Name: a model for experimental exposure of humans to larval ixodes scapularis ticks
Uploaded: 2023-12-01T13:00:00.000+00:00
Duration: 4 min 47 s
Description: Watch this Scientific Journal Video about A Model for Experimental Exposure of Humans to Larval Ixodes scapularis Ticks at JoVE.com

This research was supported by the Intramural Research Program of the NIH, National Institute of Allergy and Infectious Diseases. We thank Linden T. Hu, Sam R. Telford III, Kenneth Dardick, Carla Williams, Erin Chung, and Christina Brandeburg for their participation in the development of the procedures.

Exposing volunteers to tick bites is an experimental method and must be conducted under a clinical research protocol approved by the relevant regulatory authorities. The clinical studies (NCT01143558, NCT02446626, and NCT05036707) were approved by the respective institutional review boards, conducted under investigational device exemptions granted by the US Food and Drug Administration, and carried out in accordance with Good Clinical Practice guidelines. Additionally, these studies were registered with ClinicalTrials.gov, and written informed consent was obtained from all participants.
1. Containment dressing preparation
<ol>
	<li>To create a measurement template, photocopy the 3&#34; x 3&#34; hydrocolloid containment dressing along the side of the measurement lines (Figure 1A and Table of Materials).</li>
	<li>Cut the photocopied template along the 1&#34; line. Then, cut along the 2&#34; line, creating a 2&#34; circle with a 1&#34; center (Figure 1B).</li>
	<li>Using this template, cut the non-adhesive foam dressing to create a 2&#34; circle with a 1&#34; center (Figure 2A,B, and Table of Materials).</li>
	<li>Using the same template, cut a 2&#34; circle in the center of the 4&#34; x 4&#34; extra-thin hydrocolloid dressing (Figure 2C,D).</li>
	<li>Cut the hydrocolloid layer of the 3&#34; x 3&#34; containment dressing at the 1&#189;&#34; line.</li>
	<li>Pull back the paper liner of the hydrocolloid layer of the containment dressing to expose the adhesive. Place the cut foam dressing within the 2&#34; circle of the adhesive (Figure 3A). This will ensure that the foam dressing extends beyond the opening of the containment dressing. Replace the paper liner onto the hydrocolloid layer of the dressing&#160;(Figure 3B). 
	&#8203;NOTE: These measurements were used for studies involving up to 10 larval I. scapularis ticks. For studies involving up to 30 larval I. scapularis ticks, the template for the foam center was cut between the 1&#34; and 1&#189;&#34; markings to allow for a larger area for placement. The measurements of the containment dressing can be adapted accordingly to the study design.</li>
</ol>
2. Tick placement
<ol>
	<li>Tick storage
	<ol>
		<li>Store the larval I. scapularis ticks in a medical-grade refrigerator with the temperature set to 9-10 &#176;C.</li>
		<li>Store the ticks in a vial covered with a mesh cap to allow adequate airflow. Package the vial of ticks in a sealable plastic bag with a lightly moistened sponge to maintain humidity and prevent desiccation.</li>
	</ol>
	</li>
	<li>Placing the ticks
	<ol>
		<li>Remove the ticks from the refrigerator to allow them to warm to room temperature. The ticks can become active in as little as 15 min outside the refrigerator.</li>
		<li>Position a white sheet around the placement area to easily visualize any inadvertently dropped tick.</li>
		<li>Dampen a 4&#34;x 4&#34; gauze pad with clean water or saline and use it to clean the skin of the placement site. The area should be patted dry or allowed to air dry. 
		NOTE: When selecting the body site for tick placement, consider areas that do not restrict the volunteer&#39;s daily activities and where the integrity of the dressing can be easily monitored by the participant. Do not place ticks in areas where they could be crushed due to direct pressure (e.g., the back). Evaluate the participant&#39;s body hair, and if necessary, carefully trim excessive body hair using scissors at the site of the adhesive, avoiding damage to the skin. Avoid using a razor for hair removal, as it may increase the risk of skin injury and infection.</li>
