The authors wish to thank Nicole Hasenkampf and Amanda Tardo for technical support. We also thank Drs. Linden Hu and Adriana Marques for recommendation of the LeFlap containment device, and Dr. Lise Gern for instruction on the capillary feeding method. This work was supported by NIH/NCRR Grant 8 P20 GM103458-09 (MEE) and by the National Center for Research Resources and the Office of Research Infrastructure Programs (ORIP) of the National Institutes of Health through grant P51OD011104/P51RR000164.

An experimental outline of tick inoculation and feeding upon animals for Lyme disease research is depicted in Figure 1.
1. Inoculating Nymphal Ixodes Ticks with B. burgdorferi Using Capillary Tube-feeding
When performing manipulations with ticks, white lab coats with elastic sleeves, gloves, and disposable bouffant caps are worn.
<ol>
	<li>Our technique is a modified version of that reported by Broadwater et al. 49. Prepare capillary tubes by heating and pulling Pasteur pipets to breaking thinness using a pipet puller. Using forceps and dissecting scope, break the tips to the optimal diameter (about 0.2 mm). A tube standardized to the tick mouthpart size is used as a sizing guide. A face shield should be worn when preparing the pipets.</li>
	<li>Grow B. burgdorferi to between 2-8 x 107/ml (mid-log phase) in BSK-H medium (Sigma) containing 6% rabbit serum.</li>
	<li>Use nymphal ticks that have been stored at 23 &deg;C for 4-6 weeks post-larval molt. Place ticks onto a small 60 x 15 mm petri dish with double-sided tape on the outer bottom surface of the dish. Place the ticks ventral-side facing up.</li>
	<li>Dip the capillary tube tip into the B. burgdorferi culture tube after mixing. Place the capillary tube over the hypostome of the tick mouth parts using a dissecting scope. Use molding clay to fix the tube in place, as shown in Figure 2A.</li>
	<li>Place the petri dishes with affixed ticks inside a large clear plastic tub for an added level of containment. Wet paper towels are added to provide moisture. Place the ticks in a 37 &deg;C thermal incubator for 30 min-2 hr until defecation is apparent. This indicates that the media containing spirochetes has passed through the tick.</li>
	<li>Rest the ticks for 2-4 weeks at 23 &deg;C to allow adaptation to the tick environment before feeding them on animals.</li>
</ol>
2. Infecting Mice with B. burgdorferi by Tick
<ol>
	<li>Dilute ketamine stock 1:10 in sterile water. Anesthetize each mouse with 100 mg/kg of ketamine by intraperitoneal injection with a tuberculin syringe</li>
	<li>Once the mouse is fully anesthetized, shave the mouse from the ears to the middle back using a fine (Remington smooth &amp; silky) electric trimmer.</li>
	<li>In a white pan with no other objects nearby, transfer the nymphal ticks (that harbor B. burgdorferi) by a moistened paintbrush to the hairless area of the mouse. Alternatively, uninfected ticks can be placed on mice for xenodiagnosis of mice with suspect infection. The use of a clean, white surface for tick placement helps to ensure that any unattached ticks will be readily seen.</li>
	<li>Place the mouse in specialized caging (Allentown Caging, Allentown, PA). The caging consists of a stainless steel grill elevated from the bottom of the cage. The cage top was modified by our in-house machine shop to elevate the water bottle holder enough to allow free movement of the mouse underneath. The pan is filled with about &frac12; inch of water to trap any ticks that fall off mice (Figure 3A). To minimize the risk of hypothermia, reusable heating pads, microwaved prior to use, are placed underneath the cages until mice wake completely from anesthesia. Animals are often ataxic as they recover from anesthesia and rub against food and water trays, so these must be removed. The water level is low enough to prevent limbs of mice from submersion.</li>
	<li>Place the cage inside a tray that has been lined with Tangle trap paste (Contech, Victoria, BC, Canada) and tape to ensure the entrapment of arthropods. Mice are caged singly and observed continuously during the period of anesthesia.</li>
	<li>Within 2 hr, when mice are completely awake from the anesthesia, the food tray and water bottle are replaced to the cage. After 24 hr, the housing enrichment consisting of a plastic hut and nylabone is replaced.</li>
	<li>After 3, 4, and 5 days, check the mouse, cage and cage water for fed ticks. The cage water is sifted through a white metal pan (i.e. "panning for gold"). Rinse fed ticks in clean water and store in plastic jars (Figure 3B). On days 3 and 4, replace water in cage with clean water. On day 5, check not only the cage, but the mouse thoroughly for ticks. Usually by this point all ticks have fed and the mice can be returned to regular caging.</li>
