Thank you to Dr. Marie-Anne F&#233;lix for providing us with wild nematode strains. This work was supported by NSF under CAREER Grant 2143718 and California State University under a CSUPERB New Investigator Award to RJL, NIH under R01 AG052622 and R01 GM114139 to ERT, and by an American Heart Association Fellowship to VL.

NOTE: Nematodes may be fixed with either paraformaldehyde solution (PFA) or acetone. PFA allows for better visualization of morphology than acetone and can preserve signals from transgenic green fluorescent protein (GFP), which is destroyed by acetone. However, acetone fixation is necessary to permeabilize microsporidian spores to enable labeling this life stage. Furthermore, acetone can be more convenient than PFA because it is less toxic, and samples can be stored for several days in acetone in a -20 &#176;C freezer without the need for removing the fixative. Below are two separate protocols, using either PFA solution or acetone as a fixative. For a visualization of the protocol steps, see Figure 1.
1. FISH staining with PFA fixation
<ol>
	<li>Prepare nematodes with associated microbes
	<ol>
		<li>Grow nematodes with the desired microbe of interest on standard Nematode Growth Media (NGM) plates seeded with the appropriate food source. Incubate the nematodes at 20 &#176;C until the desired life stage is reached.</li>
		<li>Add 2 mL of M9 minimal salts media (42 mM Na2HPO4, 22 mM KH2PO4, 8.6 mM NaCl, 19 mM NH4Cl) + 0.1% Tween 20 to NGM plates containing the Caenorhabditis strain infected or colonized with the desired microbe to be visualized. 
		&#8203;NOTE: The addition of detergent is to prevent nematodes from sticking to pipettes and microfuge tubes, and to help pellet L1-L2 stage nematodes. 0.1% Triton-X can be used in place of Tween 20.</li>
		<li>Pipette up the nematodes from the plates using a glass Pasteur pipette and bulb, and transfer them to marked 1.5 mL microfuge tubes. 
		NOTE: Glass pipettes are preferred because nematodes can stick to plastic pipettes, but the addition of detergent (Tween 20 or Triton-X) can minimize this issue.</li>
		<li>Using a microcentrifuge, spin down the samples containing the colonized or infected nematodes at 2,000 x g for 60 s for L1 animals, or 500 x g for 60 s for L4 or adult animals. All subsequent centrifugation steps will be performed at the selected speed.</li>
		<li>Remove the supernatant from the microfuge tubes using a pipette. Avoid disturbing the nematode pellet by carefully removing the supernatant down to 100 &#181;L above the pellet. This can be estimated using marked 1.5 mL microfuge tubes.</li>
	</ol>
	</li>
	<li>Wash nematodes to eliminate external contamination
	<ol>
		<li>Add 1 mL of 1x PBS (137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4) + 0.1% Tween 20 (PBS-T) to microfuge tubes.</li>
		<li>Spin down the samples in a microcentrifuge at the appropriate speed (see step 1.1.4). Using a pipette, remove all but 100 &#181;L of the supernatant. This can be estimated using marked 1.5 mL microfuge tubes.</li>
		<li>Repeat the above two steps 2-3x. 
		NOTE: Three total washes are typically sufficient; however, additional washes can be performed to remove any excess external contamination.</li>
	</ol>
	</li>
	<li>Fix nematodes with PFA
	<ol>
		<li>In the fume hood, add 33 &#181;L of 16% PFA to the microfuge tube containing 100 &#181;L of supernatant above the nematode pellet obtained from step 1.2.3 for a final concentration of 4% PFA. 
		CAUTION: PFA is a carcinogen. Contact with PFA may result in skin sensitization and irritation and eye damage. PFA releases toxic fumes that may lead to respiratory irritation or sensitization. When using PFA, work in a fume hood with proper personal protective equipment and refer to appropriate safety data sheets before use.</li>
		<li>Incubate the samples containing the nematodes colonized or infected with the microbe of interest for 30-45 min at room temperature. Following incubation, store the samples in 70% ethanol at 4 &#176;C until the protocol is ready to be continued. 
		NOTE: A shorter incubation period is better for maintaining the GFP signal in transgenic strains due to the degradation of GFP by PFA over time. Longer incubation times allow the fixative to better permeabilize into the samples. It is best to determine the incubation time empirically depending on the sample.</li>
	</ol>
	</li>
	<li>Remove the PFA solution
	<ol>
		<li>Spin down the samples in a microcentrifuge at the appropriate speed (see step 1.1.4). With a pipette, remove as much of the supernatant as possible without disturbing the pellet. 
		CAUTION: The supernatant contains PFA, which is toxic. Discard the supernatant and at least the first two washes as toxic waste in a fume hood.</li>
		<li>Add 0.5 mL of PBS-T to the samples in the microfuge tubes.</li>
		<li>Follow and repeat steps 1.4.1 and 1.4.2 2-3x with PBS-T. 
		NOTE: Performing more washes will help reduce the background signal. At least four washes in total are recommended.</li>
		<li>Following the last wash, spin down the samples and remove the supernatant, leaving the pellet undisturbed.</li>
	</ol>
	</li>
	<li>Prepare hybridization buffer (HB) and wash the nematodes
	<ol>
		<li>Prepare 1 mL of HB (900 mM NaCl, 20 mM Tris pH 7.5, 0.01% SDS) per sample. 
		NOTE: HB should be prepared fresh before each use to avoid precipitation. However, a general buffer (900 mM NaCl, 20 mM Tris pH 7.5) can be made in advance and stored at room temperature until HB is needed. Prior to use, prepare 1 mL per sample of the general buffer and add SDS to a final concentration of 0.01%.</li>
		<li>Add 800 &#181;L of HB to the microfuge tubes containing the nematode pellet. Pellet the samples in the microcentrifuge (see step 1.1.4). Remove the supernatant without disturbing the pellet.</li>
	</ol>
	</li>
	<li>Hybridize the FISH probe to the desired target sequence
	<ol>
		<li>Mix 100 &#181;L per sample of prepared HB with the desired FISH probe to a final concentration of 5-10 ng/&#181;L probe. 
		NOTE: FISH probes are 15-23 -mer oligos, antisense to the SSU of the microbe of interest, and labeled with a colored fluorophore attached to the 5&#39; or 3&#39; end (see Table 1 for probes used here). Generally, stock FISH probes are stored at 1 mg/mL.</li>
		<li>Add 100 &#181;L of HB containing FISH probe to each sample. Mix by gently flicking or inverting the tubes. 
		NOTE: Different colored FISH probes may be added simultaneously to visualize multiple fluorescent signals in the same sample (see Figure 1B).</li>
		<li>Incubate the samples overnight (6-24 h) in a dry bath at 46-54 &#176;C or a thermal mixer at 46-54 &#176;C at 1,200 rpm. 
		NOTE: Incubation at 46-48 &#176;C is typically used for hybridization. However, this temperature may need to be adjusted depending on the melting temperature of the FISH probe. In general, the hybridization temperature is 4 &#176;C below the melting temperature.</li>
	</ol>
	</li>
	<li>Remove the FISH probe and wash nematodes
	<ol>
		<li>Prepare 3 mL of wash buffer (WB) (900 mM NaCl, 20 mM Tris pH 7.5, 5 mM EDTA, 0.01% SDS) per sample. 
		NOTE: WB should be prepared fresh before each use to avoid precipitation. However, a general buffer (900 mM NaCl, 20 mM Tris pH 7.5) can be made in advance and stored at room temperature until the wash buffer is needed. Prior to use, prepare 3 mL per sample of the general buffer and add EDTA to a final concentration of 5 mM and SDS to a final concentration of 0.01%.</li>
		<li>Centrifuge the samples at the appropriate speed (see step 1.1.4). Remove the HB using a pipette, while being careful to leave the nematode pellet undisturbed.</li>
		<li>Add 1 mL of prepared WB to each sample.</li>
		<li>Centrifuge the samples at the appropriate speed (see step 1.1.4). Remove the WB using a pipette, while being careful to leave the pellet undisturbed.</li>
		<li>Add 1 mL of prepared WB to each sample.</li>
		<li>Incubate the samples for 1 h at 48-56 &#176;C in a dry bath (or thermal mixer at 48-56 &#176;C at 1200 rpm). If incubating in a dry bath, gently invert the tubes every 15-20 min. 
		NOTE: In general, 48 &#176;C is used as the standard wash temperature; however, this temperature may need to be adjusted if there is high background signal. The wash temperature is often 2 &#176;C higher than the hybridization temperature. Incubation time in WB can be shortened to 30 min to further reduce background, after which, steps 1.7.4 and 1.7.5 should be repeated. This is to be followed by one (for bacteria) or two (for microsporidia sporoplasms) 30 min incubation periods at 48 &#176;C.</li>
		<li>Centrifuge the samples at the appropriate speed (see step 1.1.4). Using a pipette, remove the wash buffer, while being careful to leave the pellet undisturbed.</li>
		<li>Add 100-500 &#181;L of PBS-T to each of the samples. At this point, the samples can be stored in PBS-T at 4 &#176;C for up to a week until the protocol is ready to be continued.</li>
	</ol>
	</li>
	<li>Mount the nematodes
	<ol>
		<li>Centrifuge the samples at the appropriate speed (see step 1.1.4). Remove as much PBS-T as possible without disturbing the nematode pellet.</li>
		<li>(Optional) Add 20 &#181;L of antifade mounting medium with DAPI (Table of Materials) to the samples.</li>
		<li>Load a 20 &#181;L pipettor with a 200 &#181;L pipette tip and use scissors to cut the tip of the pipette off to allow larger nematodes to be pipetted.</li>
		<li>With the cut pipette tip, transfer 5-10 &#181;L of the pellet onto a microscope slide. Cover with a 22 x 22 coverslip. To store the slides, seal the edges of the coverslip with nail polish and keep them in a dark box at 4 &#176;C until ready for further use.</li>
	</ol>
	</li>
</ol>
2. FISH staining with acetone fixation
<ol>
	<li>Prepare nematodes with associated microbes as described in step 1.1.</li>
	<li>Wash nematodes to eliminate external contamination as described in step 1.2.</li>
	<li>Fix nematodes with acetone
	<ol>
		<li>Remove supernatant without disturbing the pellet and add 1 mL of lab-grade acetone to the sample. 
		CAUTION: Acetone is a highly flammable liquid and vapor. Although it can be purchased over the counter as nail polish remover, it is important to remember that acetone causes serious eye irritation, and may cause drowsiness or dizziness.</li>
		<li>Incubate the samples containing nematodes colonized or infected with the microbe of interest for 15 min at room temperature. Following incubation, the samples can be stored in acetone at -20 &#176;C for up to 2 weeks until the protocol is ready to be continued. 
		NOTE: Do not use this protocol with transgenic C. elegans strains expressing GFP (or its allelic mutated forms) if it is desired to maintain the fluorescence, as acetone destroys this signal.</li>
	</ol>
	</li>
	<li>Remove the acetone
	<ol>
		<li>Spin down the samples in a microcentrifuge at the appropriate speed (see step 1.1.4). With a pipette, remove the supernatant without disturbing the pellet. 
		CAUTION: The supernatant contains acetone. Discard the supernatant and at least the first two washes as toxic waste in a fume hood.</li>
		<li>Add 0.5 mL of PBS-T to the samples in the microfuge tubes.</li>
		<li>Follow and repeat steps 2.4.1 and 2.4.2 2-4x with PBS-T. 
		NOTE: Performing more washes will help reduce the background signal. It is recommended to perform four washes in total.</li>
		<li>After the last wash, spin down the samples and remove the supernatant, ensuring that the pellet is undisturbed.</li>
	</ol>
	</li>
	<li>Prepare hybridization buffer (HB) and wash the nematodes as described in step 1.5.</li>
	<li>Hybridize the FISH probe to the desired target sequence as described in step 1.6.</li>
	<li>Remove FISH probe and wash nematodes
	<ol>
		<li>Prepare 1.1 mL of wash buffer (WB) (900 mM NaCl, 20 mM Tris pH 7.5, 5 mM EDTA, 0.01% SDS) per sample. 
		NOTE: WB should be prepared fresh before each use to avoid precipitation. However, a general buffer (900 mM NaCl, 20 mM Tris pH 7.5) can be made in advance and stored at room temperature until a wash buffer is needed. Prior to use, prepare 1.1 mL per sample of the general buffer and add EDTA to a final concentration of 5 mM and SDS to a final concentration of 0.01%.</li>
		<li>Centrifuge the samples at the appropriate speed (see 1.1.4). Remove the HB using a pipette, while being careful to leave the nematode pellet undisturbed.</li>
		<li>Add 100 &#181;L of prepared WB to each sample.</li>
		<li>Centrifuge the samples at the appropriate speed (see 1.1.4). Remove the WB using a pipette, while being careful to leave the pellet undisturbed.</li>
		<li>Add 1 mL of prepared WB to each sample.</li>
		<li>Incubate the samples for 1 h at 48-56 &#176;C in a dry bath (or thermal mixer at 48-56 &#176;C at 1,200 rpm). If incubating in a dry bath, gently invert the tubes every 15-20 min. 
		NOTE: In general, 48 &#176;C is used as the standard wash temperature; however, this temperature may need to be adjusted if there is high background signal. The wash temperature is often 2 &#176;C higher than the hybridization temperature.</li>
		<li>Centrifuge the samples at the appropriate speed (see step 1.1.4). Using a pipette, remove the wash buffer, while being careful to leave the pellet undisturbed.</li>
		<li>Add 100-500 &#181;L of PBS-T to each of the samples. At this point, the samples can be stored in PBS-T at 4 &#176;C for up to a week until the protocol is ready to be continued.</li>
	</ol>
	</li>
	<li>Mount the nematodes as described in step 1.8.</li>
</ol>

