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Protein extraction from C. elegans using sea sand

Shamol Saha1, Julien Dusonchet1 and Benjamin Wolozin1
1Departments of Pharmacology and Neurology, Boston University School of Medicine, Boston MA

Correspondence to: Shamol Saha (saha@bu.edu)

Making mini scale extracts from C. elegans for visualizing proteins and RNA has been difficult and inefficient due to the nematode’s sturdy outer shell, the cuticle.  Boiling and sonication have been the methods of choice with some success, but both have their limitations. Boiling nematodes irreversibly denatures proteins, while sonication produces low, inconsistent yields. Even an extended period of sonication leaves many intact and halved nematodes in the extract, which renders biochemical experiments rather challenging and erratic. Boiling nematodes in SDS sample buffer provides an efficient way to visualize proteins on SDS-page protein gels, but certain proteins do not enter the gel due to the heat treatment. Here, we show a quick and efficient method of worm grinding with clean sea sand. The method is fast and high-quality extracts can be obtained in an Eppendorf tube for protein gel and RNA extractions.  If desired, the sand can be separated from the extract and the clean extract can be obtained in a minimum amount of time.

Method:  The method is optimized for 40 to 100 μL of clean worms.  The worms should be totally cleaned from the bacteria with two to three washes with double distilled water (0.01% Triton X-100) in a clean Eppendorf tube prior to lysis. Extra water should be gently removed to make sure that addition of the 2X sample buffer yields the appropriate dilution of buffer for sand grinding. A small scoop of 30 to 40 μL volume of washed clean sand (Fisher Chemicals catalog #14808-60-7) should be added to the worm pellet. 2X LDS* (Lithium Dodecyl Sulphate) sample buffer with protease inhibitors is added (equal to worm volume) to the tube and transferred to an ice-cold metal block; other sample buffers are also acceptable. Immediately after transfer, lysing of the worms is performed. An ice-chilled pestle (glass or plastic) is inserted into the bottom of the microfuge tube and rotated fifty to seventy five circular strokes until very little resistance is felt.  At this point, the extract can be flash frozen and handled at a later time-point. For a smaller volume of extract, it is rather important to separate the extract from the sand to maximize the yield. This can be easily performed by filtering the sand extract through a small microfuge tube column with a small needle-size hole in its bottom. The hole should be covered with 100uL of clean glass beads (425-600 μm, Sigma G8772) and be placed on top of another empty clean microfuge tube. The stack of tubes can be assembled in a 15 ml blue cap tube. Once the sand extract is ready, the tube should be put on top the tube that has glass beads at its bottom. A clean injection needle can now be pushed through the bottom of the tube with sand extract until a hole is made. Centrifugation at 2K RPM for 1 to 2 minutes provides an extract that is virtually free from sand particles.

For RNA preparation, washed worms are harvested in Trizol solution (Life Technologies), at which stage they can be snap-frozen and stored at -80 0C for RNA extraction at a later time-point.  Worms are ground in Trizol with sea sand and pestle as described above. After filtering of the sand, RNA is extracted using the PureLink RNA Mini Kit (Life Technologies) and eluted in RNAse-free ddH20.  Extra caution should be taken to make sure that sand, tubes, solutions and tools are totally RNAse free. The sand should be baked at 240 °C for at least 4 hours and plastic RNAse-free pestles should be purchased.

*2X LDS sample buffer: 100 mM Na-Tricine pH 7.8, 100 mM DTT, 14% W/V Glycerol, 4% LDS, 0.05% CHAPS and 0.002% Bromphenol Blue

Acknowledgements:  This work was supported by funding to BW from NIH (NS060872 and ES020395).

Figures



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Figure 1: Basic tools required for making extract


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Figure 1: A schematic diagram showing design of the column for removal of sand from the samples


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Figure 1: A. A SDS page gel showing Actin and Lgg1:mCherry fusion protein from two different transgenic nematodes (lanes 1 and 2). B. Total RNA extracted from n=100 day 5 adult worms with sand/pestle method. Visualized on 1% agarose gel.

Editor’s note:
Articles submitted to the Worm Breeder’s Gazette should not be cited in bibliographies. Material contained here should be treated as personal communication and be cited as such and only with the consent of the author.

