Isolation and Identification of Nematode‐Infecting Microsporidia

Nematodes are naturally infected by the fungal‐related pathogen microsporidia. These ubiquitous eukaryotic parasites are poorly understood, despite infecting most types of animals. Identifying novel species of microsporidia and studying them in an animal model can expedite our understanding of their infection biology and evolution. Nematodes present an excellent avenue for pursuing such work, as they are abundant in the environment and many species are easily culturable in the laboratory. The protocols presented here describe how to isolate bacterivorous nematodes from rotting substrates, screen them for microsporidia infection, and molecularly identify the nematode and microsporidia species. Additionally, we detail how to remove environmental contaminants and generate a spore preparation of microsporidia from infected samples. We also discuss potential pitfalls and provide suggestions on how to mitigate them. These protocols allow for the identification of novel microsporidia species, which can serve as an excellent starting point for genomic analysis, determination of host specificity, and infection characterization. © 2024 The Authors. Current Protocols published by Wiley Periodicals LLC.

widespread nature, much remains unknown about their infection biology.This, coupled with their host reliance, has made the development of effective treatment strategies challenging.Recently, Caenorhabditis elegans and other free-living terrestrial nematodes were found to harbor microsporidia infection in the wild.Although microsporidia only grow inside host cells, infections in nematodes are almost always horizontally transferred, and large amounts of spores are generated.This allows the infection to be propagated, and spores can be frozen for preservation.To date, C. elegans has been used to uncover various aspects of microsporidia infection and the host response to infection (Balla et al., 2015(Balla et al., , 2016;;Estes et al., 2011;Luallen et al., 2016;Reddy et al., 2017Reddy et al., , 2019;;Szumowski et al., 2014Szumowski et al., , 2016;;Tamim El Jarkass et al., 2022;Troemel et al., 2008;Willis et al., 2021).
Sampling of wild nematodes has revealed microsporidia species with the ability to infect various tissues, including the intestine, muscle, and epidermis (Luallen et al., 2016;Wadi et al., 2023;Zhang et al., 2016).These sampling efforts have also revealed microsporidia species with varying host specificity patterns, ranging from specialists that infect only one or two closely related species to generalists that can infect many species of nematodes, including those from different genera.These studies highlight the diversity of microsporidia within the natural habitats of nematodes.Identifying novel species of microsporidia allows for the characterization and understanding of genome evolution, infection dynamics, and host specificity.The most common method for identifying microsporidia infecting bacterivorous nematodes has been light microscopy, with which multiple stages of the parasite life cycle can be identified (Luallen et al., 2016;Troemel et al., 2008;Zhang et al., 2016).This technique relies on mounting a few animals onto a glass slide and looking for microsporidia meronts or spores.The advantages of this approach are that nematodes do not need to be fixed, both proliferative and spore stages can be observed, and fluorescent microscopy is not needed.However, identifying signs of infection can be challenging for beginners, and microsporidia from low-infection populations may be missed.An alternative approach is to use the fluorescent dye Direct Yellow 96 (DY96), which stains the chitin-containing microsporidia spores (Wadi et al., 2023).
Here, we provide multiple protocols to identify, characterize, and propagate microsporidia infection in wild bacterivorous nematodes (Fig. 1).In Basic Protocol 1, we discuss how to identify environmental nematode sources and the steps involved in propagating these nematodes in the laboratory.Support Protocol 1 describes the steps required for generating Escherichia coli OP50, the standardized nematode laboratory food source.Once nematodes are identified, Basic Protocol 2 describes the steps involved in screening these samples for microsporidia infection using the fluorescent dye DY96.Basic Protocol 3 provides a framework to identify the host and pathogen species.Antibiotic cleanup of samples as well as generating a spore preparation is covered in Basic Protocol 4. In Support Protocol 2, we discuss how to generate a bleach-synchronized population of nematodes, which is critical for the steps outlined in Basic Protocol 4.

STRATEGIC PLANNING
Sampling (Basic Protocol 1) should be prioritized during a time of year that will yield the desired nematodes.This will depend on the geographic location; in Toronto, Ontario, nematodes are most abundant in September through November.
Prior to beginning Basic Protocol 1, ensure that E. coli OP50 and 6-and 10-cm seeded nematode growth medium (NGM) plates are prepared (see Support Protocol 1).
Order the oligonucleotides required for PCR amplification (Basic Protocol 3) ahead of time.
Figure 1 Workflow of steps described in Basic Protocols 1 to 4. The first part is collecting samples containing nematodes and is outlined in Basic Protocol 1. Isolated nematodes are then screened for the presence of microsporidia (Basic Protocol 2).Once microsporidia spores are detected, molecular identification of the nematode and parasite species is performed (Basic Protocol 3).Lastly, sample cleanup and spore preparations are conducted (Basic Protocol 4).
NOTE: Follow all the appropriate safety guidelines and regulations for the use and handling of pathogenic microorganisms.Nematode-infecting microsporidia are considered Biosafety Level-1 (BSL-1) organisms, as they have never been observed to infect any other animals besides nematodes (Murareanu et al., 2021).Environmental bacteria or fungi that are inadvertently isolated along with nematodes may be considered BSL-1 or BSL-2 organisms depending on the organization where this work is performed.
NOTE: Follow all relevant safety procedures for field work involved in isolating nematodes.Ensure that if any permits or permissions to sample nematodes are needed, these are obtained before isolation occurs.

GATHERING SAMPLES
In this protocol, we describe the steps involved in identifying and isolating wild samples.The protocol begins with instructions to identify natural substrates in the environment that have a high likelihood of containing nematodes.We describe how to sample these substrates and how to handle them in the laboratory.We also describe how to scan samples for the presence of nematodes and how to discard those containing undesired animals such as mites.The samples that are screened in the laboratory may reveal no, few, or many nematodes and will certainly contain environmental microbes such as bacteria and fungi.Although the goal of this protocol is to identify nematodes that are infected with microsporidia, the nematode strains themselves can be useful to the research community.In particular, the Caenorhabditis species C. elegans, C. briggsae, and C. tropicalis can be sent to CaeNDR, the Caenorhabditis Natural Diversity Resource (https:// caendr.org/ ) (Crombie et al., 2024).Materials 6-cm NGM plates seeded with 10× E. coli OP50 (see Support Protocol 1) Likely substrates for nematodes Sealable sandwich bags (Ziploc® or VWR, 2. Identify areas with likely substrates for nematodes, which are pieces of fruit in an advanced stage of rotting that should be malleable and not dry (Fig. 2A).
Examples of areas include parks and orchards.In our experience, we most often find nematodes in rotting apples, pears, and black walnuts.This list is not exhaustive, so other fruit substrates and plant materials may also yield nematodes.
Several method articles have been published that provide additional details on where to find and how to collect free-living terrestrial nematodes (Barriere & Felix, 2014;Crombie et al., 2022;Gimond et al., 2022).
3. Invert a sealable sandwich bag and collect a sample that is ∼6 cm in diameter.Using the free hand, re-invert and seal the bag.Make note of the substrate, geographical location, and coordinates of the sample.Give each sample a unique identifier (e.g., code or number) and write this identifier on the bag using a marker.Repeat until desired number of samples are obtained, keeping samples in separate bags.
The sandwich bag will serve as a protective glove, avoiding contact with the rotting substrate but providing an effective sample storage solution.Taking note of the substrate and sample location can provide an effective means of identifying those that were most successful.This can help inform subsequent sampling excursions and allow for multi-year comparisons if desired.

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Current Protocols 4. While wearing gloves, remove the lid from a 6-cm seeded NGM plate (see step 1) and place the sample on the plate.Spread the sample out across the plate, replace plate lid, and wrap with parafilm.Store right side up in a plastic storage container with a lid at room temperature for 18 to 48 hr (Fig. 2B).
The storage container should not be placed in an incubator containing sterile nematode strains, as the wild sample likely contains undesired contaminants that may spread.If a designated sampling incubator is available, this can be used.Within 18 to 48 hr, nematodes should have had enough time to crawl from the substrate onto the plate.