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R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Natural+variation+in+the+roles+of+C.+elegans+autophagy+components+during+microsporidia+infection.">Natural variation in the roles of C. elegans autophagy components during microsporidia infection.</a> PloS One. 14 (4), 0216011 (2019).</li><li>Dirksen, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+native+microbiome+of+the+nematode+Caenorhabditis+elegans:+gateway+to+a+new+host-microbiome+model.">The native microbiome of the nematode Caenorhabditis elegans: gateway to a new host-microbiome model.</a> BMC Biology. 14 (1), 38 (2016).</li><li>Fu, R., Gong, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+cell+analysis+linking+ribosomal+(r)DNA+and+rRNA+copy+numbers+to+cell+size+and+growth+rate+provides+insights+into+molecular+protistan+ecology.">Single cell analysis linking ribosomal (r)DNA and rRNA copy numbers to cell size and growth rate provides insights into molecular protistan ecology.</a> The Journal of Eukaryotic Microbiology. 64 (6), 885-896 (2017).</li><li>Willis, A. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+parental+transcriptional+response+to+microsporidia+infection+induces+inherited+immunity+in+offspring.">A parental transcriptional response to microsporidia infection induces inherited immunity in offspring.</a> Science Advances. 7 (19), (2021).</li><li>Cuomo, C. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microsporidian+genome+analysis+reveals+evolutionary+strategies+for+obligate+intracellular+growth.">Microsporidian genome analysis reveals evolutionary strategies for obligate intracellular growth.</a> Genome Research. 22 (12), 2478-2488 (2012).</li><li>Reinke, A. W., Balla, K. M., Bennett, E. J., Troemel, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+microsporidia+host-exposed+proteins+reveals+a+repertoire+of+rapidly+evolving+proteins.">Identification of microsporidia host-exposed proteins reveals a repertoire of rapidly evolving proteins.</a> Nature Communications. 8 (1), 14023 (2017).</li><li>Rivera, D. E., La&#382;eti&#263;, V., Troemel, E. R., Luallen, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RNA+fluorescence+in+situ+hybridization+(FISH)+to+visualize+microbial+colonization+and+infection+in+the+Caenorhabditis+elegans+intestines.">RNA fluorescence in situ hybridization (FISH) to visualize microbial colonization and infection in the Caenorhabditis elegans intestines.</a> bioRxiv. , (2022).</li></ol>

The authors have no conflicts of interest.