Using magnetism to move C. elegans

Peter Wawrzusin1, Yicong Wu1, Peter Winter1 and Hari Shroff1
1Section on High Resolution Optical Imaging, National Institute of Biomedical Imaging and Bioengineering, National Institutes of Health, Bethesda MD

Correspondence to: Peter Wawrzusin (peter.wawrzusin@nih.gov)

C. elegans stands out as a powerful model organism for many reasons including its simple nervous system and relative optical transparency. Hence, techniques that enable researchers to manipulate the movement and position of C. elegans for the purposes of imaging and for behavioral studies are useful. We have developed a novel method of repositioning the nematode that involves feeding the worms magnetic beads mixed with bacteria, and then using a neodymium magnet to move the worms at will.

Methods:

  1. Remove 25 μl of 2.5% w/v 1 μM paramagnetic iron oxide microparticles (Spherotech, AMX-10-10) from the stock solution.
  2. Use a neodymium magnet (Radio Shack, 640-1895) to separate magnetic particles from buffer, pipet out buffer containing sodium azide, and replace with 25 μl of desired buffer (e.g., M9).
  3. Mix magnetic particles with 25 μL of liquid culture OP50 bacteria (1:1 dilution).
  4. Seed this mixture onto an LB agar plate. Pick or chunk a small number of worms onto the plate.
  5. Let the worms grow on the plate at 16 oC for 7 days [wait longer and your efficiency of obtaining useable worms will increase].
  6. Using a stereoscope, ensure that you can see the reddish-tinted iron oxide beads throughout the digestive track of the worm (especially near the head and tail regions). Pick the appropriate worms onto an enclosed aqueous medium (e.g., onto a coverslip inserted into a chamber topped with M9 buffer).
  7. Place the neodymium magnet directly underneath the coverslip (preferably contacting it) at the location that you prefer the nematodes to move to (note that worm may not respond to magnet at distances greater than 25 mm from the center of the magnet).

Upon visual inspection, beads appeared confined to the digestive tract and no negative impact was observed on the development of worms grown on plates with magnetic beads (normal lifespan). The main caveat for this technique is that a long incubation time is required for a satisfactory percentage of worms to consume enough magnetic beads to be sufficiently affected by external magnets. After a 7 day incubation period, ~33% of worms picked at random respond to the magnetic field. Following 14 days of incubation, this percentage increases to ~75%. One can both increase this efficiency and decrease the incubation time by selecting only for worms that have an obvious reddish tint of iron oxide beads throughout their digestive tract.

Note that at shorter incubation times (1-3 days), one should only expect to find adult worms that have consumed a sufficient number of beads. Unfortunately, waiting longer for the nematodes to consume more beads leaves the user with a significantly higher percentage of worms having entered the dauer stage.

We also considered microinjecting beads into the worms or binding them to the surface coat of the worms. The first suggestion was undesirable because of its invasive nature. Binding the magnetic beads to the surface coat of the worm either via ionic bonding (Concanavalin A binding to abundant proteoglycans) or covalent antibody attachment (worm surface coat has the O-glycosylated target for antibody M-38) would be less invasive. However, we speculate that the bond forces holding the beads to the worm would be orders of magnitude less than the forces generated by the living organism’s muscles. Feeding the beads to the worms seemed like the most promising method due to C. elegans’ indiscriminant suction powered eating machinery, the technique’s minimally invasive nature, and the fact that previously nematodes have successfully been fed 0.8 µM latex beads to study food transport through the pharynx (Avery and Shtonda, 2003).

This technology can be used to direct one or more worms to a particular field of view. Microfluidic devices provide the same functionality but take more effort to create, and also introduce a ceiling layer on top of the worm which may be problematic for certain imaging techniques such as inverted selective plane illumination microscopy, in which water dipping objectives are immersed in the aqueous medium from above, to minimize optical aberrations. Another potential use includes behavioral analysis in which researchers need to physically move the worms. With future optimization, this technique may enable other applications.

Figures



Figure 1:By mounting a camera (Thorlabs CMOS, 1545M) onto a stereoscope (Leica, MZ10F), we recorded an adult nematode’s motion as it was pulled toward and held above the magnet.

