the PAG), we prefer to use the pRF4 (rol-6(su1006)) plasmid as the co-injection marker. It allows for easy visualization of GRASP puncta, as the roller worm is slightly rotated when mounted on a slide for imaging purposes. We co-inject 5 plasmids, two split GFP plasmids, two cytoplasmic markers and a co-injection marker. Alternatively, the GRASP construct could be injected into an already integrated cytoplasmic marker strain, so the marker and GRASP do not need to be on the same array.

3) Initial split GFP concentrations should be minimal and aimed at 10 ng/mcL. If same promoters are used for the cytoplasmic markers, the latter should be injected in extremely low concentrations, to avoid competition for transcription resources. All the heritable F2 worms are maintained for initial live-imaging analysis. As many transgenic lines as possible should be screened to uncover ones with synaptic GFP puncta localization. Imaging should be carried out as described below. Positive transgenic lines are those with discrete GFP puncta between the two cells. If expression is too high, synapses are saturated with GFP and the split-GFP interaction is irreversible. Transgenic animals generated by microinjection carry large extrachromosomal arrays incorporating a high copy number of transgenes, which might cause overexpression and germ line silencing. In addition, the inherit­ance of extrachromosomal arrays is unstable and can create genetic mosaics. We integrate the extrachromosomal arrays into C. elegans chromosomes for stable inheritance, using γ-irradiation. In some cases integrated GRASP strains underwent silencing. Growing these strains at 25C for a couple generations disilenced these arrays.

4) Transgenics lines with good signals are often hard to find. Concentration of split-GFP plasmids should be gradually increased, either of both plasmids or of one of them. A common solution is to swap the split GFP fragment between the pre- and the post-synaptic cells. Alternatively, different neuronal promoters should be tried.

5) GRASP imaging conditions: Compared to our Zeiss Imager Z1 standard compound epifluorescence microscope, we have observed much better signals on a pinhole based confocal microscope (LSM880). Epi-fluorescent microscope images contain too much out-of-focus signals. Puncta were quantified by scanning the original full Z-stack for distinct dots in the area where the processes of the two neurons overlap.

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Flowchart_ver2

Figure 1: Figure 1. Pipeline for generating transgenic GRASP lines

References

Feinberg, E.H., Vanhoven, M.K., Bendesky, A., Wang, G., Fetter, R.D., Shen, K., Bargmann, C.I., 2008. GFP Reconstitution Across Synaptic Partners (GRASP) defines cell contacts and synapses in living nervous systems. Neuron 57, 353-363.  PubMed

Oren-Suissa, M., Bayer, E.A., Hobert, O., 2016. Sex-specific pruning of neuronal synapses in Caenorhabditis elegans. Nature 533, 206-211.  PubMed

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An updated fosmid recombineering toolkit

Baris Tursun1,2, Luisa Cochella1,3, Stefanie Seelk2, Ev Yemini 1, Ines Carrera1,4, Nikos Stefanakis1,5, Kelly Howell1, Dylan Rahe1, Sze Yen Kerk1, Oliver Hobert1
1 Department of Biological Sciences, Department of Biochemistry and Molecular Biophysics, Columbia University, New York, USA
2 Max Delbrück Centrum, BIMSB, Berlin, Germany
3 Research Institute of Molecular Pathology (IMP), Vienna Biocenter (VBC), Vienna, Austria.
4 Worm Biology Laboratory, Institut Pasteur de Montevideo, Montevideo, Uruguay.
5 Rockefeller University, New York, USA
Correspondence to: Oliver Hobert (or38@columbia.edu)

A readily available fosmid library covers ~90% of all worm genes. We have previously described a pipeline for manipulating the sequence of fosmid clones via a recombineering strategy in E.coli (Tursun et al., 2009, PLoS One. 2009;4(3):e4625). Recombineered fosmids permit expression of fluorescent reporters or genetically altered protein products, in transgenic worms, under the control of most, if not all, endogenous regulatory sequences of the gene of interest. We report here more than a dozen new cassettes for fosmid recombineering using our pipeline. These cassettes allow for a number of different types of tagging for biochemical and microscopy purposes among others, all listed in Table 1.

In light of the advent of CRISPR/Cas9-mediated genome engineering, it is important to point out the persistent value of fosmid-based transgenesis: (a) Single copy fluorescent proteins generated by CRISPR/Cas9 or miniMos can be difficult to visualize, especially if the gene has low expression. The multicopy nature of transgenic arrays can increase the intensity of the reporter gene signal to a detectable level. (b) Some genes remain difficult to target by CRISPR, either for technical or biological reasons (e.g. adding a tag generates a hypomorphic allele that might not provide enough gene function in single copy). In our hands, fosmid recombineering in bacteria is fast and always successful. Finally, (c) fosmid-based reporter transgenes provide a fast and straightforward tool for both rescue of mutant phenotypes and subsequent mosaic analysis.

