The authors wish to thank Eugenio Vivas, Kurt Heungens, Eric Martens, Charles Cowles, Darby Sugar, Eric Stabb, and Todd Ciche for their contributions to the development of this protocol and tools used. KEM and JMC were supported by National Institutes of Health (NIH) National Research Service Award T32 (AI55397 "Microbes in Health and Disease"). JMC was supported by a National Science Foundation (NSF) Graduate Research Fellowship. This work was supported by grants from the National Science Foundation (IOS-0920631 and IOS- 0950873).

1. Construction of a Fluorescent Bacterial Strain via Conjugation
<ol>
	<li>Grow the recipient strain (symbiont to be examined) and donor strain overnight. The donor strain, usually Escherichia coli, should be capable of donating DNA through conjugation and should be transformed with a plasmid (Table 2) that carries a gene encoding a fluorescent protein. Depending on the plasmid, a conjugation helper strain may also be required. If so, this strain should also be grown overnight. The donor strain and helper strain should be grown with antibiotics to select for maintenance of the plasmid.</li>
	<li>Subculture the donor, helper, and recipient strains into nutrient rich growth media lacking antibiotics in a 1:100 ratio of culture to medium.</li>
	<li>Grow the cultures at the temperature appropriate for each strain until they reach mid-log stage growth (OD600~0.6). For Xenorhabdus and E. coli this stage of growth is reached approximately 3 to 4 hr after subculturing. This may vary depending on the strain, or in some cases the plasmid, used.</li>
	<li>Mix the strains in a microcentrifuge tube and centrifuge for 2 min at 17,900 x g (13,000 rpm in most microfuges). The proportion of donor and recipient that yields best results will vary for different combinations, and may have to be empirically determined. For a Xenorhabdus recipient and E. coli donor, we use a 3:1 or 1:1 ratio. If a helper strain is needed (e.g. E. coli), it should be added in the same ratio as the donor.</li>
	<li>Decant the supernatant.</li>
	<li>Re-suspend the cell pellet in 30 &mu;l of fresh media.</li>
	<li>Spot the suspension onto a nutrient rich media plate without any antibiotics, and allow the spot to dry.</li>
	<li>Incubate the plate, inverted, overnight at a temperature optimal for the recipient bacterium and permissible for the donor (and helper, if applicable). The plate should be incubated at least 18 hr.</li>
	<li>Scrape up the spot and streak for single colonies on a selective antibiotic plate. The antibiotics should select against the donor and for the recipient containing the plasmid. One or two additional isolation streaks may be necessary to isolate a pure culture.</li>
	<li>Ensure the resulting colonies are the recipient symbiont, and not the donor strain, by screening for the presence of recipient specific phenotypes, such as colony morphology, or polymerase chain reaction detection of recipient-specific genes. Screen the colonies for presence of the plasmid by polymerase chain reaction of a known plasmid sequence and fluorescence. The colonies should fluoresce under the correct wavelength corresponding to the fluorescent protein.</li>
</ol>
2. Production of Axenic Nematode Eggs
<ol>
	<li>Grow 5 ml of the natural bacterial symbiont overnight. To start a culture, bacteria may be inoculated into Lysogeny Broth (LB) or other culture media directly from a freezer stock or from a streak plate.</li>
	<li>Spread 600 &mu;l of the bacterial culture onto 10 mm Lipid Agar (LA) plates 29. Eight to ten plates will yield enough nematodes for most species.</li>
	<li>Incubate in the dark without moisture at 25 &deg;C for two days.</li>
	<li>Add 5,000 infective juvenile nematodes, or other stages depending on the nematode being used, in 500 &mu;l of media to the bacterial lawns. Incubate the plates at 25 &deg;C for 3 days.</li>
	<li>Check your plates for the presence of adult nematodes and eggs. Place 20 &mu;l of water onto a microscope slide. Using a sterile stick scrape-up a small amount of nematodes from the bacterial lawn. Place the stick into the water and allow the nematodes to swim off. Look at your slide under low magnification. For more information on appearance see the discussion and Figure 2.</li>
	<li>If eggs and females are visible, continue; otherwise, incubate the plates at 25 &deg;C and check for proper development every 6-12 hr. When females contain eggs, place a few ml of water on the surface of the plates. Gently swirl the plates and pour the water into a 50 ml conical tube. Nematodes should come off of the plates and no longer be visible on the plate surface. Several rinses may be necessary.</li>
	<li>Allow the nematodes to settle to the bottom of the tube.</li>
	<li>Rinse nematodes. Pipette off the excess water from the top of the tube. Refill with clean water and allow adult nematodes to settle. Pipette off the excess water.</li>
	<li>Fill the conical tubes with egg solution. Once egg solution is added the timing of all steps with egg solution must proceed exactly as described for maximum egg yield. Incubate the tubes while gently shaking for exactly 10 min. Most nematodes should be dissolved by the end of the incubation.</li>
	<li>Immediately centrifuge the conical tubes at 1250 x g for exactly 10 min (this includes bringing the centrifuge up to speed but not down to 0 x g. Use the brake).</li>
	<li>Immediately decant the supernatant.</li>
	<li>Immediately re-suspend the pellet with egg solution by pipetting.</li>
	<li>Immediately fill conical tube with egg solution and mix well by inverting 3-5 times. Then immediately spin as in 2.10.</li>
	<li>Immediately decant the supernatant.</li>
	<li>Re-suspend the pellet in LB by pipetting, transfer to a 15 ml conical tube for improved pelleting during wash steps. LB is standard for Steinernema spp. Other buffers (e.g. a minimal salts medium) may be used depending on the nematode and bacterium, keeping in mind the buffer must not harm either the nematode or bacterium.</li>
