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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

This protocol describes a 96-well disruption of individual bacterially colonized Caenorhabditis elegans following cold paralysis and surface bleaching to remove external bacteria. The resulting suspension is plated on agar plates to allow accurate, medium-throughput quantification of bacterial load in large numbers of individual worms.

Abstract

The nematode Caenorhabditis elegans is a model system for host-microbe and host-microbiome interactions. Many studies to date use batch digests rather than individual worm samples to quantify bacterial load in this organism. Here it is argued that the large inter-individual variability seen in bacterial colonization of the C. elegans intestine is informative, and that batch digest methods discard information that is important for accurate comparison across conditions. As describing the variation inherent to these samples requires large numbers of individuals, a convenient 96-well plate protocol for disruption and colony plating of individual worms is established.

Introduction

Heterogeneity in host-microbe associations is observed ubiquitously, and variation between individuals is increasingly recognized as a contributing factor in population-level processes from competition and coexistence1 to disease transmission2,3,4. In C. elegans, "hidden heterogeneity" within isogenic populations has been observed repeatedly, with sub-populations of individuals showing distinct phenotypes in heat shock response5,6, ageing, and lifespan7,8,9,10,11, and many other aspects of physiology and development12. Most analyses that attempt to identify sub-population structure provide evidence for two sub-populations in experimental populations of isogenic, synchronized worms5,7,8, though other data suggest the possibility of within-population distributions of traits rather than distinct groups7,12,13. Of relevance here, substantial heterogeneity in intestinal populations is observed even within isogenic populations of worms colonized from a shared source of microbes13,14,15,16, and this heterogeneity can be concealed by the batch digest measurements that are widely used17,18,19,20 for bacterial quantification in the worm.

This work presents data suggesting a need for greater reliance on single-worm measurements in host-microbe association, as well as protocols for increasing accuracy and throughput in single-worm disruption. These protocols are designed to facilitate mechanical disruption of large numbers of individual C. elegans for quantification of viable bacterial load, while providing better repeatability and lower effort per sample than pestle-based disruption of individual worms. A recommended gut-purging step, where worms are permitted to feed on heat-killed E. coli prior to the preparation for disruption, is included to minimize contributions from recently ingested and other transient (non-adhered) bacteria. These protocols include a cold-paralysis method for cleaning the cuticle with a low-concentration surface bleach treatment; surface bleaching can be used as a preparatory step in single-worm disruption or as a method for preparing live, externally germ-free worms. This surface-bleaching method is sufficient to remove a wide range of external microbes, and cold treatment provides an alternative to conventional levamisole-based paralysis; while levamisole will be preferred for cold-sensitive experiments, cold paralysis minimizes contributions to hazardous waste streams and allows rapid resumption of normal activity. While the full protocol describes a laboratory experiment where worms are colonized with known bacteria, the procedures for cleaning worms and single-worm disruption can readily be applied to worms isolated from wild samples or colonized in microcosm experiments. The protocols described here produces live bacteria extracted from the worm intestine, suitable for plating and quantification of colony forming units (CFUs) in individual worms; for sequencing-based intestinal community analysis, subsequent cell lysis and nucleic acid extraction steps should be added to these protocols.

Protocol

Worms used in these experiments were obtained from the Caenorhabditis Genetic Center, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440). Bristol N2 is the wild-type. DAF-2/IGF mutants daf-16(mu86) I (CGC CF1038) and daf-2(e1370) III (CGC CB1370) are used to illustrate differences in intestinal bacterial load.

HT115(DE3) E. coli carrying the pos-1 RNAi vector is from the Ahringer library21. The MYb collection of C. elegans native gut bacteria22 was obtained from the Schulenburg lab. Salmonella enterica LT2 (ATCC 700720) attB:GFP-KmR is from this lab23. Pseudomonas mosselii was isolated in this lab. Staphylococcus aureus MSSA Newman pTRKH3-mGFP was obtained from the LaRock lab at Emory University.

All worm buffers and media are prepared according to previously published literature24 with minor modifications (see Supplementary File 1).

1. Preparation of synchronized sterile C. elegans

NOTE: In this section, step-by-step procedures are described for generating a synchronized population of reproductively sterile adult worms. Feeding on pos-1 RNAi plates is used here to prevent production of progeny because this interference is embryonic lethal; L1 larvae raised to adulthood on pos-1 RNAi develop into egg-laying hermaphrodites, but these eggs are inviable25. The RNAi feeding protocol is as in the "Reverse genetics" chapter of Wormbook26.