Wild C. elegans are naturally associated with a variety of microbes. Researchers can use RNA FISH to detect and identify these microbes as well as gain insight into their localization in the context of a whole animal. Microbes with desirable or interesting phenotypes can be identified through this method and then isolated for further characterization and sequencing. The abundance of numerous bacterial isolates from wild C. elegans may also be quantified via RNA FISH29. By using the protocol described here, it is also possible to observe known microorganisms inside their hosts and learn more about their interactions. Importantly, Orsay virus and microsporidia are obligate parasites and cannot be cultured independently of the host, so FISH is the standard visualization tool. Colonization or infection can also be quantified through RNA FISH using nematodes grown on plates seeded with a desired culturable bacteria of interest. In addition to staining microorganisms in C. elegans intestine, this protocol can be used for other nematode strains like C. tropicalis or Oscheius tipulae19,23.
The main advantage of the FISH protocol is that it offers a simple, quick, and robust method to stain microbes associated with C. elegans. Images produced from FISH staining have a high signal-to-noise ratio, which is achieved by utilizing FISH probes that target the abundant RNA of the SSU within the sample. Because there are typically 30x or higher levels of rRNA than rDNA, most of the signal from FISH staining with probes that target rRNA is due to rRNA rather than rDNA30. Furthermore, RNA FISH makes it possible to see infection or colonization within the context of the whole animal. This visualization is facilitated through co-staining host nuclei with DAPI and/or using fluorescent-marked strains of C. elegans to better highlight the localization of infection or colonization within the sample. For example, microsporidian-specific FISH was used to determine the tissue tropism of Nematocida displodere by using a panel of C. elegans strains with GFP expression in different tissues20. Additionally, this protocol is amenable to changes that allow for researchers to determine the ideal conditions suitable for their specific needs (e.g., adjusting the fixation period, increasing the hybridization temperature).
One critical step in the FISH protocol is fixing the samples. The incubation period following the addition of the fixative is necessary to allow time for the agent to permeabilize the sample. Longer incubation times are not ideal for samples containing transgenic fluorescent proteins due to protein degradation by PFA over time. For samples containing GFP, it is imperative to determine the optimal fixation time to allow for permeabilization, while still maintaining the GFP signal.
FISH can be used to stain for bacteria, viruses, or microsporidia in C. elegans. However, the best type of fixative agent used for FISH depends on the sample and downstream requirements. This protocol presents a PFA solution as the primary fixative agent to stain bacteria and viruses. However, PFA is not sufficient for the visualization of microsporidian spores as it cannot penetrate the spore wall. For visualization of spores, acetone should be used instead. Although, PFA fixation is efficient for FISH labeling of other life stages of microsporidia, including sporoplasms, meronts, and sporonts. Other major differences are seen between acetone fixation and PFA fixation; acetone is more convenient because samples can be quickly stored in the freezer after adding, without the need for washing. However, acetone quickly kills any existing GFP in a transgenic host. PFA is the preferred fixative if it is important to preserve some physiological structures in the host, as acetone-fixed animals appear to be more degraded, making identification of some tissues more difficult. Because the samples are fixed, this FISH protocol does not allow for live imaging of host-microbe interactions in vivo. However, a pulse-chase infection time course followed by FISH staining of samples at various time points can allow one to see some dynamics of microbial infection19,20,31.
Another critical step throughout the protocol is the thorough washing of the samples before and after hybridization. Before hybridization, when collecting the worms into the microfuge tubes, excess bacteria, or other microbes from the NGM plates can be carried with the worm sample. Three washes with PBS-T are standard; however, more washes may be necessary to sufficiently eliminate external microorganisms, especially when using heavily contaminated, wild-isolated C. elegans. When viewing the mounted samples after FISH, there may be some residual FISH probe that produces large amounts of signal in the background of the sample. The wash temperature and the number of washes are important to remove the excess and non-specifically bound probe. To reduce the background fluorescence, it is possible to perform two or three washes with 1 mL of WB every 30 min, instead of one wash with 1 mL of WB for an hour. Different FISH probes may require different wash temperatures. Typically, the wash temperature is 2 &#176;C above the hybridization temperature, but this can be increased if there is too much background fluorescence (high noise).
The FISH protocol utilizes fluorescent probes designed to target species-specific microbial RNA, but FISH probes can be designed for other high-copy transcripts. Other FISH probes may have different melting temperatures, so incubation steps may need to be performed at a higher or lower temperature than described. FISH staining can identify the spatial distribution of microbial colonization or infection within the host, allowing for the characterization of host-microbe and microbe-microbe interactions. One limitation is that&#160;only a few conventional fluorophores can be used simultaneously, which&#160;reduces the number of different microorganisms that can be detected via FISH at the same time. This&#160;limits its use for complex microbiome studies in C. elegans. However, multicolor rRNA-targeted FISH utilizes probes labeled with non-canonical fluorophores that can increase the number of distinct microbial group labels15. Another limitation is that it is difficult to distinguish between closely related species, especially bacteria, that have SSU sequences that are highly similar. However, the extreme sequence divergence between microsporidia species helps to facilitate their differentiation with this protocol (Figure 3)32,33.
Overall, this FISH protocol describes a technique to detect microorganisms within C. elegans. It allows researchers to use a transparent and genetically tractable model system to detect and quantify colonization and infection within the context of an intact animal, as well as identify unique microbial behavior or morphology within the host. A preprint version of this manuscript was posted during review34.

