We thank members of the Department of Biological Sciences at George Washington University for critical reading of the manuscript. All graphical figures were made using BioRender. Research in the I. E., J. H., and D. O&#39;H. laboratories have been supported by George Washington University and Columbian College of Arts and Sciences facilitating funds and Cross-Disciplinary Research Funds.

1. Production of symbiotic nematode infective juveniles
<ol>
	<li>Cover a Petri dish (10 cm) with a piece of filter paper and add approximately 10-15 Galleria mellonella larvae (Figure 1A).</li>
	<li>Using a pipette, dispense 2 mL of water containing about 25-50 IJs per 10 &#181;L suspension onto the waxworms. Store the Petri dish in a cabinet at room temperature.</li>
	<li>Depending on the moisture of the filter paper, add 1-2 mL of water every 2 days. Waxworms i...

<ol>
	<li>Lacey, L. 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=Insect+pathogens+as+biological+control+agents:+Back+to+the+future.">Insect pathogens as biological control agents: Back to the future.</a> Journal of Invertebrate Pathology. 132, 1-41 (2015).</li><li>Ozakman, Y., Eleftherianos, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nematode+infection+and+antinematode+immunity+in+Drosophila.">Nematode infection and antinematode immunity in Drosophila.</a> Trends in Parasitology. 37 (11), 1002-1013 (2021).</li><li>Kenney, E., Hawdon, J. M., O'Halloran, D. M., Eleftherianos, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Secreted+virulence+factors+from+Heterorhabditis+bacteriophora+highlight+its+utility+as+a+model+parasite+among+Clade+V+nematodes.">Secreted virulence factors from Heterorhabditis bacteriophora highlight its utility as a model parasite among Clade V nematodes.</a> International Journal of Parasitology. 51 (5), 321-325 (2021).</li><li>Bobardt, S. D., Dillman, A. R., Nair, M. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+two+faces+of+nematode+infection:+Virulence+and+immunomodulatory+molecules+from+nematode+parasites+of+mammals,+insects+and+plants.">The two faces of nematode infection: Virulence and immunomodulatory molecules from nematode parasites of mammals, insects and plants.</a> Frontiers in Microbiology. 11, 2983 (2020).</li><li>Castillo, J. C., Reynolds, S. E., Eleftherianos, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Insect+immune+responses+to+nematode+parasites.">Insect immune responses to nematode parasites.</a> Trends in Parasitology. 27 (12), 537-547 (2011).</li><li>Eleftherianos, I., Heryanto, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transcriptomic+insights+into+the+insect+immune+response+to+nematode+infection.">Transcriptomic insights into the insect immune response to nematode infection.</a> Genes. 12 (2), 202 (2021).</li><li>Ciche, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+biology+and+genome+of+Heterorhabditis+bacteriophora.">The biology and genome of Heterorhabditis bacteriophora.</a> WormBook. , 1-9 (2007).</li><li>Stock, S. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Partners+in+crime:+symbiont-assisted+resource+acquisition+in+Steinernema+entomopathogenic+nematodes.">Partners in crime: symbiont-assisted resource acquisition in Steinernema entomopathogenic nematodes.</a> Current Opinion in Insect Science. 32, 22-27 (2019).</li><li>Cao, M., Schwartz, H. T., Tan, C. -. H., Sternberg, P. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+entomopathogenic+nematode+Steinernema+hermaphroditum+is+a+self-fertilizing+hermaphrodite+and+a+genetically+tractable+system+for+the+study+of+parasitic+and+mutualistic+symbiosis.">The entomopathogenic nematode Steinernema hermaphroditum is a self-fertilizing hermaphrodite and a genetically tractable system for the study of parasitic and mutualistic symbiosis.</a> Genetics. 220 (1), (2021).</li><li>Goodrich-Blair, H., Clarke, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mutualism+and+pathogenesis+in+Xenorhabdus+and+Photorhabdus:+two+roads+to+the+same+destination.">Mutualism and pathogenesis in Xenorhabdus and Photorhabdus: two roads to the same destination.</a> Molecular Microbiology. 64 (2), 260-268 (2007).</li><li>Abd-Elgawad, M. M. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photorhabdus+spp.:+An+overview+of+the+beneficial+aspects+of+mutualistic+bacteria+of+insecticidal+nematodes.">Photorhabdus spp.: An overview of the beneficial aspects of mutualistic bacteria of insecticidal nematodes.</a> Plants. 10 (8), 1660 (2021).</li><li>Dreyer, J., Malan, A. P., Dicks, L. M. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bacteria+of+the+genus+Xenorhabdus,+a+novel+source+of+bioactive+compounds.">Bacteria of the genus Xenorhabdus, a novel source of bioactive compounds.</a> Frontiers in Microbiology. 9, 3177 (2018).</li><li>Hallem, E. A., Rengarajan, M., Ciche, T. A., Sternberg, P. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nematodes,+bacteria,+and+flies:+a+tripartite+model+for+nematode+parasitism.">Nematodes, bacteria, and flies: a tripartite model for nematode parasitism.</a> Current Biology. 17 (10), 898-904 (2007).</li><li>Castillo, J. C., Shokal, U., Eleftherianos, 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+novel+method+for+infecting+Drosophila+adult+flies+with+insect+pathogenic+nematodes.">A novel method for infecting Drosophila adult flies with insect pathogenic nematodes.</a> Virulence. 3 (3), 339-347 (2012).</li><li>Castillo, J. C., Shokal, U., Eleftherianos, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immune+gene+transcription+in+Drosophila+adult+flies+infected+by+entomopathogenic+nematodes+and+their+mutualistic+bacteria.">Immune gene transcription in Drosophila adult flies infected by entomopathogenic nematodes and their mutualistic bacteria.</a> Journal of Insect Physiology. 59 (2), 179-185 (2013).</li><li>Eleftherianos, I., Joyce, S., Ffrench-Constant, R. H., Clarke, D. J., Reynolds, 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=Probing+the+tri-trophic+interaction+between+insects,+nematodes+and+Photorhabdus.">Probing the tri-trophic interaction between insects, nematodes and Photorhabdus.</a> Parasitology. 137 (11), 1695-1706 (2010).</li><li>Nielsen-LeRoux, C., Gaudriault, S., Ramarao, N., Lerelcus, D., Givaudan, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=How+the+insect+pathogen+bacteria+Bacillus+thuringiensis+and+Xenorhabdus/Photorhabdus+occupy+their+hosts.">How the insect pathogen bacteria Bacillus thuringiensis and Xenorhabdus/Photorhabdus occupy their hosts.</a> Current Opinion in Microbiology. 15 (3), 220-231 (2012).</li><li>Waterfield, N. R., Ciche, T., Clarke, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photorhabdus+and+a+host+of+hosts.">Photorhabdus and a host of hosts.</a> Annual Review of Microbiology. 63, 557-574 (2009).</li><li>Ozakman, Y., Eleftherianos, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immune+interactions+between+Drosophila+and+the+pathogen+Xenorhabdus.">Immune interactions between Drosophila and the pathogen Xenorhabdus.</a> Microbiological Research. 240, 126568 (2020).</li><li>Yadav, S., Shokal, U., Forst, S., Eleftherianos, I. <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+method+for+generating+axenic+entomopathogenic+nematodes.">An improved method for generating axenic entomopathogenic nematodes.</a> BMC Research Notes. 8 (1), 1-6 (2015).</li><li>Mitani, D. K., Kaya, H. K., 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=Comparative+study+of+the+entomopathogenic+nematode,+Steinernema+carpocapsae,+reared+on+mutant+and+wild-type+Xenorhabdus+nematophila.">Comparative study of the entomopathogenic nematode, Steinernema carpocapsae, reared on mutant and wild-type Xenorhabdus nematophila.</a> Biological Control. 29 (3), 382-391 (2004).</li><li>McMullen, J. G., Stock, S. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+and+in+vitro+rearing+of+entomopathogenic+nematodes+(Steinernematidae+and+Heterorhabditidae).">In vivo and in vitro rearing of entomopathogenic nematodes (Steinernematidae and Heterorhabditidae).</a> Journal of Visualized Experiments. (91), e52096 (2014).</li></ol>