References

Avery L. and Shtonda B. (2003). Food transport in the C. elegans pharynx. J. Exp. Biol. 206, 2441-2457. PubMed

Live worm respiration in 24-well format

Beverley M. Dancy1, Ernst-Bernhard Kayser1, Margaret M. Sedensky1 and Philip G. Morgan1
1Center for Developmental Therapeutics, Seattle Children’s Research Institute, Seattle WA

Correspondence to: Beverley M. Dancy (beverley.m.dancy@gmail.com)

This is a protocol for the determination of oxygen consumption rate (OCR) in living C. elegans using the Seahorse apparatus with 24-well plates. OCR provides a measurement of respiration, which can be altered by genetic or pharmacologic manipulation of the mitochondrial respiratory chain. Although the Seahorse was originally designed for use with ex vivo cells, it can readily be used in vivo with living C. elegans by following our simple procedure,2 described here for the analysis of four samples side-by-side.

Synchronize each of the four samples of worms. We allow 200 adults to lay eggs for 4 h on a 10 cm plate by picking 200 adults to a 10 cm plate, kill the adults, then study the F1s as day 1 adults. Program the Seahorse Bioscience XF24 Extracellular Flux Analyzer with the following protocol: 1- calibrate probes; 2- loop 10 times; 3- mix 2 min; 4- time delay 2 min; 5- measure 2 min; 6- loop end. Furthermore, the internal heater should be switched off to allow the instrument to be close to room temperature. Hydrate the probes in the extracellular flux assay kit by adding 1 ml of Seahorse calibrant solution to each well and incubating overnight at 37 oC . Load this into the Seahorse on the day of the experiment for calibration.

Rinse the worms off the plate and wash three times using gravity separation in M9 and resuspend each washed sample in 2 ml. Get a rough estimate of the concentration of worms by pipetting 5 drops of 20 µl each per sample onto an empty dish and counting under the dissecting scope. There should be about 2 worms per drop. In the Seahorse assay plate, place 500 µl M9 into four blank wells. At least 30 min after the worms were first rinsed, pipet the four worm samples into a total of 500 ul M9 in each of the other wells with between 25 and 150 worms per well and five replicates of each sample. We use a low retention tip cut to create a wider opening, and found that picking worms into the wells with a platinum wire leads to unstable results.

When prompted by the instrument, replace the calibration plate with the worm plate. At the end of the run, count the actual number of worms per well; for this purpose, we capture images on a camera-fitted dissecting scope. Average the background-corrected OCR (pMol O2/min) across the 10 time-points for each well, then normalize the OCR to the actual number of worms in each well. Calculate the average and standard deviation of OCR per worm across the five replicates for each sample. A moderate amount of variation is expected between time-points, wells, plates, and days: we therefore use a minimum of 10 time points of 5 wells, and on at least 2 plates and/or days, to get a meaningful result.

1 For a more detailed protocol with additional tips, please contact Beverley at Beverley.M.Dancy@gmail.com.

2 We acknowledge other labs using Seahorse for C. elegans who provided some tips when we began this work, including Laurent Mouchiroud in Johan Auwerx’s lab at Ecole Polytechnique Fédérale de Lausanne, David Raizen at University of Pennsylvania, and Paul Brookes at University of Rochester Medical Center.

3 We thank José Antonio Mari (ja_mari@yahoo.com) for putting together the graphics in Figure 1.

Figures



Figure 1: Overview of the Seahorse workflow. The four worm samples (e.g., different mutants, drug treatments, food sources) are represented by navy, yellow, green, and pink colors. The blank (no worm) wells are clear.3


Figure 2: OCR is linearly related to the number of worms per well. In this experiment N2 were synchronized by picking the parents, grown at 20 oC on NGM+OP50, assayed at day 1 of adulthood, and measured in 16 wells of one Seahorse plate in M9 at 22 oC. The average oxygen consumption rate was 13.8 ± 2.5 picomoles per minute per worm. The average across 57 wells (on 6 Seahorse plates, on 5 days) was 10.2 ± 4.6.

