The authors wish to thank past members of the Goodrich-Blair lab: Samantha Orchard, Kimberly Cowles, Erin Herbert-Tran, Greg Richards, Megan Menard, and Youngjin Park for their contributions to the development of this protocol. This work was funded by the National Science Foundation grant IOS-0950873 and the National Institutes of Health NRSA fellowship FAI084441Z.

1. Insect Egg Sterilization and Rearing
<ol>
	<li>Prepare diet by first autoclaving 15 g of the provided agar in 900-1,000 ml H2O. Immediately after autoclaving, mix with 166 g wheat germ diet and blend well in a laboratory blender. Pour into a dish (or dishes) to cool, then transfer diet to aluminum foil, wrap tightly, and store at 4 &deg;C.</li>
	<li>Upon arrival, sterilize M. sexta eggs with 250 ml of 0.6% bleach solution for 2-3 min in a glass filter holder and vacuum flask apparatus with a 90 mm filter paper, stirring occasionally.</li>
	<li>Turn on vacuum to drain bleach solution and wash eggs 3-4 times with 250 ml sterile distilled H2O per wash.</li>
	<li>Transfer eggs to fresh 90 mm filter paper placed in a Petri plate lid and allow them to dry until they no longer stick together (about 20-30 min).</li>
	<li>Transfer eggs to the bottom of plastic containers with insect diet, placing approximately 40 eggs in each container (Figure 1A). Eggs are separated from diet to prevent moisture-induced fungal contamination. Maintain eggs at 26 &deg;C with a 16 hr light: 8 hr dark photoperiod. Eggs will lighten to a yellow-white color prior to hatching.</li>
	<li>When hatching is complete, carefully transfer approximately 25 larvae each to new containers with diet on the bottom (Figure 1B). It is important to pick up each larva by the long black horn using forceps to avoid harming them during their fragile early developmental stages. Incubate for 2 days as above. While larval death is unusual at this stage, some dead or, more likely, developmentally stunted larvae may be observed. Such insects should be excluded from further study and sacrificed by freezing.</li>
	<li>To avoid cannibalistic behaviors, transfer larvae to individual containers with small pieces of food (Figure 1C) and incubate as above. Monitor insect development, replacing food and cleaning feces out of containers every other day until they undergo the third larval molt (usually 6-9 days after hatching). This is the 4th instar larval stage, characterized by the appearance of black hook-like crochets on each proleg and increased prominence of stripes along the insect body (Figure 1D). The larvae may vary considerably in size, but typically weigh 0.15-0.4 g upon entering the 4th instar stage. Larvae of similar size should be chosen for injection.</li>
</ol>
2. Preparation of Bacteria for Injection
<ol>
	<li>Grow the bacterial strain(s) to be tested according to standard procedures to the growth phase you wish to test.</li>
	<li>Pellet 500 &mu;l of each bacterial strain to be tested by spinning for 2 min in a microcentrifuge at 13,000 rpm at room temperature.</li>
	<li>Resuspend cells in 1 ml sterile 1x phosphate buffered saline (PBS) and pellet again as above.</li>
	<li>Resuspend cells in 0.5 ml sterile 1x PBS and measure the optical density (OD) of the suspension. Dilute all suspensions to the same OD, if necessary. Sterile growth media may also be used for cell resuspension and dilution in place of 1x PBS, if desired.</li>
</ol>
3. Injection of 4th Instar Larvae
<ol>
	<li>Prepare serial 10-fold dilutions of the first bacterial strain in 6 wells of a 96-well microtiter plate in sterile 1x PBS, using a fresh pipette tip for each dilution (Figure 2A). The final volume of cell suspension in each well should exceed the volume required for injection (10 &mu;l per insect) plus 20 &mu;l to plate the inoculum (see steps 3.2 and 3.7).</li>
	<li>Spot 10 &mu;l each for 5 dilutions (10-2 through 10-6) along the top of a Petri plate containing an appropriate growth medium. Tilt the plate so that the spots spread to the center (Figure 2B).</li>
	<li>Remove insect diet and feces and place insects in containers on ice for approximately 5 min.</li>
	<li>Sterilize the syringe needle by rinsing 3 times each in 2 tubes of 70-100% ethanol, followed by one tube of sterile H2O (Figure 2C). The volume of liquid must be sufficient to submerge the needle.</li>
	<li>Quantitate the dilute cell suspensions under a microscope using a hemocytometer. Depending on the desired inoculum, draw 10 &mu;l from the appropriate microtiter well (Figure 2D).</li>
	<li>Swab the insect surface with 95% ethanol, and inject the 10 ml cell suspension into the insect at an angle (less than 45&deg;) behind the one of the abdominal prolegs. Take care not to puncture the gut and inject the contents just underneath the epidermal layer (Figure 2E). While some loss of hemolymph (clear, green-blue liquid) is normal, particulate matter and yellow liquid emerging from the injection site are indicative of gut puncture. If this occurs, sacrifice the insect and obtain a new larva to replace it.</li>
	<li>Repeat step 3.6 for each remaining insect in the first injection group. Needle sterilization is not necessary between each insect injected with the same dilution and strain of bacteria unless contamination occurs. Alternatively, a repeating dispenser may be used for greater consistency in injection volume among samples.</li>
	<li>After injecting each insect with the first strain of bacteria, repeat step 3.2, this time spotting cell suspensions in the middle of the plate and letting them flow toward the bottom (Figure 2F). Incubate plates at appropriate temperature and count colonies to enumerate the inoculum. This second plating step is used to verify that the samples were not contaminated during the injection process, as this would result in disparate colony numbers between the first and second platings.</li>
	<li>Repeat steps 3.1 through 3.8 for each strain of bacteria in the experiment. Finally, inject 3-5 insects with sterile 1x PBS using a sterile needle as a negative control group. Incubate insects at 26 &deg;C with a 16 hr light: 8 hr dark photoperiod.</li>
	<li>Monitor insect survival over time after injection. Insect death is characterized as a lack of voluntary movement upon stimulation. Insects often exhibit appetite loss, diarrhea (watery, yellow excrement), and/or water loss ("sweating") during infection prior to death; these characteristics may be worth noting as well.</li>
</ol>