Understanding the molecular basis of entomopathogenic nematode infection and insect anti-nematode immunity requires the separation of the parasites from the mutualistically associated bacteria13,15,16. The entomopathogenic nematodes H. bacteriophora and S. carpocapsae live together with the Gram-negative bacteria P. luminescens and X. nematophila, respectively17. Both b...

Research on entomopathogenic nematodes (EPN) has intensified over the past few years due primarily to the utility of these parasites in integrated pest management strategies and their involvement in basic biomedical research1,2. Recent studies have established EPN as model organisms in which to examine the nematode genetic components that are activated during the different stages of the infection process. This information provides critical clues on the nature and number of molecules secreted by the parasites to alter host physiology and destabilize the insect innate immune response3,4. Simultaneously, this knowledge is commonly supplemented by novel details on the type of insect host immune signaling pathways and the functions they regulate to restrict the entry and spread of the pathogens5,6. Understanding these processes is crucial for envisioning both sides of the dynamic interplay between EPN and their insect hosts. Better appreciation of EPN-insect host relationship will undoubtedly facilitate similar studies with mammalian parasitic nematodes, which can lead to the identification and characterization of infection factors that interfere with the human immune system.
The EPN nematodes Heterorhabditis sp. and Steinernema sp. can infect a wide range of insects, and their biology has been intensely studied previously. The two nematode parasites differ in their mode of reproduction with Heterorhabditis being self-fertilized and Steinernema undergoing amphimictic reproduction, although recently S. hermaphroditum was shown to reproduce by self-fertilization of hermaphrodites or through parthenogenesis7,8,9. Another difference between Heterorhabditis and Steinernema nematodes is their symbiotic mutualism with two distinct genera of Gram-negative bacteria, Photorhabdus and Xenorhabdus, respectively, which are both potent pathogens of insects. These bacteria are found in the free-living and non-feeding infective juvenile (IJ) stage of the EPN, which detect susceptible hosts, gain access to the insect hemocoel where they release their associated bacteria that replicate rapidly, and colonize insect tissues. Both the EPN and their bacteria produce virulence factors that disarm insect defenses and impair homeostasis. Following insect death, the nematode IJs develop to become adult EPN and complete their life cycle. A new cohort of IJs formed in response to food deprivation and overcrowding within the insect cadaver finally emerges in the soil to hunt suitable hosts9,10,11,12.
Here, an efficient protocol for maintaining, amplifying and genetically manipulating EPN nematodes is described. In particular, the protocol outlines the replication of symbiotic H. bacteriophora and S. carpocapsae IJs, the generation of axenic nematode IJs, the production of H. bacteriophora hermaphrodites for microinjection, the preparation of the dsRNA, and the microinjection technique. These methods are essential for understanding the molecular basis of nematode pathogenicity and host anti-nematode immunity.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Agarose</td>
			<td>VWR</td>
			<td>97062-244</td>
			<td></td>
		</tr>
		<tr>
			<td>Ambion Megascript T7 Kit</td>
			<td>Thermo Fisher Scientific</td>
			<td>AM1333</td>
			<td></td>
		</tr>
		<tr>
			<td>Ampicillin</td>
			<td>Fisher Scientific</td>
			<td>611770250</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell culture flask T25</td>