		<li>Once the skin is dry, remove the adhesive liner from the hydrocolloid side of the prepared dressing and firmly affix it to the skin at the placement site, ensuring that the mesh layer remains open.</li>
		<li>Break the wooden swab (see Table of Materials) in half to expose jagged ends.</li>
		<li>Carefully open the vial of ticks.</li>
		<li>Using the jagged end of the wooden swab, transfer active ticks from the vial and place them onto the skin within the containment dressing (Figure 4A). Have an additional team member monitor the tick placement site to ensure the ticks remain inside the containment dressing. Use adhesive tape to trap ticks that escape from the vial during this process. 
		NOTE: Breaking a wooden swab and using the jagged end is effective for tick manipulation and transfer without causing damage to the ticks. A small brush with soft bristles in a light color also works for transferring ticks from the container to the skin site.</li>
		<li>Once the specified number of ticks for the study is placed, remove the adhesive liner from the mesh layer of the containment dressing and close it over the opening.</li>
		<li>After ensuring that the adhesive is securely attached, apply the prepared 4&#34; x 4&#34; extra-thin hydrocolloid dressing over the containment dressing for additional security (Figure 4B). 
		&#8203;NOTE: Additional information regarding the pathogen-free I. scapularis tick colony and the duration of the tick exposure procedures used in the cited studies can be found in the methods sections of the published studies19,22.</li>
	</ol>
	</li>
</ol>
3. Site care
<ol>
	<li>Provide a water barrier cover and 2&#34; hypoallergenic tape (see Table of Materials) to the research participant.</li>
	<li>Provide instructions for caring for the tick containment site.
	<ol>
		<li>Periodically check the dressing and monitor the edges for secure adhesion to the skin. Apply additional tape to reinforce the dressing edges as needed.</li>
		<li>Do not scratch the dressing site, and refrain from opening or removing the dressing during the study period.</li>
		<li>Avoid bathing or soaking in water. Also, avoid hot showers and limit the duration of showering. Before showering, protect the containment dressing with the provided water barrier cover and remove it after completing the shower. Gently pat the dressing dry, and inspect it to ensure it remains secure.</li>
		<li>Avoid activities that may lead to heavy perspiration and compromise the dressing&#39;s integrity (e.g., aerobic exercise, extended hikes).</li>
	</ol>
	</li>
</ol>
4. Tick removal
<ol>
	<li>Collection vials preparation
	<ol>
		<li>Label the tick collection vials with the desired identifiers (e.g., study code, date).</li>
		<li>If preservation of live ticks is necessary, puncture holes in the cap of the vials with a 20 G needle. Use a layer of nylon or polyester mesh along with the cap to seal the vial. Place a small piece of moist sponge in a sealable bag with the vial of ticks to maintain humidity.</li>
	</ol>
	</li>
	<li>Removing the ticks
	<ol>
		<li>Place white sheets around the bandage site and ensure a collection basin is within reach.</li>
		<li>Using alcohol wipes, carefully remove the bandage from the skin.</li>
		<li>Once fully removed, place the bandage in the collection basin and examine it for the presence of detached ticks.</li>
		<li>Assess the placement site for attached ticks. Ticks that have fed to repletion will detach (Figure 5A). For the remaining attached ticks, use fine-tip forceps for removal (Figure 5B).</li>
		<li>Clean the skin with alcohol wipes or soap and water.</li>
	</ol>
	</li>
</ol>