  1. Before synchronizing worms, ensure that fresh 10 cm NGM + pos-1 RNAi plates are available. Plates can be prepared fresh from concentrated induced liquid culture (+Amp +IPTG) or inoculated as lawns on NGM + 100 µg/mL ampicillin + 1 mM IPTG and allowed to grow at 25 °C in the dark for 1 day27.
    NOTE: Carbenicillin (25 µg/mL) is often used instead of ampicillin on RNAi plates. Ampicillin is less expensive but less stable; if using ampicillin, plates should be seeded immediately once dry and used as soon as possible (can be stored for <1 week at 4 °C)27. The high antibiotic concentration recommended here will help to ensure adequate selection.
  2. Start with several (typically two to four) NGM plates with large populations of gravid hermaphrodites. Isolate eggs using bleach-NaOH synchronization24.
    1. Wash worms off agar plates using 2 mL of sterile ddH2O per plate. Distribute the liquid evenly into 1.5 mL microcentrifuge tubes (one tube per plate or hermaphrodites).
    2. Spin down for ~5 s in a benchtop minicentrifuge (2,000 x g) to pull adults to the bottom of the tubes. Pipette off the supernatant and discard.
    3. Wash with 1 mL of sterile ddH2O; spin down as before and discard the supernatant.
    4. Repeat the previous step to reduce remaining bacterial debris.
    5. Resuspend the contents of each tube in 1 mL of sterile ddH2O. Add to each tube 130 µL of commercial bleach (8.25% sodium hypochlorite) and 130 µL of 5 N sodium hydroxide (NaOH, final concentration 0.5 N).
    6. Vortex tubes vigorously for at least 10-15 s every 2 min until adult bodies have broken up. Do not allow bleach-NaOH treatment to go longer than 5 min to avoid killing eggs.
    7. Spin in a minicentrifuge for 30-60 s at 2,000 x g to pellet the eggs. Pipette off the supernatant and discard. There may or may not be a visible pellet, this is normal.
    8. Add 1 mL of M9 worm buffer and spin for 30-60 s at 2000 x g. Discard the supernatant.
    9. Repeat the rinse step (1.2.8) 5x to thoroughly remove bleach-NaOH mixture, removing as much of the supernatant as possible without disturbing the egg pellet.
    10. Transfer eggs to 10 mL of M9 worm buffer in a 50 mL conical tube or 30 mL culture tube with cap. If using conical tubes, leave the lid unscrewed slightly and use a bit of tape to keep it secure. Incubate with shaking overnight (16 h) at 25 °C and 200 RPM to allow the larvae to hatch.
  3. Transfer synchronized L1 larvae to RNAi plates to grow to adulthood.
    1. Add 2 mL of sterile M9 buffer + 0.01% Triton X-100 (henceforth M9TX-01) to each L1 tube and transfer the entire volume (12 mL) to a 15 mL screw-top conical tube.
    2. Place 15 mL tubes with L1 worms at 4 °C for 10 min to slow the larval movement.
    3. Spin down 15 mL conical tubes in a large tabletop centrifuge (1,500 x g at 4 °C for 3 min; acceleration and deceleration should be no higher than 80% of maximum).
    4. Carefully pipette off the supernatant without disturbing the L1 pellet. Discard the supernatant.
    5. Add 12 mL of cold M9TX-01 to each tube. Repeat the centrifugation. Carefully pipette off and discard the supernatant. Each tube should have ~200 µL remaining.
    6. Rinse a 200 µL pipette tip in M9TX-01 to keep worms from sticking to the plastic, then use this tip to resuspend the worm pellet. Transfer resuspended worms to prepared pos-1 plates by pipetting drops of liquid onto the bacterial lawn.
    7. Incubate the plates at 25 °C until the first day of adulthood.
      NOTE: If growing worms on pos-1 RNAi plates, worms MUST feed ad libitum on the RNAi bacteria until they have fully transitioned to adulthood to ensure high penetrance of the embryonic-lethal phenotype. Check the plates at 24 and 48 h. If the plates appear starved or nearly starved, the worms will need to be moved to fresh plates to finish growing into full-sized adults. To avoid depleting plates before worms are grown, aim to add 250-500 L1 larvae to each 10 cm RNAi plate.
  4. Harvest adults and clear intestinal E. coli to create germ-free worms.
    1. Rinse adult worms from plates using 5 mL of M9TX-01 per plate. Transfer buffer + worms to a 15 mL conical tube and allow adults to settle to the bottom of the tube.
    2. Rinse adults in changes of 10 mL of fresh M9TX-01 buffer until no visible bacterial turbidity remains (typically 1-2x). Tubes can be centrifuged at 700 x g for 30 s to pellet worms, or adults can be allowed to settle by gravity.
    3. Perform one additional wash with 10 mL of M9TX-01 to reduce external bacteria.
    4. Transfer worms to 50 mL conical tubes or 30 mL culture tubes containing 5 mL S Medium + 2x heat-killed E. coli OP50 (~5 x 109 killed cells/mL) + 200 µg/mL of gentamycin + 50 µg/mL of chloramphenicol. If using conical tubes, leave the lid unscrewed slightly and use a bit of tape to keep it secure. Use glass pipettes or rinse plastic pipettes in M9TX-01 to keep the worms from sticking.
    5. Incubate adults at 25 °C with shaking at 200 RPM for 24-48 h to produce germ-free adults.
      NOTE: If the worms are to remain in antibiotics for >24 h, more heat-killed OP50 may have to be added to ensure that the worms have an adequate food source. Check tubes at 24 h and supplement with heat-killed OP50 if turbidity is visibly reduced.
  5. Sucrose wash adults according to the Wormbook protocols24 to obtain clean, reproductively sterile, synchronized adult-only stocks for bacterial colonization.
    1. Ensure that cold volumes of 60% sucrose, M9 worm buffer, and M9TX-01 are ready for the use. For simplicity, these can be left at 4 °C the night before.
    2. For each sample to be washed, create a labeled 15 mL conical tube containing 8 mL of M9TX-01 and set aside on ice. These will be needed in step 1.5.10.
    3. Add 5 mL of M9TX-01 to each 50 mL tube containing L1 larvae. Transfer the entire volume (now 10 mL) to an empty 15 mL screw-top conical tube and allow adults to settle to the bottom of the tube.
    4. Carefully pipette off the supernatant and discard.
    5. Add 10 mL M9TX-01 to each tube and move tubes to an ice bucket for 5-10 min.
      NOTE: Starting at this point, worms and all buffers should be kept on ice.
    6. Use the "fast temp" setting to cool a large tabletop centrifuge to 4 °C.
    7. Add 10 mL of cold M9TX-01 to each tube to rinse off any remaining debris. Let worms settle on ice; remove the supernatant and discard.
    8. Sucrose float: Add 5 mL of cold M9 buffer and 5 mL of cold 60% sucrose solution to each tube, mixing thoroughly. Then, carefully float 1 mL of cold M9 buffer on top of the sucrose-buffer mixture in each tube. Do not mix after the float has been added.
      CAUTION: Move quickly for the next few steps-worms can desiccate if exposed to high concentrations of sucrose for too long!
    9. Centrifuge at 1500 x g for 3 min at 4 °C. Live adult worms will be at the interface of the M9 and the sucrose, approximately 1 mL from the top of the tube.
    10. Use a glass 5 mL serological pipette to transfer the worm layer to prepared 15 mL conical tubes with cold M9TX-01 (from step 1.5.2). Be very careful to get the layer of live worms without pipetting up too much of the sucrose.
    11. If necessary, add M9TX-01 to get equal volumes of 10-12 mL/tube. Centrifuge at 1500 x g at 4 °C for 1 min, then pipette off the supernatant. Worms can be returned to room temperature at this point.
    12. Repeat the wash step 1.5.11 twice, reducing the speed to 700 x g at 4 °C and time to 30 s.