	<li>Fill the tube with LB and spin as in 2.10.</li>
	<li>Decant the supernatant, and re-suspend in LB.</li>
	<li>Rinse the nematodes a total of 3 times by repeating 2.14 and 2.15.</li>
	<li>Dilute the re-suspended nematode eggs to an appropriate volume. There should be at least 10 eggs per microliter. Eggs can be stored in a 6-cm Petri dish in 5 ml LB for at least four days by adding antibiotics that inhibit growth of colonizing bacteria and wrapping the plate in parafilm. The eggs will need to be washed once in 15 ml LB (as in 2.13 and 2.14) prior to inoculating onto bacterial lawns. Note that eggs will hatch during this time and may not be synchronized developmentally when used in colonization assays.</li>
</ol>
3. Co-cultivation Assay with Fluorescent Bacteria
<ol>
	<li>Grow fluorescent bacteria overnight, selecting for the plasmid if necessary.</li>
	<li>Spread 600 &mu;l of the bacterial culture onto 10 mm Lipid Agar plates with a sterile stick. Incubate at 25 &deg;C for 2 days.</li>
	<li>Place 500-5,000 axenic nematode eggs in a total of 100 to 400 &mu;l (optimally 2,000 nematodes in 200 &mu;l) onto each lipid agar plate.</li>
	<li>Incubate in the dark without moisture at 25 &deg;C until IJs, or other stages of interest, form. IJs will be visible as a fuzzy white ring on the edge of the plate. To look at other life stages, see Protocol 4.</li>
	<li>Place the plates in a modified White trap30. Remove the lid of the lipid agar plate and place the bottom into the bottom of an empty 100 mm x 20 mm Petri dish. Fill the large Petri dish with water, or other buffer appropriate to your nematode, to surround the small plate. The water level should be approximately half the height of the small plate.</li>
	<li>Incubate the water trap until progeny IJs have emerged into the buffer.</li>
	<li>Infective juvenile nematodes can be stored in water in a tissue culture flask.</li>
</ol>
4. Collection of Early Life Stages for Screening
<ol>
	<li>Use steps 3.1 through 3.4 to set up the assay.</li>
	<li>Incubate the plates until the desired life stages are present. Daily visual inspection may be necessary to determine timing. To inspect the plates, use the procedure described in Protocol 2.5. For S. carpocapsae eggsgrown with X. nematophila on LA plates, gravid females will be present in 4-5 days, juveniles will be present in 7 days, and infective juveniles will start to be produced by 15-17 days.</li>
	<li>When the desired life stages are present, collect your sample. Fill a well of a 96 well plate with 200 &mu;l of PBS or other appropriate buffer. Gently scrape off some of the bacterial lawn that contains nematodes. A pea-sized amount (~50 mg) will provide at least several hundred nematodes once F1 progeny are produced, but more may be necessary for earlier time points. Place the stick into the well containing buffer.</li>
	<li>Incubate for 30 sec to a minute. Remove the stick and scrape off the remaining residue. Use the stick to remove any agar from the well. The nematodes should be visible within the buffer. Move the buffer mixture to a clean microfuge tube.</li>
	<li>Treat the nematodes with levamisole or other paralyzing agent. Dissolve a few grains of levamisole into 30 &mu;l of water. Add 1-2 &mu;l of this mix for each 50 &mu;l of sample. After levamisole treatment the nematodes will be non-viable.</li>
	<li>If samples will be viewed at a later date, fix as described in this step. If not, skip to step 4.6. Add 200 &mu;l of fixative solution containing 1X buffer and 4% paraformaldehyde. Mix gently, pipetting may cause delicate life stages to lyse. Cover with foil, and incubate at room temperature on a shaker on low for at least 18 hr.</li>
	<li>Place the tubes in a tube rack, and allow the nematodes to settle to the bottom of the tube.</li>
	<li>Rinse nematodes to remove background. Pipette off excess liquid and add 200 - 500 &mu;l of buffer and gently mix. Repeat. This may be repeated a few times to remove more background if desired.</li>
	<li>Allow the nematodes to settle to the bottom of the tube and re-suspend in the desired volume.</li>
	<li>If the nematodes are not fixed, screen immediately. Fixed nematodes may be stored in the refrigerator for up to two weeks. Store in the dark.</li>
</ol>
5. Screening Nematodes for Bacterial Association by Microscopy
<ol>
	<li>Select some nematodes to view under the microscope by removing a small sample of your mixture.</li>
	<li>To ensure that the nematodes are still for the picture, treat with a paralytic agent as described in Protocol 4.5. If you are not taking a picture or the nematodes are already fixed, this step may be skipped.</li>
	<li>Add 20-30 &mu;l of nematodes in water to your microscope slide and place a coverslip on top.</li>
	<li>View whole nematodes using light microscopy to ensure nematodes are in the field of view.</li>
	<li>View nematodes using the fluorescent setting on the microscope that corresponds to the fluorescence expressed by the bacteria.</li>
	<li>To identify bacterial localization, take photos of the nematode under fluorescent setting and a light microscopy setting, and then superimpose the images. The photos need to be of the same field of view. It may be necessary to try several magnifications and views of the nematode to find the bacteria.</li>
	<li>The distribution of nematode colonization can be quantified by counting a population of nematodes, scoring for the presence or absence of bacteria, and calculating the percent colonized. We recommend counting until at least 30 nematodes within the population are counted in each category (with or without colonization) to obtain a reliable value.</li>
	<li>When only a few nematodes in the population are colonized, it may be necessary to count several thousand nematodes before 30 are observed. For high-throughput counting use the following steps.</li>