	<li>Place all waste from the mouse cages, including liquids, in biohazard containers for autoclaving and disposal. Keep a log of the number of ticks placed on mice and those recovered at all times.</li>
</ol>
3. Feeding Ticks on Nonhuman Primates for Infection with B. burgdorferi or xenodiagnosis
<ol>
	<li>Prepare the tick containment device: Cut a 1 &frac34; inch diameter circle in the 3 inch x 3 inch LeFlap (flap) using a clean scalpel and the measurement guide. Use the cut-out as a template to cut circles of identical size in the Biatane foam and Duoderm. The foam is used to elevate flap over the surface of the skin and prevent possible crushing of tick. The Duoderm adds another layer of cushioning and overlays the edges of the containment device for added security from tick escape. A diagram of the containment device is depicted in Figure 4.</li>
	<li>Veterinary staff will anesthetize the animal with 5-8 mg/kg Telazol by intramuscular injection.</li>
	<li>Clip the hair of the animal using electric trimmer (Oster) equipped with size 40 blades. All areas that will be covered by the jacket are clipped: back, front, upper arms. Using shaving cream and dual-blade disposable razors, closely shave an area of approximately 25 cm vertical x 20 cm horizontal. Wipe clean with moist paper towels and blow dry with low heat to dry the skin.</li>
	<li>Place the flap on the animal's dorsum, just below the scapula, on either side of the spine. Use a marker to trace the circle in that spot. Prepare the area of skin around the circle by wiping it with SkinPrep. This removes oil in skin that could affect adhesion of glue and containment devices. Leaving about a 1 cm circumference of space around the circle, apply a layer of skin glue (SkinBond) with a width of ~4 cm.</li>
	<li>Remove the adhesive backing from the Biatane foam and affix to the skin in the appropriate spot. Animals are again anesthetized by veterinary staff with 5 mg/kg Telazol. Seal the edges with skin glue and Hypafix tape. Remove the adhesive backing from the flap and affix on top of the Biatane. Place Hypafix tape around the edges of the LeFlap, then tape down the mesh flap of the flap and place the jacket on the animal. Tape and Tangle Trap paste is applied to the floor in a perimeter surrounding the nonhuman primate caging for added security.</li>
	<li>To minimize the effects of chemicals used in Step 3.4 on tick feeding, the ticks are added 24 hr after the containment device is in place. At this point, the security of the device is also inspected and reinforced if needed. Typically, 20 unfed nymphs (4-8 weeks post-larval molt) are added to the skin within the device using a paintbrush.</li>
	<li>Remove the adhesive backing from the mesh of the flap and seal it in place. Lastly, remove the Duoderm backing to expose adhesive, and place on top of the containment device. Add a piece of Hypafix tape across the open mesh circle, and replace the jacket. The completed containment device is shown in Figure 5A.</li>
	<li>After 5 days, anesthetize the animals as above and the jackets are removed. Remove the tape first to inspect tick feeding through the mesh (Figure 5B). Carefully peel the Duoderm away from the flap.</li>
	<li>Pull back the mesh portion at the edges to provide access to the ticks. Fed ticks are frequently found near or under the foam circle (Figure 5C) and are removed and placed in clean water with a paintbrush. Remove the device once all visible fed ticks are collected (Figure 5D).</li>
</ol>
Note: Oftentimes, the containment device can simply be peeled away from the skin. If adhesion is strong and could potentially damage the skin, Unisolve solvent is applied to the area for gentle removal. The skin is wiped with isopropanol and ticks are stored at 23 &deg;C. If used for infection, the ticks can be crushed to confirm the number that contained B. burgdorferi. If used for xenodiagnosis, the ticks are kept for 1-3 weeks prior to analysis of midgut contents.

The authors declare that they have no competing financial interests.

In order to obtain ticks that harbor B. burgdorferi for downstream studies, the ticks can be: (1) fed on infected mice at the larval stage; (2) immersed in B. burgdorferi cultures at either the larval or nymphal stage 48; (3) microinjected with B. burgdorferi 47; or (4) capillary tube-fed B. burgdorferi 49. While each of these methods has its purpose, for ensuring that a large portion of the ticks to be used for infection harbor B. burgdorferi,...