Caenorhabditis elegans has emerged as a powerful model system to study innate immunity and host-microbe interactions in the intestinal epithelial cells1,2. Due to having a transparent body and only 20 intestinal cells, C. elegans represents a convenient system for monitoring the processes of microbial intestinal colonization and infection in the context of an intact organism. Nematode intestinal cells share multiple morphological and functional similarities with mammalian intestinal epithelial cells, making them a tractable in vivo model for dissection of processes that govern microbiome colonization and pathogen infection3,4,5,6.
Wild C. elegans feed on a variety of microbes that colonize and infect the intestine, and sampling of these nematodes has resulted in the discovery of viruses, eukaryotes (fungi, oomycetes), and bacteria that naturally associate with this host7,8,9,10. The Orsay virus was found infecting the intestine and is currently the only known natural virus of C. elegans9. Microsporidia are fungal-related obligate intracellular pathogens that are the most commonly found infection in wild-caught Caenorhabditis, with several species having been discovered infecting C. elegans and related nematodes8,11. Many bacteria are commonly found inhabiting the intestinal lumen of wild-caught C. elegans and several species have been established as a natural model for the C. elegans microbiome (CeMbio)6,12,13,14. Discovering and characterizing microbes that naturally colonize and/or infect C. elegans is essential to understanding the genetic mechanisms that govern these host-microbe interactions, as well as visualizing novel microbial processes that only occur in the context of an intact host animal.
After sampling, wild nematodes are screened via differential interference contrast (DIC) microscopy to look for phenotypes that are indicative of infection or colonization. For example, changes in the stereotypical granulated appearance of the intestinal cells can be associated with the presence of an intracellular parasite infection8. Specifically, the loss of the gut granules and decreased cytosolic viscosity are signs of viral infection, whereas the reorganization of gut granules into &#39;grooves&#39; may indicate infection with microsporidia in the genus Nematocida8,9. Because there is a wide variety of microbes present in wild C. elegans samples, it can be difficult to distinguish among microbes through DIC microscopy. Information regarding the spatial distribution of microbes within the host may also be difficult to detect due to the small size of many microbes15. Additionally, culturing any particular microbes of interest in vitro is not always possible, leading to difficulties in detection and/or quantification.
RNA fluorescence in situ hybridization (FISH) provides a method to fluorescently label microbes by utilizing fluorescent probes that bind to the RNA of the small ribosomal subunit (SSU) in fixed cells. If analysis of morphological characteristics suggests a particular class of microbe, FISH probes that target specific or broad classes of such microbes can be used. For example, EUB338 is considered a universal probe for the bacterial SSU and is commonly used to detect a wide range of bacteria16. The protocol described here uses single-stranded DNA probes that are end-labeled with a fluorophore and specifically designed to be complementary to the target SSU of the microbe of interest, although there are previously designed probes available16. The main advantage of targeting the SSU of microbes is the relatively large abundance of this RNA, which typically comprises 80%-90% of all RNA in the cell, leading to staining with a very high signal-to-noise ratio17. Probes can also be designed to target RNA to detect viruses, like the Orsay virus9,18, which are often present in very high copies in infected cells if the virus is actively replicating.
Depending on the results with known probes, it may be necessary to obtain further sequence information to design more specific probes for species confirmation in situ. A common approach is to use universal primers against conserved regions of the SSU (16S for bacteria and 18S for eukaryotes) to amplify (via PCR) regions that are more divergent8. Using this sequence information, probes with more species-specificity can be designed. These FISH probes can then enable the identification of microbes in a culture-independent manner8. Additionally, RNA FISH can give insight into unique morphological colonization and infection characteristics, including filamentation or tissue localization patterns19,20. Different colored FISH probes can be used simultaneously, which allows for visual distinction between microbes in wild nematode samples, as well as observation of microbe-microbe dynamics inside a host15,20. Furthermore, RNA FISH staining can be applied to host-pathogen interaction studies where infection and colonization of a known species can be easily quantified manually or through automated approaches to provide insights on pathogen load, for example, in comparing C. elegans mutants that have either increased or decreased resistance to infection21.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>10% SDS</td>
			<td>Invitrogen</td>
			<td>AM9822</td>
			<td></td>
		</tr>
		<tr>
			<td>Acetone</td>
			<td>Fisher Scientific</td>
			<td>A-11-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Antifade mounting serum with DAPI (Vectashield)</td>
			<td>Vectalab</td>
			<td>NC9524612</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA</td>
			<td>Fisher Scientific</td>
			<td>S311-500</td>
			<td></td>
		</tr>
		<tr>
			<td>FISH probes (see Table 1)</td>
			<td>LGC Biosearch Technologies</td>
			<td></td>
			<td>FISH probes were commercially purchased via custom oligonucleotide synthesis</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Fisher Scientific</td>
			<td>P217</td>
			<td></td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>Fisher Scientific</td>
			<td>P-286</td>
			<td></td>
		</tr>
		<tr>
			<td>Na2HPO4</td>
			<td>Fisher Scientific</td>
			<td>S375-500</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Fisher Scientific</td>
			<td>S-671</td>
			<td></td>
		</tr>
		<tr>
			<td>NH4Cl</td>
			<td>Fisher Scientific</td>
			<td>A-661</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Electron Microscopy Science</td>
			<td>50-980-487</td>
			<td>CAUTION: PFA is a carcinogen. Handle appropriately</td>
		</tr>
		<tr>
			<td>Thermal mixer</td>
			<td>Eppendorf</td>
			<td>5384000020</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris base</td>
			<td>Fisher Scientific</td>
			<td>BP152</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Fisher Scientific</td>
			<td>BP-151</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween-20</td>
			<td>Fisher Scientific</td>
			<td>BP337-500</td>
			<td></td>
		</tr>
	</tbody>
</table>

rna fluorescence in situ hybridization (fish) to visualize microbial colonization and infection in caenorhabditis elegans intestines

The intestines of wild Caenorhabditis nematodes are inhabited by a variety of microorganisms, including gut microbiome bacteria and pathogens, such as microsporidia and viruses. Because of the similarities between Caenorhabditis elegans and mammalian intestinal cells, as well as the power of the C. elegans system, this host has emerged as a model system to study host intestine-microbe interactions in vivo. While it is possible to observe some aspects of these interactions with bright-field microscopy, it is difficult to accurately classify microbes and characterize the extent of colonization or infection without more precise tools. RNA fluorescence in situ hybridization (FISH) can be used as a tool to identify and visualize microbes in nematodes from the wild or to experimentally characterize and quantify infection in nematodes infected with microbes in the lab. FISH probes, labeling the highly abundant small subunit ribosomal RNA, produce a bright signal for bacteria and microsporidian cells. Probes designed to target conserved regions of ribosomal RNA common to many species can detect a broad range of microbes, whereas targeting divergent regions of the ribosomal RNA is useful for narrower detection. Similarly, probes can be designed to label viral RNA. A protocol for RNA FISH staining with either paraformaldehyde (PFA) or acetone fixation is presented. PFA fixation is ideal for nematodes associated with bacteria, microsporidia, and viruses, whereas acetone fixation is necessary for the visualization of microsporida spores. Animals were first washed and fixed in paraformaldehyde or acetone. After fixation, FISH probes were incubated with samples to allow for the hybridization of probes to the desired target. The animals were again washed and then examined on microscope slides or using automated approaches. Overall, this FISH protocol enables detection, identification, and quantification of the microbes that inhabit the C. elegans intestine, including microbes for which there are no genetic tools available.