5
. Generate a spreadsheet containing information on the sample substrate and location (see Table S1 in the Supporting Information for an example).As more information becomes available about these samples, update accordingly.
We recommend keeping a detailed log to keep track of sample history.Desired information can include the contaminants present, the presence/absence of nematodes, or anything else deemed useful.
6. Remove as much of the substrate as possible and wipe any condensation off the lid.
Try to remove any insects or other invertebrates, if large and visible.Wrap the plate in parafilm and incubate at room temperature until screening the plates for nematodes (see step 7).
Common insects include fruit flies and their larvae, caterpillars, beetles, and earthworms.These animals are visible without a microscope and can be removed using a Kimwipe.
7. Screen the parafilmed plates using a dissecting microscope with transmitted light and look for the presence of nematodes.
Depending on the contamination present on a plate, visually identifying nematodes may be difficult.Fungal hyphae can make this challenging, as can thick lawns of bacteria.If nematodes are not readily seen, look for nematode tracks in the E. coli OP50 (Fig. 2C).If nematodes are not observed, the plate can be screened once more in a few days.If nematodes are still not observed, the sample can be discarded.Remember to annotate the sample spreadsheet, indicating the presence/absence of nematodes and their characteristics.If samples are discarded, make note of the reason why.
8. Characterize nematodes by making note of their appearance and the presence/absence of embryos and/or males.Keep an eye out for mites and set these samples aside (Figs. 2C and 3A).(Barriere & Felix, 2014).
There can be a range of mite sizes on a plate, indicating a heavily infested population.In addition, not all mites look the same.Zoom in and slowly and carefully scan the plates.Mites are readily visible under laboratory dissecting microscopes and have a domed back and eight little legs, unlike insects.Scan the sides and the lid of the plates, as mites can crawl around.If a plate contains a single mite, it may be possible to remove the mite or to chunk a portion of the sample to a new plate.In our experience, however, plates often contain more than a single mite.Therefore, discarding the entire plate is recommended.9. Place a sealable plastic bag inside a plastic beaker (Fig. 3B).Place all the parafilmwrapped plates containing mites into the plastic bag and seal it (Fig. 3B).Microwave the beaker for 1 min, until the agar has liquified.Toss the sealed bag into a biosafety bin liner and immediately discard the liner as biohazardous waste.

GENERATING 10× AND 40× Escherichia coli OP50 AND SEEDING NGM PLATES
This protocol describes the steps involved in generating 10× and 40× E. coli OP50 stocks.The 10× OP50 is used for seeding NGM plates, whereas 40× E. coli OP50 is used in spore preparations to provide a large amount of food in a small volume.We also describe how to seed and store NGM plates.

Materials
E. coli OP50-1 (available from the CGC) 10-cm LB agar plate (see recipe) Sterile 100-ml bottle containing 100 ml sterile LB (see recipe) Two sterile 2-L flasks containing 1 L sterile LB (see recipe) 6% (v/v) bleach (sodium hypochlorite; Fisher, SS290-1) 6-cm unseeded NGM plates (see recipe) 10-cm unseeded NGM plates (see recipe; optional) 37°C incubator 21°C and 37°C shaking incubator (Eppendorf TM Innova TM S44i shaker or equivalent) 500-ml conical tubes, sterile (Fisher Scientific, C431123) The plate should not be incubated >24 hr, as colonies will get too large and become difficult to pick.Store the plate at 4°C for a maximum of 1 month.Use fresh plates to generate OP50 when possible.
2. Working near a flame, pick a single E. coli OP50-1 colony into a sterile 100-ml bottle containing 100 ml sterile LB.
3. Cap the bottle and shake to ensure even distribution throughout the medium.
4. Pour 50 ml LB from step 3 into each of two sterile 2-L flasks containing 1 L sterile LB.
Ensure that this is done near a flame.Every 1 L LB will get split into two 500-ml conical tubes.Label each flask with a letter (e.g., A or B) and each 500-ml conical tube with the aliquot number (e.g., if from flask A: A1 or A2).
7. Using a scale, ensure that conical tubes are the same weight.Use a serological pipet to transfer volume from one conical tube to the other if required to balance out the weights.
This step is important to ensure that samples are balanced when spinning in the centrifuge.Only transfer volumes between conical tubes with contents that originate from the same flask.For example, transfer only from A1 to A2 and never from A1 to B2.
8. Spin conical tubes on 50-ml centrifuge tube cushions for 10 min at 3500 × g. 9. Pour 45 ml supernatant from the 500-ml conical tubes into 50-ml conical tubes, keeping track of the sample origins.
For example, 45 ml from the 500 ml conical labeled B2 would be poured into a 50 ml conical labeled B2.
10. Pour the remaining 450 ml supernatant into the corresponding culture flask from step 6.
This flask will ultimately be decontaminated using bleach.
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Current Protocols 11.Pour the 45 ml from step 9 back into the corresponding 500-ml conical tube containing the cell pellet.
By doing this, the bacteria are concentrated to 10×, as the original 500-ml culture is being brought down to 50 ml.
Do not dispose of the labeled 50-ml conical tubes, as they will be reused in step 13.
12. Use a serological pipet to resuspend the bacteria or shake the conical tubes by placement in the shaking incubator at 21°C and 180 rpm for ∼20 min.
13. Transfer the resuspended 50-ml culture into the corresponding 50-ml conical tube used in step 9.
14. Label this aliquot with "10× OP50", the date, and the initials of the person who prepared it.
Until the aliquots are confirmed to be free from contamination, we recommend not using them.
16. Decontaminate the 2-L flasks containing bacterial supernatants (see step 10) by adding 6% bleach to a final concentration of 1% for ≥30 min and then discard.20.Discard any contaminated aliquots and store the clean aliquots for ≤1 month at 4°C.

22.
Remove the supernatant until 12.5 ml remains and resuspend the cell pellet.
Approximately 37.5 ml supernatant should be removed.This pellet will be even more difficult to resuspend than the 10× pellet was in step 12.
Alternatively, by resuspending the cell pellet from step 9 in 12.5 ml supernatant, a 40× aliquot can be generated.However, this will be difficult to resuspend, and sample loss may occur as a small volume is placed in a large conical tube.We recommend that the steps listed in this section are followed instead.
25.For 10-cm plates, spread the culture using an ethanol-and flame-sterilized metal spreader.
Ensure that the flamed spreader cools down prior to spreading to prevent killing of the E. coli cells.
The E. coli on 6-cm NGM plates does not need to be spread.2. Sterilize a metal spatula by dipping it in 95% ethanol and carefully placing into the flame of a Bunsen burner.Once the spatula cools down, chunk 1 4 of the plate onto a new 6-cm seeded NGM plate.To do this, use the spatula to cut through the agar, separating the chunk from the rest of the plate.Then, push the spatula under the chunk, lifting it out of the plate.Finally, put the chunk over the recipient plate and invert the spatula so that the chunk is now face down on the new plate.Wash off the animals on the remaining 3 4 of the plate with 700 μl M9 + 0.1% Tween-20 into a microcentrifuge tube.CAUTION: Ethanol is flammable.
The 1 4 sector will serve as a source to continue to propagate both the nematodes and the microsporidia infection.As the agar will contain potential spores shed by the nematodes, it serves as a reservoir for both the infection and the population.Exercise caution when chunking to ensure that enough nematodes are being transferred.This is especially critical when the population is composed of males and females.
When washing animals off the plate, it is best to aim toward the remaining 3 4 of the plate.This ensures that nematodes do not end up in the liquid between the agar and the plate, maximizing nematode recovery.If lots of contamination is present on the plate, the plate may need to be washed multiple times to dislodge all the nematodes.
It is optional to keep the washed plate or discard it.If kept, seal it with parafilm and store upright with the expectation that this plate will very likely develop fungal mold.
3. Place the microcentrifuge tubes in a tube rack on the bench to promote gravity settling.Once a nematode pellet is clearly visible, remove the supernatant but avoid disrupting the pellet.
Avoid spinning the samples in a tabletop centrifuge at this point to prevent carrying over contamination.By gravity settling the animals, most of the bacterial and fungal contaminants will not settle as fast, allowing efficient removal.This can take up to 5 min.
4. Wash the samples with 700 μl M9 + 0.1% Tween-20, invert to mix, and gravity settle.Repeat this step as many times as necessary until the sample supernatant is clear.Then, add 700 μl acetone to the pellet.
Alternatively, after 10 min of incubation in acetone, store samples at −20°C to be processed later.
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There may be instances where even after multiple rounds of washing, some contamination persists.Proceed with the next steps.
At this point, the nematodes will die and may appear as large cloudy clumps under the dissecting microscope.If any residual bacterial or fungal material is present, it will also form clumps.Samples can be stored ≤1 year at −20°C in acetone.
5. Centrifuge the samples for 30 s at 6010 × g.