In 2011, Lyme disease was the 6th most common Nationally Notifiable disease in North America (<a href="http://www.cdc.gov/lyme/stats/index.html" target="_blank">http://www.cdc.gov/lyme/stats/index.html</a>). B. burgdorferi is a versatile microbe, both genetically and antigenically (reviewed in 1). Its genetic constitution includes a large (&gt;900 kB) chromosome and up to 21 plasmids (12 linear, 9 circular), with plasmid content varying among isolates. Much is to be learned about this spirochete, as over 90% of the plasmid open reading frames are unrelated to any known bacterial sequences 2,3 . B. burgdorferi presents a wide variety of antigens as potential targets of host immunity. However, an untreated infection often persists. The interaction of spirochetes with the tick milieu and the vertebrate host environment necessitates adaptation by B. burgdorferi throughout the infection process. Several plasmid-encoded genes are known to be differentially expressed in response to changes in temperature, pH, cell density and even stage of the tick life cycle 4-8.
The study of B. burgdorferi adaptation throughout its enzootic cycle, and host responses following infection by the natural route relies on the ability to feed ticks on appropriate animal models. Such studies are met with the technical challenges of generating ticks that harbor B. burgdorferi, and ensuring the efficient transmission and/or feeding of ticks on the model host. In addition, the containment and recovery of infected ticks is essential. Among the models used are mice and nonhuman primates, each of which serves as a valuable tool in Lyme disease research. As with the white-footed mouse, which is a natural reservoir host for B. burgdorferi, the laboratory mouse is a highly susceptible host that supports persistent infection by B. burgdorferi 9. Following infection of disease-susceptible mice, such as the C3H strain, the spirochetes disseminate to multiple tissues, including the skin, bladder, muscles, joints and heart. Inflammatory responses to the infection lead to diseased heart and joint tissue. While the spirochetes persist in this host and remain infectious, inflammatory lesions may become intermittent, not unlike the process in humans. The mouse model has thus provided much information on B. burgdorferi-induced pathology, including arthritis and carditis and host immune responses 10-12. From the perspective of the pathogen, certain genes differentially expressed during mammalian infection have been characterized, as have some necessary for transmission from the tick vector 13-21.
Though several animal species have been used to study Lyme disease 22, rhesus macaques most closely mimic the multi-organ character of human disease 23. Unlike other animal models, the breadth of disease manifestations such as erythema migrans, carditis, arthritis, and neuropathy of the peripheral and central nervous systems are observed in macaques. In mice, the reservoir host for B. burgdorferi, disease varies by mouse strain and age 24, while the early and late-disseminated manifestations are uncommon 9. In addition, other rodents, lagomorphs, and canines all fail to exhibit neurological disease from B. burgdorferi infection 25. Importantly, macaques exhibit signs that are characteristic of all three phases of Lyme borreliosis, namely, early-localized, early-disseminated, and late-stage Lyme disease 26-28. Erythema migrans (EM) is thought to occur in 70-80% of human cases 29, and is also seen in rhesus macaques 28,30 . Following infection, the spirochetes disseminate from the site of inoculation to multiple organs. Spirochetal DNA has been detected in skeletal muscles, heart, bladder, peripheral nerve and plexus, as well as in the central nervous system (cerebrum, brainstem and cerebellum, spinal cord, and dura mater) 31.
Tick feeding on mice has been utilized by us and other research teams for propagation of tick colonies, in reservoir competence studies 32-36 and in studies of B. burgdorferi pathogenesis 37-40. This technique has also been used for xenodiagnosis and testing of vaccine efficacy in mice 41-44. We have fed Ixodes ticks on nonhuman primates for model development 28, a study of vaccine efficacy 45, and for xenodiagnosis in the assessment of persistence post-antibiotic treatment 46. Ticks that harbor B. burgdorferi can be maintained in a natural enzootic cycle by feeding larvae on infected mice and using the nymphs for studies, as the spirochetes are transmitted through the life stages. In this report, we instruct on how to generate ticks infected with wild type or mutant B. burgdorferi, using capillary tube-feeding. This can also be accomplished by microinjection 47 and by immersion 48. The purpose of artificial introduction of B. burgdorferi into ticks can be to study mutant strains whose transmissibility is unknown, to generate a group of ticks with a high infection rate, and to reduce potential for error by maintaining a clean and otherwise uninfected tick colony. In addition, we demonstrate tick feeding on mice and nonhuman primates, so as to assure containment and recovery of replete ticks. The use of tick feeding is essential for future studies of immune responses to B. burgdorferi infection, potential Lyme vaccine efficacy, and xenodiagnosis for detection of occult infections.