			<td>Fisher Scientific</td>
			<td>156367</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell culture flask T75</td>
			<td>Fisher Scientific</td>
			<td>156499</td>
			<td></td>
		</tr>
		<tr>
			<td>ChoiceTaq Mastermix</td>
			<td>Denville Scientific</td>
			<td>C775Y42</td>
			<td></td>
		</tr>
		<tr>
			<td>Corn oil</td>
			<td>VWR</td>
			<td>470200-112</td>
			<td></td>
		</tr>
		<tr>
			<td>Corn syrup</td>
			<td>MP Biomedicals/VWR</td>
			<td>IC10141301</td>
			<td></td>
		</tr>
		<tr>
			<td>Culture tube 10 mL</td>
			<td>Fisher Scientific</td>
			<td>14-959-14</td>
			<td></td>
		</tr>
		<tr>
			<td>Eppendorf Femtotips Microloader Tips</td>
			<td>Eppendorf</td>
			<td>E5242956003</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Millipore-Sigma</td>
			<td>E7023</td>
			<td></td>
		</tr>
		<tr>
			<td>Falcon tube 50 mL</td>
			<td>Fisher Scientific</td>
			<td>14-432-22</td>
			<td></td>
		</tr>
		<tr>
			<td>Femtojet Microinjector</td>
			<td>Eppendorf</td>
			<td>5252000021</td>
			<td></td>
		</tr>
		<tr>
			<td>Filter paper</td>
			<td>VWR</td>
			<td>28320-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Galleria mellonella waxorms</td>
			<td>Petco</td>
			<td>-</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass coverslip</td>
			<td>Fisher Scientific</td>
			<td>12-553-464</td>
			<td>50 x 24 mm</td>
		</tr>
		<tr>
			<td>Halocarbon Oil 700</td>
			<td>Sigma</td>
			<td>H8898</td>
			<td></td>
		</tr>
		<tr>
			<td>Inoculating loop</td>
			<td>VWR</td>
			<td>12000-806</td>
			<td></td>
		</tr>
		<tr>
			<td>Kanamycin</td>
			<td>VWR</td>
			<td>97062-956</td>
			<td></td>
		</tr>
		<tr>
			<td>Kwik-Fil Borosilicate Glass Capillaries</td>
			<td>World Precision Instruments</td>
			<td>1B100F-3</td>
			<td>1.0 mm</td>
		</tr>
		<tr>
			<td>LB Agar</td>
			<td>Fisher Scientific</td>
			<td>BP1425-500</td>
			<td>LB agar miller powder 500 g</td>
		</tr>
		<tr>
			<td>LB Broth</td>
			<td>Fisher Scientific</td>
			<td>BP1426-500</td>
			<td>LB broth miller powder 500 g</td>
		</tr>
		<tr>
			<td>Leica DM IRB Inverted Research Microscope</td>
			<td>Microscope Central</td>
			<td>-</td>
			<td></td>
		</tr>
		<tr>
			<td>MacConkey medium</td>
			<td>Millipore-Sigma</td>
			<td>M7408-250G</td>
			<td></td>
		</tr>
		<tr>
			<td>MEGAclear Transcription Clean-Up Kit</td>
			<td>Thermo Fisher Scientific</td>
			<td>AM1908</td>
			<td></td>
		</tr>
		<tr>
			<td>Microcentrifuge tube</td>
			<td>VWR</td>
			<td>76332-064</td>
			<td>1.5 ml</td>
		</tr>
		<tr>
			<td>NanoDrop 2000 Spectrophotometer</td>
			<td>Thermo Fisher Scientific</td>
			<td>ND-2000</td>
			<td></td>
		</tr>
		<tr>
			<td>Needle syringe</td>
			<td>VWR</td>
			<td>BD305155</td>
			<td>22G</td>
		</tr>
		<tr>
			<td>Nutrient broth</td>
			<td>Millipore-Sigma</td>
			<td>70122-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>VWR</td>
			<td>52858-076</td>
			<td></td>
		</tr>
		<tr>
			<td>Partitioned Petri dish</td>
			<td>VWR</td>
			<td>490005-212</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>VWR</td>
			<td>97062-732</td>
			<td>Buffer PBS tablets biotech grade 200 tab</td>
		</tr>
		<tr>
			<td>PCR primers</td>
			<td>Azenta</td>
			<td>-</td>
			<td></td>
		</tr>
		<tr>
			<td>Pestle</td>
			<td>Millipore-Sigma</td>
			<td>BAF199230001</td>
			<td>Bel-Art Disposable Pestle</td>
		</tr>
		<tr>
			<td>Petri dish 6 cm</td>
			<td>VWR</td>
			<td>25384-092</td>
			<td>60 x 15 mm</td>
		</tr>
		<tr>
			<td>Petri dish 10 mm</td>
			<td>VWR</td>
			<td>10799-192</td>
			<td>35 x 10 mm</td>
		</tr>
		<tr>
			<td>Proteose Peptone #3</td>
			<td>Thermo Fisher Scientific</td>
			<td>211693</td>
			<td></td>
		</tr>
		<tr>
			<td>Yeast extract</td>
			<td>Millipore-Sigma</td>
			<td>Y1625</td>
			<td></td>
		</tr>
	</tbody>
</table>