6.
Remove the supernatant and discard in an appropriate liquid waste container.
7. Add 500 μl DY96 solution, wrap the tubes in tinfoil, and rotate the samples for 30 min in the dark.
Samples can be attached to any rotator.This promotes sample mixing in the tube.
8. Centrifuge samples for 30 s at 6010 × g and discard supernatant.
9. Add 10 to 20 μl cold EverBrite Mounting Medium and pipet up and down gently to evenly mix.
EverBrite is a mounting medium containing DAPI.
This solution is viscous and requires gentle pipetting to ensure the correct volume is used.
10. Load 10 μl sample onto a glass microscope slide and place a coverslip on top.
The DY96-stained samples can be stored in a slide box for ≤2 years at 4°C in the dark.
11. Image the samples using a fluorescence microscope with GFP and DAPI filters.
Characterizing infected nematode samples 12. Screen samples using a 10× or 20× objective and the GFP channel to identify microsporidia.

Microsporidia will appear as many green rod-or bean-shaped spores. See Figures 4 and 5 for detailed images of expected results.
Although microsporidia can often be identified at a lower magnification, using a higher magnification via a 40× or 63× objective can help confirm the presence of microsporidia.DY96 stains chitin, which is present in the embryos of nematodes, and other fungal material.Therefore, the presence of fluorescence alone is not sufficient to identify microsporidia.
13. Identify the fraction of adults that contain spores by counting about 100 to 200 adults.Count adult worms that contain spores throughout the body and not just in the intestinal lumen (Figs. 4 and 5).

This indicates the prevalence of the infection and can help inform the size of the chunk required to continually propagate the population (e.g., more than 1 4 in step 2 if a small fraction of infected animals is present). Continual propagation of infected samples can be better maintained if the density of the infection on the plate is monitored. Typically, only adults or later-stage larvae have been infected long enough for the microsporidia to sporulate, so only these larger animals should be counted. Adult populations with >10%
of animals displaying microsporidia spores can usually be propagated to maintain the infection.If only a small percentage of the adult population displays spores, it may be difficult to propagate the infection.As DY96 only stains the spore stage, the absence of fluorescence signal does not mean that the worms are not infected with microsporidia.
Discard any uninfected samples, making note of this in the sample spreadsheet.The Caenorhabditis species C. elegans, C. briggsae, and C. tropicalis can be sent to CaeNDR, the Caenorhabditis Natural Diversity Resource (https:// caendr.org/ ) (Crombie et al., 2024).14.Characterize the spore morphology by determining the size of the spores present.
To do this, take Z-stack images of individual infected worms using a 40× or 63× objective.Determine the plane at the top of the worm where spores are in focus (this will be the starting height) and determine the plane at the bottom of the worm where there are no spores in focus (this is the ending height).Determine the exposure time that allows the brightest signal possible without any saturation at any of the Zplanes.After capturing the Z-stack, create a maximum intensity projection.Using this image, measure the length and width of at least 50 spores from at least two infected animals and determine the mean length and width.Only measure spores that are not touching other spores and that appear to be parallel to the plane of the image.
The preferred approach for obtaining microscopy images is one that limits background fluorescence, such as using a confocal microscope, an apotome, or deconvolution.
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Current Protocols For some species of microsporidia, there are distinct classes of spore size (Murareanu et al., 2021;Wadi et al., 2023).In this case, measure the values for these different classes individually (Fig. 5D).Spores from Nematocida species exhibit a range of sizes, from 1.3 to 2.4 μm in length and 0.5 to 1.0 μm in width.Spores from Pancytospora and Enteropsectra species can be longer, up to 4 μm in length.15.Characterize the infection properties by assessing the tissues infected and if the spores form large clusters.
Microsporidia infections in nematodes are often intestinal, but some species can infect other tissues.The intestine in acetone-fixed nematodes mounted on slides can often appear both medial and lateral, making it difficult to distinguish the intestine from the Tamim El Jarkass and Reinke

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Current Protocols muscle and epidermis.To circumvent this, we determine if spores are present either anterior or posterior to the digestive tract (Fig. 5B).If anterior, this is evidence that the species can proliferate in non-intestinal tissues.Take note if spores form large, distinct clusters (Fig. 5C).

These infection properties are useful for determining what genus or species of microsporidia is present.
16.If the infection is not very prevalent in the population, chunk a densely populated plate and propagate at different temperatures (e.g., 15°C and 25°C) to promote spread, followed by determination of whether any improvements have occurred (see steps 2 to 15).
Some species of microsporidia display very particular growth requirements.For example, Nematocida displodere cannot grow at ≥20°C (Luallen et al., 2016).

Determining if a microsporidia species can infect C. elegans
17. Chunk about 50 large animals from the infected plates generated in step 2 after about 5 to 6 days and add about 20 ERT54 L1 animals to a 10-cm seeded NGM plate.Incubate this plate at 21°C or room temperature.
The ERT54 strain carries an mCherry marker in the pharynx, allowing this strain of C. elegans to be differentiated from the unknown nematodes.The ERT54 strain also contains a promoter driving expression of GFP, which turns on during microsporidia infection.
18.After ∼1 day, determine what percentage of mCherry-positive animals also display GFP fluorescence using a fluorescent dissecting microscope with GFP and RFP filters.After 4 days, fix and stain with DY96 (as in steps 4 to 10) and determine what proportion of mCherry-positive adult animals have microsporidia spores in them.
The GFP reporter monitors initial infection, so it is possible that GFP is induced but no spores are seen in the animals.This suggests that the microsporidia species can invade C. elegans but cannot replicate within it.Alternatively, it is also possible for the GFP reporter to not turn on but for spores to be observed after 4 days.ERT54 can be infected with a C. elegans-infecting species such as Nematocida parisii as a positive control.Alternatively, for an example of how the GFP reporter should appear, see Reddy et al. (2017).Having a plate of ERT54 animals grown on E. coli OP50 can serve as a negative control for reporter fluorescence.
Freezing of samples 19.To freeze down animals that contain microsporidia infection, first chunk the infected plate onto a 10-cm seeded NGM plate as in step 2. Monitor the plate for a few days.
Once lots of L1 larvae with little E. coli OP50 are visible on the plate, wash the animals into four sterile cryotubes using 5 ml freezing medium.
A single populated 10-cm plate is washed into four individual cryotubes.Three are kept for long-term storage, and the last is used to ensure that viable nematodes can be recovered post-freezing.The L1 stage is most resilient to freezing, making almost-starved or freshly starved plates the most ideal.Washing the worms off the plates will also wash off any microbes.This is desirable, as microsporidia spores will likely be on the plates.
20.If C. elegans (ERT54) can be infected with microsporidia, then freeze these animals as well.Freeze down all nematode samples regardless of infection status if they will be sent to nematode repositories.Place the tubes in a Styrofoam tube rack and place at −80°C.
The Styrofoam ensures that the samples are frozen slowly, maximizing sample viability post-thaw.
21. Thaw the fourth vial ≥24 hr post-freezing to ensure nematodes are successfully recovered.
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Current Protocols Plating the entire volume on a 10-cm seeded NGM plate should result in the recovery of nematodes.This can be seen as animals thrashing in the liquid or via tracks in the food.After a few days, contaminating bacteria or fungi that were in the original sample will likely start to appear.