For analyzing microbiome bacteria, specific and universal FISH probes to bacterial 16S were utilized on wild-isolated animals. Wild Caenorhabditis tropicalis strain (JU1848) was sampled from the Nouragues forest near a small river in the French Guiana from rotting palm tree fruits22. Under the differential interference contrast (DIC) microscope, this nematode strain was found to be colonized with a bacterium that appears to directionally adhere to the intestinal epithelium (Figure 2A). JU1848 was then selectively cleaned to eliminate other microbial contaminants and enrich for the desired adhering bacterium23. Using the universal PCR method, the bacterium was identified as a new species in the Alphaproteobacteria class. A FISH probe labeled with Cal Fluor Red 610 was then designed specifically to the 16S rRNA sequence of this bacterium to allow fluorescent visualization of colonization within C. tropicalis (Figure 2B). A universal 16S rRNA FISH probe capable of binding many species of bacteria (EUB338) was labeled with 6-carboxyfluorescin (FAM) and was also added to this sample. The green and red fluorescent signals overlap completely, suggesting that most bacteria colonizing the intestines are the adhering Alphaproteobacteria bacterium. These animals were fixed in PFA before staining.
For analyzing experimental infection in the lab with intracellular pathogens of known identity, Orsay virus and microsporidian-specific FISH probes were utilized on C. elegans with a wild-type background. The Orsay virus is a positive-strand RNA virus from the Nodaviridae family, and the only natural viral pathogen found in C. elegans. The bipartite RNA genome of the Orsay virus consists of RNA1 and RNA2 segments, and FISH probes targeting both of these segments have been developed (Figure 3A,B)9,18. In the intestine, viral RNA is sensed by RIG-I homolog DRH-124, which is required for activation of the transcriptional defense program named the Intracellular Pathogen Response (IPR)25,26,27. The transcription of antiviral IPR genes is at least partially controlled by the ZIP-1 transcription factor21. Here, the expression of ZIP-1::GFP is seen localized in the intestinal nuclei of cells that show positive Orsay virus FISH staining in the cytoplasm (Figure 3A)21. Multiple animals stained with Orsay-specific FISH are shown to indicate the strength of this signal for easy quantification (Figure 3B). Animals shown in Figure 3A,B were fixed in PFA.
The microsporidian parasite named Nematocida parisii, meaning nematode-killer from Paris, is an obligate intracellular pathogen of the intestine. Several FISH probes that label 18S rRNA of N. parisii have been used, including fluorescently tagged MicroA and MicroB&#160;probes. Multiple animals stained with MicroB FISH are shown to indicate the strength of this signal for easy quantification (Figure 3C). Additionally, C. elegans is infected by other closely related microsporidia. Co-infection of N2 with N. parisii and the related N. ausubeli can be distinguished using this FISH protocol by designing species-specific FISH probes that compete against each other for binding to a divergent region on the 18S rRNA (Figure 3D)28. In this example, the N. parisii FISH probe has perfect base-pairing to the N. parisii 18S rRNA, but a 7 bp mismatch to N. ausubeli 18S rRNA. The converse is true for the N. ausubeli probe. As such, each species-specific FISH probe will outcompete for binding to the cognate species 18S over the non-cognate species. Additionally, the use of DAPI to stain nuclei allows for better localization of the infection in the context of the whole animal, especially for the intestine which has large, easily identifiable nuclei. Figure 3C,D contains animals that were fixed in PFA. Later infections with N. parisii result in the development of meronts into spores. To visualize N. parisii spores, the animals must be fixed in acetone as it penetrates the spore wall better than PFA (Figure 3E,F)8. The resulting FISH staining demonstrates the small and large rod-shaped structures, that likely correspond with N. parisii spores, which are stained with N. parisii-specific probes in red.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63980/63980fig1v2.jpg" /> 
Figure 1: Visual representation of FISH protocol. Created with Biorender.com. <a href="https://www.jove.com/files/ftp_upload/63980/63980fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63980/63980fig02.jpg" /> 
Figure 2: FISH staining of wild C. tropicalis JU1848 strain colonized with adhering bacteria in the intestines. (A) Nomarski image depicting thousands of thin bacilli bacteria (bac) directionally binding to the intestine (in) of JU1848, creating a hair-like phenotype within the lumen (lu). This figure panel is adapted from Morgan, E. et al. (2021)23. (B) FISH staining of JU1848, fixed in PFA, using a red labeled probe (b002_16S_A-CF610) designed to target the 16S rRNA sequence of the adhering bacterium (top) and a green-labeled universal FISH probe (EUB338-FAM) designed to target the 16S of bacteria (bottom). DAPI staining of host nuclei is shown in blue. See Table 1 for probe sequences. <a href="https://www.jove.com/files/ftp_upload/63980/63980fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63980/63980fig03.jpg" /> 
Figure 3: FISH staining of C. elegans infected with intracellular pathogens. (A,B) FISH staining of C. elegans expressing ZIP-1::GFP and infected with the Orsay virus, which were fixed with PFA before staining to preserve the GFP signal. Orsay 1 Red and Orsay 2 Red probes were used for pathogen staining. (A) The composite image consists of merged red and green fluorescent channels. Nuclear ZIP-1::GFP expression is induced upon Orsay virus infection and is shown in green. Autofluorescence from the gut granules is shown in yellow and indicated with yellow arrows. Dotted lines outline the nematode body. Scale bar = 25 &#181;m. (B) The composite image consists of merged red fluorescent and DIC channels. Scale bar = 200 &#181;m. (C,D) FISH staining of wild-type C. elegans infected with microsporidia that were fixed in PFA. (C) FISH staining of wild-type C. elegans infected with N. parisii and fixed in PFA. MicroB-CF610 probe was used for pathogen staining. The composite image consists of merged red fluorescent and DIC channels. Scale bar = 100 &#181;m. (D) FISH staining of wild-type C. elegans co-infected with N. parisii and N. ausubeli in the intestine. The two pathogens were co-stained using a pair of specific FISH probes that compete for binding to the same region of the 18S rRNA. N. parisii was stained using MicroF-CF610 (red) and N. ausubeli was stained using MicroSp1A-FAM (green). DAPI staining of host nuclei is seen in blue. Scale bar = 25 &#181;m. (E) FISH staining with acetone-fixed wild-type C. elegans infected with N. parisii spores. MicroA-CF610 (red) was used for staining (red). Scale bar = 15 &#181;m. (F) Nomarski image depicting N. parisii spores seen in (E). Scale bar = 15 &#181;m. In (E) and (F), the small and large rod-shaped structures are labeled with small and large arrows, respectively, that correspond to N. parisii spores. See Table 1 for probe sequences. The image shown in (A) is adapted from La&#382;eti&#263;, V. et al. (2022)21. Images shown in (B) and (C) are adapted from Reddy, K. C. et al. (2019)26. Images shown in (E) and (F) are adapted from Troemel, E. R. et al. (2008)8. <a href="https://www.jove.com/files/ftp_upload/63980/63980fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Probe name</td>
			<td>Probe specificity</td>
			<td>Probe fluorophore</td>
			<td>Probe sequence</td>
		</tr>
		<tr>
			<td>EUB338-FAM</td>
			<td>Bacterial 16S (universal)</td>
			<td>5&#39; 6-fluorescein (FAM)</td>
			<td>GCTGCCTCCCGTAGGAGT</td>
		</tr>
		<tr>
			<td>b002_16S_A-CF610</td>
			<td>Alphaproteobacteria 16S</td>
			<td>Cal Fluor Red 610 (CF610)</td>
			<td>TGTACCGACCCTTAACGTTC</td>
		</tr>
		<tr>
			<td>Orsay1 Red</td>
			<td>Orsay virus RNA1</td>
			<td>Cal Fluor Red 610 (CF610)</td>
			<td>GACATATGTGATGCCGAGAC</td>
		</tr>
		<tr>
			<td>Orsay2 Red</td>
			<td>Orsay virus RNA2</td>
			<td>Cal Fluor Red 610 (CF610)</td>
			<td>GTAGTGTCATTGTAGGCAGC</td>
		</tr>
		<tr>
			<td>MicroA-CF610</td>
			<td>Nematocida parisii 18S</td>
			<td>Cal Fluor Red 610 (CF610)</td>
			<td>CTCTGTCCATCCTCGGCAA</td>
		</tr>
		<tr>
			<td>MicroB-CF610</td>
			<td>Nematocida parisii 18S</td>
			<td>Cal Fluor Red 610 (CF610)</td>
			<td>CTCTCGGCACTCCTTCCTG</td>
		</tr>
		<tr>
			<td>MicroF-CF610</td>
			<td>Nematocida parisii 18S</td>
			<td>Cal Fluor Red 610 (CF610)</td>
			<td>AGACAAATCAGTCCACGAATT</td>
		</tr>
		<tr>
			<td>MicroSp1A-FAM</td>
			<td>Nematocida ausubeli 18S</td>
			<td>5&#39; 6-fluorescein (FAM)</td>
			<td>CAGGTCACCCCACGTGCT</td>
		</tr>
	</tbody>
</table>
Table 1: List of FISH probe sequences. All FISH probes were commercially purchased with the fluorophore attached to the 5&#39; end (via custom oligonucleotide synthesis; see Table of Materials) and the oligonucleotides were purified by reverse-phase HPLC.