<table border="1">
		<tbody>
		 <tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Reagent</td>
		 </tr>
		 <tr>
			<td>BSK-H</td>
			<td>Sigma</td>
			<td>B-8291</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Ketamine HCl</td>
			<td></td>
			<td></td>
			<td></td>
		 </tr>
		 <tr>
			<td>Tangle Trap coating Paste</td>
			<td>Ladd research</td>
			<td>T-131</td>
			<td></td>
		 </tr>
		 <tr>
			<td>SkinPrep</td>
			<td>Allegro Medical Supplies</td>
			<td>177364</td>
			<td></td>
		 </tr>
		 <tr>
			<td>LeFlap, 3" x 3"</td>
			<td>Monarch Labs</td>
			<td></td>
			<td></td>
		 </tr>
		 <tr>
			<td>Hypafix tape</td>
			<td>Allegro Medical Supplies</td>
			<td>191523</td>
			<td></td>
		 </tr>
		 <tr>
			<td>SkinBond</td>
			<td>Allegro Medical Supplies</td>
			<td>554536</td>
			<td></td>
		 </tr>
		 <tr>
			<td>UniSolve</td>
			<td>Allegro Medical Supplies</td>
			<td>176640</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Biatane Foam, adhesive 4"x4"</td>
			<td>Coloplast</td>
			<td>3420</td>
			<td></td>
		 </tr>
		 <tr>
			<td>DuoDerm CGF Dressing - 4" x 4", (3/4)" adhesive border </td>
			<td>Convatec</td>
			<td>187971 </td>
			<td></td>
		 </tr>
		 <tr>
			<td>Nonhuman primate jackets with flexible 2" back panels; add drawstrings at top and bottom</td>
			<td>Lomir Biomedical Inc.</td>
			<td></td>
			<td></td>
		 </tr>
		 <tr>
			<td></td>
			<td></td>
			<td></td>
			<td>EQUIPMENT </td>
		 </tr>
		 <tr>
			<td>Pipet puller</td>
			<td>David Kopf Instruments</td>
			<td>Model 700C</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Dark field microscope</td>
			<td>Leitz Wetzlar</td>
			<td>Dialux</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Dissecting microscope</td>
			<td>Leica</td>
			<td>Zoom 2000</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Mouse caging</td>
			<td>Allentown caging</td>
			<td></td>
			<td></td>
		 </tr>
		</tbody>
 </table>

feeding of ticks on animals for transmission and xenodiagnosis in lyme disease research

Transmission of the etiologic agent of Lyme disease, Borrelia burgdorferi, occurs by the attachment and blood feeding of Ixodes species ticks on mammalian hosts. In nature, this zoonotic bacterial pathogen may use a variety of reservoir hosts, but the white-footed mouse (Peromyscus leucopus) is the primary reservoir for larval and nymphal ticks in North America. Humans are incidental hosts most frequently infected with B. burgdorferi by the bite of ticks in the nymphal stage. B. burgdorferi adapts to its hosts throughout the enzootic cycle, so the ability to explore the functions of these spirochetes and their effects on mammalian hosts requires the use of tick feeding. In addition, the technique of xenodiagnosis (using the natural vector for detection and recovery of an infectious agent) has been useful in studies of cryptic infection. In order to obtain nymphal ticks that harbor B. burgdorferi, ticks are fed live spirochetes in culture through capillary tubes. Two animal models, mice and nonhuman primates, are most commonly used for Lyme disease studies involving tick feeding. We demonstrate the methods by which these ticks can be fed upon, and recovered from animals for either infection or xenodiagnosis.

Following the completion of capillary feeding, the ticks are typically rested at 23 &deg;C for 2-3 weeks before they are fed on animals for transmission. Using the capillary-feeding technique, we have found that over 90% of the fed ticks harbor B. burgdorferi. The percentage of positive ticks is determined by washing ticks in peroxide and ethanol, then crushing them in sterile PBS with a microfuge tube-shaped pestle. The midgut contents spilled into the PBS are fixed on slides and stained with an anti-Borrel...