DNA EXTRACTION, PCR AMPLIFICATION, AND SEQUENCING TO IDENTIFY NEMATODE AND MICROSPORIDIA SPECIES
In this protocol, we discuss how to extract DNA from infected samples (Basic Protocol 2) and provide the PCR parameters and the primer sequences used for amplification.As the nematodes are infected, lysis and DNA extraction of these animals result in the release of host and pathogen DNA.PCR amplification involves the use of either Caenorhabditisspecific or nematode-specific PCR primers that amplify the 18S rRNA of the host.For microsporidia, there are multiple primer pairs that can be used, with amplification of the 18S rRNA being the most common.

Materials
Oligonucleotides (IDT; see There should be one tube per sample.If desired, a higher volume can be used, but be sure to scale up the number of animals being picked in step 3.
3. Pick about 20 large adult animals from an infected sample into the same PCR tube containing the lysis buffer using a sterile worm pick.
Include a tube without worms as a negative control.Uninfected C. elegans can be used as a positive control for amplifying nematode DNA, and if C. elegans infected with a known species of microsporidia is available, that can also be used as a positive control.Aim to pick large, slow-moving animals from infected samples.These animals likely harbor large amounts of infection and will provide enough DNA template for amplification.
Keep in mind that it is typically not possible to know for sure which animals harbor infection by visually looking at the animals, especially when the infection rates are low.In picking older animals, we assume that they have been around long enough to harbor microsporidia.In heavily infected populations, animals display decreased mobility, which can be an indicator of infection.
4. Place the strip tubes into a thermocycler and run the following worm lysis program: 65°C for 60 min 95°C for 15 min.
An optional hold at 4°C can be added if desired.For long-term storage, place at −20°C.The annealing temperature "X" will depend on the primer pair being used.Consult Table 1.The exact thermocycling conditions will depend on the DNA polymerase being used, and some optimization may be required.

Prepare a PCR master mix by multiplying the amounts of PCR components in
Assessing PCR amplicons via DNA gel electrophoresis 7. Prepare a 1% agarose gel.
This gel can be prepared up to 1 week in advance and stored at 4°C.
8. Load 10 μl DNA ladder onto gel.Mix 1 μl of 6× loading dye with 5 μl of each sample and then load the entire volume.Run the gel at 80 V for 50 min.
Now 45 μl should be remaining in the sample.
9. Image the gel using a gel imaging apparatus.Assess the success of the PCR reaction by the presence of bands of the correct size, as outlined in step 1. 11. Assess the quality and concentration of the purified amplicon using a spectrophotometer.
If the DNA quality or purity is low, poor sequencing quality may result, and the data therefore will not be able to be reliably interpreted.
12. Prepare and submit samples for Sanger sequencing by following the instructions of the facility that will perform the sequencing.
The primers used for PCR amplification can also be used for sequencing.There will be two aliquots of the same amplicon, one receiving the forward primer and one receiving the reverse primer.
13. Trim any regions of poor sequencing quality.Align the sequences obtained from the forward and reverse primers using software such as ApE (https:// www.frontiersin.org/ articles/ 10.3389/ fbinf.2022.818619/full) and fill in any sequencing gaps to generate a single sequence.
Remove regions that are not being defined as individual bases.Matches to microsporidia sequences with ≥99% identity are likely a strain of a previously identified species, and those with <99% are potentially a novel species (de Albuquerque & Haag, 2022).For nematodes, the highest percentage match to the 18S rDNA sequence can be used to determine the genus of the isolated species.Within Caenorhabditis, ITS2 sequences with less than two or three nucleotide differences can be used to determine the species (Barriere & Felix, 2014).For definitive identification of the nematode species, crosses can be performed using known species available from the CGC (https:// cgc.umn.edu/ ).

REMOVAL OF CONTAMINATING MICROBES AND PREPARATION OF MICROSPORIDIA SPORES
Once infected samples of interest are identified (Basic Protocol 3), they can be prioritized for the removal of contaminating bacteria using antibiotics.Although bleaching is an excellent way to destroy contaminating bacteria, bleach will also destroy the microsporidia present within the nematodes.In contrast, antibiotics are a selective way to remove contaminating bacteria.Once the samples are cleared of bacterial contamination, any residual fungal contamination can be dealt with by chunking the contaminated fraction of the plate away if the fungus is filamentous (e.g., mold).Once a clean population of nematodes containing the microsporidia of interest is obtained, these samples can then be used to generate a spore preparation.This preparation involves using the nematodes as a factory to produce large quantities of dormant microsporidia spores.These spores are then aliquoted and stored frozen for use in subsequent experiments.

NOTE:
It is crucial that proper sterile technique is utilized and that filter tips are used at every step in this protocol.Any contaminated plates must immediately be discarded to prevent spread of contamination.

Antibiotic cleanup
About 4 to 5 days before cleanup 1. Chunk the environmental sample for cleanup onto a 10-cm seeded NGM plate (see Basic Protocol 2, step 2) and grow until densely populated.
As an alternative, the environmental sample can be chunked onto a 10-cm seeded antibiotic plate to reduce the initial amount of bacterial contamination.
Cleanup day 2. Sterilize the benchtop and micropipets using 70% ethanol.Assess the contaminants present on the plate, making note of the properties and colors of bacterial colonies.
It is critical that no contamination is acquired during the cleanup and spore preparation process.Being aware of the contamination present prior to cleanup helps with assessing if any new contaminants may have arisen in the process or if certain contaminants are more difficult to remove.Some contaminating bacteria may be invisible or indistinguishable from E. coli OP50, making the identification of contaminants more challenging.
4. Place tubes in a bead disruptor and vortex at 3000 rpm for 5 min.
This step breaks open the worm bodies, releasing the microsporidia spores from the host.Note that the microsporidia spores are still viable at this stage.
5. Let the beads settle and transfer supernatant to a clean tube.Store the supernatant for ≥1 hr at −80°C.
This step ensures that there are no live worms, as removing contaminating bacteria from live animals is more difficult.The supernatant can be stored ≤1 month at −80°C and thawed at room temperature prior to use.
6. Place the entire supernatant on a 6-cm unseeded antibiotic NGM plate.Incubate the plate for 24 to 48 hr at 21°C.This gives the antibiotics some time to act on bacterial contaminants.Worm bodies will be visible, but all nematodes will be dead.We use a combination of antibiotics to target a wider diversity of bacterial species.This is critical, as we do not know the identity of the bacterial species and to which antibiotics they may be resistant.
7. Bleach-synchronize a population of animals and hatch overnight (see Support Protocol 2).
The nematode host of interest can be used, but bleach synchronization may require some optimization depending on the host species used.These animals will be the host in which the infection is propagated.If C. elegans can be infected by the microsporidia identified, we strongly recommend propagation in this host.
8. Add 400 μl of 10× E. coli OP50 and 1000 synchronized L1 animals to the plate from step 6.Dry the plate either on a bench or in a laminar flow cabinet.
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Current Protocols The synchronized animals will serve as the new host for infection propagation.The antibiotic cocktail will act on the bacterial contaminants as well as the E. coli OP50 food source.Therefore, monitor plates to ensure they do not run out of food.Additional OP50 can be added as necessary.
9. Parafilm and incubate the plate for ∼4 days at the appropriate temperature in a sealed plastic box.
This temperature will most likely be 21°C, but if an alternate temperature was deemed to result in higher infection rates, it should be used.We recommend waiting for ≥4 days before continuing to ensure that the infection has had adequate time to spread.
10. Chunk the animals onto a new 6-cm seeded antibiotic NGM plate (see Basic Protocol 2, step 2) and incubate for several days.
This step serves as a second round of exposure of the sample to antibiotics.
11. Chunk the animals onto a 10-cm seeded NGM plate (no antibiotics) (see Basic Protocol 2, step 2) and incubate for several days.Using the remaining animals on the plate, determine the percentage of the adult population containing microsporidia spores, as described in Basic Protocol 2, steps 12 and 13.
If low amounts of infection are observed, repeat the process, but increase the amount of time at step 9 to ≥7 days.For species with intestinal infections, this usually is not necessary, as the spores are continually defecated.For epidermal-and muscle-infecting species, the spores are thought to only be released upon the worms either dying or bursting open, so the additional time allows for this to happen and for the larvae to become infected.
12. Assess the degree of contamination present on the plate.If the sample is deemed to be clean, proceed to using it to generate a spore preparation.If not, use the plate generated in step 10 and repeat steps 2 to 11 until no bacterial contamination is detected.
The cleanup process may take several rounds before a sample is contaminant free.This treatment only targets bacterial contaminants, which are generally harder to clean up than fungal counterparts.In the case of fungal contamination, keeping a close eye on the plates daily will help with identifying the early stages of filamentous fungal growth.Areas of the plate far from the fungal contamination can then be chunked onto a new plate.
13. Freeze down several clean aliquots of the sample, as described in Basic Protocol 2, steps 19 to 21.