Watch this Scientific Journal Video about RNA Fluorescence in situ Hybridization FISH to Visualize Microbial Colonization and Infection in Caenorhabditis elegans Intestines at JoVE.com

RNA Fluorescence in situ Hybridization FISH to Visualize Microbial Colonization and Infection in Caenorhabditis elegans Intestines

Intestinal microbes, including extracellular bacteria and intracellular pathogens like the Orsay virus and microsporidia (fungi), are often associated with wild Caenorhabditis nematodes. This article presents a protocol for detecting and quantifying microbes that colonize and/or infect C. elegans nematodes, and for measuring pathogen load after controlled infections in the lab.

RNA Fluorescence in situ Hybridization (FISH) to Visualize Microbial Colonization and Infection in Caenorhabditis elegans Intestines

The intestines of wild Caenorhabditis nematodes are inhabited by a variety of microorganisms, including gut microbiome bacteria and pathogens, such as ...

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Immunology and Infection

3.7K Views.  San Diego State University. Intestinal microbes, including extracellular bacteria and intracellular pathogens like the Orsay virus and microsporidia (fungi), are often associated with wild Caenorhabditis nematodes. This article presents a protocol for detecting and quantifying microbes that colonize and/or infect C. elegans nematodes, and for measuring pathogen load after controlled infections in the lab.

Video: RNA Fluorescence in situ Hybridization FISH to Visualize Microbial Colonization and Infection in Caenorhabditis elegans Intestines

Combination of Adhesive-tape-based Sampling and Fluorescence in situ Hybridization for Rapid Detection of Salmonella on Fresh Produce

This protocol describes a simple approach for adhesive-tape-based sampling of tomato and other fresh produce surfaces, followed by on-tape fluorescence in situ hybridization (FISH) for rapid culture-independent detection of Salmonella spp. Cell-charged tapes can also be placed face-down on selective agar for solid-phase enrichment prior to detection. Alternatively, low-volume liquid enrichments (liquid surface miniculture) can be performed on the surface of the tape in non-selective broth, followed by FISH and analysis via flow cytometry. To begin, sterile adhesive tape is brought into contact with fresh produce, gentle pressure is applied, and the tape is removed, physically extracting microbes present on these surfaces. Tapes are mounted sticky-side up onto glass microscope slides and the sampled cells are fixed with 10% formalin (30 min) and dehydrated using a graded ethanol series (50, 80, and 95%; 3 min each concentration). Next, cell-charged tapes are spotted with buffer containing a Salmonella-targeted DNA probe cocktail and hybridized for 15 - 30 min at 55&deg;C, followed by a brief rinse in a washing buffer to remove unbound probe. Adherent, FISH-labeled cells are then counterstained with the DNA dye 4',6-diamidino-2-phenylindole (DAPI) and results are viewed using fluorescence microscopy. For solid-phase enrichment, cell-charged tapes are placed face-down on a suitable selective agar surface and incubated to allow in situ growth of Salmonella microcolonies, followed by FISH and microscopy as described above. For liquid surface miniculture, cell-charged tapes are placed sticky side up and a silicone perfusion chamber is applied so that the tape and microscope slide form the bottom of a water-tight chamber into which a small volume (&le; 500 &mu;L) of Trypticase Soy Broth (TSB) is introduced. The inlet ports are sealed and the chambers are incubated at 35 - 37&deg;C, allowing growth-based amplification of tape-extracted microbes. Following incubation, inlet ports are unsealed, cells are detached and mixed with vigorous back and forth pipetting, harvested via centrifugation and fixed in 10% neutral buffered formalin. Finally, samples are hybridized and examined via flow cytometry to reveal the presence of Salmonella spp. As described here, our "tape-FISH" approach can provide simple and rapid sampling and detection of Salmonella on tomato surfaces. We have also used this approach for sampling other types of fresh produce, including spinach and jalape&ntilde;o peppers.

Combination of Adhesive-tape-based Sampling and Fluorescence in situ Hybridization for Rapid Detection of Salmonella on Fresh Produce

This protocol describes a simple adhesive-tape-based approach for sampling of tomato and other fresh produce surfaces, followed by rapid whole cell detection of Salmonella using fluorescence in situ hybridization (FISH).

This protocol describes a simple approach for adhesive-tape-based sampling of tomato and other fresh produce surfaces, followed by on-tape fluorescence in ...

Locked Nucleic Acid Flow Cytometry-fluorescence in situ Hybridization (LNA flow-FISH): a Method for Bacterial Small RNA Detection

Fluorescence in situ hybridization (FISH) is a powerful technique that is used to detect and localize specific nucleic acid sequences in the cellular environment. In order to increase throughput, FISH can be combined with flow cytometry (flow-FISH) to enable the detection of targeted nucleic acid sequences in thousands of individual cells. As a result, flow-FISH offers a distinct advantage over lysate/ensemble-based nucleic acid detection methods because each cell is treated as an independent observation, thereby permitting stronger statistical and variance analyses. These attributes have prompted the use of FISH and flow-FISH methods in a number of different applications and the utility of these methods has been successfully demonstrated in telomere length determination1,2, cellular identification and gene expression3,4, monitoring viral multiplication in infected cells5, and bacterial community analysis and enumeration6.
Traditionally, the specificity of FISH and flow-FISH methods has been imparted by DNA oligonucleotide probes. Recently however, the replacement of DNA oligonucleotide probes with nucleic acid analogs as FISH and flow-FISH probes has increased both the sensitivity and specificity of each technique due to the higher melting temperatures (Tm) of these analogs for natural nucleic acids7,8. Locked nucleic acid (LNA) probes are a type of nucleic acid analog that contain LNA nucleotides spiked throughout a DNA or RNA sequence9,10. When coupled with flow-FISH, LNA probes have previously been shown to outperform conventional DNA probes7,11 and have been successfully used to detect eukaryotic mRNA12 and viral RNA in mammalian cells5.
Here we expand this capability and describe a LNA flow-FISH method which permits the specific detection of RNA in bacterial cells (Figure 1). Specifically, we are interested in the detection of small non-coding regulatory RNA (sRNA) which have garnered considerable interest in the past few years as they have been found to serve as key regulatory elements in many critical cellular processes13. However, there are limited tools to study sRNAs and the challenges of detecting sRNA in bacterial cells is due in part to the relatively small size (typically 50-300 nucleotides in length) and low abundance of sRNA molecules as well as the general difficulty in working with smaller biological cells with varying cellular membranes. In this method, we describe fixation and permeabilzation conditions that preserve the structure of bacterial cells and permit the penetration of LNA probes as well as signal amplification steps which enable the specific detection of low abundance sRNA (Figure 2).

Locked Nucleic Acid Flow Cytometry-fluorescence in situ Hybridization LNA flow-FISH: a Method for Bacterial Small RNA Detection

A novel high-throughput method is described that enables the detection and relative quantitation of small RNA and mRNA expression from single bacterial cells using locked nucleic acid probes and flow cytometry-fluorescence in situ hybridization.

Fluorescence in situ hybridization (FISH) is a powerful technique that is used to detect and localize specific nucleic acid sequences in the cellular ...