	<li>Instead of counting nematodes individually, aliquot the population into a multi-well plate (e.g. a 24-well plate). After the nematodes have settled to the bottom of the dish, each well can be scanned on the microscope and the number of colonized nematodes can be scored.</li>
	<li>The total number of nematodes in a population can be identified by serial dilutions of nematodes in the well. For example, 1 ml of nematodes can be added to a well, mixed well by stirring with a pipet tip, and 3 &mu;l can be removed and counted for quantification of the total population in the well.</li>
	<li>The percentage of colonized nematodes can be obtained by dividing the number colonized by the total number of nematodes in the well.</li>
</ol>
6. Representative Results
Example microscope images of Steinernema nematodes associated with Xenorhabdus bacteria are shown in Figure 3. To create the composite image seen in Figure 3A, a phase contrast image was overlaid with a fluorescent image. The arrow in Figure 3A indicates the bacteria present within the infective juvenile nematode (bar = 100 &mu;m). Figure 3B was constructed in a similar manner and depicts a juvenile nematode with green fluorescent protein labeled bacteria (green rods) localized throughout the nematode intestinal lumen (bar = 20 &mu;m). A population of the nematodes from two media were counted and scored for colonization by the bacterial symbiont (Table 1). For robust statistics, it is best to count at least 100 nematodes per sample with at least 30 falling in each category. As seen in Table 1, these nematodes are colonized at a level of approximately 14.6% when grown on lipid agar and 68.6% when grown on liver kidney agar. Other nematode and bacterial species have been shown to have different levels of colonization. For example X. nematophila colonizes 99% of S. carpocapsae infective juveniles (Martens 2003), and P. luminescens colonizes 26% of H. bacteriophora infective juveniles (Ciche 2003).
<img alt="Figure 1" src="/files/ftp_upload/4298/4298fig1.jpg" /> 
 Figure 1. Schematic outline of the method. A. The bacterium is engineered to express a fluorescent protein. B. Nematode eggs are isolated from adult nematodes to produce sterile nematodes. C. The sterile nematodes are co-cultivated with the fluorescent bacteria. D. The resulting life stages are viewed under a microscope to evaluate bacterial presence within the nematode.
<img alt="Figure 2" src="/files/ftp_upload/4298/4298fig2.jpg" /> 
 Figure 2. Depiction of adult females containing eggs. A. The schematic shows the general appearance of Steinernema females. Inset: DIC image of a S. feltiae gravid female. The black arrow indicates the vulva. White arrows show visible eggs. Image is at 20X magnification, and the scale bar represents 100 &mu;m. B. DIC image of a developed but unhatched S. feltiae nematode egg. Image is 40X magnification, and the scale bar represents 50 &mu;m. C. DIC image of eggs isolated from S. feltiae nematodes under 10X magnification. Scale bar is 100 &mu;m.
<img alt="Figure 3" src="/files/ftp_upload/4298/4298fig3.jpg" /> 
 Figure 3.Example microscope images of nematode-bacterial association. A. S. puntauvense nematodes were associated with their bacterial symbiont, X. bovienii, expressing GFP. The image is a composite image produced by overlaying a phase contrast image with a fluorescent image from the same field of view. The arrow indicates the fluorescent bacterial symbiont within the nematode host. Scale bar represents 100 &mu;m. B. This image shows a S. carpocapsae juvenile nematode with GFP-expressing X. nematophila localized within the nematode intestine. This image was constructed through overlaying a fluorescent image over a differential interference contrast image. The scale bar represents 20 &mu;m.
<table border="1">
	<tbody>
		<tr>
			<td>Strain</td>
			<td>Number of Nematodes With acteria</td>
			<td>Total Nematodes Counted</td>
			<td>Percent of Nematodes olonized</td>
		</tr>
		<tr>
			<td>S. puntauvense Lipid Agar</td>
			<td>30</td>
			<td>205</td>
			<td>14.60%</td>
		</tr>
		<tr>
			<td>S. puntauvense Liver Kidney Agar</td>
			<td>72</td>
			<td>105</td>
			<td>68.60%</td>
		</tr>
	</tbody>
</table>
Table 1. Example scoring of a nematode population for bacterial presence. In this experiment, axenic S. puntauvense nematodes were grown with their GFP-expressing symbiont on different growth media (lipid agar and liver kidney agar 32) to test for colonization defects. A total of at least one hundred nematodes per sample were counted and scored for the presence of bacteria. For statistical power, three experimental replicates should be counted with at least 30 nematodes falling in each category.
<img alt="Table 2" src="/files/ftp_upload/4298/4298table2.jpg" /> 
Table 2. Fluorescent protein containing plasmids. A list of potential plasmids for insertion of a fluorescent protein into the bacterial symbiont is given listed by name of plasmid. Other information included are the fluorescent protein encoded, antibiotic cassette used for plasmid maintenance, other instructions for use, the source of the plasmid. The concentration noted in parentheses is the concentration of the antibiotic used for X. nematophila. Each of these plasmids has been used successfully in either Xenorhabdus or Photorhabdus. Additional information can be obtained from the noted citations. Depending on the bacterium being tested, some plasmids may not work based upon the fluorescent protein, antibiotic selection, insertion site, or origin of replication. The plasmids listed above contain different features that may enable use in the bacterium of interest. For example, mini-Tn7-KSGFP inserts into the attTn7 site of the chromosome, while pECM20 inserts into the X. nematophila chromosome by homologous recombination. Alternatively, the pPROBE plasmids are maintained extrachromosomally, and each pPROBE plasmid has the same backbone and fluorophore but have different selectable markers or origins of replication to enable their use in a variety of taxonomic or mutant backgrounds.