Human Exposure to Larval Ticks for Tickborne Disease Research

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J., Spielman, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Duration+of+tick+attachment+and+Borrelia+burgdorferi+transmission.">Duration of tick attachment and Borrelia burgdorferi transmission.</a> J Clin Microbiol. 25 (3), 557-558 (1987).</li><li>Vora, 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=Ticks+elicit+variable+fibrinogenolytic+activities+upon+feeding+on+hosts+with+different+immune+backgrounds.">Ticks elicit variable fibrinogenolytic activities upon feeding on hosts with different immune backgrounds.</a> Sci Rep. 7, 44593 (2017).</li><li>Narasimhan, S., 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=Immunity+against+Ixodes+scapularis+salivary+proteins+expressed+within+24+hours+of+attachment+thwarts+tick+feeding+and+impairs+Borrelia+transmission.">Immunity against Ixodes scapularis salivary proteins expressed within 24 hours of attachment thwarts tick feeding and impairs Borrelia transmission.</a> PLoS One. 2 (5), 451 (2007).</li><li>Nazario, S., 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=Prevention+of+Borrelia+burgdorferi+transmission+in+guinea+pigs+by+tick+immunity.">Prevention of Borrelia burgdorferi transmission in guinea pigs by tick immunity.</a> Am J Trop Med Hyg. 58 (6), 780-785 (1998).</li><li>Krause, P. J., 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=Dermatologic+changes+induced+by+repeated+Ixodes+scapularis+bites+and+implications+for+prevention+of+tick-borne+infection.">Dermatologic changes induced by repeated Ixodes scapularis bites and implications for prevention of tick-borne infection.</a> Vector Borne Zoonotic Dis. 9 (6), 603-610 (2009).</li><li>Anderson, J. M., 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=Ticks,+Ixodes+scapularis,+feed+repeatedly+on+white-footed+mice+despite+strong+inflammatory+response:+an+expanding+paradigm+for+understanding+tick-host+interactions.">Ticks, Ixodes scapularis, feed repeatedly on white-footed mice despite strong inflammatory response: an expanding paradigm for understanding tick-host interactions.</a> Front Immunol. 8, 1784 (2017).</li><li>Rosenberg, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vital+signs:+trends+in+reported+vectorborne+disease+cases+-+United+States+and+Territories,+2004-2016.">Vital signs: trends in reported vectorborne disease cases - United States and Territories, 2004-2016.</a> MMWR Morb Mortal Wkly Rep. 67 (17), 496-501 (2018).</li><li>Paules, C. I., Marston, H. D., Bloom, M. 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Dr. Adriana Marques has a patent US 8,926,989 B2; and is an unpaid Scientific Advisor to the Global Lyme Alliance and to the American Lyme Disease Foundation. Siu Ping Turk and Aleah Eschman do not have an association that might pose a conflict of interest. The content of this publication does not necessarily reflect the views of or policies of the Department of Health and Human Services, nor does mention of trade names, commercial products, or organizations imply endorsement by the U.S. Government.

While animal studies involving exposures to ticks4,5,6,7,21 have been invaluable in increasing our understanding of host response to tickborne diseases and tick bites, these models have limitations in how well they predict the human host response. This model, which describes the methodology for exposing humans to tick bites in a controlled manner, can be easil...

Hard ticks (Ixodidae: Acari) are obligate blood-feeding ectoparasites that occur worldwide and are capable of transmitting a broad range of pathogens, including bacteria, viruses, and parasites, of major medical and veterinary importance. Ixodid ticks must remain attached to the host for days to complete a blood meal, and they have the capacity to stay attached to the skin while avoiding recognition, preventing local blood coagulation, and facilitating long-term feeding1,2,3. Animal studies have demonstrated that non-permissive hosts acquire resistance to tick bites with repeated tick exposures, which can lead to a decreased ability to transmit a pathogen, while ticks can repeatedly parasitize permissive hosts. Acquired tick resistance is dependent on the nature of the host immune response4,5,6,7.
Tickborne diseases are an increasing threat in the United States (US), with the number of reported cases more than doubling between 2004 and 20168,9. Due to climate change, the geographic ranges of different ticks continue to expand10,11. The leading tickborne diseases in the US include Lyme disease, anaplasmosis, ehrlichiosis, spotted fever rickettsiosis, babesiosis, tularemia, and Powassan virus disease8. Lyme disease, caused by infection with Borrelia burgdorferi sensu lato, is the most common tickborne disease in the US and Europe12. With approximately 476,000 individuals diagnosed with Lyme disease annually in the US, there is both a public health and economic burden to individuals and to society13,14,15.
Ixodes scapularis (the black-legged or deer tick) is the primary vector for Lyme disease, as well as anaplasmosis, babesiosis, Borrelia miyamotoi disease, and Powassan virus disease in the US. Other medically important tick species in the US include Amblyomma americanum (Lone star tick), Dermacentor variabilis (American dog tick), Ixodes pacificus (Western black-legged tick), Dermacentor andersoni (Rocky Mountain wood tick), Ixodes cookei (Groundhog tick), Dermacentor occidentalis (Pacific Coast tick), Rhipicephalus sanguineus (brown dog tick), and Amblyomma maculatum (Gulf Coast tick)16.
Developing a method to expose research volunteers to tick bites supports studies using the natural vector to search for evidence of infection, a procedure known as xenodiagnosis17,18,19,20,21, and to learn more about immunity induced by exposure to ticks, which can contribute to the discovery of an anti-tick vaccine5,6,7. The procedure described here was developed and used in the first human research study using laboratory-reared Ixodes scapularis larva for xenodiagnosis of B. burgdorferi infection after antibiotic therapy (NCT01143558), published in 201419. The system has been successfully used in a phase 2 study investigating whether a positive xenodiagnosis correlated with the persistence of symptoms after antibiotic treatment of Lyme disease (NCT02446626) and in a study exploring the host response to tick bites (NCT05036707).
This procedure protocol describes the process for creating the containment dressing, the tick placement procedure, and the tick removal procedure, as well as the site care needed to maintain the containment dressing. The details regarding the pathogen-free I. scapularis tick colony and tick exposure procedures used for the above-cited studies have been previously described19,22. This methodology offers a flexible research tool that can be adapted to studying different aspects of the human host response to tick bites, the effectiveness of tick prevention medications, as well as Lyme disease and other tickborne illnesses.