Watch this Scientific Journal Video about Feeding of Ticks on Animals for Transmission and Xenodiagnosis in Lyme Disease Research at JoVE.com

Feeding of Ticks on Animals for Transmission and Xenodiagnosis in Lyme Disease Research

Lyme disease is the most commonly-reported vector-borne disease in North America. The causative agent, Borrelia burgdorferi is a spirochete bacterium transmitted by Ixodid ticks. Transmission and detection of infection in animal models is optimized by the use of tick feeding, which we describe here.

Transmission of the etiologic agent of Lyme disease, Borrelia burgdorferi, occurs by the  attachment and blood feeding of Ixodes species ticks on ...

feeding-ticks-on-animals-for-transmission-xenodiagnosis-lyme-disease

Research

JoVE Journal

Immunology and Infection

13.1K Views.  Tulane University Health Sciences Center. Lyme disease is the most commonly-reported vector-borne disease in North America. The causative agent, Borrelia burgdorferi is a spirochete bacterium transmitted by Ixodid ticks. Transmission and detection of infection in animal models is optimized by the use of tick feeding, which we describe here.

Video: Feeding of Ticks on Animals for Transmission and Xenodiagnosis in Lyme Disease Research

RNA Interference in Ticks

Ticks are obligate hematophagous ectoparasites of wild and domestic animals and humans, and are considered to be second worldwide to mosquitoes as vectors of human diseases1 and the most important vectors affecting cattle industry worldwide2. Ticks are classified in the subclass Acari, order Parasitiformes, suborder Ixodida and are distributed worldwide from Arctic to tropical regions3. Despite efforts to control tick infestations, these ectoparasites remain a serious problem for human and animal health4,5.
RNA interference (RNAi)6 is a nucleic acid-based reverse genetic approach that involves disruption of gene expression in order to determine gene function or its effect on a metabolic pathway. Small interfering RNAs (siRNAs) are the effector molecules of the RNAi pathway that is initiated by double-stranded RNA (dsRNA) and results in a potent sequence-specific degradation of cytoplasmic mRNAs containing the same sequence as the dsRNA trigger7-9. Post-transcriptional gene silencing mechanisms initiated by dsRNA have been discovered in all eukaryotes studied thus far, and RNAi has been rapidly developed in a variety of organisms as a tool for functional genomics studies and other applications10.
RNAi has become the most widely used gene-silencing technique in ticks and other organisms where alternative approaches for genetic manipulation are not available or are unreliable5,11. The genetic characterization of ticks has been limited until the recent application of RNAi12,13. In the short time that RNAi has been available, it has proved to be a valuable tool for studying tick gene function, the characterization of the tick-pathogen interface and the screening and characterization of tick protective antigens14. Herein, a method for RNAi through injection of dsRNA into unfed ticks is described. It is likely that the knowledge gained from this experimental approach will contribute markedly to the understanding of basic biological systems and the development of vaccines to control tick infestations and prevent transmission of tick-borne pathogens15-19.

A method for RNA interference (RNAi) by injection of dsRNA into unfed ticks is described. RNAi is the most widely used gene-silencing technique in ticks where the use of other methods of genetic manipulation has been limited.

Ticks are obligate hematophagous ectoparasites of wild and domestic animals and humans, and are considered to be second worldwide to mosquitoes as vectors ...

Methods for Rapid Transfer and Localization of Lyme Disease Pathogens Within the Tick Gut

Lyme disease is caused by infection with the spirochete pathogen Borrelia burgdorferi, which is maintained in nature by a tick-rodent infection cycle 1. A tick-borne murine model 2 has been developed to study Lyme disease in the laboratory. While na&iacute;ve ticks can be infected with B. burgdorferi by feeding them on infected mice, the molting process takes several weeks to months to complete. Therefore, development of more rapid and efficient tick infection techniques, such as a microinjection-based procedure, is an important tool for the study of Lyme disease 3,4. The procedure requires only hours to generate infected ticks and allows control over the delivery of equal quantities of spirochetes in a cohort of ticks. This is particularly important as the generation of B. burgdorferi infected ticks by the natural feeding process using mice fails to ensure 100% infection rate and potentially results in variation of pathogen burden amongst fed ticks. Furthermore, microinjection can be used to infect ticks with B. burgdorferi isolates in cases where an attenuated strain is unable to establish infection in mice and thus can not be naturally acquired by ticks 5. This technique can also be used to deliver a variety of other biological materials into ticks, for example, specific antibodies or double stranded RNA 6. In this article, we will demonstrate the microinjection of nymphal ticks with in vitro-grown B. burgdorferi. We will also describe a method for localization of Lyme disease pathogens in the tick gut using confocal immunofluorescence microscopy.