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Microbiol. 33, 878-880 (1977).</li><li>P&#233;chy-Tarr, M., Bruck, D. J., Maurhofer, M., Fischer, E., Vogne, C., Henkels, M. D., Donahue, K. M., Grunder, J., Loper, J. E., Keel, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+analysis+of+a+novel+gene+cluster+encoding+an+insect+toxin+in+plant-associated+strains+of+Pseudomonas+fluorescens.">Molecular analysis of a novel gene cluster encoding an insect toxin in plant-associated strains of Pseudomonas fluorescens.</a> Environ. Microbiol. 10, 2368-2386 (2008).</li><li>Nu&#241;ez-Valdez, M. E., Calder&#243;n, M. A., Aranda, E., Hern&#225;ndez, L., Ram&#237;rez-Gama, R. M., Lina, L., Rodr&#237;guez-Segura, Z., Guti&#233;rrez Mdel, C., Villalobos, F. 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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dissecting+the+immune+response+to+the+entomopathogen+Photorhabdus.">Dissecting the immune response to the entomopathogen Photorhabdus.</a> Trends Microbiol. 18, 552-560 (2010).</li><li>Herbert, E. 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=Friend+and+foe:+the+two+faces+of+Xenorhabdus+nematophila.">Friend and foe: the two faces of Xenorhabdus nematophila.</a> Nat. Rev. Microbiol. 5, 634-646 (2007).</li><li>D'Argenio, D. A., Gallagher, L. A., Berg, C. A., Manoil, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+as+a+model+host+for+Pseudomonas+aeruginosa+Infection.">Drosophila as a model host for Pseudomonas aeruginosa Infection.</a> J. Bacteriol. 183, 1466-1471 (2001).</li><li>Waterfield, N., Dowling, A., Sharma, S., Daborn, P. J., Potter, U., Ffrench-Constant, R. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oral+toxicity+of+Photorhabdus+luminescens+W14+toxin+complexes+in+Escherichia+coli.">Oral toxicity of Photorhabdus luminescens W14 toxin complexes in Escherichia coli.</a> Appl. Environ. Microbiol. 67, 5017-5024 (2001).</li><li>Park, Y., Herbert, E. E., Cowles, C. E., Cowles, K. N., Menard, M. L., Orchard, S. 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E., Richards, G. R., Martens, E. C., 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=The+global+regulator+Lrp+contributes+to+mutualism,+pathogenesis+and+phenotypic+variation+in+the+bacterium+Xenorhabdus+nematophila.">The global regulator Lrp contributes to mutualism, pathogenesis and phenotypic variation in the bacterium Xenorhabdus nematophila.</a> Cell. Microbiol. 9, 1311-1323 (2007).</li><li>Cowles, K. N., 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=Expression+and+activity+of+a+Xenorhabdus+nematophila+haemolysin+required+for+full+virulence+towards+Manduca+sexta+insects.">Expression and activity of a Xenorhabdus nematophila haemolysin required for full virulence towards Manduca sexta insects.</a> Cell. Microbiol. 7, 209-219 (2005).</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> Mol. Microbiol. 64, 260-268 (2007).</li><li>Eleftherianos, I., Baldwin, H., ffrench-Constant, R. H., 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=Developmental+modulation+of+immunity:+changes+within+the+feeding+period+of+the+fifth+larval+stage+in+the+defence+reactions+of+Manduca+sexta+to+infection+by+Photorhabdus.">Developmental modulation of immunity: changes within the feeding period of the fifth larval stage in the defence reactions of Manduca sexta to infection by Photorhabdus.</a> J. Insect Physiol. 54, 309-318 (2008).</li><li>Kavanagh, K., Reeves, E. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Exploiting+the+potential+of+insects+for+in+vivo+pathogenicity+testing+of+microbial+pathogens.">Exploiting the potential of insects for in vivo pathogenicity testing of microbial pathogens.</a> FEMS Microbiol. Rev. 28, 101-112 (2004).</li></ol>