Spore preparation
Stage 1: Setting up source plates 14.Spray the bench with 70% ethanol to sterilize.
15. Thaw an aliquot of an infected population (see step 13) or chunk a densely populated contaminant-free plate (see step 12) evenly onto three 10-cm seeded NGM plates.
Ensure that only samples already cleaned of contaminating bacteria are used here, or else the spore preparation will contain a variety of other microbes.
It is best if all three 10-cm seeded NGM plates receive an equal volume or chunk size (1/3 sector each).
16. Dry the plates on a bench or in a laminar flow cabinet and incubate at 21°C until the population has crashed out.

This can take between 4 and 7 days. It is best to monitor these plates daily. The key is to have the infected population get sick enough to yield large amounts of infection but still generate large animals. A "crashed-out" population has many large but slow-moving
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Current Protocols animals, growing at a slower pace than uninfected animals.The extent of infection is enough that few additional progeny will be generated.Most of the intestinal-infecting isolates will generate this "crashed out" phenotype.The epidermal-and muscle-infecting species often do not cause the animals to appear sick, and the population will consume all the available food and starve.As described above, this is beneficial, as it allows for infected animals to die, releasing spores so that larvae can become infected.
The growth dynamics will depend on the nematode host being used and the culturing conditions.The recommended time frames provided here assume a C. elegans host grown at 21°C.
Stage 1, continued: 18 to 24 hr before using source plates 17.Bleach-synchronize a population of animals (see Support Protocol 2) to be used in step 19.Hatch overnight.
These plates do not have to be freshly poured and seeded but should not be older than 1 month.It is crucial that these plates are not too dry and the agar is not peeling off the edges.This would interfere with downstream steps.
Stage 2: Expanding the infected population 19.The next day, add 2640 L1 animals from step 17 to 6600 μl of 40× E. coli OP50 for every 24 harvesting plates.
The 40× E. coli OP50 should not be older than 1 month and must be verified to be contaminant free prior to use.One master tube containing the required number of L1 animals and the total volume of E. coli for all plates can be generated.Remember to overestimate the required volume to account for pipetting errors.For non-intestinal-infecting species that do not cause severe fitness defects, L1 animals do not need to be added in this step, as the progeny from the infected animals are sufficient to produce spores.
20. Pipet 250 μl of the nematode-E.coli mixture onto each of the 72 plates from step 18.
This will provide each plate with plentiful food and about 100 L1 worms.The L1 animals being added, in addition to the progeny of the infected animals, will expand the microsporidia population.
21. Chunk each of the three source plates into 24 sectors (pie slices) using a metal spatula dipped in ethanol and flamed as in Basic Protocol 2, step 2. Place 1/24 of a sector upside down on top of the nematode-E.coli mix.
It may be a little tricky to get evenly sized sectors, but if there is a source of worms on each slice, this should be sufficient.The additional L1 animals added in step 19 will help propagate the infection.
22. Store the plates right side up in plastic boxes with a lid.Incubate at 21°C for about 4 to 7 days.
23. Screen the plates every 2 days for contamination, discarding any plates that display dark bacterial colonies or fungal mold.

24.
Monitor the population density by looking at the number and size of animals on the plates.
Harvesting plates should ideally contain large animals that have consumed almost all the food on the plate.This point is usually achieved by day 6 but can vary.Keeping a close eye is important to avoid starving the population and missing the harvesting point.
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Current Protocols Stage 3: Harvesting animals 25.Pool the animals from about 5 to 6 plates together into a single sterile 15-ml conical tube by washing animals off plates using 2 to 3 ml sterile M9 per plate.
Try to remove as many worms as possible from the plates to maximize spore yield.If many animals remain on the plates, additional washes can be performed.It is expected that these washes will contain bacteria, which will be removed in the following steps.

27.
Remove the supernatant and wash the worms with 10 ml sterile M9.
The objective is to remove all traces of bacteria from the supernatant.
28. Repeat steps 26 and 27 until the supernatants are clear.
This can take anywhere from 3 to 6 washes.
29. Centrifuge tubes for 30 s at 7197 × g and remove the supernatant down to 1 ml.

30.
Transfer the 1 ml into a sterile 2-ml microcentrifuge tube.Transfer 50 μl into a separate microcentrifuge tube for contamination checks (see steps 32 and 33).
There will be two microcentrifuge tubes per pooled set.
This step is not optional.The samples need to be checked for contamination (see steps 32 and 33) prior to proceeding with the rest of the protocol.This step ensures that worms are non-viable prior to thawing the sample for contamination testing.
Incubate each dilution at 21°C and 25°C for 5 days.
The dilution allows for the identification of contaminants present in low quantities.Incubation at two different temperatures allows for the identification of colonies that grow only at one or the other.
33. Assess the plates under a dissecting microscope with transmitted light, looking for any colonies that do not look like OP50.
E. coli OP50 is pale yellow and is almost white.Anything that looks different from this is considered contamination.If contaminants are present, the corresponding 1-ml aliquot in the freezer (see step 31) should be discarded and should not be used in downstream steps.
Stage 5: Break up worms and filter lysate 34.Thaw uncontaminated 1-ml aliquots from step 31 and add ∼500 μl sterile 2.0-mm zirconia beads.Vortex in a bead disruptor at 3000 rpm for 5 min.
This step has the identical purpose as step 4, bursting open worm bodies to release dormant spores.Unlike in step 4, we are not expecting to release any bacterial contamination from the worm bodies.

35.
Transfer the supernatants into a sterile 50-ml conical tube.Wash the beads twice more with 1 ml sterile M9 and add supernatants to the same tube.
At this point, all the spores will be pooled into a single conical tube.

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The syringe will need to be replaced every 2 to 3 ml as it will become clogged with worm carcasses.This can be felt as resistance when attempting to pass more volume through the filter.Replace the syringe as necessary.
37. Aliquot the filtered lysate into desired volumes in microcentrifuge tubes and store at −80°C.Reserve some unfrozen lysate for step 38.
We recommend 50-, 100-, and 200-μl aliquots.Repeatedly pipet the spore mixture up and down prior to aliquoting to ensure an equal resuspension of spores in solution.
Stage 6: Measure spore concentration 38.Determine the concentration of spores by combining 10 μl spore prep lysate (see step 37) and 90 μl spore-counting solution in a new microcentrifuge tube.Mix by pipetting to ensure spores are in solution.Add 4 μl stained spores onto a CELL-VU TM slide by putting the drop on the inner edge of the circle.Place the coverslip so that the edge just covers the drop of spores and then carefully slide the coverslip so that the circle is fully covered.
See the manufacturer's instructions for further guidance on using CELL-VU TM slides appropriately.
39. Using a fluorescence microscope with a 20× objective, count the total number of green spores in 10 grids.Divide the number of spores counted by 200 to determine the concentration of the spore preparation lysate in millions of spores per μl.
The concentration of spores can be helpful in determining in what volumes to aliquot spores but does not always inform on the infectivity of the spore lot.The concentration of spores typically falls between 0.05 and 0.5 million spores per μl.
Stage 7: Quality control for contamination 40.Thaw one aliquot of the spore prep from step 37 to test for contamination as described in steps 32 and 33.