The complete toolkit can be requested from the Hobert lab (or38@columbia.edu).

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Figure 1:

References

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Setting up simple and cost-effective gene editing for nematodes

Yongquan Shen1 and Ronald E Ellis1
1 Department of Molecular Biology, Rowan University-SOM, Stratford, NJ 08084
Correspondence to: Ronald E Ellis PhD (ellisre@rowan.edu)
Genome editing with CRISPR/Cas9 or TALENs has become an essential tool for working with nematodes [3,4,5]. Plasmids expressing Cas9 and a custom sgRNA can be used for gene editing in many situations, particularly for C. elegans [2]. However, this approach is less effective for C.briggase and some male-female species. Furthermore, some edits will be difficult if a good Cas9 target is not located nearby.Our approach is to use Cas9 protein complexes for injections [4] when Cas9 target sites are available, and TALENs for other situations [5]. The best targets for Cas9 end in a 3’-GGnGG sequence [3], which is infrequent in worms. Although satisfactory sites can usually be found for gene knockouts, they might not be available for site-specific editing.

If possible, we use the native CRISPR/Cas9 system, which relies on Cas9 protein with both a universal RNA (tracrRNA) and a specific guide RNA (crRNA). These RNAs are bought from Dharmacon, and the same tracrRNA can be used for every injection. Although commercial Cas9 is also available, it is expensive. Thus, Paix and Seydoux developed a protocol for Cas9 production in E. coli [4]. We modified their procedure to simplify the Cas9 purification. Briefly, the plasmid NM2973 was transformed into NEB Nico-21 competent cells (#C2529H), and Cas9 expression was induced by adding IPTG to the culture media. We purified Cas9 under native conditions, using a Qiagen Fast-Start Kit (#30600). Following instructions, the bacterial pellet was resuspended in 10 ml of lysis buffer, and cooled on ice for 30 minutes. The cell lysate supernatant flowed through the Fast-Start Ni-NTA column by gravity, and 4 ml of washing buffer was used to wash the column twice. Next, Cas9 was eluted with 2 ml of elution buffer. Finally, the Cas9 eluate was centrifuged with a centricon filter (#UFC905024) to concentrate the protein. We obtained 2.5 mg of Cas9 from 1L of culture, and made 2 µl aliquots at a concentration of 10 µg/µl for storage at -80°C.

Home-made Cas9 was tested to measure it efficiency. First, we performed a cleavage assay to confirm that it had endonuclease activity. A mixture of Cas9, mouse SLC guide RNA and a PCR fragment from the SLC gene were digested at 37°C for 1 hour. The result showed that our Cas9 completely cleaved the SLC fragment as shown in Figure 1. Second, we used the Cas9 to knockout the cbr-dpy-5 gene in vivo. The synthetic cbr-dpy-5 crRNA and tracrRNA were incubated with Cas9 at 37°C for 15 minutes, and microinjected into the gonads of C. briggase hermaphrodites. Among all the F1 progeny, 25% produced homozygous Dpy children. A similar injection using commercial Cas9 (PNA Bio.) gave 18% affected F1s. Sequencing confirmed that the cbr-dpy-5 mutation was a 51 bp deletion near the PAM site of its target. Thus, home-made Cas9 has high efficiency and allows rapid and inexpensive gene editing.

If the desired target lacks a nearby 3’-GGnGG, TALENs can be used for precise gene editing. Since the only constraint for TALEN binding is that the first nucleotide in each target sequence is a T, sites exist almost everywhere in the genome. The plasmids to synthesize a pair of TALEN mRNAs can easily be built in a week, using a procedure developed by Cermak et al [1], with the destination vector pRE189 for expressing nematode mRNAs [5]. A Golden Gate TALEN kit is available from Addgene (#1000000024) and mRNA kits from Ambion (#AM1344, #AM1908).

Acknowledgments: We thank Dr. Geraldine Seydoux for the NM2973 plasmid. This work was supported by NIH grant to Dr. Ronald E Ellis.