A modified leaving assay to study pathogen avoidance

Alexandra Anderson1 and Rachel McMullan1
1Department of Life Science, Imperial College London, London, UK

Correspondence to: Rachel McMullan (r.mcmullan@imperial.ac.uk)

C. elegans constantly encounter pathogenic threats and the ability to recognize and avoid these threats is crucial to survival. C. elegans have evolved a variety of immediate and learned behavioral responses to minimize pathogen exposure (Yook and Hodgkin, 2007; McMullan et al., 2012; and others). Behavioral responses to pathogens are commonly investigated using a food choice assay where animals are given a choice between their standard laboratory food, OP50, and the pathogen of interest (Yook and Hodgkin, 2007; McMullan et al. 2012). The results of this type of assay integrate both the known attractive effect of OP50 and the attractive/repellent effect of the pathogen under investigation. In order to examine only potential repellent cues, we have adapted the male leaving assay (Lipton et al., 2004) to analyze the response of naïve animals upon exposure to Microbacterium nematophilum, a gram positive bacteria which binds to and colonizes the rectal opening of C. elegans.

Method: 90 mm NGM assay plates were prepared using 13 ml NGM in order to make animals tracks easier to observe. Bacterial cultures were grown to OD600 = 1.0 before plating 20 µl in the center of the assay plate.

Single synchronized young adult hermaphrodites were transferred onto individual plates. 10 plates were prepared for each bacterial lawn to be tested.

Plates were kept at 20 oC and examined at 1 hr intervals to determine if the animal was a leaver, meaning that the animal or its tracks can been seen 1 cm or less from the edge of the plate. Animals determined to have left were not scored again at later time points

Results: Animals placed on plates seeded with OP50 show little leaving behaviour over 24 hrs. An avirulent M. nematophilum strain (Akimkina et al., 2006) induces a limited leaving response, whilst virulent M. nematophilum induces a rapid response over 24 hrs (Figure 1).

Using this assay we tested the role of the cyclic nucleotide gated ion channel TAX-2/TAX-4 in this behaviour. This pathway has been shown to be required for wild type responses to M. nematophilum using a food choice assay (Yook and Hodgkin 2007). Avoidance of M. nematophilum depends on TAX-2/TAX-4, since mutations in either TAX-2 or TAX-4 abrogate the leaving response (Figure 2). In wild-type animals TAX-2/TAX-4 is expressed in AWC, AFD, ASE, ASG, ASJ, AQR, BAG, ASK, ASI, AWB, and PQR (Coburn and Bargmann, 1996). The allele tax-2(p694) has a deletion in the promoter region and first exon of tax-2 abolishing its expression in ASE, AQR, AFD, and BAG (Coburn and Bargmann, 1996). In contrast to the response to M. nematophilum in a food choice assay (Yook and Hodgkin, 2007), tax-2(p694) animals show a defective leaving response particularly at longer time points (Figure 2), suggesting these four cells are important for the normal leaving response to this pathogen.

This simple behavioural assay could be adapted to investigate the response to numerous pathogens and removes the influence of the attractive cue from OP50 in food choice assays.

Acknowledgements: We thank Arantza Barrios (University College London) and Richard Poole (University College London) for suggestions and assistance in setting up this assay. We thank Jonathan Hodgkin for M. nematophilum strains.

Figures



Figure 1: Leaving assay response to OP50, avirulent M. nematophilum and virulent M. nematophilum over 24 hrs. Results are average of the mean of three independent experiments, n=30.

Leaving assay Figure 2

Figure 2: Leaving assay response of tax-2/tax-4 mutants to virulent M. nematophilum over 24 hrs. Results are average of the mean of five independent experiments, n=50

References

Akimkina T, Yook K, Curnock S, and Hodgkin J. (2006). Genome characterization, analysis of virulence and transformation of Microbacterium nematophilum, a coryneform pathogen of the nematode Caenorhabditis elegans. FEMS Microbiol. Lett. 264, 145–151. PubMed

Coburn CM, and Bargmann CI. (1996). A putative cyclic nucleotide-gated channel is required for sensory development and function in C. elegans. Neuron 17, 695–706. PubMed

Lipton J, Kleemann G, Ghosh R, Lints R, and Emmons SW. (2004). Mate searching in Caenorhabditis elegans: a genetic model for sex drive in a simple invertebrate. J. Neurosci. 24, 7427-7434. PubMed

McMullan R, Anderson A, and Nurrish S. (2012). Behavioral and immune responses to infection require Gαq-RhoA signaling in C. elegans. PLoS Pathog 8, e1002530. PubMed

Yook, K. and Hodgkin J. (2007). Mos1 mutagenesis reveals a diversity of mechanisms affecting response of Caenorhabditis elegans to the bacterial pathogen Microbacterium nematophilum. Genetics 175, 681-697. PubMed