Use of Artificial Sputum Medium to Test Antibiotic Efficacy Against Pseudomonas aeruginosa in Conditions More Relevant to the Cystic Fibrosis Lung

There is growing concern about the relevance of in vitro antimicrobial susceptibility tests when applied to isolates of P. aeruginosa from cystic fibrosis (CF) patients. Existing methods rely on single or a few isolates grown aerobically and planktonically. Predetermined cut-offs are used to define whether the bacteria are sensitive or resistant to any given antibiotic1. However, during chronic lung infections in CF, P. aeruginosa populations exist in biofilms and there is evidence that the environment is largely microaerophilic2. The stark difference in conditions between bacteria in the lung and those during diagnostic testing has called into question the reliability and even relevance of these tests3.
 Artificial sputum medium (ASM) is a culture medium containing the components of CF patient sputum, including amino acids, mucin and free DNA. P. aeruginosa growth in ASM mimics growth during CF infections, with the formation of self-aggregating biofilm structures and population divergence4,5,6. The aim of this study was to develop a microtitre-plate assay to study antimicrobial susceptibility of P. aeruginosa based on growth in ASM, which is applicable to both microaerophilic and aerobic conditions.
 An ASM assay was developed in a microtitre plate format. P. aeruginosa biofilms were allowed to develop for 3 days prior to incubation with antimicrobial agents at different concentrations for 24 hours. After biofilm disruption, cell viability was measured by staining with resazurin. This assay was used to ascertain the sessile cell minimum inhibitory concentration (SMIC) of tobramycin for 15 different P. aeruginosa isolates under aerobic and microaerophilic conditions and SMIC values were compared to those obtained with standard broth growth. Whilst there was some evidence for increased MIC values for isolates grown in ASM when compared to their planktonic counterparts, the biggest differences were found with bacteria tested in microaerophilic conditions, which showed a much increased resistance up to a &gt;128 fold, towards tobramycin in the ASM system when compared to assays carried out in aerobic conditions.
 The lack of association between current susceptibility testing methods and clinical outcome has questioned the validity of current methods3. Several in vitro models have been used previously to study P. aeruginosa biofilms7, 8. However, these methods rely on surface attached biofilms, whereas the ASM biofilms resemble those observed in the CF lung9 . In addition, reduced oxygen concentration in the mucus has been shown to alter the behavior of P. aeruginosa2 and affect antibiotic susceptibility10. Therefore using ASM under microaerophilic conditions may provide a more realistic environment in which to study antimicrobial susceptibility.

Use of Artificial Sputum Medium to Test Antibiotic Efficacy Against Pseudomonas aeruginosa in Conditions More Relevant to the Cystic Fibrosis Lung

Current diagnostic antimicrobial susceptibility testing relies on the planktonic growth of isolates in nutrient rich, aerobic conditions. Here, we employ an alternative artificial sputum medium to study antimicrobial susceptibility of Pseudomonas aeruginosa biofilms under both aerobic and microaerophilic conditions more representative of the cystic fibrosis lung.

There  is growing concern about the relevance of in vitro antimicrobial  susceptibility tests when applied to isolates of P. aeruginosa from  cystic ...

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Fluorescent in situ hybridization (FISH) is a technique routinely used by many laboratories to determine the chromosomal position of DNA and RNA probes. One important application of this method is the development of high-quality physical maps useful for improving the genome assemblies for various organisms. The natural banding pattern of polytene and mitotic chromosomes provides guidance for the precise ordering and orientation of the genomic supercontigs. Among the three mosquito genera, namely Anopheles, Aedes, and Culex, a well-established chromosome-based mapping technique has been developed only for Anopheles, whose members possess readable polytene chromosomes 1. As a result of genome mapping efforts, 88% of the An. gambiae genome has been placed to precise chromosome positions 2,3 . Two other mosquito genera, Aedes and Culex, have poorly polytenized chromosomes because of significant overrepresentation of transposable elements in their genomes 4, 5, 6. Only 31 and 9% of the genomic supercontings have been assigned without order or orientation to chromosomes of Ae. aegypti 7 and Cx. quinquefasciatus 8, respectively. Mitotic chromosome preparation for these two species had previously been limited to brain ganglia and cell lines. However, chromosome slides prepared from the brain ganglia of mosquitoes usually contain low numbers of metaphase plates 9. Also, although a FISH technique has been developed for mitotic chromosomes from a cell line of Ae. aegypti 10, the accumulation of multiple chromosomal rearrangements in cell line chromosomes 11 makes them useless for genome mapping. Here we describe a simple, robust technique for obtaining high-quality mitotic chromosome preparations from imaginal discs (IDs) of 4th instar larvae which can be used for all three genera of mosquitoes. A standard FISH protocol 12 is optimized for using BAC clones of genomic DNA as a probe on mitotic chromosomes of Ae. aegypti and Cx. quinquefasciatus, and for utilizing an intergenic spacer (IGS) region of ribosomal DNA (rDNA) as a probe on An. gambiae chromosomes. In addition to physical mapping, the developed technique can be applied to population cytogenetics and chromosome taxonomy/systematics of mosquitoes and other insect groups.

Fluorescent in situ Hybridization on Mitotic Chromosomes of Mosquitoes

Among the three mosquito genera, namely Anopheles, Aedes, and Culex, physical genome mapping techniques were established only for Anopheles, whose members possess readable polytene chromosomes. For the genera of Aedes and Culex, however, cytogenetic mapping remains challenging because of the poor quality of polytene chromosomes. Here we present a universal protocol for obtaining high-quality preparations of mitotic chromosomes and an optimized FISH protocol for all three genera of mosquitoes.

Fluorescent in situ hybridization (FISH) is a technique routinely used by many laboratories to determine the chromosomal position of DNA and RNA probes. ...

Fluorescence in situ Hybridizations (FISH) for the Localization of Viruses and Endosymbiotic Bacteria in Plant and Insect Tissues

Fluorescence in situ hybridization (FISH) is a name given to a variety of techniques commonly used for visualizing gene transcripts in eukaryotic cells and can be further modified to visualize other components in the cell such as infection with viruses and bacteria. Spatial localization and visualization of viruses and bacteria during the infection process is an essential step that complements expression profiling experiments such as microarrays and RNAseq in response to different stimuli. Understanding the spatiotemporal infections with these agents complements biological experiments aimed at understanding their interaction with cellular components. Several techniques for visualizing viruses and bacteria such as reporter gene systems or immunohistochemical methods are time-consuming, and some are limited to work with model organisms&#160;and involve complex methodologies. FISH that targets RNA or DNA species in the cell is a relatively easy and fast method for studying spatiotemporal localization of genes and for diagnostic purposes. This method can be robust and relatively easy to implement when the protocols employ short hybridizing, commercially-purchased probes, which are not expensive. This is particularly robust when sample preparation, fixation, hybridization, and microscopic visualization do not involve complex steps. Here we describe a protocol for localization of bacteria and viruses in insect and plant tissues. The method is based on simple preparation, fixation, and hybridization of insect whole mounts and dissected organs or hand-made plant sections, with 20 base pairs short DNA probes conjugated to fluorescent dyes on their 5&#39; or 3&#39; ends. This protocol has been successfully applied to a number of insect and plant tissues, and can be used to analyze expression of mRNAs or other RNA or DNA species in the cell.

Fluorescence in situ Hybridizations FISH for the Localization of Viruses and Endosymbiotic Bacteria in Plant and Insect Tissues

We describe here a simple fluorescence in situ hybridization (FISH) method for the localization of viruses and bacteria in insect and plant tissues. This protocol can be extended for the visualization of mRNA in whole mount and microscopic sections.

Fluorescence in situ hybridization (FISH) is a name given to a variety of techniques commonly used for visualizing gene transcripts in eukaryotic cells ...