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E., Goodrich-Blair, H. <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+Xenorhabdus+nematophila+genes+required+for+mutualistic+colonization+of+Steinernema+carpocapsae+nematodes.">Identification of Xenorhabdus nematophila genes required for mutualistic colonization of Steinernema carpocapsae nematodes.</a> Mol. Microbiol. 45, 1337-1353 (2002).</li><li>Goetsch, M., Owen, H., Goldman, B., Forst, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+the+PixA+inclusion+body+protein+of+Xenorhabdus+nematophila.">Analysis of the PixA inclusion body protein of Xenorhabdus nematophila.</a> J. Bacteriol. 188, 2706-2710 (2006).</li><li>Bhasin, A., Chaston, J. M., Goodrich-Blair, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mutational+Analyses+Reveal+Overall+Topology+and+Functional+Regions+of+NilB,+a+Bacterial+Outer+Membrane+Protein+Required+for+Host+Association+in+a+Model+of+Animal-Microbe+Mutualism.">Mutational Analyses Reveal Overall Topology and Functional Regions of NilB, a Bacterial Outer Membrane Protein Required for Host Association in a Model of Animal-Microbe Mutualism.</a> J. Bacteriol. 194, 1763-1776 (2012).</li><li>Ciche, T. A., Goffredi, S. K., Reddy, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Methods in General and Molecular Microbiology. , 394-419 (2007).</li><li>Goodrich-Blair, H., Clarke, D., Grewal, P. S., Ciche, T. A., Stock, S. P., Boemare, N., Vandenberg, J., Glazar, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Insect Pathogens, Molecular Approaches and Techniques. , 239-269 (2009).</li><li>Stock, S. P., Goodrich-Blair, H., Lacey, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Manual of Techniques in Invertebrate Pathology. , 375-425 (2012).</li><li>Martens, E. C., Gawronski-Salerno, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Xenorhabdus+nematophila+requires+an+intact+iscRSUA-hscBA-fdx+locus+to+colonize+Steinernema+carpocapsae+nematodes.">Xenorhabdus nematophila requires an intact iscRSUA-hscBA-fdx locus to colonize Steinernema carpocapsae nematodes.</a> J. Bacteriol. 185, 3678-3682 (2003).</li><li>Vivas, E. I., Goodrich-Blair, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Xenorhabdus+nematophilus+as+a+model+for+host-bacterium+interactions:+rpoS+is+necessary+for+mutualism+with+nematodes.">Xenorhabdus nematophilus as a model for host-bacterium interactions: rpoS is necessary for mutualism with nematodes.</a> J. Bacteriol. 183, 4687-4693 (2001).</li><li>White, G. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+method+for+obtaining+infective+nematode+larvae+from+cultures.">A method for obtaining infective nematode larvae from cultures.</a> Science. 66, 302-303 (1927).</li><li>Ciche, T. A., Ensign, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=For+the+insect+pathogen+Photorhabdus+luminescens,+which+end+of+a+nematode+is+out.">For the insect pathogen Photorhabdus luminescens, which end of a nematode is out.</a> Appl. Environ. Microbiol. 69, 1890-1897 (2003).</li><li>Sicard, M., Brun, N. L. e. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+native+Xenorhabdus+on+the+fitness+of+their+Steinernema+hosts:+contrasting+types+of+interaction.">Effect of native Xenorhabdus on the fitness of their Steinernema hosts: contrasting types of interaction.</a> Parasitol. Res. 91, 520-524 (2003).</li><li>Lambertsen, L., Sternberg, C., Molin, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mini-Tn7+transposons+for+site-specific+tagging+of+bacteria+with+fluorescent+proteins.">Mini-Tn7 transposons for site-specific tagging of bacteria with fluorescent proteins.</a> Environ. Microbiol. 6, 726-732 (2004).</li><li>Miller, W. G., Leveau, J. H., Lindow, S. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improved+gfp+and+inaZ+broad-host-range+promoter-probe+vectors.">Improved gfp and inaZ broad-host-range promoter-probe vectors.</a> Mol. Plant Microbe Int. 13, 1243-1250 (2000).</li><li>Teal, T. K., Lies, D. P., Wold, B. J., Newman, D. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatiometabolic+stratification+of+Shewanella+oneidensis+biofilms.">Spatiometabolic stratification of Shewanella oneidensis biofilms.</a> Appl. Environ. Microbiol. 72, 7324-7330 (2006).</li><li>Bao, Y., Lies, D. P., Fu, H., Roberts, G. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+improved+Tn7-based+system+for+the+single-copy+insertion+of+cloned+genes+into+chromosomes+of+Gram-negative+bacteria.">An improved Tn7-based system for the single-copy insertion of cloned genes into chromosomes of Gram-negative bacteria.</a> Gene. 109, 167-168 (1991).</li><li>Woodring, J. L., Kaya, H. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Steinernematid+and+Heterorhabditid+Nematodes:+A+Handbook+of+Biology+and+Techniques.">Steinernematid and Heterorhabditid Nematodes: A Handbook of Biology and Techniques.</a> Southern Cooperative Series Bulletin 331. , (1988).</li></ol>