2. Feeding worms on live bacteria in liquid culture

NOTE: This protocol is used to colonize worms with laboratory-grown bacteria in well-mixed conditions in liquid culture (Supplementary Figure 1). Worms can be colonized with individual isolates from pure culture (e.g., pathogens such as Enterococcus faecium28,29) or mixtures of isolates (e.g., minimal microbiome communities14).

  1. Start with sucrose washed synchronized adult worms from protocol step 1.5 in a 15 mL conical tube. Wash the worms once in 12 mL of S buffer and discard the supernatant.
  2. Resuspend the washed worms in the volume of S medium needed for the experiment. Consider the volume of experimental conditions, the number of conditions over which worms will be split, and the final concentrations of worms and bacteria.
    NOTE: Feeding in worms varies with bacterial availability30 and worms can be stressed by crowding31. For colonization in liquid culture, <1000 worms/mL and >107 CFU/mL are recommended; 1011 CFU/mL is considered "ad libitum" feeding density on E. coli32.
  3. Spin down bacterial cultures. Pour off the supernatant; aspiration or pipetting can be used to remove the supernatant for bacteria that form loose pellets.
    NOTE: For cultures >5 mL, transfer to 15 mL tubes and spin at ~2800 x g in a large tabletop centrifuge for 8-10 min. Cultures <5 mL can be transferred to 1.5 mL tubes and centrifuged at 9000 x g for 1-2 min in a small tabletop centrifuge. Highly motile bacteria (e.g., many species of Pseudomonas) may need to be chilled at 4 °C for 10-15 min to facilitate formation of a stable pellet, and it may be better to centrifuge at 4 °C.
  4. Resuspend bacterial cultures in one volume of S buffer and centrifuge again to pellet. Remove and discard the supernatant as before.
  5. Resuspend bacterial cultures in S medium at the desired density for the experiment, plus any antibiotics for selection. The antibiotics to be used, if any, will depend on the resistance profile of the bacteria used for colonization.
  6. Using a pipette tip coated in M9TX-01, pipette worms gently up and down until worms are thoroughly resuspended in S medium, then transfer to tubes or plate wells for bacterial colonization.
  7. Add bacterial suspension to each worm culture to reach the desired bacterial concentration and final volume.
  8. If using a multi-well plate for colonization, cover the plate with a sterile 96-well gas-permeable sealing membrane.
  9. Incubate with shaking at 200 RPM to prevent bacteria from settling during incubation.

3. Mechanical disruption of individual worms in a 96-well format

NOTE: This section describes a 96-well plate format protocol for mechanical disruption of individual bacterially colonized C. elegans. The first steps in the protocol (3.1-3.8) describe a method for purging non-adhered bacteria from the worm intestine and cleaning the exterior of the worms using cold paralysis and surface-bleaching. These steps will produce clean, live adult worms that can be mechanically disrupted for quantification of bacterial contents (3.8-end) or used for further experiments (Supplementary Figure 1). This protocol can be adapted to quantify bacteria in worms colonized in liquid culture (Section 2), on agar plates, or from natural or microcosm soil.