culturing and genetically manipulating entomopathogenic nematodes

Entomopathogenic nematodes in the genera Heterorhabditis and Steinernema are obligate parasites of insects that live in the soil. The main characteristic of their life cycle is the mutualistic association with the bacteria Photorhabdus and Xenorhabdus, respectively. The nematode parasites are able to locate and enter suitable insect hosts, subvert the insect immune response, and multiply efficiently to produce the next generation that will actively hunt new insect prey to infect. Due to the properties of their life cycle, entomopathogenic nematodes are popular biological control agents, which are used in combination with insecticides to control destructive agricultural insect pests. Simultaneously, these parasitic nematodes represent a research tool to analyze nematode pathogenicity and host anti-nematode responses. This research is aided by the recent development of genetic techniques and transcriptomic approaches for understanding the role of nematode secreted molecules during infection. Here, a detailed protocol on maintaining entomopathogenic nematodes and using a gene knockdown procedure is provided. These methodologies further promote the functional characterization of entomopathogenic nematode infection factors.

To assess the status of H. bacteriophora nematodes that have gone through the axenization, the presence or absence of P. luminescens bacterial colonies in IJs was determined. To do this, a pellet of approximately 500 IJs that had been previously surface sterilized and homogenized in PBS was collected. The positive control treatment consisted of a pellet of approximately 500 IJs from the nematode culture containing symbiotic P. luminescens bacteria. The pellets of axenized and positive control n...

Watch this Scientific Journal Video about Culturing and Genetically Manipulating Entomopathogenic Nematodes at JoVE.com

Culturing and Genetically Manipulating Entomopathogenic Nematodes

Entomopathogenic nematodes live in symbiosis with bacteria and together they successfully infect insects by undermining their innate immune system. To promote research on the genetic basis of nematode infection, methods for maintaining and genetically manipulating entomopathogenic nematodes are described.

Entomopathogenic nematodes in the genera Heterorhabditis and Steinernema are obligate parasites of insects that live in the soil. The main characteristic ...

An efficient protocol for maintaining and amplifying entomopathogenic nematode is essential for understanding the molecular basis of nematode pathogenicity and host anti-nematode immunity. This protocol enables us to produce a high number of symbiotic and axenic Heterorhabditis bacteriophora and Steinernema carpocapsae IJs using insect host, Galleria mellonella. Begin by covering the Petri dish with a piece of filter paper and add approximately 10 to 15 Galleria mellonella larvae.Using a pipette, dispense two milliliters of water containing about 25 to 50 infective juveniles per 10 microliters of suspension onto the wax worms. And store the Petri dish in a cabinet at room temperature. Depending on the moisture of the filter paper, add one to two milliliters of water every two days.Wax worms infected with infective juveniles usually die within 48 hours. Prepare White's water traps approximately 10 days after the wax worms are infected with the infective juveniles. Carefully transfer the dead insects onto the unsoaked part of the filter paper.Use tap water to fill the bottom Petri dish and place a cap of a 15 milliliter tube as a spacer for ventilation. Use tap water instead of demineralized or deionized water to avoid nematode aggregation, and ensure that the water level in the water trap reaches approximately half the height of the Petri dish. When the water in the small Petri dish turns cloudy due to nematodes, use a pipette to move the new generation of infective juveniles into a T25 or a T75 cell culture flask.Later, add tap water up to 40%volume or until the appropriate density is reached. And store the cell culture flask horizontally to avoid nematode congestion. Repeat transferring of infective juveniles into cell culture flasks and the addition of water until the infective juveniles stop emerging from the insect carcasses in approximately three to five days.Using an inoculation loop, scrape several flakes of a frozen culture of Photorhabdus temperata Ret16 onto a MacConkey plate and streak for single colonies. Incubate the plate for two to three days at 28 degrees Celsius. After incubation, inoculate 10 milliliters of LB broth with a colony on MacConkey agar in a 50 milliliter tube and incubate the culture overnight at 28 degrees Celsius in a shaking incubator.After incubation, transfer 100 microliters of overnight culture from a 50 milliliter tube into a new microcentrifuge tube, then wash 100 the overnight culture with 900 microliters of single strength PBS by centrifusion, in a 1.5 milliliter microcentrifuge tube, at 17, 900 times g. Decant the supernatant. Next, dilute the culture 10 times in single strength PBS and leave the tube on ice.Now, immerse the wax worms into a 70%ethanol solution. After drying the insects with a paper towel, place them in a 50 milliliter tube. Place the tube on ice for 20 minutes to immobilize the wax worms.Use filter paper to cover a Petri dish's top and bottom halves. Use the Petri dish lid covered with filter paper as a base support for injection. Moist in the filter paper of the bottom Petri dish and transfer it on ice to help the injected wax worms recover.Pipette 50 microliters of ice cold bacteria on a piece of Parafilm then prepare a 22 gauge refiner needle syringe and press the plunger gently to remove the air at the tip of the needle. Keeping a wax worm close to the posterior end under the stereoscope. Inject bacterial culture into the dorsal side of the thorax.Preferably at the junction between two segments just underneath the cuticle parallel to the wax worm to minimize internal damage and pay attention to how the larval body starts bulging. Next, transfer the injected insects to the recovery Petri dish. And once all the insects are injected and placed in the recovery dish, place the Petri dish in the dark.Wax worms succumb two days after injection and appear brick-red after approximately three to four days. However, brown-colored insects indicate unsuccessful bacterial infection. At seven days post-infection, transfer the insects with the characteristic brick-red color onto a fresh filter paper lined Petri dish, and repeat the production of symbiotic nematode infective juveniles.To begin surface sterilization, collect sufficient symbiotic, Heterorhabditis bacteriophora and candidate axenic infective juveniles by centrifugation. To make a pellet in a 1.5 milliliter centrifuge tube, add 500 microliters of 5%bleach to the pellet in each microcentrifuge tube and invert the tube. After incubating for 10 minutes, centrifuge at 17, 900 times g for one minute and remove the supernatant.Then wash the pellet with one milliliter of sterile water. Centrifuge it, remove the supernatant and repeat this procedure four more times. The graph shows the results of assessing the status of Heterorhabditis bacteriophora nematodes that have gone through axenization.Heterorhabditis bacteriophora from the stock culture carried symbiotic Photorhabdus luminescens cells. However, nematodes devoid of their associated Photorhabdus luminescens bacteria were axenic. To maintain the reproducibility of your experiment, always streak freshly from bacterial stock culture from a fresh colony and take out the overnight culture before it reaches stationary face.Only use the brick-red Galleria wax worm for White's water trap and discard the rest. After this procedure, the nematodes can be used to investigate the interaction with insect immune system. This technique paves the way for researcher to explore the molecular basis of nematode parasitism and host anti-nematode immunity.