No conflicts of interest declared.

The protocol described here provides a method for the optical detection of bacteria within a nematode host (Figure 1). This method takes advantage of the optical transparency of nematodes and the ability to fluorescently label bacteria, enabling in vivo analysis of bacteria within the nematode host (Figure 3). Specifically, this approach identifies bacterial localization within its host. By counting a nematode population and scoring for bacterial presence, the frequency of bacte...

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>Lipid Agar 
 (sterile)</td>
 <td></td>
 <td></td>
 <td>8 grams nutrient broth, 15 grams agar, 5 grams yeast extract, 890 ml water, 10 ml 0.2 g/ml MgCl2. 6H20, 96 ml corn syrup solution*, 4 ml corn oil* 
 Stir media while pouring plates 
 *add sterile ingredient after autoclaving </td>
 </tr>
 <tr>
 <td>Corn Syrup Solution 
 (sterile)</td>
 <td></td>
 <td></td>
 <td>7 ml corn syrup, 89 ml water 
 mix and autoclave</td>
 </tr>
 <tr>
 <td>Egg Solution</td>
 <td></td>
 <td></td>
 <td>16.6 ml 12% sodium hypochlorite, 5 ml 5M KOH, 80 ml water</td>
 </tr>
 <tr>
 <td>Lysogeny Broth 
 (sterile)</td>
 <td></td>
 <td></td>
 <td>5 grams yeast extract, 10 grams tryptone, 5 grams salt, 1 L water 
 mix and autoclave</td>
 </tr>
 <tr>
 <td>Microfuge</td>
 <td>Fisher</td>
 <td>13-100-675</td>
 <td>Any microfuge that holds microfuge tubes will work</td>
 </tr>
 <tr>
 <td>Centrifuge</td>
 <td>Beckman</td>
 <td>366802 </td>
 <td>Large table top centrifuge that holds 15 ml and 50 ml conical tubes</td>
 </tr>
 <tr>
 <td>Sterile 60 mm X 15 mm Petri Dish</td>
 <td>Fisher</td>
 <td>0875713</td>
 <td></td>
 </tr>
 <tr>
 <td>50 ml centrifuge tubes</td>
 <td>Fisher</td>
 <td>05-539-6</td>
 <td></td>
 </tr>
 <tr>
 <td>15 ml centrifuge tubes</td>
 <td>Fisher</td>
 <td>05-531-6</td>
 <td></td>
 </tr>
 <tr>
 <td>Sterile 100 mm X 20 mm Petri Dish</td>
 <td>Fisher</td>
 <td>0875711Z</td>
 <td>Deeper than standard Petri dishes</td>
 </tr>
 <tr>
 <td>24-well plate</td>
 <td>Greiner Bio-One</td>
 <td>662000-06</td>
 <td></td>
 </tr>
 <tr>
 <td>Microscope</td>
 <td></td>
 <td></td>
 <td>The microscope needs florescent capabilities compatible with your fluorophore </td>
 </tr>
 <tr>
 <td>Paraformaldehyde</td>
 <td>Electron Microscopy Sciences</td>
 <td>15710</td>
 <td></td>
 </tr>
 <tr>
 <td>PBS 
 (sterile)</td>
 <td></td>
 <td></td>
 <td>8 g NaCL 
 0.2 g KCL 
 1.44 g Na2HPO4 
 0.24 g KH2PO4 
 1 L water Adjust to a pH of 7.4 and water to 1 L and autoclave</td>
 </tr>
 <tr>
 <td>Microfuge tubes</td>
 <td>Fisher</td>
 <td>05-408-138</td>
 <td>2 ml or 1.5 ml tubes</td>
 </tr>
 <tr>
 <td>Shaker</td>
 <td></td>
 <td></td>
 <td>Any shaker that causes the liquid to gently move will work</td>
 </tr>
 <tr>
 <td>Diaminopimelic acid</td>
 <td>Sigma</td>
 <td>D-1377</td>
 <td>If needed, supplement media to a concentration or 1 mM</td>
 </tr>
 </tbody>
</table>