  1. Place an aliquot of M9TX-01 on ice to chill (4-5x the number of samples in mL).
  2. Prepare an aliquot of M9TX-01 + bleach (6% sodium hypochlorite, 1:1000 or 1:2000 v/v, 1 mL per sample + 1 mL extra) and place on ice to chill. This aliquot will be used in step 3.8.
  3. Prepare 96-well plates for serial dilution of disrupted worm samples.
    1. Obtain sterile 300 µL capacity 96-well plates with lids; this protocol uses one dilution plate per 12 worms digested.
    2. Use a 96-well multichannel pipettor to fill rows B-D of each 96 well plate (300 µL capacity) with 180 µL of 1x PBS buffer. Leave the top row empty. Rows B-D will become 10x serial dilutions of the worm digests [0.1x, 0.01x, 0.01x].
    3. Set plates aside. Dilution plates will be used in step 3.13.
  4. Resuspend each worm sample in 1 mL of M9TX-01 in a 1.5 mL microcentrifuge tube.
  5. Spin tubes briefly (2-3 s) in a low speed minicentrifuge (2,000 x g) at 25 °C to pellet adults. Pipette off the supernatant and discard, being sure not to disturb the worm pellet.
  6. Using the centrifugation settings in step 3.5, rinse worms twice with 1 mL of M9TX-01, then once with 1 mL of M9 worm buffer, to reduce external bacteria.
  7. Purge non-adhered bacteria from the worm intestine.
    1. Resuspend each sample of worms in 1 mL of S medium + 2x heat-killed OP50 in a culture tube.
    2. Incubate at 25 °C for 20-30 min to allow passage of any non-adhered bacteria from the gut.
      NOTE: This will also purge any extracellular fluorescent protein from the lumen and allow clearer visualization of labeled bacteria adhered to the intestinal epithelium, particularly when acid-fast fluorophores (e.g., mCherry, dsRed) are used.
  8. Surface bleach worms to clear external bacteria.
    1. Rinse purged worms twice with 1 mL of cold M9TX-01 and discard the supernatant.
    2. Allow tubes to chill for 10 min on ice (preferred) or at 4 °C. This will paralyze worms and prevent ingestion of bleach.
      NOTE: Other protocols use a chemical paralysis agent such as levamisole; this is an established approach33 which requires addition of a hazardous waste stream.
    3. Add 1 mL of ice-cold M9 worm buffer + unscented bleach (8.25% sodium hydroxide, 1:1000 or 1:2000 v/v) to each tube. Allow tubes to sit on ice (preferred) or at 4 °C for at least 10 min to kill external bacteria.
      NOTE: Do not exceed 1:1000 concentration of bleach. Even in paralyzed worms, mortality can result.
    4. Pipette off bleach buffer and discard; return tubes to ice to ensure worms do not resume pumping until bleach is cleared.
    5. Add 1 mL of cold M9TX-01 to each tube. Spin for ~5 s in a minicentrifuge (2,000 x g at 25 °C); return tubes to ice. Remove the supernatant and discard.
    6. Repeat this rinse step with another 1 mL of cold M9TX-01, discarding as much of the supernatant as possible.
      NOTE: If using worms for further experiments, skip the permeabilization step (Protocol 3.9) and instead transfer freshly surface-bleached adults to ice-cold buffer in a 6 cm Petri dish and separate worms into experimental conditions as in Protocol 3.10. Keep worms cold to prevent motility from resuming but work quickly - keeping worms for >30 min on ice can potentially result in <100% resumption of normal activity34.
  9. Chemical permeabilization of worm cuticle with sodium dodecyl sulfate and dithiothrietol (0.25% SDS + 300 mM DTT) (based on35)
    CAUTION: DTT is a reducing agent and irritant. Wear PPE and work in a fume hood when handling dry stocks or solutions. A hazardous waste stream is required.
    1. In the fume hood, prepare enough SDS/DTT solution to allow 100 µL for each sample. For 1 mL, to 965 µL of M9 worm buffer or M9TX-01 in a 1.5 mL microcentrifuge tube, add 5 µL of 5% (w/v) SDS and 30 µL of 1M DTT.
      NOTE: 1 M DTT solution (aqueous) should be prepared fresh or stored in aliquots at -20 °C to ensure potency. Aliquots should be sized to be used up in two to three experiments to avoid excessive freeze-thaw cycling.
    2. Move microcentrifuge tubes containing surface-bleached worms to a room-temperature tube rack. Each tube should contain worms in ~20 µL of buffer.
    3. Add 100 µL of SDS/DTT solution to each worm sample. Dispose of any remaining SDS/DTT solution in the appropriate hazardous waste stream.
    4. Allow the treatment to proceed for up to 8 min on the bench to partially break down the resistant cuticle of the adult worms. Worms will die and settle to the bottom of the tube during this time.
    5. After permeabilization time is up, carefully pipette off the SDS/DTT supernatant and dispose of it in an appropriate SDS/DTT hazardous waste stream.
    6. Add 1 mL of M9TX-01 to each tube. Spin briefly in a table-top centrifuge to pellet the worms or allow worms to settle by gravity to the bottom of the tubes, then draw off the supernatant and dispose in an SDS/DTT hazardous waste stream.
    7. Resuspend worms in 1 mL M9 worm buffer + 0.1% Triton X-100 until ready to use.
  10. Separate worms into a deep 96-well plate with silicon carbide grit for mechanical disruption. Prepare the 96-well disruption plate as under.
    1. Obtain a sterile 2 mL deep-well 96-well plate and a matching silicon 96-well plate cover.