culturing-and-genetically-manipulating-entomopathogenic-nematodes

Research

JoVE Journal

Immunology and Infection

High and Low Throughput Screens with Root-knot Nematodes Meloidogyne spp.

Root-knot nematodes (genus Meloidogyne) are obligate plant parasites. They are extremely polyphagous and considered one of the most economically important plant parasitic nematodes. The microscopic second-stage juvenile (J2), molted once in the egg, is the infective stage. The J2s hatch from the eggs, move freely in the soil within a film of water, and locate root tips of suitable plant species. After penetrating the plant root, they migrate towards the vascular cylinder where they establish a feeding site and initiate feeding using their stylets. The multicellular feeding site is comprised of several enlarged multinuclear cells called 'giant cells' which are formed from cells that underwent karyokinesis (repeated mitosis) without cytokinesis. Neighboring pericycle cells divide and enlarge in size giving rise to a typical gall or root knot, the characteristic symptom of root-knot nematode infection. Once feeding is initiated, J2s become sedentary and undergo three additional molts to become adults. The adult female lays 150-250 eggs in a gelatinous matrix on or below the surface of the root. From the eggs new infective J2s hatch and start a new cycle. The root-knot nematode life cycle is completed in 4-6 weeks at 26-28&deg;C.
Here we present the traditional protocol to infect plants, grown in pots, with root-knot nematodes and two methods for high-throughput assays. The first high-throughput method is used for plants with small seeds such as tomato while the second is for plants with large seeds such as cowpea and common bean. Large seeds support extended seedling growth with minimal nutrient supplement. The first high throughput assay utilizes seedlings grown in sand in trays while in the second assay plants are grown in pouches in the absence of soil. The seedling growth pouch is made of a 15.5 x 12.5cm paper wick, folded at the top to form a 2-cm-deep trough in which the seed or seedling is placed. The paper wick is contained inside a transparent plastic pouch. These growth pouches allow direct observation of nematode infection symptoms, galling of roots and egg mass production, under the surface of a transparent pouch. Both methods allow the use of the screened plants, after phenotyping, for crossing or seed production. An additional advantage of the use of growth pouches is the small space requirement because pouches are stored in plastic hanging folders arranged in racks.

High and Low Throughput Screens with Root-knot Nematodes Meloidogyne spp.

Two distinct methods to screen plants with root-knot nematodes are described. The described approaches include high-throughput screens with nematodes in a nondestructive manner facilitating the use of these plants in breeding programs.

Root-knot nematodes (genus Meloidogyne) are obligate plant parasites. They are extremely polyphagous and considered one of the most economically important ...

Parasite Induced Genetically Driven Autoimmune Chagas Heart Disease in the Chicken Model

The Trypanosoma cruzi acute infections acquired in infancy and childhood seem asymptomatic, but approximately one third of the chronically infected cases show Chagas disease up to three decades or later. Autoimmunity and parasite persistence are competing theories to explain the pathogenesis of Chagas disease 1, 2. To separate roles played by parasite persistence and autoimmunity in Chagas disease we inoculate the T. cruzi in the air chamber of fertilized eggs. The mature chicken immune system is a tight biological barrier against T. cruzi and the infection is eradicated upon development of its immune system by the end of the first week of growth 3. The chicks are parasite-free at hatching, but they retain integrated parasite mitochondrial kinetoplast DNA (kDNA) minicircle within their genome that are transferred to their progeny. Documentation of the kDNA minicircle integration in the chicken genome was obtained by a targeted prime TAIL-PCR, Southern hybridizations, cloning, and sequencing 3, 4. The kDNA minicircle integrations rupture open reading frames for transcription and immune system factors, phosphatase (GTPase), adenylate cyclase and phosphorylases (PKC, NF-Kappa B activator, PI-3K) associated with cell physiology, growth, and differentiation 3, 5-7, and other gene functions. Severe myocarditis due to rejection of target heart fibers by effectors cytotoxic lymphocytes is seen in the kDNA mutated chickens, showing an inflammatory cardiomyopathy similar to that seen in human Chagas disease. Notably, heart failure and skeletal muscle weakness are present in adult chickens with kDNA rupture of the dystrophin gene in chromosome 1 8. Similar genotipic alterations are associated with tissue destruction carried out by effectors CD45+, CD8&gamma;&delta;+, CD8&alpha; lymphocytes. Thus this protozoan infection can induce genetically driven autoimmune disease.