visualizing bacteria in nematodes using fluorescent microscopy

Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on earth, nematodes and bacteria form a wide array of symbiotic associations that range from beneficial to pathogenic 1-3. One such association is the mutually beneficial relationship between Xenorhabdus bacteria and Steinernema nematodes, which has emerged as a model system of symbiosis 4. Steinernema nematodes are entomopathogenic, using their bacterial symbiont to kill insects 5. For transmission between insect hosts, the bacteria colonize the intestine of the nematode's infective juvenile stage 6-8. Recently, several other nematode species have been shown to utilize bacteria to kill insects 9-13, and investigations have begun examining the interactions between the nematodes and bacteria in these systems 9.
We describe a method for visualization of a bacterial symbiont within or on a nematode host, taking advantage of the optical transparency of nematodes when viewed by microscopy. The bacteria are engineered to express a fluorescent protein, allowing their visualization by fluorescence microscopy. Many plasmids are available that carry genes encoding proteins that fluoresce at different wavelengths (i.e. green or red), and conjugation of plasmids from a donor Escherichia coli strain into a recipient bacterial symbiont is successful for a broad range of bacteria. The methods described were developed to investigate the association between Steinernema carpocapsae and Xenorhabdus nematophila 14. Similar methods have been used to investigate other nematode-bacterium associations 9,15-18and the approach therefore is generally applicable.
The method allows characterization of bacterial presence and localization within nematodes at different stages of development, providing insights into the nature of the association and the process of colonization 14,16,19. Microscopic analysis reveals both colonization frequency within a population and localization of bacteria to host tissues 14,16,19-21. This is an advantage over other methods of monitoring bacteria within nematode populations, such as sonication 22or grinding 23, which can provide average levels of colonization, but may not, for example, discriminate populations with a high frequency of low symbiont loads from populations with a low frequency of high symbiont loads. Discriminating the frequency and load of colonizing bacteria can be especially important when screening or characterizing bacterial mutants for colonization phenotypes 21,24. Indeed, fluorescence microscopy has been used in high throughput screening of bacterial mutants for defects in colonization 17,18, and is less laborious than other methods, including sonication 22,25-27and individual nematode dissection 28,29.

Watch this Scientific Journal Video about Visualizing Bacteria in Nematodes using Fluorescent Microscopy at JoVE.com

Visualizing Bacteria in Nematodes using Fluorescent Microscopy

To study the mutualism between Xenorhabdus bacteria and Steinernema nematodes, methods were developed to monitor bacterial presence and location within nematodes. The experimental approach, which can be applied to other systems, entails engineering bacteria to express the green fluorescent protein and visualizing, using fluorescence microscopy bacteria within the transparent nematode.

 
Symbioses, the living together of two or more organisms, are widespread throughout all kingdoms of life. As two of the most ubiquitous organisms on ...

visualizing-bacteria-in-nematodes-using-fluorescent-microscopy

Research

JoVE Journal

Biology

18.7K Views.  University of Wisconsin-Madison. Per studiare il mutualismo tra Xenorhabdus Batteri e Steinernema, i metodi sono stati sviluppati per monitorare la presenza di batteri e posizione all'interno nematodi. L'approccio sperimentale, che può essere applicata ad altri sistemi, comporta batteri ingegneria per esprimere la proteina fluorescente verde e visualizzazione, utilizzando batteri microscopia a fluorescenza all'interno del nematode trasparente.