Lyme disease research studies often require generation of ticks infected with the pathogen Borrelia burgdorferi, a process that typically takes several weeks. Here we demonstrate a microinjection-based tick infection procedure that can be accomplished within hours. We also demonstrate an immunofluorescence method for in situ localization of B. burgdorferi within ticks.

Lyme disease is caused by infection with the spirochete pathogen Borrelia burgdorferi, which is maintained in nature by a tick-rodent infection cycle 1.  ...

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 ...

Swab Sampling Method for the Detection of Human Norovirus on Surfaces

Human noroviruses are a leading cause of epidemic and sporadic gastroenteritis worldwide. Because most infections are either spread directly via the person-to-person route or indirectly through environmental surfaces or food, contaminated fomites and inanimate surfaces are important vehicles for the spread of the virus during norovirus outbreaks.
We developed and evaluated a protocol using macrofoam swabs for the detection and typing of human noroviruses from hard surfaces. Compared with fiber-tipped swabs or antistatic wipes, macrofoam swabs allow virus recovery (range 1.2-33.6%) from toilet seat surfaces of up to 700 cm2. The protocol includes steps for the extraction of the virus from the swabs and further concentration of the viral RNA using spin columns. In total, 127 (58.5%) of 217 swab samples that had been collected from surfaces in cruise ships and long-term care facilities where norovirus gastroenteritis had been reported tested positive for GII norovirus by RT-qPCR. Of these 29 (22.8%) could be successfully genotyped. In conclusion, detection of norovirus on environmental surfaces using the protocol we developed may assist in determining the level of environmental contamination during outbreaks as well as detection of virus when clinical samples are not available; it may also facilitate monitoring of effectiveness of remediation strategies.

A macrofoam based sampling methodology was developed and evaluated for the detection and quantification of norovirus on environmental hard surfaces.

Human noroviruses are a leading cause of epidemic and sporadic gastroenteritis worldwide. Because most infections are either spread directly via the ...

Protocols for Investigating the Host-tissue Distribution, Transmission-mode, and Effect on the Host Fitness of a Densovirus in the Cotton Bollworm

Many novel viruses have been discovered in animal hosts using next-generation sequencing technologies. Previously, we reported a mutualistic virus, Helicoverpa armigera densovirus (HaDV2), in a lepidopteran species, the cotton bollworm, Helicoverpa armigera (Hubner). Here, we describe the protocols that are currently used to study the effect of HaDV2 on its host. First, we establish a HaDV2-free cotton bollworm colony from a single breeding pair. Then, we orally inoculate some neonate larval offspring with HaDV2-containing filtered liquid to produce two colonies with the same genetic background: one HaDV2-infected, the other uninfected. A protocol to compare life table parameters (e.g., larval, pupal, and adult periods and fecundity) between the HaDV2-infected and -uninfected individuals is also presented, as are the protocols for determining the host-tissue distribution and transmission efficiency of HaDV2. These protocols would also be suitable for investigating the effects of other orally transmitted viruses on their insect hosts, lepidopteran hosts in particular.

Here, we present a protocol to investigate the host-tissue distribution, transmission mode, and effect on the host fitness of a densovirus within a lepidopteran species, the cotton bollworm. This protocol can also be used for studying the interaction between other orally-transmitted viruses and their insect hosts.

Many novel viruses have been discovered in animal hosts using next-generation sequencing technologies. Previously, we reported a mutualistic virus, ...