      NOTE: It is important to use plates that are compatible with the 96-well adaptors for the tissue disruptor. Tiny differences in external dimensions make the difference between a plate that can be removed from the adaptors and one that cannot.
    2. Using a sterile scoop spatula, add a small amount of sterile 36-grit silicon carbide to each well of the plate that will receive a worm. Use enough grit to barely cover the bottom of the well (about 0.2 g per well). Excessive material will make it difficult to get a pipette tip to the bottom of the well when retrieving the contents.
    3. Add 180 µL of M9 worm buffer to each well.
    4. Label the columns or rows to indicate where each sample will go, then cover the plate loosely with the silicon 96-well plate cover.
  11. Transfer individual worms to the 96-well plate for disruption.
    1. Move permeabilized worms carefully to a small (35 or 60 mm) Petri dish containing sufficient M9TX-01 to fill the dish to a depth of ~1 cm.
      NOTE: If a large number of worms are present, it may not be feasible to transfer the entire sample as the liquid will become crowded and it will be difficult to pipette individual worms.
    2. Using a dissecting microscope or other low-magnification device, pipette off individual worms in 20 µL volumes, and transfer these worms to individual wells of the 96-well plate.
      NOTE: It is best to harvest only freshly killed worms. Avoid worms with a rigid linear shape, as these worms may have been dead for some time. Try to take worms that are curved or S-shaped, with normal gross physiology and an intact gut.
    3. After transferring each volume, make sure that the selected worm was actually ejected into the well. To do this, pipette up 20 µL of M9TX-01 from a clear area of the Petri dish and release the full volume back into the dish; this will normally eject the worm if it is stuck to the pipette. If the worm was stuck, remove 20 µL from the well and try the transfer again.
    4. Once all worms have been transferred, cover the 96-well plate with a sheet of commercially available flexible paper-backed sealing film (2 x 2 squares), making sure that the paper-backed side of the sealing film is facing down onto the sample wells. Be careful not to stretch the sealing film too thin, or it will be very difficult to remove later.
    5. Place the silicon sealing mat lightly on top of the flexible sealing film; do not press the cover down into the wells at this time.
    6. Move the plate to 4 °C to chill for 30-60 min. This will prevent over-heating during disruption, which can damage the samples.
      NOTE: This is a break point in the protocol. In most cases, the plate can be left at 4°C for up to 4 h before grinding. Do not leave the worms overnight, as this will change the bacterial counts.
  12. Load 96-well plates onto a tissue disruptor to break up worm tissues and release intestinal bacteria.
    NOTE: (Optional) If using an odd number of 96-well plates for digests, it is necessary to prepare a counterweight before proceeding. Use an empty deep 96-well plate and fill wells with water until it weighs the same as the first plate.
    1. Press the silicon sealing mat down firmly into the wells to create a seal, making sure the lid lies flat across the entire surface of the plate.
      NOTE: If the flexible sealing film is too thick after stretching, it will be difficult to secure the silicon lid such that it is lying flat in all wells. This will result in an insufficient seal and well-to-well contamination during shaking.
    2. Secure plates in the tissue disruptor using the 96-well plate adaptors. Shake plates for 1 min at 30 Hz, then rotate plates 180° and shake again for 1 min. This will help ensure even disruption in all wells of the plate.
    3. Tap plates firmly on the bench two or three times to dislodge any grit from the flexible sealing film.
    4. Using a large centrifuge with two 96-well plate adaptors, spin the plates down at 2400 x g for 2 min to gather all material to the bottom of the wells.
    5. Remove the silicon lid and carefully pull off the flexible sealing film.
      NOTE: If the flexible sealing film sticks in any of the wells, use a 200 µL pipette tip to remove it. This is common when the flexible sealing film was stretched too thin.
  13. Serially dilute worm digest samples in 300 µL in 96-well plates.
    1. Using a multi-well pipettor set to 200 µL, pipette up and down several times slowly and carefully to re-mix the contents of the wells, then draw off as much of the liquid as possible. Transfer this liquid to the top rows of the 96-well plates prepared in step 3.3.
    2. Using a 96-well pipettor set to 20 µL, remove this volume of liquid from the top row and dispense into row B. Pipette up and down 8-10x to mix. Discard tips.
    3. Repeat step 3.13.2, starting from the 0.1x samples in row B to create 0.01x dilution samples in row C.
    4. Repeat step 3.13.2 again, going from row C to row D.
    5. Plate onto solid agar for bacterial quantification. For mono-colonized worms, it is generally sufficient to plate 10-20 µL drops of each dilution [1x-0.001x] on agar plates. For multi-species colonization, plate each dilution separately by pipetting 100 µL onto a 10 cm agar plate; spread immediately using glass plating beads.