A simple method for quantifying M. nematophilum clearance from the rectal opening of C. elegans

Alexandra Anderson1, Shinon Kang1 and Rachel McMullan1
1Department of Life Sciences, Imperial College London. London, UK

Correspondence to: Rachel McMullan (r.mcmullan@imperial.ac.uk)

In its natural environment C. elegans encounters many bacterial pathogens and is capable of mounting a conserved immune response to protect itself from these infections. Microbacterium nematophilum (M. nematophilum) is a coryneform gram positive bacteria which binds to the post anal cuticle and colonises the rectal opening of C. elegans. This leads to a chronic, non-lethal infection characterised by gut distension, constipation and an innate immune response that includes swelling of the post-anal region known as the deformed anal region (DAR) phenotype (Hodgkin et al., 2000). M. nematophilum binding to the rectal opening can be visualised using the nucleic acid stain SYTO13 (Invitrogen), which shows green fluorescence when excited at 488-491 nm (Figure 1). Several groups including our own have used this nucleic acid stain to detect the presence of M. nematophilum colonisation in the rectal opening of animals that display the bacterially unswollen (BUS) phenotype following infection (Figure 1) (Nicholas and Hodgkin, 2004; McMullan et al., 2012).

An important mechanism of the host defence system to combat and resolve infections is the ability to clear pathogens. We have utilised the easy detection of SYTO13 to follow the ability of C. elegans to clear M. nematophilum from its rectal opening. By scoring the number of animals in a population that show green fluorescence in the rectal opening at 30 minute time points we can see a decrease in M. nematophilum adherence over time (Figure 2). This reflects loss of M. nematophilum attachment rather than fluorescent decay or loss only of the SYTO13 stain, as re-incubation of the animals with fresh SYTO stain at a 90 minute time point results in the same number of animals with green fluorescence.

Bus mutants which still show pathogen binding, such as mutants in the EGL-30–UNC-73–RHO-1 pathway (McMullan et al., 2012) show a corresponding decrease in their ability to clear the SYTO13 positive M. nematophilum from the rectal opening (Figure 2).

This is the first demonstration of the positive role of the DAR phenotype in aiding pathogen clearance during infection and confirms that the DAR phenotype acts as part of the protective response. This method of quantifying M. nematophilum clearance could be applied to other pathogens where GFP labelled variants cannot be generated making it a simple method to assay the effect of mutations on an animal’s ability to clear infection.

Method: Adult animals were transferred from OP50 plates to infection plates seeded with 10% M. nematophilum and the progeny grown to L4-adult stage.

SYTO13 labeling was performed as described previously (Nicholas and Hodgkin, 2004) except that after a 60 minute incubation with SYTO13 10-20 µl of settled, stained worms were transferred to NGM plates lacking food. A ring of 150 mM Copper Sulphate, 2% SDS was used to prevent animals escaping from the plates.

The number of animals colonized by SYTO13 positive M. nematophilum was scored using a Nikon SMZ1500 microscope with GFP filter.

Acknowledgements:We thank Stephen Nurrish (University College London) for suggestions in setting up this assay. We thank Jonathan Hodgkin for M. nematophilum strains.

Figures



Figure 1: Syto13 staining of the rectal opening in wildtype and DAR defective unc-73(ce362) and mpk-1(ku1) animals showing the presence of adhered M. nematophilum. Arrows denote the rectal opening. Scale bar = 50 μm


Figure 2: Quantitation of pathogen clearance. After 90mins wild-type DAR animals showed significantly faster M. nematophilum clearance than Dar-defective mutants (n=3, 30-50 animals were scored in triplicate in each independent assay)

References

Hodgkin J, Kuwabara P, and Corneliussen B. (2000). A novel bacterial pathogen, Microbacterium nematophilum, induces morphological change in the nematode C. elegans. Curr. Biol. 10, 1615–1618. PubMed

McMullan R, Anderson A, and Nurrish S. (2012). Behavioral and immune responses to infection require Gαq-RhoA signaling in C. elegans. PLoS Pathog 8, e1002530. PubMed

Nicholas H, and Hodgkin J. (2004). The ERK MAP kinase cascade mediates tail swelling and a protective response to rectal infection in C. elegans. Curr. Biol. 14, 1256–1317. PubMed

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