No conflicts of interest declared.

The direct injection of M. sexta larvae with entomopathogenic bacteria, as described here, serves as a simple and effective means to analyze bacterial virulence. The method is also highly adaptable to suit different experimental subjects and/or conditions. Bacteria can be prepared in various ways prior to injection. In the case of X. nematophila, wild type cells grown in nutrient-rich Luria-Bertani (LB) medium to mid-log phase are typically the most virulent, killing most or all insects within 30 hr sub...

<table border="1">
	<tbody>
		<tr>
			<td>Reagent</td>
			<td>Company</td>
			<td>Catalogue number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>90 mm filter paper</td>
			<td>Whatman</td>
			<td>1001 090</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Glass filter holder</td>
			<td>Millipore</td>
			<td>XX1004700</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Manduca sexta eggs</td>
			<td>Carolina Biological Supply</td>
			<td>143880</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>Gypsy Moth Diet + agar</td>
			<td>MP Biomedicals</td>
			<td>0296029301</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>5.5 oz. plastic containers and lids</td>
			<td>Solo Cup Company</td>
			<td>URC55-0090 Pl4-0090</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>1 oz. plastic containers and lids</td>
			<td>DART Container Corporation</td>
			<td>100PC 100PCL25</td>
			<td>&nbsp;</td>
		</tr>
		<tr>
			<td>1x PBS</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>137 mm NaCl, 2.7 mM KCl, 8 mM Na2HPO4, 1.46 mM KH2PO4, pH 7.4</td>
		</tr>
		<tr>
			<td>Syringe</td>
			<td>Hamilton</td>
			<td>80208</td>
			<td>30 gauge, 0.375" length, point style 2</td>
		</tr>
	</tbody>
</table>