Assessment of the Cytotoxic and Immunomodulatory Effects of Substances in Human Precision-cut Lung Slices

Respiratory diseases in their broad diversity need appropriate model systems to understand the underlying mechanisms and enable development of new therapeutics. Additionally, registration of new substances requires appropriate risk assessment with adequate testing systems to avoid the risk of individuals being harmed, for example, in the working environment. Such risk assessments are usually conducted in animal studies. In view of the 3Rs principle and public skepticism against animal experiments, human alternative methods, such as precision-cut lung slices (PCLS), have been evolving. The present paper describes the ex vivo technique of human PCLS to study the immunomodulatory potential of low-molecular-weight substances, such as ammonium hexachloroplatinate (HClPt). Measured endpoints include viability and local respiratory inflammation, marked by altered secretion of cytokines and chemokines. Pro-inflammatory cytokines, tumor necrosis factor alpha (TNF-&#945;), and interleukin 1 alpha (IL-1&#945;) were significantly increased in human PCLS after exposure to a sub-toxic concentration of HClPt. Even though the technique of PCLS has been substantially optimized over the past decades, its applicability for the testing of immunomodulation is still in development. Therefore, the results presented here are preliminary, even though they show the potential of human PCLS as a valuable tool in respiratory research.

In view of the 3Rs principle, respiratory models as alternatives to animal studies are evolving. Especially for risk assessment of respiratory substances, there is a lack of appropriate assays. Here, we describe the use of human precision-cut lung slices for the assessment of airborne substances.

Respiratory diseases in their broad diversity need appropriate model systems to understand the underlying mechanisms and enable development of new ...

Collection and Processing of Lymph Nodes from Large Animals for RNA Analysis: Preparing for Lymph Node Transcriptomic Studies of Large Animal Species

Large animals (both livestock and wildlife) serve as important reservoirs of zoonotic pathogens, including Brucella, Mycobacterium bovis, Salmonella, and E. coli, and are useful for the study of pathogenesis and/or spread of the bacteria in natural hosts. With the key function of lymph nodes in the host immune response, lymph node tissues serve as a potential source of RNA for downstream transcriptomic analyses, in order to assess the temporal changes in gene expression in cells over the course of an infection. This article presents an overview of the process of lymph node collection, tissue sampling, and downstream RNA processing in livestock, using cattle (Bos taurus) as a model, with additional examples provided from the American bison (Bison bison). The protocol includes information about the location, identification, and removal of lymph nodes from multiple key sites in the body. Additionally, a biopsy sampling methodology is presented that allows for a consistency of sampling across multiple animals. Several considerations for sample preservation are discussed, including the generation of RNA suitable for downstream methodologies like RNA-sequencing and RT-PCR. Due to the long delays inherent in large animal vs. mouse time course studies, representative results from bison and bovine lymph node tissues are presented to describe the time course of the degradation in this tissue type, in the context of a review of previous methodological work on RNA degradation in other tissues. Overall, this protocol will be useful to both veterinary researchers beginning transcriptome projects on large animal samples and to molecular biologists interested in learning techniques for in vivo tissue sampling and in vitro processing.

This protocol provides an overview of procedures for the isolation of RNA for the transcriptomic profiling of lymph node tissues from large animals, including steps in the identification and excision of lymph nodes from livestock and wildlife, sampling approaches to provide consistency across multiple animals, and considerations plus representative results for the post-collection preservation and processing for RNA analysis.

Large animals (both livestock and wildlife) serve as important reservoirs of zoonotic pathogens, including Brucella, Mycobacterium bovis, Salmonella, and ...

Sexual Transmission of American Trypanosomes from Males and Females to Naive Mates

American trypanosomiasis is transmitted to humans by triatomine bugs through the ingestion of contaminated food, by blood transfusions or accidently in hospitals and research laboratories. In addition, the Trypanosoma cruzi infection is transmitted congenitally from a&#160;chagasic mother to her offspring, but the male partner&#39;s contribution to in utero contamination is unknown. The findings of nests and clumps of amastigotes and of trypomastigotes in the theca cells of the ovary, in the goniablasts and in the lumen of seminiferous tubules suggest that T. cruzi infections are sexually transmitted. The research protocol herein presents the results of a family study population showing parasite nuclear DNA in the diploid blood mononuclear cells and in the haploid gametes of human subjects. Thus, three independent biological samples collected one year apart confirmed that T. cruzi infections were sexually transmitted to progeny. Interestingly, the specific T. cruzi antibody was absent in the majority of family progeny that bore immune tolerance to the parasite antigen. Immune tolerance was demonstrated in chicken refractory to T. cruzi after the first week of embryonic growth, and chicks hatched from the flagellate-inoculated eggs were unable to produce the specific antibody.&#160;Moreover, the instillation of the human semen ejaculates intraperitoneally or into the vagina of naive mice yielded T. cruzi amastigotes in the epididymis, seminiferous tubule, vas deferens and uterine tube with an absence of inflammatory reactions in the immune privileged organs of reproduction. The breeding of T. cruzi-infected male and female mice with naive mates resulted in acquisition of the infections, which were later transmitted to the progeny. Therefore, a robust education, information and communication program that involves the population and social organizations is deemed necessary to prevent Chagas disease.