<table><tbody><tr><td>20 G needle</td><td>Any brand</td><td></td><td>For puncturing the vial cap.</td></tr><tr><td>3&quot; x 3&quot; containment dressing</td><td>Monarch Labs Names</td><td>LeFlap</td><td>https://www.monarchlabs.com/ordering</td></tr><tr><td>4&quot; x 4&quot; extra-thin hydrocolloid dressing</td><td>ConvaTec</td><td>DuoDerm</td><td>https://www.convatec.com/products/advanced-wound-care/brand-names/pc-wound-duoderm-granluflex/duoderm-extra-thin-dressing/</td></tr><tr><td>4&quot; x 4&quot; gauze</td><td>Monarch Labs Names</td><td></td><td>For cleaning skin</td></tr><tr><td>Clean water or saline</td><td></td><td></td><td>For cleaning skin</td></tr><tr><td>Moisture barrier (e.g. 7&quot; x 7&quot;)</td><td>AquaGuard</td><td>TIDI</td><td>For showering, ttps://www.tidiproducts.com/product-listing/aquaguard-shower-cover-sheets&amp;nbsp;</td></tr><tr><td>Non-adhesive foam dressing</td><td>Coloplast</td><td>Biatain</td><td>https://www.coloplast.us/biatain-non-adhesive-en-us.aspx</td></tr><tr><td>Roll of 2&quot; hypoallergenic tape</td><td>Monarch Labs Names</td><td>Durapore</td><td>For reinforcing containment dressing.</td></tr><tr><td>Roll of adhesive tape</td><td></td><td></td><td>For trapping ticks</td></tr><tr><td>Vials for collection (e.g. cryovials)</td><td>Ependorf</td><td>ECC200</td><td></td></tr></tbody></table>

a model for experimental exposure of humans to larval ixodes scapularis ticks

Tickborne diseases are a significant public health problem in the United States and worldwide. Ticks are obligate blood-feeding arthropods; an ixodid tick must remain attached to the skin of the host and complete its multi-day feeding process to acquire its blood meal. Exposing animals to ticks is a common practice for studying host responses to tick bites and tickborne diseases. We developed the procedure, conducted the first human research study, and published the findings on exposing human volunteers to uninfected larval Ixodes scapularis ticks. This article describes the methodology used to construct the containment dressing, how to apply and secure the ticks to the host, how to maintain the dressing, and how to remove the ticks from the host. Exposing volunteers to tick bites is an experimental procedure and must be performed under a clinical research protocol approved by the appropriate regulatory authorities. This method allows for translational research to better understand the human response to tick bites and foster the development of diagnostics, prevention, and therapies for tickborne diseases.

The study demonstrated that the procedure is safe and well-tolerated, with the primary adverse event being mild itching at the site of the bites, observed in 58% of the procedures. There were no serious adverse events related to the procedure when using clean laboratory-reared larval I. scapularis ticks19. In the 43 procedures conducted, the mean percentage of recovery of attached ticks compared to placed ticks was 45% &#177; 27% (SD), with a median percentage of 40% (<strong class="xfig"...