The inoculation of Trypanosoma cruzi in fertile eggs prior to incubation renders the parasite kDNA minicircle integration in embryo cells genome. Crossbreeding reveals the vertical transfer of the mutations to progeny. The kDNA integrates into coding regions at several chromosomes and the chickens die with an inflammatory autoimmune heart disease.

The Trypanosoma cruzi acute infections acquired in infancy and childhood seem asymptomatic, but approximately one third of the chronically infected cases ...

TransFLP &#x2014; A Method to Genetically Modify Vibrio cholerae Based on Natural Transformation and FLP-recombination

Several methods are available to manipulate bacterial chromosomes1-3. Most of these protocols rely on the insertion of conditionally replicative plasmids (e.g. harboring pir-dependent or temperature-sensitive replicons1,2). These plasmids are integrated into bacterial chromosomes based on homology-mediated recombination. Such insertional mutants are often directly used in experimental settings. Alternatively, selection for plasmid excision followed by its loss can be performed, which for Gram-negative bacteria often relies on the counter-selectable levan sucrase enzyme encoded by the sacB gene4. The excision can either restore the pre-insertion genotype or result in an exchange between the chromosome and the plasmid-encoded copy of the modified gene. A disadvantage of this technique is that it is time-consuming. The plasmid has to be cloned first; it requires horizontal transfer into V. cholerae (most notably by mating with an E. coli donor strain) or artificial transformation of the latter; and the excision of the plasmid is random and can either restore the initial genotype or create the desired modification if no positive selection is exerted. Here, we present a method for rapid manipulation of the V. cholerae chromosome(s)5 (Figure 1). This TransFLP method is based on the recently discovered chitin-mediated induction of natural competence in this organism6 and other representative of the genus Vibrio such as V. fischeri7. Natural competence allows the uptake of free DNA including PCR-generated DNA fragments. Once taken up, the DNA recombines with the chromosome given the presence of a minimum of 250-500 bp of flanking homologous region8. Including a selection marker in-between these flanking regions allows easy detection of frequently occurring transformants.
This method can be used for different genetic manipulations of V. cholerae and potentially also other naturally competent bacteria. We provide three novel examples on what can be accomplished by this method in addition to our previously published study on single gene deletions and the addition of affinity-tag sequences5. Several optimization steps concerning the initial protocol of chitin-induced natural transformation6 are incorporated in this TransFLP protocol. These include among others the replacement of crab shell fragments by commercially available chitin flakes8, the donation of PCR-derived DNA as transforming material9, and the addition of FLP-recombination target sites (FRT)5. FRT sites allow site-directed excision of the selection marker mediated by the Flp recombinase10.

TransFLP &amp;#x2014; A Method to Genetically Modify &lt;em&gt;Vibrio cholerae&lt;/em&gt; Based on Natural Transformation and FLP-recombination

A quick method to modify the genome of V. cholerae is described. These modifications include the deletion of single genes, gene clusters and genomic islands as well as the integration of short sequences (e.g. promoter elements or affinity-tag sequences). The method is based on the natural transformation and FLP-recombination.

Several methods are available to manipulate bacterial chromosomes1-3. Most of these protocols rely on the insertion of conditionally replicative plasmids ...

Clinical Application of Sleeping Beauty and Artificial Antigen Presenting Cells to Genetically Modify T Cells from Peripheral and Umbilical Cord Blood

The potency of clinical-grade T cells can be improved by combining gene therapy with immunotherapy to engineer a biologic product with the potential for superior (i) recognition of tumor-associated antigens (TAAs), (ii) persistence after infusion, (iii) potential for migration to tumor sites, and (iv) ability to recycle effector functions within the tumor microenvironment. Most approaches to genetic manipulation of T cells engineered for human application have used retrovirus and lentivirus for the stable expression of CAR1-3. This approach, although compliant with current good manufacturing practice (GMP), can be expensive as it relies on the manufacture and release of clinical-grade recombinant virus from a limited number of production facilities. The electro-transfer of nonviral plasmids is an appealing alternative to transduction since DNA species can be produced to clinical grade at approximately 1/10th the cost of recombinant GMP-grade virus. To improve the efficiency of integration we adapted Sleeping Beauty (SB) transposon and transposase for human application4-8. Our SB system uses two DNA plasmids that consist of a transposon coding for a gene of interest (e.g. 2nd generation CD19-specific CAR transgene, designated CD19RCD28) and a transposase (e.g. SB11) which inserts the transgene into TA dinucleotide repeats9-11. To generate clinically-sufficient numbers of genetically modified T cells we use K562-derived artificial antigen presenting cells (aAPC) (clone #4) modified to express a TAA (e.g. CD19) as well as the T cell costimulatory molecules CD86, CD137L, a membrane-bound version of interleukin (IL)-15 (peptide fused to modified IgG4 Fc region) and CD64 (Fc-&gamma; receptor 1) for the loading of monoclonal antibodies (mAb)12. In this report, we demonstrate the procedures that can be undertaken in compliance with cGMP to generate CD19-specific CAR+ T cells suitable for human application. This was achieved by the synchronous electro-transfer of two DNA plasmids, a SB transposon (CD19RCD28) and a SB transposase (SB11) followed by retrieval of stable integrants by the every-7-day additions (stimulation cycle) of &gamma;-irradiated aAPC (clone #4) in the presence of soluble recombinant human IL-2 and IL-2113. Typically 4 cycles (28 days of continuous culture) are undertaken to generate clinically-appealing numbers of T cells that stably express the CAR. This methodology to manufacturing clinical-grade CD19-specific T cells can be applied to T cells derived from peripheral blood (PB) or umbilical cord blood (UCB). Furthermore, this approach can be harnessed to generate T cells to diverse tumor types by pairing the specificity of the introduced CAR with expression of the TAA, recognized by the CAR, on the aAPC.