4. Cleaning silicon carbide grit for re-use

NOTE: This procedure is used to clean and sterilize the grinding material, silicon carbide grit, for re-use after experiments. This protocol should be followed in its entirety before first use, as silicon carbide grit is an industrial product and does not come pre-sterilized. Si-carbide grit (3.2 g/cc) is a dense, rough-edged material that works efficiently to disrupt tough samples. However, the particles can wear down over repeated use and should be replaced when wear becomes apparent. Fortunately, the material is inexpensive, and the sizes typically sold (~1 lb) are sufficient for many experiments.

  1. After removing samples for plating, add 10% bleach solution to all wells of the 96-well plate and let sit for at least 10 min.
  2. Remove the bulk of the grit by rapidly inverting the 96-well plate over a small high-sided tray or empty P1000 pipette tip container large enough to catch all the contents. The grit will sink immediately to the bottom of the tray. Pour off the bleach solution into a sink.
  3. Re-fill the 96-well plate with tap water and invert into the same tray to rinse out remaining grit. Pour the water off into the sink.
  4. Repeat one to three more times with tap water until plate is completely clear of grit.
  5. Rinse grit 2x in tap water, filling the tray each time.
    NOTE: The 96-well deep-well plate can be washed in a laboratory dishwasher, covered securely with foil, and autoclaved with other reusable plastics. Grit does not need to be washed immediately and can be set aside at this point. Used grit is usually accumulated from multiple experiments before washing and autoclaving.
  6. Wash grit in a solution of laboratory detergent for 30 min, agitating occasionally by swirling or mixing with a metal spatula.
  7. Rinse away all traces of detergent in several (8-10) changes of tap water, then rinse 2x with distilled water.
  8. Spread grit in an open tray, such as a shallow polypropylene autoclave tray, and dry at 40-70 °C for several hours.
    NOTE: If the grit is clumpy when dry, it was not cleaned or rinsed sufficiently. Repeat the cleaning protocol starting at step 4.6, adding additional rinses in step 4.7.
  9. Distribute clean, dry grit into screw-top autoclavable glass bottles to a maximum depth of 5-6 cm. Autoclave on pre-vacuum cycle for 30 min to sterilize.

Results

Bleach sterilization of live worms
Surface-bleached worms are effectively free of external bacteria until motility returns and excretion resumes. Under the conditions used here, rapid extinction of bacteria in buffer is observed (Figure 1A-C, Supplementary Figure 2, Video 1) without disturbing the gut-associated bacteria in cold-paralyzed worms (Figure 1D-F, Video 2). Thes...

Discussion

Here data are presented on the advantages of single-worm quantification of bacterial load in C. elegans, along with a 96-well disruption protocol to allow the rapid and consistent acquisition of large data sets of this type. As compared with existing methods33, these protocols allow higher-throughput measurement of intestinal microbial communities in the worm.

This approach has plating as a rate-limiting step and is not truly "high-throughput". Large-ob...

Disclosures

The authors have no conflicts of interest.

Acknowledgements

The authors would like to acknowledge H. Schulenberg and C. LaRock for their generous sharing of bacterial strains used in these experiments. This work was supported by funding from Emory University and NSF (PHY2014173).

Materials

NameCompanyCatalog NumberComments
96-well flat-bottom polypropylene plates, 300 uLEvergreen Labware290-8350-03F
96-well plate sealing mat, silicon, square wells (AxyMat)AxygenAM-2ML-SQ
96-well plates, 2 mL, square wellsAxygenP-2ML-SQ-C-S
96-well polypropylene plate lidsEvergreen Labware290-8020-03L
AgarFisher Scientific443570050
Bead mill adapter set for 96-well platesQIAGEN119900Adapter plates for use with two 96-well plates on the TissueLyser II
Bead mill tissue homogenizer (TissueLyser II)QIAGEN85300Mechanical homogenizer for medium to high-throughput sample disruption
BioSorterUnion BiometricaBy quotationLarge object sorter equipped with a 250 micron focus for C. elegans
Bleach, commercial, 8.25% sodium hypochloriteClorox
Breathe-Easy 96-well gas permeable sealing membraneDiversified BiotechBEM-1Multiwell plate gas permeable polyurethane membranes. Thin sealing film is permeable to O2, CO2, and water vapors and is UV transparent down to 300 nm. Sterile, 100/box.
Calcium chloride dihydrateFisher ScientificAC423525000
CholesterolVWRAAA11470-30
Citric acid monohydrateFisher ScientificAC124910010
Copper (II) sulfate pentahydrateFisher ScientificAC197722500
Corning 6765 LSE Mini MicrocentrifugeCorning COR-6765
Disodium EDTAFisher Scientific409971000
DL 1,4 Dithiothreitol, 99+%, for mol biology, DNAse, RNAse and Protease free, ACROS OrganicsFisher Scientific327190010
Eppendorf 1.5 mL microcentrifuge tubes, naturalEppendorf
Eppendorf 5424R microcentrifugeEppendorf540600064024-place refrigerated benchtop microcentrifuge
Eppendorf 5810R centrifuge with rotor S-4-104Eppendorf226270403L benchtop centrifuge with adaptors for 15-50 mL tubes and plates
Eppendorf plate bucket (x2), for Rotor S-4-104Eppendorf22638930
Ethanol 100%Fisher ScientificBP2818500
Glass beads, 2.7 mmLife Science ProductsLS-79127
Glass beads, acid-washed, 425-600 µmSigmaG877-500G
Glass plating beadsVWR76005-124
Hydrochloric acidVWRBDH7204-1
Iron (II) sulfate heptahydrateFisher Scientific423731000
Kimble Kontes pellet pestle motorDWK Life Sciences749540-0000
Kimble Kontes polypropylene pellet pestles and microtubes, 0.5 mLDWK Life Sciences749520-0590
Leica DMi8 motorized inverted microscope with motorized stageLeica11889113
Leica LAS X Premium softwareLeica11640687
Magnesium sulfate heptahydrateFisher ScientificAC124900010
Manganese(II) chloride tetrahydrateVWR470301-706
PARAFILM M flexible laboratory sealing filmAmcorPM996
PeptoneFisher ScientificBP1420-500
Petri dishes, round, 10 cmVWR25384-094
Petri dishes, round, 6 cmVWR25384-092
Petri dishes, square, 10 x 10 cmVWR10799-140
Phospho-buffered saline (1X PBS)Gold BioP-271-200
Polypropylene autoclave tray, shallowFisher Scientific13-361-10
Potassium hydroxideFisher ScientificAC134062500
Potassium phosphate dibasicFisher ScientificBP363-1
Potassium phosphate monobasicFisher ScientificBP362-1
R 4.1.3/RStudio 2022.02.0 build 443R Foundationn/a
Scoop-type laboratory spatula, metalVWR470149-438
Silicon carbide 36 gritMJR Tumblersn/aBlack extra coarse silicon carbide grit. Available in 0.5-5 lb sizes from this vendor.
Sodium dodecyl sulfateFisher ScientificBP166-100
Sodium hydroxideVWRBDH7247-1
Sodium phosphate dibasic anhydrousFisher ScientificBP332-500
Sodum chlorideFisher ScientificBP358-1
SucroseFisher ScientificAC419760010
Tri-potassium citrate monohydrateFisher ScientificAC611755000
Triton X-100Fisher ScientificBP151-100
Zinc sulfate heptahydrateFisher ScientificAC205982500