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Figure 1: Cas9 Cleavage assay in vitro

References

[1] Cermak T, Doyle EL, Christian M, Wang L, Zhang Y, Schmidt C, Baller JA, Somia NV, Bogdanove AJ, and Voytas DF. (2011) Efficient design and assembly of custom TALEN and other TAL effector-based constructs for DNA targeting. Nucleic Acids Res. 39(12):1-11  PubMed

[2] Dickinson DJ, And Goldstein B. (2016) CRISPR-based methods for Caenorhabditis elegans genome engineering. Genetics. 202(3):885-901  PubMed

[3] Farbound B, and MeyerBJ. (2015) Dramatic enhancement of genome editing by CRISPR-Cas9 through improved guide RNA design. Genetics. 199(4):959-971  PubMed

[4] Paix A, Kolkmann A, Rasoloson D, and Seydoux G. (2015) High efficiency, homology-directed genome editing in Caenorhabditis elegans using CRISPR-Cas9 ribonucleoprotein complexes. Genetics. 201(1):47-54  PubMed

[5] Wei Q, Shen Y, Chen X. Shifman Y. and Ellis RE. (2014) Rapid creation of forward genetics tools for C. briggase using TALENs: lessons from nonmodel organisms. Mol. Biol. Evol. 31(2):468-473  PubMed

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Analysis of mitochondrial sodium, magnesium, calcium, manganese, and iron in wild-type (N2) C. elegans

Laura L. Maurer1, Chuanjia Jiang2, Heileen Hsu-Kim2, Joel N. Meyer1
1 Nicholas School of the Environment, Duke University, Durham NC
2 Department of Civil & Environmental Engineering, Duke University, Durham NC
Correspondence to: Laura L. Maurer (laura.kubik@duke.edu)
Mitochondria are dynamic organelles that require a controlled balance of essential ions to perform critical functions within cells. The content of these elements can both influence the effect of and be influenced by different stressors, including aging (Klang et al., 2014) and exposure to toxic chemicals (Sahu et al., 2013). This article provides reference concentrations of five essential ions present in the mitochondrial fraction of young adult N2 C. elegans. Briefly, 50,000 N2s were grown to young adult stage for mitochondrial isolation. Mitochondria were isolated from whole nematodes by dounce homogenization and differential centrifugation into a crude mitochondrial fraction as described (Kayser et al., 2004) with minor modifications. Elemental analysis of mitochondrial fractions was performed by inductively coupled plasma mass spectrometry (ICP-MS). Samples were prepared by oven drying (95°C overnight) and HNO3 digestion (90°C for 4 h), followed by dilution with MilliQ water to a final HNO3 concentration of 3% prior to elemental analysis by ICP-MS. Results were normalized to protein content (BCA assay). Sodium (Na) was present in the highest concentrations compared to other elements (significantly higher than calcium (Ca), magnesium (Mg), iron (Fe), and manganese (Mn); One-way ANOVA with Tukey’s post-hoc test, *p≤0.05; means are denoted above each bar). Mitochondrial Na concentrations are significantly higher than Ca concentrations in mammalian cells (Murphy and Eisner, 2009), and mitochondria can be a sink for magnesium (Kubota et al., 2005) as well as iron in mammals. It should be noted, however, that measured mitochondrial ion concentrations are influenced greatly by the method used for isolation; caution should be exercised when interpreting relative ion concentrations. These data serve as reference concentrations for mitochondria isolated from young adult N2 utilizing the cited methods.

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ng_ug protein Mito Fraction N2

References

Kayser, E.B., Morgan, P.G., and Sedensky, M.M. (2004). Mitochondrial complex I function affects halothane sensitivity in Caenorhabditis elegans. Anesthesiology 101, 365-372.  PubMed

Klang, I.M., Schilling, B., Sorensen, D.J., Sahu, A.K., Kapahi, P., Andersen, J.K., Swoboda, P., Killilea, D.W., Gibson, B.W., and Lithgow, G.J. (2014). Iron promotes protein insolubility and aging in C. elegans. Aging (Albany NY) 6, 975-991.  PubMed

Kubota, T., Shindo, Y., Tokuno, K., Komatsu, H., Ogawa, H., Kudo, S., Kitamura, Y., Suzuki, K., and Oka, K. (2005). Mitochondria are intracellular magnesium stores: investigation by simultaneous fluorescent imagings in PC12 cells. Biochim Biophys Acta 1744, 19-28.  PubMed

Murphy, E., and Eisner, D.A. (2009). Regulation of intracellular and mitochondrial sodium in health and disease. Circ Res 104, 292-303.  PubMed

Sahu, S.N., Lewis, J., Patel, I., Bozdag, S., Lee, J.H., Sprando, R., and Cinar, H.N. (2013). Genomic analysis of stress response against arsenic in Caenorhabditis elegans. PLoS One 8, e66431.  PubMed

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