rearing and injection of manduca sexta larvae to assess bacterial virulence

Manduca sexta, commonly known as the tobacco hornworm, is considered a significant agricultural pest, feeding on solanaceous plants including tobacco and tomato. The susceptibility of M. sexta larvae to a variety of entomopathogenic bacterial species1-5, as well as the wealth of information available regarding the insect's immune system6-8, and the pending genome sequence9 make it a good model organism for use in studying host-microbe interactions during pathogenesis. In addition, M. sexta larvae are relatively large and easy to manipulate and maintain in the laboratory relative to other susceptible insect species. Their large size also facilitates efficient tissue/hemolymph extraction for analysis of the host response to infection.
The method presented here describes the direct injection of bacteria into the hemocoel (blood cavity) of M. sexta larvae. This approach can be used to analyze and compare the virulence characteristics of various bacterial species, strains, or mutants by simply monitoring the time to insect death after injection. This method was developed to study the pathogenicity of Xenorhabdus and Photorhabdus species, which typically associate with nematode vectors as a means to gain entry into the insect. Entomopathogenic nematodes typically infect larvae via natural digestive or respiratory openings, and release their symbiotic bacterial contents into the insect hemolymph (blood) shortly thereafter10. The injection method described here bypasses the need for a nematode vector, thus uncoupling the effects of bacteria and nematode on the insect. This method allows for accurate enumeration of infectious material (cells or protein) within the inoculum, which is not possible using other existing methods for analyzing entomopathogenesis, including nicking11 and oral toxicity assays12. Also, oral toxicity assays address the virulence of secreted toxins introduced into the digestive system of larvae, whereas the direct injection method addresses the virulence of whole-cell inocula.
The utility of the direct injection method as described here is to analyze bacterial pathogenesis by monitoring insect mortality. However, this method can easily be expanded for use in studying the effects of infection on the M. sexta immune system. The insect responds to infection via both humoral and cellular responses. The humoral response includes recognition of bacterial-associated patterns and subsequent production of various antimicrobial peptides7; the expression of genes encoding these peptides can be monitored subsequent to direct infection via RNA extraction and quantitative PCR13. The cellular response to infection involves nodulation, encapsulation, and phagocytosis of infectious agents by hemocytes6. To analyze these responses, injected insects can be dissected and visualized by microscopy13, 14.

A representative example of an insect mortality assay is depicted in Figure 3. In this experiment, insects were injected with approximately 50 colony forming units (CFU) of either wild type (ATCC19061) or an attenuated mutant strain (lrp13) of Xenorhabdus nematophila grown to mid-log phase (n=6 insects per strain). Insects were observed for approximately 72 hr, and the percent of injected insects still alive at each timepoint recorded. In this case, the attenuated strain exhi...

Watch this Scientific Journal Video about Rearing and Injection of Manduca sexta Larvae to Assess Bacterial Virulence at JoVE.com

Rearing and Injection of Manduca sexta Larvae to Assess Bacterial Virulence

The method described here utilizes direct injection of entomopathogenic bacteria into the hemocoel of Manduca sexta insect larvae. M. sexta is a commercially available and well-studied insect. Thus, this method represents a simple approach to analyzing host-bacterial interactions from the perspective of one or both partners.

Rearing and Injection of Manduca sexta Larvae to Assess Bacterial Virulence

Manduca sexta, commonly known as the tobacco hornworm, is considered a significant agricultural pest, feeding on solanaceous plants including tobacco and ...

rearing-injection-manduca-sexta-larvae-to-assess-bacterial

Research

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Immunology and Infection

12.6K Views.  University of Wisconsin-Madison. Метод, описанный здесь используется непосредственный впрыск энтомопатогенных бактерий в hemocoel из Manduca Sexta Личинками насекомых. M. Sexta Является коммерчески доступным и хорошо изученных насекомых. Таким образом, этот метод представляет собой про?...