References

  1. Armitage, D. W., Jones, S. E. How sample heterogeneity can obscure the signal of microbial interactions. The ISME Journal. 13 (11), 2639-2646 (2019).
  2. Stephenson, J., et al. Host heterogeneity affects both parasite transmission to and fitness on subsequent hosts. Philosophical Transactions of the Royal Society B: Biological Sciences. 372 (1719), 20160093 (2017).
  3. VanderWaal, K. L., Ezenwa, V. O. Heterogeneity in pathogen transmission: mechanisms and methodology. Functional Ecology. 30 (10), 1606-1622 (2016).
  4. Dwyer, G., Elkinton, J. S., Buonaccorsi, J. P. Host heterogeneity in susceptibility and disease dynamics: tests of a mathematical model. The American Naturalist. 150 (6), 685-707 (1997).
  5. Wu, D., Rea, S. L., Yashin, A. I., Johnson, T. E. Visualizing hidden heterogeneity in isogenic populations of C. elegans. Experimental Gerontology. 41 (3), 261-270 (2006).
  6. Yashin, A. I., et al. Heat shock changes the heterogeneity distribution in populations of Caenorhabditis elegans does it tell us anything about the biological mechanism of stress response. The Journals of Gerontology Series A: Biological Sciences and Medical Sciences. 57 (3), 83-92 (2002).
  7. Zhao, Y., et al. Two forms of death in ageing Caenorhabditis elegans. Nature Communications. 8 (1), 1-8 (2017).
  8. Eckley, D. M., et al. Molecular characterization of the transition to mid-life in Caenorhabditis elegans. AGE. 35 (3), 689-703 (2012).
  9. Rea, S. L., Wu, D., Cypser, J. R., Vaupel, J. W., Johnson, T. E. A stress-sensitive reporter predicts longevity in isogenic populations of Caenorhabditis elegans. Nature Genetics. 37 (8), 894-898 (2005).
  10. Kinser, H. E., Mosley, M. C., Plutzer, I. B., Pincus, Z. Global, cell non-autonomous gene regulation drives individual lifespan among isogenic C. elegans. eLife. , (2021).
  11. Churgin, M. A., et al. Longitudinal imaging of Caenorhabditis elegans in a microfabricated device reveals variation in behavioral decline during aging. eLife. 6, 26652 (2017).
  12. Perez, M. F., Francesconi, M., Hidalgo-Carcedo, C., Lehner, B. Maternal age generates phenotypic variation in Caenorhabditis elegans. Nature. 552 (7683), 106-109 (2017).
  13. Baeriswyl, S., et al. Modulation of aging profiles in isogenic populations of Caenorhabditis elegans by bacteria causing different extrinsic mortality rates. Biogerontology. 11 (1), 53 (2009).
  14. Taylor, M., Vega, N. M. Host immunity alters community ecology and stability of the microbiome in a Caenorhabditis elegans model. mSystems. 6 (2), 00608-00620 (2021).
  15. Diaz, S. A., Restif, O. Spread and transmission of bacterial pathogens in experimental populations of the nematode Caenorhabditis elegans. Applied and Environmental Microbiology. 80 (17), 5411-5418 (2014).
  16. Twumasi-Boateng, K., Berg, M., Shapira, M. Automated separation of C. elegans variably colonized by a bacterial pathogen. Journal of Visualized Experiments: JoVE. (85), e51090 (2014).
  17. Ortiz, A., Vega, N. M., Ratzke, C., Gore, J. Interspecies bacterial competition regulates community assembly in the C. elegans intestine. The ISME Journal. 15 (7), 2131-2145 (2021).
  18. Berg, M., et al. TGFβ/BMP immune signaling affects abundance and function of C. elegans gut commensals. Nature Communications. 10 (1), 604 (2019).
  19. Portal-Celhay, C., Blaser, M. J. Competition and resilience between founder and introduced bacteria in the Caenorhabditis elegans gut. Infection and Immunity. 80 (3), 1288-1299 (2012).
  20. Scott, E., Holden-Dye, L., O'Connor, V., Wand, M. E. Intra strain variation of the effects of gram-negative ESKAPE pathogens on intestinal colonization, host viability, and host response in the model organism Caenorhabditis elegans. Frontiers in Microbiology. 10, 3113 (2020).