Clinical Application of Sleeping Beauty and Artificial Antigen Presenting Cells to Genetically Modify T Cells from Peripheral and Umbilical Cord Blood

T cells expressing a CD19-specific chimeric antigen receptor (CAR) are infused as investigational treatment of B-cell malignancies in our first-in-human gene therapy trials. We describe genetic modification of T cells using the Sleeping Beauty (SB) system to introduce CD19-specific CAR and selective propagation on designer CD19+ artificial antigen presenting cells.

The potency of clinical-grade T cells can be improved by combining gene therapy with immunotherapy to engineer  a biologic product with the potential for ...

Culturing Microglia from the Neonatal and Adult Central Nervous System

Microglia are the resident macrophage-like cells of the central nervous system (CNS) and, as such, have critically important roles in physiological and pathological processes such as CNS maturation in development, multiple sclerosis, and spinal cord injury. Microglia can be activated and recruited to action by neuronal injury or stimulation, such as axonal damage seen in MS or ischemic brain trauma resulting from stroke. These immunocompetent members of the CNS are also thought to have roles in synaptic plasticity under non-pathological conditions. We employ protocols for culturing microglia from the neonatal and adult tissues that are aimed to maximize the viable cell numbers while minimizing confounding variables, such as the presence of other CNS cell types and cell culture debris. We utilize large and easily discernable CNS components (e.g. cortex, spinal cord segments), which makes the entire process feasible and reproducible. The use of adult cells is a suitable alternative to the use of neonatal brain microglia, as many pathologies studied mainly affect the postnatal spinal cord. These culture systems are also useful for directly testing the effect of compounds that may either inhibit or promote microglial activation. Since microglial activation can shape the outcomes of disease in the adult CNS, there is a need for in vitro systems in which neonatal and adult microglia can be cultured and studied.

We outline methods for the efficient and quick isolation/culture of viable microglia from the neonatal cerebral cortex and adult spinal cord. The dissection and plating of cortical microglia can be accomplished within 90 minutes, with the subsequent microglial harvest taking place ~ 10 days following the initial dissection.

Microglia are the resident macrophage-like cells of the central nervous system (CNS) and, as such, have critically important roles in physiological and ...

Culturing and Maintaining Clostridium difficile in an Anaerobic Environment

Clostridium difficile is a Gram-positive, anaerobic, sporogenic bacterium that is primarily responsible for antibiotic associated diarrhea (AAD) and is a significant nosocomial pathogen. C. difficile is notoriously difficult to isolate and cultivate and is extremely sensitive to even low levels of oxygen in the environment. Here, methods for isolating C. difficile from fecal samples and subsequently culturing C. difficile for preparation of glycerol stocks for long-term storage are presented. Techniques for preparing and enumerating spore stocks in the laboratory for a variety of downstream applications including microscopy and animal studies are also described. These techniques necessitate an anaerobic chamber, which maintains a consistent anaerobic environment to ensure proper conditions for optimal C. difficile growth. We provide protocols for transferring materials in and out of the chamber without causing significant oxygen contamination along with suggestions for regular maintenance required to sustain the appropriate anaerobic environment for efficient and consistent C. difficile cultivation.

Culturing and Maintaining Clostridium difficile in an Anaerobic Environment

Clostridium difficile is a pathogenic bacterium that is a strict anaerobe and causes antibiotic associated diarrhea (AAD). Here, methods for isolating, culturing and maintaining C. difficile vegetative cells and spores are described. These techniques necessitate an anaerobic chamber, which requires regular maintenance to ensure proper conditions for optimal C. difficile cultivation.

Clostridium  difficile is a Gram-positive, anaerobic, sporogenic bacterium  that is primarily responsible for antibiotic associated diarrhea (AAD) and is ...