Video: Rearing and Injection of Manduca sexta Larvae to Assess Bacterial Virulence

Saliva, Salivary Gland, and Hemolymph Collection from Ixodes scapularis Ticks

Ticks are found worldwide and afflict humans with many tick-borne illnesses. Ticks are vectors for pathogens that cause Lyme disease and tick-borne relapsing fever (Borrelia spp.), Rocky Mountain Spotted fever (Rickettsia rickettsii), ehrlichiosis (Ehrlichia chaffeensis and E. equi), anaplasmosis (Anaplasma phagocytophilum), encephalitis (tick-borne encephalitis virus), babesiosis (Babesia spp.), Colorado tick fever (Coltivirus), and tularemia (Francisella tularensis) 1-8. To be properly transmitted into the host these infectious agents differentially regulate gene expression, interact with tick proteins, and migrate through the tick 3,9-13. For example, the Lyme disease agent, Borrelia burgdorferi, adapts through differential gene expression to the feast and famine stages of the tick's enzootic cycle 14,15. Furthermore, as an Ixodes tick consumes a bloodmeal Borrelia replicate and migrate from the midgut into the hemocoel, where they travel to the salivary glands and are transmitted into the host with the expelled saliva 9,16-19.
As a tick feeds the host typically responds with a strong hemostatic and innate immune response 11,13,20-22. Despite these host responses, I. scapularis can feed for several days because tick saliva contains proteins that are immunomodulatory, lytic agents, anticoagulants, and fibrinolysins to aid the tick feeding 3,11,20,21,23. The immunomodulatory activities possessed by tick saliva or salivary gland extract (SGE) facilitate transmission, proliferation, and dissemination of numerous tick-borne pathogens 3,20,24-27. To further understand how tick-borne infectious agents cause disease it is essential to dissect actively feeding ticks and collect tick saliva. This video protocol demonstrates dissection techniques for the collection of hemolymph and the removal of salivary glands from actively feeding I. scapularis nymphs after 48 and 72 hours post mouse placement. We also demonstrate saliva collection from an adult female I. scapularis tick.

Saliva, Salivary Gland, and Hemolymph Collection from Ixodes scapularis Ticks

The collection of infected tick hemolymph, salivary glands, and saliva is important to study how tick-borne pathogens cause disease. In this protocol we demonstrate how to collect hemolymph and salivary glands from feeding Ixodes scapularis nymphs. We also demonstrate saliva collection from female I. scapularis adults.

Ticks are found worldwide and afflict humans with many tick-borne illnesses. Ticks are vectors for pathogens that cause Lyme disease and tick-borne ...

Monitoring the Assembly of a Secreted Bacterial Virulence Factor Using Site-specific Crosslinking

This article describes a method to detect and analyze dynamic interactions between a protein of interest and other factors in vivo. Our method is based on the amber suppression technology that was originally developed by Peter Schultz and colleagues1. An amber mutation is first introduced at a specific codon of the gene encoding the protein of interest. The amber mutant is then expressed in E. coli together with genes encoding an amber suppressor tRNA and an amino acyl-tRNA synthetase derived from Methanococcus jannaschii. Using this system, the photo activatable amino acid analog p-benzoylphenylalanine (Bpa) is incorporated at the amber codon. Cells are then irradiated with ultraviolet light to covalently link the Bpa residue to proteins that are located within 3-8 &#197;. Photocrosslinking is performed in combination with pulse-chase labeling and immunoprecipitation of the protein of interest in order to monitor changes in protein-protein interactions that occur over a time scale of seconds to minutes. We optimized the procedure to study the assembly of a bacterial virulence factor that consists of two independent domains, a domain that is integrated into the outer membrane and a domain that is translocated into the extracellular space, but the method can be used to study many different assembly processes and biological pathways in both prokaryotic and eukaryotic cells. In principle interacting factors and even specific residues of interacting factors that bind to a protein of interest can be identified by mass spectrometry.

This article illustrates the use of pulse-chase radio labeling in combination with site-specific photocrosslinking to monitor interactions between a protein of interest and other factors in E. coli. Unlike traditional chemical cross-linking methods, this approach generates high resolution &#8220;snapshots&#8221; of an ordered assembly pathway in a living cell.