The Trypanosoma cruzi agent of Chagas disease produces long-lasting asymptomatic infections that abruptly develop into clinically recognized pathology. The following research protocol describes a short-run family-based epidemiological study to unravel the T. cruzi infection transmitted sexually from parent to progeny.

American trypanosomiasis is transmitted to humans by triatomine bugs through the ingestion of contaminated food, by blood transfusions or accidently in ...

Highly Efficient Transfection of Primary Macrophages with In Vitro Transcribed mRNA

Macrophages are phagocytic cells specialized in detecting molecules of non-self origin. To this end, they are equipped with a large array of pattern recognition receptors (PRRs). Unfortunately, this also makes macrophages particularly challenging to transfect as the transfection reagent and the transfected nucleic acids are often recognized by the PRRs as non-self. Therefore, transfection often results in macrophage activation and degradation of the transfected nucleic acids or even in suicide of the macrophages. Here, we describe a protocol that allows highly efficient transfection of murine primary macrophages such as peritoneal macrophages (PM) and bone marrow-derived macrophages (BMDM) with mRNA in vitro transcribed from DNA templates such as plasmids. With this simple protocol, transfection rates of about 50-65% for PM and about 85% for BMDM are achieved without cytotoxicity or immunogenicity observed. We describe in detail the generation of mRNA for transfection from DNA constructs such as plasmids and the transfection procedure.

Macrophages, especially primary macrophages, are challenging to transfect as they specialize in detecting molecules of non-self origin. We describe a protocol that allows highly efficient transfection of primary macrophages with mRNA generated from DNA templates such as plasmids.

Macrophages are phagocytic cells specialized in detecting molecules of non-self origin. To this end, they are equipped with a large array of pattern ...

Standard Membrane Feeding Assay for the Detection of Plasmodium falciparum Infection in Anopheles Mosquito Vectors

Malaria remains one of the most devastating diseases worldwide and, to date, the African region is still responsible for 94% of all cases worldwide. This parasitic disease requires a protozoan parasite, an Anopheles mosquito vector, and a vertebrate host. The Anopheles genus comprises more than 500 species, of which 60 are known as vectors of the parasite. The Plasmodium parasite genus consists of 250 species, and 48 of these are involved in disease transmission. Furthermore, the Plasmodium falciparum parasite has contributed toward an estimated 99.7% of malaria cases in sub-Saharan Africa in recent years. 
Gametocytes form part of the sexual stage of the parasite and are ingested by the female mosquito upon feeding on an infected human host. Further development of the parasite within the mosquito is enhanced by favorable environmental conditions in the midgut of the mosquito. Here, the fusion of the female and male gametes takes place, and the motile ookinetes originate. The ookinetes enter the midgut epithelium of the mosquito, and mature ookinetes form oocysts, which, in turn, produce motile sporozoites. These sporozoites migrate to the mosquito's salivary glands and are injected as a mosquito takes a blood meal. 
For drug discovery purposes, mosquitoes were artificially infected with gametocyte-infected blood in the standard membrane feeding assay (SMFA). To detect infection within the mosquito and/or to assess the efficacy of antimalarial compounds, the midguts of the female mosquitoes were removed post infection and were stained with mercurochrome. This method was used to enhance the visual detection of oocysts under the microscope for the accurate determination of oocyst prevalence and intensity.

Standard Membrane Feeding Assay for the Detection of Plasmodium falciparum Infection in Anopheles Mosquito Vectors

The standard membrane feeding assay (SMFA) is regarded as the gold standard for the assessment and identification of potential antimalarial compounds. This artificial feeding system is used to infect mosquitoes to further evaluate the effects of such compounds on the intensity and prevalence of the Plasmodium falciparum parasite.

Malaria remains one of the most devastating diseases worldwide and, to date, the African region is still responsible for 94% of all cases worldwide. This ...