Selective Cleaning of Wild Caenorhabditis Nematodes to Enrich for Intestinal Microbiome Bacteria

Caenorhabditis elegans (C. elegans)&#160;has proven to be an excellent model for studying host-microbe interactions and the microbiome, especially in the context of the intestines. Recently, ecological sampling of wild Caenorhabditis nematodes has discovered a diverse array of associated microbes, including bacteria, viruses, fungi, and microsporidia. Many of these microbes have interesting colonization or infection phenotypes that warrant further study, but they are often unculturable. This protocol presents a method to enrich the desired intestinal microbes in C. elegans and related nematodes and reduce the presence of the many contaminating microbes adhering to the cuticle. This protocol involves forcing animals into the dauer stage of development and using a series of antibiotic and detergent washes to remove external contamination. As dauer animals have physiological changes that protect nematodes from harsh environmental conditions, any intestinal microbes will be protected from these conditions. But, for enrichment to work, the microbe of interest must be maintained when animals develop into dauers. When the animals leave the dauer stage, they are singly propagated into individual lines. F1 populations are then selected for desired microbes or infection phenotypes and against visible contamination. These methods will allow researchers to enrich unculturable microbes in the intestinal lumen, which make up part of the natural microbiome of C. elegans and intracellular intestinal pathogens. These microbes can then be studied for colonization or infection phenotypes and their effects on the host fitness.

Selective Cleaning of Wild Caenorhabditis Nematodes to Enrich for Intestinal Microbiome Bacteria

Wild Caenorhabditis nematodes are associated with many microbes, often in the gut lumen or infecting the intestine. This protocol details a method to enrich unculturable microbes colonizing the intestine, taking advantage of the resistance of the dauer cuticle.

Caenorhabditis elegans (C. elegans)&#160;has proven to be an excellent model for studying host-microbe interactions and the microbiome, especially in the ...

Economical and Efficient Protocol for Isolating and Culturing Bone Marrow-derived Dendritic Cells from Mice

The demand for dendritic cells (DCs) is gradually increasing as immunology research advances. However, DCs are rare in all tissues. The traditional method for isolating DCs primarily involves inducing bone marrow (BM) differentiation into DCs by injecting large doses (&#62;10 ng/mL) of granulocyte-macrophage colony-stimulating factor/interleukin-4 (GM-CSF/IL-4), making the procedure complex and expensive. In this protocol, using all BM cells cultured in 10 ng/mL GM-CSF/IL-4 medium, after 3-4 half-culture exchanges, up to 2.7 x 107 CD11c+ cells&#160;(DCs) per mouse (two femurs) were harvested with a purity of 80%-95%. After 10 days in culture, the expression of CD11c, CD80, and MHC II increased, whereas the number of cells decreased. The number of cells peaked after 7 days of culture. Moreover, this method only took 10 min to harvest all bone marrow cells, and a high number of DCs were obtained after 1 week of culture.

Here, we present an economical and efficient method to isolate and generate high-purity bone marrow-derived dendritic cells from mice after 7 days of culture with 10 ng/mL GM-CSF/IL-4.

The demand for dendritic cells (DCs) is gradually increasing as immunology research advances. However, DCs are rare in all tissues. The traditional method ...

Culturing Lymphocytes in Simulated Microgravity Using a Rotary Cell Culture System

Given the current limitations of conducting biological research in space, a few options exist for subjecting cell culture to simulated microgravity (SMG) on Earth. These options vary in their methods, principles, and suitability for use with suspension cell culture. Here, a cell culture method is described for subjecting lymphocytes to simulated microgravity using a commercially available rotary cell culture system, also known as a 2D clinostat or a rotating wall vessel (RWV) device. This cell culture method utilizes the principle of time-averaged gravity vector nullification to simulate microgravity by rotating the cells on a horizontal axis. The cells cultured in this system can be harvested and utilized in many different experimental assays to assess the effects of simulated microgravity on cellular function and physiology. The culturing technique may vary slightly depending on the cell type or line that is used, but the method described here may be applied to any suspension-type cell culture.

This is a step-by-step guide for using a commercially available rotary cell culture system to culture lymphocytes in simulated microgravity using specialized disposable culture vessels. This culturing method may be applied to any suspension-type cell culture.

Given the current limitations of conducting biological research in space, a few options exist for subjecting cell culture to simulated microgravity (SMG) ...

Generating Genetically Modified Plasmodium berghei Sporozoites

Malaria is a deadly disease caused by the parasite Plasmodium and is transmitted through the bite of female Anopheles mosquitoes. The sporozoite stage of Plasmodium deposited by mosquitoes in the skin of vertebrate hosts undergoes a phase of mandatory development in the liver before initiating clinical malaria. We know little about the biology of Plasmodium development in the liver; access to the sporozoite stage and the ability to genetically modify such sporozoites are critical tools for studying the nature of Plasmodium infection and the resulting immune response in the liver. Here, we present a comprehensive protocol for the generation of transgenic Plasmodium berghei sporozoites. We genetically modify blood-stage P. berghei and use this form to infect Anopheles mosquitoes when they take a blood meal. After the transgenic parasites undergo development in the mosquitoes, we isolate the sporozoite stage of the parasite from the mosquito salivary glands for in vivo and in vitro experimentation. We demonstrate the validity of the protocol by generating sporozoites of a novel strain of P. berghei expressing the green fluorescent protein (GFP) subunit 11 (GFP11), and show how it could be used to investigate the biology of liver-stage malaria.

Generating Genetically Modified Plasmodium berghei Sporozoites

Malaria is transmitted through inoculation of the sporozoite stage of Plasmodium by infected mosquitoes. Transgenic Plasmodium has allowed us to understand the biology of malaria better and has contributed directly to malaria vaccine development efforts. Here, we describe a streamlined methodology to generate transgenic Plasmodium berghei sporozoites.

Malaria is a deadly disease caused by the parasite Plasmodium and is transmitted through the bite of female Anopheles mosquitoes. The sporozoite stage of ...