This article describes a method to detect and analyze dynamic interactions between a protein of interest and other factors in vivo. Our method is based on ...

Generation and Multi-phenotypic High-content Screening of Coxiella burnetii Transposon Mutants

Invasion and colonization of host cells by bacterial pathogens depend on the activity of a large number of prokaryotic proteins, defined as virulence factors, which can subvert and manipulate key host functions. The study of host/pathogen interactions is therefore extremely important to understand bacterial infections and develop alternative strategies to counter infectious diseases. This approach however, requires the development of new high-throughput assays for the unbiased, automated identification and characterization of bacterial virulence determinants. Here, we describe a method for the generation of a GFP-tagged mutant library by transposon mutagenesis and the development of high-content screening approaches for the simultaneous identification of multiple transposon-associated phenotypes. Our working model is the intracellular bacterial pathogen Coxiellaburnetii, the etiological agent of the zoonosis Q fever, which is associated with severe outbreaks with a consequent health and economic burden. The obligate intracellular nature of this pathogen has, until recently, severely hampered the identification of bacterial factors involved in host pathogen interactions, making of Coxiella the ideal model for the implementation of high-throughput/high-content approaches.

Generation and Multi-phenotypic High-content Screening of Coxiella burnetii Transposon Mutants

Coxiella burnetii is an obligate intracellular Gram-negative bacterium responsible for the zoonotic disease Q fever. Here we describe methods for the generation of Coxiella fluorescent transposon mutants as well as the automated identification and analysis of the resulting internalization, replication and cytotoxic phenotypes.

Invasion and colonization of host cells by bacterial pathogens depend on the activity of a large number of prokaryotic proteins, defined as virulence ...

Applying an Inducible Expression System to Study Interference of Bacterial Virulence Factors with Intracellular Signaling

The technique presented here allows one to analyze at which step a target protein, or alternatively a small molecule, interacts with the components of a signaling pathway. The method is based, on the one hand, on the inducible expression of a specific protein to initiate a signaling event at a defined and predetermined step in the selected signaling cascade. Concomitant expression, on the other hand, of the gene of interest then allows the investigator to evaluate if the activity of the expressed target protein is located upstream or downstream of the initiated signaling event, depending on the readout of the signaling pathway that is obtained. Here, the apoptotic cascade was selected as a defined signaling pathway to demonstrate protocol functionality. Pathogenic bacteria, such as Coxiella burnetii, translocate effector proteins that interfere with host cell death induction in the host cell to ensure bacterial survival in the cell and to promote their dissemination in the organism. The C. burnetii effector protein CaeB effectively inhibits host cell death after induction of apoptosis with UV-light or with staurosporine. To narrow down at which step CaeB interferes with the propagation of the apoptotic signal, selected proteins with well-characterized pro-apoptotic activity were expressed transiently in a doxycycline-inducible manner. If CaeB acts upstream of these proteins, apoptosis will proceed unhindered. If CaeB acts downstream, cell death will be inhibited. The test proteins selected were Bax, which acts at the level of the mitochondria, and caspase 3, which is the major executioner protease. CaeB interferes with cell death induced by Bax expression, but not by caspase 3 expression. CaeB, thus, interacts with the apoptotic cascade between these two proteins.

The method described here is used to induce the apoptotic signaling cascade at defined steps in order to dissect the activity of an anti-apoptotic bacterial effector protein. This method can also be used for inducible expression of pro-apoptotic or toxic proteins, or for dissecting interference with other signaling pathways.

The technique presented here allows one to analyze at which step a target protein, or alternatively a small molecule, interacts with the components of a ...