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

The obligate intracellular bacterium Chlamydia elicits a great burden on global public health. C.&#160;trachomatis is the leading bacterial cause of sexually transmitted infection and also the primary cause of preventable blindness in the world. An essential determinant for successful infection of host cells by Chlamydia is the bacterium's ability to manipulate host cell signaling from within a novel, vacuolar compartment called the inclusion. From within the inclusion, Chlamydia acquire nutrients required for their 2-3 day developmental growth, and they additionally secrete a panel of effector proteins onto the cytosolic face of the vacuole membrane and into the host cytosol. Gaps in our understanding of Chlamydia biology, however, present significant challenges for visualizing and analyzing this intracellular compartment. Recently, a reverse-imaging strategy for visualizing the inclusion using GFP expressing host cells was described. This approach rationally exploits the intrinsic impermeability of the inclusion membrane to large molecules such as GFP. In this work, we describe how GFP- or mCherry-expressing host cells are generated for subsequent visualization of chlamydial inclusions. Furthermore, this method is shown to effectively substitute for costly antibody-based enumeration methods, can be used in tandem with other fluorescent labels, such as GFP-expressing Chlamydia, and can be exploited to derive key quantitative data about inclusion membrane growth from a range of Chlamydia species and strains.

Using Fluorescent Proteins to Visualize and Quantitate Chlamydia Vacuole Growth Dynamics in Living Cells

A live cell fluorescent protein based method for illuminating cellular vacuoles (inclusions) containing Chlamydia is described. This strategy enables rapid, automated determination of Chlamydia infectivity in samples and can be used to quantitatively investigate inclusion growth dynamics.

The obligate intracellular bacterium Chlamydia elicits a great burden on global public health. C.&#160;trachomatis is the leading bacterial cause of ...

Evaluating Virulence and Pathogenesis of Aeromonas Infection in a Caenorhabditis elegans Model

The human pathogen Aeromonas has been clinically shown to cause gastroenteritis, wound infections, septicemia, and urinary tract infections. Most human diseases have been reported to be associated with four species of bacteria: Aeromonas dhakensis, Aeromonas hydrophila, Aeromonas veronii, and Aeromonas caviae. The model organism Caenorhabditis elegans is a bacterivore that provides an excellent infection model by which to study the bacterial pathogenesis of Aeromonas. Here, we introduce three different experiments to study Aeromonas infection using a C. elegans model, including survival, liquid toxicity, and muscle necrosis assays. The results of the three methods determining the virulence of Aeromonas were consistent. A. dhakensis was shown to be the most toxic among the 4 major Aeromonas species causing clinical infections. These methods are shown to be a convenient way to evaluate the toxicity among and within Aeromonas species and contribute to our understanding of the pathogenesis of Aeromonas infection.

Evaluating Virulence and Pathogenesis of Aeromonas Infection in a Caenorhabditis elegans Model

Here, we introduce three different experiments to study Aeromonas infection in C. elegans. Using these convenient methods, it is easy to evaluate the toxicity among and within Aeromonas species.

The human pathogen Aeromonas has been clinically shown to cause gastroenteritis, wound infections, septicemia, and urinary tract infections. Most human ...

Quantitative Fluorescence In Situ Hybridization (FISH) and Immunofluorescence (IF) of Specific Gene Products in KSHV-Infected Cells

Mechanistic insight arrives from careful study and quantification of specific RNAs and proteins. The relative locations of these biomolecules throughout the cell at specific times can be captured with fluorescence in situ hybridization (FISH) and immunofluorescence (IF). During lytic herpesvirus infection, the virus hijacks the host cell to preferentially express viral genes, causing changes in cell morphology and behavior of biomolecules. Lytic activities are centered in nuclear factories, termed viral replication compartments, which are discernable only with FISH and IF. Here we describe an adaptable protocol of RNA FISH and IF techniques for Kaposi's sarcoma-associated herpesvirus (KSHV)-infected cells, both adherent and in suspension. The method includes steps for the development of specific anti-sense oligonucleotides, double RNA FISH, RNA FISH with IF, and quantitative calculations of fluorescence intensities. This protocol has been successfully applied to multiple cell types, uninfected cells, latent cells, lytic cells, time-courses, and cells treated with inhibitors to analyze the spatiotemporal activities of specific RNAs and proteins from both the human host and KSHV.

Quantitative Fluorescence In Situ Hybridization FISH and Immunofluorescence IF of Specific Gene Products in KSHV-Infected Cells

We describe a protocol utilizing fluorescence in situ hybridization (FISH) to visualize multiple herpesviral RNAs within lytically infected human cells, either in suspension or adherent. This protocol includes quantification of fluorescence producing a nucleocytoplasmic ratio and can be extended for simultaneous visualization of host and viral proteins with immunofluorescence (IF).

Mechanistic insight arrives from careful study and quantification of specific RNAs and proteins. The relative locations of these biomolecules throughout ...

Visualization of SARS-CoV-2 using Immuno RNA-Fluorescence In Situ Hybridization

This manuscript provides a protocol for in situ hybridization chain reaction (HCR) coupled with immunofluorescence to visualize severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) RNA in cell line&#160;and three-dimensional (3D) cultures of human airway epithelium. The method allows highly specific and sensitive visualization of viral RNA by relying on HCR initiated by probe localization. Split-initiator probes help amplify the signal by fluorescently labeled amplifiers, resulting in negligible background fluorescence in confocal microscopy. Labeling amplifiers with different fluorescent dyes facilitates the simultaneous recognition of various targets. This, in turn, allows the mapping of the infection in tissues to better understand viral pathogenesis and replication at the single-cell level. Coupling this method with immunofluorescence may facilitate better understanding of host-virus interactions, including alternation of the host epigenome and immune response pathways. Owing to sensitive and specific HCR technology, this protocol can also be used as a diagnostic tool. It is also important to remember that the technique may be modified easily to enable detection of any RNA, including non-coding RNAs and RNA viruses that may emerge in the future.

Here, we describe a simple method that combines RNA fluorescence in situ&#160;hybridization (RNA-FISH) with immunofluorescence to visualize severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2) RNA. This protocol may increase understanding of the molecular characteristics of SARS-CoV-2 RNA-host interactions at a single-cell level.

This manuscript provides a protocol for in situ hybridization chain reaction (HCR) coupled with immunofluorescence to visualize severe acute respiratory ...

Visualization of Gut Microbiota-host Interactions via Fluorescence In Situ Hybridization, Lectin Staining, and Imaging

Measuring the localization of microbes within their in vivo context is an essential step in revealing the functional relationships between the microbiota and the vertebrate gut. The spatial landscape of the gut microbiota is tightly controlled by physical features - intestinal mucus, crypts, and folds - and is affected by host-controlled properties such as pH, oxygen availability, and immune factors. These properties limit the ability of commensal microbes and pathogens alike to colonize the gut stably. At the micron-scale, microbial organization determines the close-range interactions between different microbes as well as the interactions between microbes and their host. These interactions then affect large-scale organ function and host health.
This protocol enables the visualization of the gut microbiota spatial organization from distances between cells to organ-wide scales. The method is based on fixing gut tissues while preserving intestinal structure and mucus properties. The fixed samples are then embedded, sectioned, and stained to highlight specific bacterial species through fluorescence in situ hybridization (FISH). Host features, such as mucus and host cell components, are labeled with fluorescently labeled lectins. Finally, the stained sections are imaged using a confocal microscope utilizing tile-scan imaging at high magnification to bridge the micron to centimeter length scales. This type of imaging can be applied to intestinal sections from animal models and biopsies from human tissues to determine the biogeography of the microbiota in the gut in health and disease.

Visualization of Gut Microbiota-host Interactions via Fluorescence In Situ Hybridization, Lectin Staining, and Imaging

This streamlined protocol details a workflow to detect and image bacteria in complex tissue samples, from fixing the tissue to staining microbes with fluorescent in situ hybridization.

Measuring the localization of microbes within their in vivo context is an essential step in revealing the functional relationships between the microbiota ...