Watch this Scientific Journal Video about A Model for Experimental Exposure of Humans to Larval Ixodes scapularis Ticks at JoVE.com

Author Spotlight: Controlled Human Exposure Model for Tick Research and Lyme Disease Studies

This article presents the methodology for exposing humans to larval Ixodes scapularis for clinical research. The technique is relatively simple, tolerable by the research volunteers, and can be modified according to experimental needs. Such research involving human subjects must be conducted under clinical study protocols approved by the appropriate regulatory authorities.

A Model for Experimental Exposure of Humans to Larval Ixodes scapularis Ticks

Tickborne diseases are a significant public health problem in the United States and worldwide. Ticks are obligate blood-feeding arthropods; an ixodid tick ...

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Immunology and Infection

446 Views.  National Institutes of Health. This article presents the methodology for exposing humans to larval Ixodes scapularis for clinical research. The technique is relatively simple, tolerable by the research volunteers, and can be modified according to experimental needs. Such research involving human subjects must be conducted under clinical study protocols approved by the appropriate regulatory authorities.

Video: Author Spotlight: Controlled Human Exposure Model for Tick Research and Lyme Disease Studies

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 ...

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.

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 ...

Rearing Ixodes scapularis, the Black-legged Tick: Feeding Immature Stages on Mice

Ixodes scapularis, the vector of Lyme disease, is one of the most important disease vectors in the eastern and Midwestern United States. This species is a three host tick that requires a blood meal from a vertebrate host for each development stage, and the adult females require a blood meal for reproduction. Larval ticks attach to their host for 3 - 5 days for feeding and drop off the host when fully engorged. This dependency on several different hosts and the lengthy attachment time for engorgement complicates tick rearing in the laboratory setting. However, to understand tick biology and tick-pathogen interactions, the production of healthy, laboratory-reared ticks is essential. Here, we demonstrate a simple, cost-effective protocol for immature tick feeding on mice. We modified the existing protocols for decreased stress on mice and increased tick feeding success and survival by using disposable cages without mesh bottoms to avoid contact of ticks with water contaminated with mice urine and feces.

Rearing Ixodes scapularis, the Black-legged Tick: Feeding Immature Stages on Mice

The production of healthy laboratory-reared ticks is essential to studies on tick biology, and tick-pathogen interactions. Here we demonstrate a simple protocol for immature tick feeding that is cost-effective and less stressful to mice.

Ixodes scapularis, the vector of Lyme disease, is one of the most important disease vectors in the eastern and Midwestern United States. This species is a ...

Disclosing Hemolymph Collection and Inoculation of Metarhizium Blastospores into Rhipicephalus Microplus Ticks Towards Invertebrate Pathology Studies

Ticks are obligate hematophagous ectoparasites and Rhipicephalus microplus has great importance in veterinary medicine because it causes anemia, weight loss, depreciation of the animals&#39; leather and also can act as a vector of several pathogens. Due to the exorbitant costs to control these parasites, damage to the environment caused by the inappropriate use of chemical acaricides, and the increased resistance against traditional parasiticides, alternative control of ticks, by the use of entomopathogenic fungi, for example, has been considered an interesting approach. Nevertheless, few studies have demonstrated how the tick&#39;s immune system acts to fight these entomopathogens. Therefore, this protocol demonstrates two methods used for entomopathogen inoculation into engorged females and two techniques used for hemolymph collection and hemocytes harvesting. Inoculation of pathogens at the leg insertion in the tick female&#39;s body allows evaluation of females biologic parameters unlike the inoculation between the scutum and capitulum, which frequently damages Gen&#233;&#39;s organ. Dorsal hemolymph collection yielded a higher volume recovery than collection through the legs. Some limitations of tick hemolymph collection and processing include i) high rates of hemocytes&#39; disruption, ii) hemolymph contamination with disrupted midgut, and iii) low hemolymph volume recovery. When hemolymph is collected through the leg cutting, the hemolymph takes time to accumulate at the leg opening, favoring the clotting process. In addition, fewer hemocytes are obtained in the collection through the leg compared to the dorsal collection, even though the first method is considered easier to be performed. Understanding the immune response in ticks mediated by entomopathogenic agents helps to unveil their pathogenesis and develop new targets for tick control. The inoculation processes described here require very low technological resources and can be used not only to expose ticks to pathogenic microorganisms. Similarly, the collection of tick hemolymph may represent the first step for many physiological studies.