Zebrafish as a Model to Assess the Teratogenic Potential of Nitrite

High nitrate levels in the environment may&#160;result in congenital defects or miscarriages in humans. Presumably, this is due to the conversion of nitrate to nitrite by gut and salivary bacteria. However, in other mammalian studies, high nitrite levels do not cause birth defects, although they can lead to poor reproductive outcomes. Thus, the teratogenic potential of nitrite is not clear. It would be useful to have a vertebrate model system to easily assess teratogenic effects of nitrite or any other chemical of interest. Here, we demonstrate the utility of zebrafish (Danio rerio) to screen compounds for toxicity and embryonic defects. Zebrafish embryos are fertilized externally and have rapid development, making them a good model for teratogenic studies. We show that increasing the time of exposure to nitrite negatively affects survival. Increasing the concentration of nitrite also adversely affects survival, whereas nitrate does not. For embryos that survive nitrite exposure, various defects can occur, including pericardial and yolk sac&#160;edema, swim bladder noninflation, and craniofacial malformation. Our results indicate that the zebrafish is a convenient system for studying the teratogenic potential of nitrite. This approach can easily be adapted to test other chemicals for their effects on early vertebrate development.

Exposure to teratogens may cause birth defects. Zebrafish are useful for determining the teratogenic potential of chemicals. We demonstrate the utility of zebrafish by exposing embryos to various levels of nitrite and also at different times of exposure. We show that nitrite can be toxic and cause severe developmental defects.

High nitrate levels in the environment may&#160;result in congenital defects or miscarriages in humans. Presumably, this is due to the conversion of ...

The Madagascar Hissing Cockroach as an Alternative Non-mammalian Animal Model to Investigate Virulence, Pathogenesis, and Drug Efficacy

Many aspects of innate immunity are conserved between mammals and insects. An insect, the Madagascar hissing cockroach from the genus Gromphadorhina, can be utilized as an alternative animal model for the study of virulence, host-pathogen interaction, innate immune response, and drug efficacy. Details for the rearing, care and breeding of the hissing cockroach are provided. We also illustrate how it can be infected with bacteria such as the intracellular pathogens Burkholderia mallei, B. pseudomallei, and B. thailandensis. Use of the hissing cockroach is inexpensive and overcomes regulatory issues dealing with the use of mammals in research. In addition, results found using the hissing cockroach model are reproducible and similar to those obtained using mammalian models. Thus, the Madagascar hissing cockroach represents an attractive surrogate host that should be explored when conducting animal studies.

We present a protocol to utilize the Madagascar hissing cockroach as an alternative non-mammalian animal model to conduct bacterial virulence, pathogenesis, drug toxicity, drug efficacy, and innate immune response studies.

Many aspects of innate immunity are conserved between mammals and insects. An insect, the Madagascar hissing cockroach from the genus Gromphadorhina, can ...

Identification of Virulence Markers of Mycobacterium abscessus for Intracellular Replication in Phagocytes

What differentiates Mycobacterium abscessus from other saprophytic mycobacteria is the ability to resist phagocytosis by human macrophages and the ability to multiply inside such cells. These virulence traits render M. abscessus pathogenic, especially in vulnerable hosts with underlying structural lung disease, such as cystic fibrosis, bronchiectasis or tuberculosis. How patients become infected with M. abscessus remains unclear. Unlike many mycobacteria, M. abscessus is not found in the environment but might reside inside amoebae, environmental phagocytes that represent a potential reservoir for M. abscessus. Indeed, M. abscessus is resistant to amoebal phagocytosis and the intra-amoeba life seems to increase M. abscessus virulence in an experimental model of infection. However, little is known about M. abscessus virulence in itself. To decipher the genes conferring an advantage to M. abscessus intracellular life, a screening of a M. abscessus transposon mutant library was developed. In parallel, a method of RNA extraction from intracellular Mycobacteria after co-culture with amoebae was developed. This method was validated and allowed the sequencing of whole M. abscessus transcriptomes inside the cells; providing, for the first time, a global view on M. abscessus adaptation to intracellular life. Both approaches give us an insight into M. abscessus virulence factors that enable M. abscessus to colonize the airways in humans.