BLEACH-SYNCHRONIZING NEMATODES
This protocol describes the use of a sodium hypochlorite and sodium hydroxide mixture (bleach solution) to isolate embryos from adult nematodes.In doing so, the population becomes synchronized as the embryos hatch into M9 in the absence of food.Thus, all nematodes are unable to continue development until placed on a food source.

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Remember to prepare double the volume of bleach solution per sample and to account for pipetting errors (e.g., two plates to bleach means 4 ml bleach solution should be prepared, plus an extra 1 ml should be prepared to account for pipetting error).Always prepare this solution fresh.
2. Wash densely populated but unstarved nematodes off plate and into a sterile 1.7-ml microcentrifuge tube with 1 ml sterile M9.
It is critical that the plate being used for bleach synchronization has many large adult animals carrying embryos.Note that depending on the nematode species used, some may yield fewer embryos (e.g., C. briggsae do not hold their embryos internally as long as C. elegans).
3. Allow nematodes to gravity settle, generating a worm pellet.
This can take anywhere from 2 to 5 min.A centrifuge could be used to speed up the process.The advantage of gravity settling is that any residual bacteria do not settle as fast as the nematodes.
This will also remove the residual bacteria that have not settled.
5. Add 1 ml bleach solution from step 1 to the nematode pellet and set a timer for 1 min.
Invert the tube a few times to ensure mixing.
Carefully monitor the nematodes in the microcentrifuge tube.With time, they will become less mobile, and thrashing activity will decrease.

7.
Remove supernatant and add another 1 ml bleach solution.
8. Observe the samples under a dissecting microscope with transmitted light, looking for populations where ∼80% of the carcasses have dissolved and released embryos into the environment.
This can take anywhere from 1 to 5 min.Keeping a close eye is encouraged.Avoid waiting too long, until when all carcasses have dissolved and embryo clumps have begun to form; this could indicate that the sample is overbleached and may not yield sufficient L1 animals.
This step should be done quickly to ensure that the residual bleach solution is diluted out.This ensures that released embryos are not exposed to bleach longer than necessary.
11. Repeat steps 9 and 10 two more times.
A total of three wash steps in M9 should be performed.
12. Add 1 ml sterile M9 to the worm pellet and transfer to a sterile 15-ml conical tube containing 4 ml sterile M9.
13. Place tube on a rotisserie rotator overnight at 21°C.
Embryos begin to hatch overnight but cannot begin development, as there is no food present in the conical tube.L1 animals should be used within 20 hr of bleach synchronization to prevent starvation.
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Current Protocols 14. Observe the tube under the dissecting microscope.
L1 animals should appear to be thrashing.
If large clumps of embryos appear, this indicates that the sample was overbleached.
15. Spin down the tube for 30 s at 1400 × g.

16.
Concentrate the worm pellet down to 1 ml.
The L1 pellet will be soft and is easily perturbed.We do not recommend pouring out the supernatant.Instead, use a serological pipet.
17. Resuspend the pellet and estimate the concentration of nematodes present by aliquoting 5 μl onto a 6-cm seeded NGM plate and counting the number of thrashing L1s.
The nematodes will gravity settle, so always invert the tube 2 to 3 times prior to taking an aliquot.We recommend taking the average of three 5-μl aliquots for increased accuracy.

REAGENTS AND SOLUTIONS
Agarose, 1% 0.5 g ultra-pure agarose (Invitrogen, LS16500500) 50 ml 1× TAE (diluted from 50× TAE; see recipe) Microwave for 1 min Add 2.5 μl 10,000× SYBR® Safe DNA Gel Stain (Invitrogen, S33102) Pour mixture into gel cast with comb and let solidify Wrap in plastic wrap followed by tinfoil Store ≤1 week at 4°C Volumes should be adjusted to the size and dimensions of the gel-casting equipment used.

Antibiotic NGM plates
50 mg/ml carbenicillin 25 mg/ml kanamycin 12.5 mg/ml tetracycline 37.5 mg/ml chloramphenicol (resuspended in 95% ethanol) 100 mg/ml cefotaxime 50 mg/ml gentamycin For generating unseeded plates, create antibiotic mixture by adding 12.5 μl of each antibiotic solution except for cefotaxime and gentamycin, for which 25 μl should be added, to 300 μl M9 (see recipe) For generating seeded plates, centrifuge 1 ml 10× E. coli OP50 (see Support Protocol 1) for 1 min at 7197 × g, remove all but 300 μl of supernatant, resuspend pellet, and add bacteria to 100 μl antibiotic mixture Add the 400 μl solution to the top of an 6-cm unseeded NGM plate and gently rock plate to spread liquid evenly Dry in laminar flow cabinet or on bench Store ≤1 week at 4°C

Bleach solution
125 μl sodium hypochlorite (6% bleach; Fisher, SS290-1; store at 4°C in the dark) 200 μl 1 N NaOH (Fisher, SS266-1; store at 4°C) 675 μl MilliQ H 2 O Prepare fresh immediately before use, store for maximum of 1 hr at room temperature Each sample will require 2 ml bleach solution, so scale up as required.

Background Information
Microsporidia are a major threat to many commercially and ecologically important animals (Stentiford et al., 2013(Stentiford et al., , 2016)).They are prevalent in the environment and can cause disease in the immunocompromised (Han & Weiss, 2017;Ruan et al., 2021).Microsporidia are currently known to infect about 1500 animal species, though this is likely a large underestimation of the true diversity of microsporidia (Larsen et al., 2017;Murareanu et al., 2021).Despite their widespread nature, many aspects of their infection biology remain enigmatic.Given their extreme host dependance, studying microsporidia within a whole-animal model is advantageous and can reveal host-specific aspects of infection.Approximately 15 years ago, microsporidia were found to naturally infect C. elegans (Tecle & Troemel, 2022;Troemel et al., 2008).The discovery of the nematode-infecting species N. parisii provided an excellent opportunity to study this genetically intractable parasite in a genetically tractable host.Additionally, studying microsporidia infection in nematodes offers several advantages.First, nematodes can be readily sampled from the environment and screened for the presence of microsporidia.In fact, such efforts have already identified at least 15 species of nematode-infecting microsporidia, many of which can also infect C. elegans (Luallen et al., 2016;Reinke et al., 2017;Wadi et al., 2023;Zhang et al., 2016).Second, many species of nematodes can be easily cultured in the laboratory under the same conditions.Third, many nematodes have short generation times and can be cultured at high densities, allowing for infection of large populations of animals.Lastly, and particularly for C. elegans, the many gene editing and manipulation tools available can circumvent the lack of those in microsporidia (Reinke & Troemel, 2015).As multiple species of microsporidia infect C. elegans, this has become a powerful system for understanding evolution of transcriptional responses and host resistance mechanisms and for identifying microsporidia inhibitors with broad specificity (Mok et al., 2023;Murareanu et al., 2022;Wan et al., 2022).
There are thousands of known free-living terrestrial species of nematodes, though this is likely an underestimation (Schratzberger et al., 2019;Smythe et al., 2019).Many species of terrestrial free-living nematodes are often found in microbially rich environments, including decomposing fruits and stems, as discussed in Basic Protocol 1 (Barriere & Felix, 2014).The geographical location of sampling will dictate the nematode species isolated, as will the time of year.For example, C. elegans are found in temperate regions primarily in the fall (Frézal & Félix, 2015;Schulenburg & Félix, 2017).From our own sampling efforts in southern Ontario, C. elegans does not appear to be present, but we have had success identifying other Caenorhabditis species, as well as those from the genera Oscheius, Pristionchus, and Panagrellus.Microsporidia can infect many different animal tissues, but the intestine or muscle/epidermis is the primary site of infection in nematodes (Murareanu et al., 2021).
Species of the genera Nematocida, Enterospectra, and Pancytospora have been found to infect various Caenorhabditis and related nematodes.The spores of many Nematocida species have been characterized and can be divided into two main groups.Those that form two classes of spore sizes are characteristic of the Parisii group, whereas those forming a single spore size and large spore clusters are more likely to be part of the Displodere group.Species in the Parisii group have so far only been found to infect the intestine, whereas some species in the Displodere group have been observed to infect the muscle and epidermis.There are currently respectively seven Tamim El Jarkass and Reinke