Video: Visualizing Bacteria in Nematodes using Fluorescent Microscopy

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle synapses, we developed a larval tissue preparation that had features of live intact larvae, but also had good optical properties.  This new preparation also allowed for access of perfusates to the synapse.  We used fly larvae, immersed them in artificial hemolymph, and relaxed their normal rhythmic body contractions by chilling them. Next we dissected off the posterior segments of each animal and with a blunt insect pin pushed the mouth parts backward through the body cavity.  This everted the larval body wall, like turning a sock inside-out.  We completed the dissection with ultra-fine dissection scissors and thus exposed the visceral side of the body wall muscles. The glial structures at the NMJ expressed membrane targeted GFP under the control of glial specific promoters. The post-synaptic membrane, the SSR (Subsynaptic Reticula) in muscle expressed synaptically targeted dsRed. We needed to acutely label the motor neuron terminals, the third part of the synapse. To do this we applied primary antibodies to HRP, conjugated to a far-red emitting flurophore.  To test for dye diffusion properties into the perisynaptic space between the motor neuron terminals and the SSR, we applied a solution of large Dextran molecules conjugated to far-red emitting flurophore and collected images.

Visualizing the Live Drosophila Glial-neuromuscular Junction with Fluorescent Dyes

We described structural features of the Glia-neuromuscular synapses in a novel Inside-out tissue preparation of live fly larvae using fluorescent dyes with confocal microscopy. We labeled live neuron terminals with fluorescent primary antibodies to HRP, and also visualized the perisynaptic space with fluorescent Dextrans.

Our project identified GFP labeled glial structures at the developing larval fly neuromuscular synapse. To look at development of live glial-nerve-muscle ...

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular level. To date the shear complexity of biological systems has caused precise single-molecule experimentation to be far too demanding, instead focusing on studies of single systems using relatively crude bulk ensemble-average measurements. However, many important processes occur in the living cell at the level of just one or a few molecules; ensemble measurements generally mask the stochastic and heterogeneous nature of these events. Here, using advanced optical microscopy and analytical image analysis tools we demonstrate how to monitor proteins within a single living bacterial cell to a precision of single molecules and how we can observe dynamics within molecular complexes in functioning biological machines. The techniques are directly relevant physiologically. They are minimally-perturbative and non-invasive to the biological sample under study and are fully attuned for investigations in living material, features not readily available to other single-molecule approaches of biophysics. In addition, the biological specimens studied all produce fluorescently-tagged protein at levels which are almost identical to the unmodified cell strains ("genomic encoding"), as opposed to the more common but less ideal approach for generating significantly more protein than would occur naturally ('plasmid expression'). Thus, the actual biological samples which will be investigated are significantly closer to the natural organisms, and therefore the observations more relevant to real physiological processes.

Visualizing Single Molecular Complexes In Vivo Using Advanced Fluorescence Microscopy

Here we demonstrate the protocols for performing single-molecule fluorescence microscopy on living bacterial cells to enable functional molecular complexes to be detected, tracked and quantified.

Full insight into the mechanisms of living cells can be achieved only by investigating the key processes that elicit and direct events at a cellular ...

Visualizing Single-molecule DNA Replication with Fluorescence Microscopy

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA template is attached to a functionalized glass coverslip and replicated extensively after introduction of replication proteins and nucleotides (Figure 1). The growing product double-strand DNA (dsDNA) is extended with laminar flow and visualized by using an intercalating dye. Measuring the position of the growing DNA end in real time allows precise determination of replication rate (Figure 2). Furthermore, the length of completed DNA products reports on the processivity of replication. This experiment can be performed very easily and rapidly and requires only a fluorescence microscope with a reasonably sensitive camera.


This protocol demonstrates a simple single-molecule fluorescence microscopy technique for visualizing DNA replication by individual replisomes in real time.

We describe a simple fluorescence microscopy-based real-time method for observing DNA replication at the single-molecule level. A circular, forked DNA ...

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy (f3D-SIM)

Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the diffraction of light allowing super-resolution of live samples. There are currently three main types of super-resolution techniques &#8211; stimulated emission depletion (STED), single-molecule localization microscopy (including techniques such as PALM, STORM, and GDSIM), and structured illumination microscopy (SIM). While STED and single-molecule localization techniques show the largest increases in resolution, they have been slower to offer increased speeds of image acquisition. Three-dimensional SIM (3D-SIM) is a wide-field fluorescence microscopy technique that offers a number of advantages over both single-molecule localization and STED. Resolution is improved, with typical lateral and axial resolutions of 110 and 280 nm, respectively and depth of sampling of up to 30 &#181;m from the coverslip, allowing for imaging of whole cells. Recent advancements (fast 3D-SIM) in the technology increasing the capture rate of raw images allows for fast capture of biological processes occurring in seconds, while significantly reducing photo-toxicity and photobleaching. Here we describe the use of one such method to image bacterial cells harboring the fluorescently-labelled cytokinetic FtsZ protein to show how cells are analyzed and the type of unique information that this technique can provide.

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy f3D-SIM

Spatiotemporal information about dynamic proteins inside live cells is crucial for understanding biology. A type of super-resolution microscopy called fast 3D-structured illumination microscopy (f3D-SIM) reveals unique information about the cytokinetic Z ring in bacteria: both its bead-like appearance and the rapid dynamics of FtsZ within the ring.

Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

Visualizing Stromule Frequency with Fluorescence Microscopy

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but whose functions remain mysterious. Alongside growing attention on the role of chloroplasts in coordinating plant responses to stress, interest in stromules and their relationship to chloroplast signaling dynamics has increased in recent years, aided by advances in fluorescence microscopy and protein fluorophores that allow for rapid, accurate visualization of stromule dynamics. Here, we provide detailed protocols to assay stromule frequency in the epidermal chloroplasts of Nicotiana benthamiana, an excellent model system for investigating chloroplast stromule biology. We also provide methods for visualizing chloroplast stromules in vitro by extracting chloroplasts from leaves. Finally, we outline sampling strategies and statistical approaches to analyze differences in stromule frequencies in response to stimuli, such as environmental stress, chemical treatments, or gene silencing. Researchers can use these protocols as a starting point to develop new methods for innovative experiments to explore how and why chloroplasts make stromules.

Protocols to investigate the dynamics of chloroplast stromules, the stroma-filled tubules that extend from the surface of chloroplasts, are described.

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but ...

Split Green Fluorescent Protein System to Visualize Effectors Delivered from Bacteria During Infection

Bacteria, one of the most important causative agents of various plant diseases, secrete a set of effector proteins into the host plant cell to subvert the plant immune system. During infection cytoplasmic effectors are delivered to the host cytosol via a type III secretion system (T3SS). After delivery into the plant cell, the effector(s) targets the specific compartment(s) to modulate host cell processes for survival and replication of the pathogen. Although there has been some research on the subcellular localization of effector proteins in the host cells to understand their function in pathogenicity by using fluorescent proteins, investigation of the dynamics of effectors directly injected from bacteria has been challenging due to the incompatibility between the T3SS and fluorescent proteins.
Here, we describe our recent method of an optimized split superfolder green fluorescent protein system (sfGFPOPT) to visualize the localization of effectors delivered via the bacterial T3SS in the host cell. The sfGFP11 (11th &#946;-strand of sfGFP)-tagged effector secreted through the T3SS can be assembled with a specific organelle targeted sfGFP1-10OPT (1-10th &#946;-strand of sfGFP) leading to fluorescence emission at the site. This protocol provides a procedure to visualize the reconstituted sfGFP fluorescence signal with an effector protein from Pseudomonas syringae in a particular organelle in the Arabidopsis and Nicotiana benthamiana plants.

Fluorescent protein-based approaches to monitor effectors secreted by bacteria into host cells are challenging. This is due to the incompatibility between fluorescent proteins and the type-III secretion system. Here, an optimized split superfolder GFP system is used for visualization of effectors secreted by bacteria into the host plant cell.

Bacteria, one of the most important causative agents of various plant diseases, secrete a set of effector proteins into the host plant cell to subvert the ...

Visualizing Lymph Node Structure and Cellular Localization using Ex-Vivo Confocal Microscopy

Lymph nodes (LNs) are organs spread within the body, where the innate immune responses can connect with the adaptive immunity. In fact, LNs are strategically interposed in the path of the lymphatic vessels, allowing intimate contact of tissue antigens with all resident immune cells in the LN. Thus, understanding the cellular composition, distribution, location and interaction using ex vivo whole LN imaging will add to the knowledge on how the body coordinates local and systemic immune responses. This protocol shows an ex vivo imaging strategy following an in vivo administration of fluorescent-labeled antibodies that allows a very reproducible and easy-to-perform methodology by using conventional confocal microscopes and stock reagents. Through subcutaneous injection of antibodies, it is possible to label different cell populations in draining LNs without affecting tissue structures that can be potentially damaged by a conventional immunofluorescence microscopy technique.

This protocol describes a technique to image different cell populations in draining lymph nodes without alterations in the organ structure.

Lymph nodes (LNs) are organs spread within the body, where the innate immune responses can connect with the adaptive immunity. In fact, LNs are ...

Cryo-Electron Microscopic Grid Preparation for Time-Resolved Studies using a Novel Robotic System, Spotiton

The capture of short-lived molecular states triggered by the early encounter of two or more interacting particles continues to be an experimental challenge of great interest to the field of cryo-electron microscopy (cryo-EM). A few methodological strategies have been developed that support these &#34;time-resolved&#34; studies, one of which, Spotiton-a novel robotic system-combines the dispensing of picoliter-sized sample droplets with precise temporal and spatial control. The time-resolved Spotiton workflow offers a uniquely efficient approach to interrogate early structural rearrangements from minimal sample volume. Fired from independently controlled piezoelectric dispensers, two samples land and rapidly mix on a nanowire EM grid as it plunges toward the cryogen. Potentially hundreds of grids can be prepared in rapid succession from only a few microliters of a sample. Here, a detailed step-by-step protocol of the operation of the Spotiton system is presented with a focus on troubleshooting specific problems that arise during grid preparation.

The protocol presented here describes the use of Spotiton, a novel robotic system, to deliver two samples of interest onto a self-wicking, nanowire grid that mix for a minimum of 90 ms prior to vitrification in liquid cryogen.

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Subtitles

Visualizing Bacteria in Nematodes using Fluorescent Microscopy