Identification of Virulence Markers of Mycobacterium abscessus for Intracellular Replication in Phagocytes

Here, we present two protocols to study Phagocyte-Mycobacterium abscessus interactions: the screening of a transposon mutant library for bacterial intracellular deficiency and the determination of bacterial intracellular transcriptome from RNA sequencing. Both approaches provide insight into the genomic advantages and transcriptomic adaptations enhancing intracellular bacteria fitness.

What differentiates Mycobacterium abscessus from other saprophytic mycobacteria is the ability to resist phagocytosis by human macrophages and the ability ...

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

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

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

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

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

A High-throughput Shigella-specific Bactericidal Assay

Serum bactericidal assays (SBAs) measure the functional activity of antibodies and have been used for many decades. SBAs directly measure antibody killing activity by assessing the ability of antibodies in serum to bind to bacteria and activate complement. This complement activation results in the lysis and killing of the target bacteria. These assays are valuable because they go beyond quantifying antibody production to elucidate the biological functions that these antibodies have, allowing researchers to study the role that antibodies may play in preventing infection. SBAs have been used to study immune responses for many human pathogens, but there is no widely accepted methodology for Shigella at present. Historically, SBAs have been very labor-intensive, requiring many time-consuming steps to accurately quantify surviving bacteria. This protocol describes a simple, robust, and high-throughput assay that measures functional antibodies specific for Shigella in serum in vitro. The method described here offers many advantages over traditional SBAs, including the use of frozen bacterial stocks, 96 well assay plates, a micro-culture system, and automated colony-counting. All of these modifications make this assay less labor-intensive and more high-throughput. This protocol is simpler and faster to perform than traditional SBAs while still using simple technologies and readily available reagents. The protocol has been successfully applied in multiple independent laboratories and the assay is robust and reproducible. The assay can be used to assess immune responses in pre-clinical as well as clinical studies. Quantifying shigellacidal antibody titers both before and after antigen exposure (either by immunization or infection) allows for a broader understanding of how functional antibody responses are generated and their contribution to protective immunity. The development of this standardized, well-characterized assay may greatly facilitate Shigella vaccine design.

A High-throughput Shigella-specific Bactericidal Assay

Here we present a protocol to measure Shigellacidal activity of antibodies in serum. Serum is mixed with bacteria and exogenous complement, incubated, and the reaction mixture is plated on agar plates. Viable bacteria form colonies which are counted, using an automated colony enumerator, and used to determine the bactericidal titer.

Serum bactericidal assays (SBAs) measure the functional activity of antibodies and have been used for many decades. SBAs directly measure antibody killing ...

Opsonophagocytic Killing Assay to Assess Immunological Responses Against Bacterial Pathogens

A key aspect of the immune response to bacterial colonization of the host is phagocytosis. An opsonophagocytic killing assay (OPKA) is an experimental procedure in which phagocytic cells are co-cultured with bacterial units. The immune cells will phagocytose and kill the bacterial cultures in a complement-dependent manner. The efficiency of the immune-mediated cell killing is dependent on a number of factors and can be used to determine how different bacterial cultures compare with regard to resistance to cell death. In this way, the efficacy of potential immune-based therapeutics can be assessed against specific bacterial strains and/or serotypes. In this protocol, we describe a simplified OPKA that utilizes basic culture conditions and cell counting to determine bacterial cell viability after co-culture with treatment conditions and HL-60 immune cells. This method has been successfully utilized with a number of different pneumococcal serotypes, capsular and acapsular strains, and other bacterial species. The advantages of this OPKA protocol are its simplicity, versatility (as this assay is not limited to antibody treatments as opsonins), and minimization of time and reagents to assess basic experimental groups.

This opsonophagocytic killing assay is used to compare the ability of phagocytic immune cells to respond to and kill bacteria based on different treatments and/or conditions. Classically, this assay serves as the gold&#160;standard for assessing effector functions of antibodies raised against a bacterium as opsonin.

A key aspect of the immune response to bacterial colonization of the host is phagocytosis. An opsonophagocytic killing assay (OPKA) is an experimental ...