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Current Protocols and eight members of the two groups (Wadi et al., 2023;Zhang et al., 2016).Nematocida spores range from 1.3 to 2.4 μm in length and 0.5 to 1.0 μm in width.Pancytospora species are typically longer, up to 4 μm in length (Zhang et al., 2016).Therefore, characterizing spore size and tissue tropism can provide insight into the identity of the species.
Performing infection characterization in a genetically tractable host is desirable, and many species infect C. elegans in the lab, even if they were originally isolated from a different nematode species.We provide a strategy to assess if C. elegans is susceptible to infections from isolated microsporidia species by utilizing a transgenic strain carrying an infection reporter, as described in Basic Protocol 2. An immune response termed the intracellular pathogen response (IPR) is upregulated in C. elegans upon microsporidia infection.One of the most highly upregulated IPR genes is pals-5.The use of the Ppals5::GFP reporter, coupled with DY96 staining, can be used to determine if a species can infect C. elegans (Willis et al., 2022).This enables the potential study of novel microsporidia species within a genetically tractable host.
Although several primer pairs exist for amplifying molecular barcode genes in microsporidia, amplification of nematodeinfecting microsporidia is not always successful (Zhang et al., 2016).Although the small subunit 18S rRNA is the most commonly used phylogenetic marker in microsporidia, several other options exist (Bojko et al., 2022).Additionally, as genomic sequencing technology such as Nanopore DNA sequencing becomes cheaper and more available, multigene species identification that is independent of PCR will become possible (Taxt et al., 2020).The molecular characterization of nematode-infecting microsporidia is explored thoroughly in Basic Protocol 3.
The generation of a large quantity of dormant spores can enable multiple different areas of investigation and is explored in Basic Protocol 4. These spores can serve as a template for whole-genome sequencing, allowing for genomic comparison between multiple different species and assessment of microsporidian evolution.They could be used to characterize infection dynamics compared to other nematode-infecting species as well as the host response to infection (Bakowski et al., 2014;Wan et al., 2022).Lastly, as microsporidia demonstrate great host specificity, the availability of multiple different species of microsporidia that infect related nematodes can help uncover the host-specific cues that drive host specificity (Wadi et al., 2023;Willis & Reinke, 2022;Zhang et al., 2016).Given the number of species of nematode-infecting microsporidia identified to date, we have likely just scratched the surface (see Table S2 in the Supporting Information for properties of reported nematode-infecting microsporidia species).Expanding the known diversity of these species will allow us to gain insight into novel aspects of microsporidian evolution and infection mechanisms.

Basic Protocol 1
The sampling process may not yield any nematodes.The key parameters are the time of year, temperature, location, type of substrate, and level of substrate decay.In our own experience, from sampling in and around Toronto, Canada, up to ∼50% of samples collected between September and November contain nematodes.Once nematodes are identified, it is expected that the population will continue to proliferate.However, some nematode populations may fail to do so.It is important to consider the temperature and conditions from which the sample was obtained and assess if these differ drastically from the laboratory environment.It is worth considering if the nematodes are a male-female species or hermaphrodites.In the unfortunate event that the population obtained from the substrate was exclusively male or female, the population will likely not be able to be propagated.Careful observation and characterization of the population should be done as soon as nematodes are detected.
To increase the chance of detecting microsporidia in a population, allow the nematodes to grow for at least 5 to 6 days after substrate removal.As DY96 dye stains only spores, the microsporidia need to reach this stage to be observed.Although the time to sporulation will depend on the species, we have observed that most species will form spores within 96 hr after infection.Harvesting samples too soon may result in missing potential microsporidia infection.Additionally, as we propagate samples by chunking the plates, there is a higher chance that dormant spores are also present on the agar if animals had sufficient time to grow and sporulate.This is especially critical if an epidermal infection is present, where spores are shed only when old animals rupture and release new spores.This is different from intestinal-infecting species, Tamim El Jarkass and Reinke

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Once the sample is fixed, the nematodes and microsporidia are not viable and cannot be recovered.

Support Protocol 1
Working in a sterile environment to avoid contaminating the E. coli OP50 stock is critical.This also applies when pouring and seeding NGM plates.Avoid using seeded NGM plates that are older than 1 month.Always inspect plates to ensure that the agar is not peeling off the sides.

Basic Protocol 2
When screening DY96-stained animals for microsporidia, be aware that many things may fluoresce in the green channel.Use the flowchart in Figure 4A as a guide.When assessing the fraction of infected animals, do so in at least 100 animals and make sure that only the adult animals are being quantified.Younger animals will not have been around for ≥72 hr, meaning that microsporidia will not have had sufficient time to sporulate.Thus, they will appear uninfected but may harbor the intracellular stages that cannot be stained with DY96.In contrast, smaller animals full of spores may indicate heavily infected animals that succumbed to the infection, as microsporidia infection can impact host fitness in a dose-dependent manner (Willis et al., 2021).If only a small fraction of worms are infected, consider how to modify the propagation approach to ensure that the population is not bottlenecked when chunking.If only 10% of the population is infected, consider leaving animals on the plates for longer than 5 to 6 days and adding additional E. coli to the plates instead.Alternatively, chunk half the plate to propagate the population, rather than a quarter, and continually supplement the plate with additional E. coli.If the infectivity is still low, consider incubating the plates at different temperatures.One other possibility for low infectivity is that the infection is not in the native nematode host.It is possible that this microsporidia species propagates better in other nematode species.If the species can infect C. elegans, this is a convenient host in which to grow and characterize the microsporidia species.

Basic Protocol 3
When generating worm lysates, it is important to pick about 15 large, old animals into the lysis buffer.This increases the chances of picking infected animals, as they will have been around the longest.If animals are stuck on the sides of the PCR tube, this will not result in successful DNA extraction.If a sample was found to be only mildly infected, this may influence the ability to amplify microsporidiaspecific sequences.

Basic Protocol 4
When performing antibiotic cleanup, it is critical that no additional contamination is acquired.Having a clean sample will ensure that during the spore preparation, only pure microsporidia spores are isolated.Keeping a sterile work environment and using proper sterile technique should help limit this.When harvesting animals for spore preparation, it is critical that plates are densely populated with large animals.This ensures maximum spore yield.

Support Protocol 2
The bleach solution should always be prepared fresh.Densely populated plates with large, unstarved animals that have many embryos should be used.

Troubleshooting
Commonly encountered issues and their potential causes and solutions are listed in Table 3.