  21. Kamath, R. S., Martinez-Campos, M., Zipperlen, P., Fraser, A. G., Ahringer, J. Effectiveness of specific RNA-mediated interference through ingested double-stranded RNA in Caenorhabditis elegans. Genome Biology. 2 (1), (2001).
  22. Dirksen, P., et al. The native microbiome of the nematode Caenorhabditis elegans: gateway to a new host-microbiome model. BMC Biology. 14, 38 (2016).
  23. Vega, N. M., Allison, K. R., Samuels, A. N., Klempner, M. S., Collins, J. J. Salmonella typhimurium intercepts Escherichia coli signaling to enhance antibiotic tolerance. Proceedings of the National Academy of Sciences. 110 (35), 14420-14425 (2013).
  24. Stiernagle, T. Maintenance of C. elegans. WormBook. , (2006).
  25. Tabara, H., et al. The rde-1 gene, RNA interference, and transposon silencing in C. elegans. Cell. 99 (2), 123-132 (1999).
  26. Ahringer, J. Reverse genetics. WormBook. , (2006).
  27. Rual, J. -. F., et al. Toward improving Caenorhabditis elegans phenome mapping with an ORFeome-based RNAi library. Genome Research. 14 (10), 2162-2168 (2004).
  28. Revtovich, A. V., et al. Development and characterization of high-throughput Caenorhabditis elegans - Enterococcus faecium infection model. Frontiers in Cellular and Infection Microbiology. 11, 667327 (2021).
  29. Anderson, Q. L., Revtovich, A. V., Kirienko, N. V. A high-throughput, high-content, liquid-based C. elegans pathosystem. JoVE (Journal of Visualized Experiments. (137), e58068 (2018).
  30. Scholz, M., Dinner, A. R., Levine, E., Biron, D. Stochastic feeding dynamics arise from the need for information and energy. Proceedings of the National Academy of Sciences. 114 (35), 9261-9266 (2017).
  31. Wu, T., et al. Pheromones modulate learning by regulating the balanced signals of two insulin-like peptides. Neuron. 104 (6), 1095-1109 (2019).
  32. Ching, T. -. T., Hsu, A. -. L. Solid plate-based dietary restriction in Caenorhabditis elegans. Journal of Visualized Experiments: JoVE. (51), e2701 (2011).
  33. Walker, A. C., Bhargava, R., Vaziriyan-Sani, A. S., Brust, A. S., Czyz, D. M. Quantification of bacterial loads in Caenorhabditis elegans. Bio-protocol. 12 (2), 4291-4291 (2022).
  34. Manjarrez, J. R., Mailler, R. Stress and timing associated with Caenorhabditis elegans immobilization methods. Heliyon. 6 (7), 04263 (2020).
  35. Zhang, S., Banerjee, D., Kuhn, J. R. Isolation and culture of larval cells from C. elegans. PLoS ONE. 6 (4), 0019505 (2011).
  36. Garsin, D. A., et al. Long-lived C. elegans daf-2 mutants are resistant to bacterial pathogens. Science. 300 (5627), 1921 (2003).
  37. Thutupalli, S., et al. Farming and public goods production in Caenorhabditis elegans populations. Proceedings of the National Academy of Sciences. 114 (9), 2289-2294 (2017).
  38. Ly, K., Reid, S. J., Snell, R. G. Rapid RNA analysis of individual Caenorhabditis elegans. MethodsX. 2, 59-63 (2015).
  39. Johnke, J., Dirksen, P., Schulenburg, H. Community assembly of the native C. elegans microbiome is influenced by time, substrate, and individual bacterial taxa. Environmental Microbiology. 22 (4), 1265-1279 (2020).
  40. Vega, N. M., Gore, J. Stochastic assembly produces heterogeneous communities in the Caenorhabditis elegans intestine. PLOS Biology. 15 (3), 2000633 (2017).
  41. Gulyas, L., Powell, J. R. Cold shock induces a terminal investment reproductive response in C. elegans. Scientific Reports. 12 (1), 1338 (2022).
  42. Jiang, W., et al. A genetic program mediates cold-warming response and promotes stress-induced phenoptosis in C. elegans. eLife. 7, 35037 (2018).
  43. Robinson, J. D., Powell, J. R. Long-term recovery from acute cold shock in Caenorhabditis elegans. BMC Cell Biology. 17 (1), 2 (2016).

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