Understanding Results
When collecting substrates from the environment (Basic Protocol 1), those containing nematodes will be easy to distinguish, as nematodes will proliferate on the NGM agar and populate the plate.Nematode tracks in the E. coli OP50 lawn or embryos on the plate can also indicate the presence of nematodes.To monitor how long nematodes have been on plates, labeling each with the date is strongly encouraged.In parallel to this, keeping a log of sample history is very useful and can help avoid confusion when dealing with multiple samples.Table S1 in the Supporting Information provides an example of the type of information to collect on the samples.
The most critical part of this set of protocols is correctly identifying microsporidia infection in nematodes (Basic Protocol 2). Figure 4A provides a flowchart to help assess if any of the samples contain microsporidia.This flowchart assumes that the previous steps (sample propagation) have been performed as outlined.We also provide examples of non-microsporidia fungal material that is frequently observed in environmental samples.We often see yeast cells that are circular in structure and accumulate in the intestinal lumen (Fig. 4B).Fungal hyphae that contain cell walls or septa between segments can also be seen growing inside of animals (Fig. 4C).This is not seen in microsporidia.We also observe GFP material accumulating in the intestinal lumen, with no defined structure or shape, that looks like speckles (Fig. 4D).The identity of this material is unknown but may represent digested fungal material in the intestinal lumen.Fungal spores that contain septa can also be seen inside the nematode intestine.However, these are quite large and are easily visible even at 10× magnification (Fig. 4E).Large amounts of chitinous structures that outline the intestinal lumen are not microsporidia.Microsporidia spores will often be present both in the intestinal lumen and within the cells of the nematode.Epidermal-infecting species must first be ingested but should not be seen exclusively within the intestinal lumen and should also be visible in the target tissue.In the rare event that microsporidia spores are seen exclusively in the intestinal lumen, this indicates that this species may not be able to infect the current nematode host.
Figure 5 provides examples of what microsporidia look like and how to identify if parasite cells display spore clusters (Fig. 5C) and assess if there are two classes of spore sizes (Fig. 5D).We also provide examples of how microsporidia spores may appear (Figs.5E and 5F).Some spores look like rice grains (Fig. 5E) or have a short and stubby morphology (Fig. 5F).This is not an exhaustive list of what may be seen but should serve as a good starting point to help understand what to look for.Microsporidia morphology is generally not visible at 10× or 20× magnification and appears as a cluster of fluorescent material that is not restricted to the intestinal lumen.The example provided in Figure 5A depicts an intestinal-infecting species.Keep in mind that there may be microsporidia species that grow in the muscle or epidermis.These can be detected by assessing if spores can be observed anterior to the pharynx of the nematode (Fig. 5B).
PCR amplification of microsporidia SSU RNA (Basic Protocol 3) has had variable success in our experience.The presence of a clear band at ∼1300 bp with V1F and any of its accompanying reverse primers is a good indication of success.However, the lack of a PCR amplicon coupled with the lack of a positive control makes it difficult to troubleshoot this part of the protocol.Using different primer pairs can be useful in ensuring that at least one of them amplifies a microsporidia sequence.The primer pair yielding the strongest signal on a DNA gel with the least amount of nonspecific amplification is prioritized for Sanger sequencing.It is not uncommon to have difficulty amplifying microsporidian SSU RNA.Part of the problem lies in the large amount of diversity present in microsporidian genomes, making the design of a completely universal primer challenging.Whole-genome sequencing can be considered for species identification if amplification proves unsuccessful.
For antibiotic cleanup of samples (Basic Protocol 4), the lack of visible foreign microbial growth on the plate will indicate success.These samples can now be prioritized for spore preparations.A successful Tamim El Jarkass and Reinke

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Time Considerations
The entire set of protocols can be completed in about 2.5 to 4 months.Initial sample collection and screening (Basic Protocols 1 and 2) take ∼3 weeks.Molecular characterization of nematode and microsporidia species (Basic Protocol 3) can take up to 1 month if troubleshooting of PCR parameters is required.The most time-consuming part will be the antibiotic cleanup (anywhere from 2 weeks to 1.5 months) and the spore preparation (∼1 month) in Basic Protocol 4.

Figure 2
Figure 2 Overview of the sample collection process.(A) Substrates are identified in the environment.(B) Substrates are then placed on NGM plates.(C) A total of 18 to 48 hr later, plates are scanned for the presence of nematodes, embryos, or tracks.Bacterial or fungal contamination of the plate will also be noticeable.

Figure 3
Figure 3 Mite identification and disposal.(A) Microscopy images of mites found in environmental samples.(B) Setup involved in discarding mite-infested samples.

Figure 4
Figure 4 Microsporidia identification pipeline and common contaminants.(A) Flowchart to help determine if microsporidia are present within a sample.(B-E) Images of various environmental samples captured at 10×, and a 40× magnification of the designated area (blue box) within each image.Common fungal contaminants include circular yeast cells (B), fungal filaments (C), and fungal spores with septa (E).GFP material that looks like speckles may also be observed (D)."Septum" indicates the cell wall within fungal cells, dashed lines outline worms, and solid lines outline the intestinal lumen.All samples are stained with DY96, labeling chitin.

Figure 5
Figure 5 Microsporidia infection in nematodes from various environmental samples.(A) Nematode containing microsporidia.The dashed line outlines nematodes imaged at 10×, and a 40× magnification of the designated area within the image is indicated in the blue box.(B) Epidermalinfecting species can be identified by the presence of spores in the tissues neighboring the pharynx.The anterior (ant.) and posterior/terminal (post.)bulbs of the pharynx are depicted.The grinder is indicated and is housed within the posterior bulb.Spore clusters (C) and various spore sizes (small and large) (D) are depicted.(E and F) Various microsporidia morphologies are shown.All samples are stained with DY96, labeling chitin.
If the PCR was successful and only bands of the correct size are obtained, proceed with PCR purification in step 10.If bands of the correct size are seen in combination with other sizes in abundant quantities, proceed with gel extraction in step 10. 10.Purify the PCR reaction using the Monarch PCR and DNA Cleanup Kit or perform a gel extraction by running the entire remaining volume (45 μl) on a 1% agarose gel as before (see steps 7 to 9).Excise the band of the correct size using a razor blade or scalpel and image with a blue light box following the Monarch DNA Gel Extraction Kit instructions.Limit exposure of the amplified DNA to ultraviolet light to avoid introducing mutations into the DNA.

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. Screen candidate plates daily under a dissecting microscope with transmitted light and identify those with lots of large adult animals that have mostly consumed the E. coli OP50.

Table 1
Acquire oligonucleotides listed in Table1for PCR amplification.For nematodes, start with the Nematode 18S primers, and if the species is within Caenorhabditis, use the Caenorhabditis 18S to determine the species identity.For microsporidia, start with V1F paired with either 18SR1492r or 1492r.If these pairs do not provide sufficient amplification, try the other primer pairs.2. Pipet 10 μl single-worm lysis buffer into a sterile PCR strip tube for each sample.

Table 1
PCR Primers for Amplifying Nematode and Microsporidia Sequences a Indicates experimentally determined optimal annealing temperatures.All other annealing temperatures are recommended based on primer melting temperature (Tm) via the NEB Tm calculator and Phusion (https:// tmcalculator.neb.com/#!/main).bTheforward primer V1F can be used in combination with any of the subsequent reverse primers in this table.NEM_R1 was designed to potentially amplify additional Nematocida species not captured by the previous two primers.VPE_R1 was designed to potentially amplify species from the genera Vittaforma, Pancytospora, and Enterospectra.
Table 2 by the number of samples + 1. 6. Place the PCR reactions in a thermocycler using the following parameters:

Table 2
Amounts of Each Component for a 50-μl PCR Reaction These volumes are per the NEB Phusion kit instructions.Other DNA polymerases can be used, but the exact volume of each component may need to be modified per the manufacturer's instructions.Scale up the volumes according to the number of reactions being set up. a

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UseBLASTn (https:// blast.ncbi.nlm.nih.gov/Blast.cgi)toreveal the closest known sequence match.Keep track of this information in the spreadsheet generated in Basic Protocol 1.

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Always ensure that LB is sterile before use (sterile LB is clear and not cloudy).

Table 3
Troubleshooting Guide for Sample Screening and AmplificationIt is best to discard the plates rather than trying to save the samples.Microwave samples and dispose of in appropriate biosafety waste.This is most likely to happen during the first step, which is gravity settling.Ensure that when removing the supernatant, that the pellet is not disrupted.If the sample has been disrupted, place the sample back into the tube and let it settle again.Not enough sample on the slideLoad more sample on a separate slide Thawed vials of a once-infected sample are no longer infected Sample was not heavily infected to begin with and may have been bottleneckedTry to recover the other frozen aliquots.If the issue persists, try freezing another set if the plate is still being propagated.

Table 3
Troubleshooting Guide for Sample Screening and Amplification, continued Only wash nematodes off plates when there are many large, sick animals.Washing before this point may result in animals that contain intracellular stages that have not yet sporulated, resulting in low yield.Avoid exposing animals to the bleach solution for too long.Aim for ∼20% visible animal carcasses before stopping the reaction.Nematode species being bleached may not carry as many embryos internally, resulting in low yield.Some species that do not lay embryos, such as Panagrellus redivivus, are also difficult to bleach.