Disclosing Hemolymph Collection and Inoculation of Metarhizium Blastospores into Rhipicephalus Microplus Ticks Towards Invertebrate Pathology Studies

Analysis of tick hemolymph represents an important source of information on how some pathogens cause disease and how ticks immunologically respond to this infection. The present study demonstrates how to inoculate fungal propagules and collect hemolymph from Rhipicephalus microplus engorged females.

Ticks are obligate hematophagous ectoparasites and Rhipicephalus microplus has great importance in veterinary medicine because it causes anemia, weight ...

A Capsule-Based Model for Immature Hard Tick Stages Infestation on Laboratory Mice

Ticks are obligatory blood feeding parasites at all stages of development (except eggs) and are recognized as vectors of various pathogens. The use of mouse models in tick research is critical for understanding their biology and tick-host-pathogen interactions. Here we demonstrate a non-laborious technique for the feeding of immature stages of hard ticks on laboratory mice. The benefit of the method is its simplicity, short duration, and the ability to monitor or collect ticks at different time points of an experiment. In addition, the technique allows attachment of two individual capsules on the same mouse, which is beneficial for a variety of experiments where two different groups of ticks are required to feed on the same animal. The non-irritating and flexible capsule is made from easily accessible materials and minimizes the discomfort of the experimental animals. Furthermore, euthanasia is not necessary, mice recover completely after the experiment and are available for re-use.

In this study, a feeding system for nymphal and larval stages of hard tick was developed using a capsule attached to laboratory mouse. The feeding capsule is made from flexible materials and remains firmly attached to the mouse for at least one week and allows comfortable monitoring of tick feeding.

Ticks are obligatory blood feeding parasites at all stages of development (except eggs) and are recognized as vectors of various pathogens. The use of ...

Biology

Isolation of microRNAs from Tick Ex Vivo Salivary Gland Cultures and Extracellular Vesicles

Ticks are important ectoparasites that can vector multiple pathogens. The salivary glands of ticks are essential for feeding as their saliva contains many effectors with pharmaceutical properties that can diminish host immune responses and enhance pathogen transmission. One group of such effectors are microRNAs (miRNAs). miRNAs are short non-coding sequences that regulate host gene expression at the tick-host interface and within the organs of the tick. These small RNAs are transported in the tick saliva via extracellular vesicles (EVs), which serve inter-and intracellular communication. Vesicles containing miRNAs have been identified in the saliva of ticks. However, little is known about the roles and profiles of the miRNAs in tick salivary vesicles and glands. Furthermore, the study of vesicles and miRNAs in tick saliva requires tedious procedures to collect tick saliva. This protocol aims to develop and validate a method for isolating miRNAs from purified extracellular vesicles produced by ex vivo organ cultures. The materials and methodology needed to extract miRNAs from extracellular vesicles and tick salivary glands are described herein.

Isolation of microRNAs from Tick Ex Vivo Salivary Gland Cultures and Extracellular Vesicles

The present protocol describes the isolation of microRNAs from tick salivary glands and purified extracellular vesicles. This is a universal procedure that combines commonly used reagents and supplies. The method also allows the use of a small number of ticks, resulting in quality microRNAs that can be readily sequenced.

Ticks are important ectoparasites that can vector multiple pathogens. The salivary glands of ticks are essential for feeding as their saliva contains many ...

Embryo Injection Technique for Gene Editing in the Black-Legged Tick, Ixodes scapularis

Ticks can transmit various viral, bacterial, and protozoan pathogens and are therefore considered vectors of medical and veterinary importance. Despite the growing burden of tick-borne diseases, research on ticks has lagged behind insect disease vectors due to challenges in applying genetic transformation tools for functional studies to the unique biology of ticks. Genetic interventions have been gaining attention to reduce mosquito-borne diseases. However, the development of such interventions requires stable germline transformation by injecting embryos. Such an embryo injection technique is lacking for chelicerates, including ticks. Several factors, such as an external thick wax layer on tick embryos, hard chorion, and high intra-oval pressure, are some&#160;obstacles that previously prevented embryo injection protocol development in ticks. The present work has overcome these obstacles, and an embryo injection technique for the black-legged tick, Ixodes scapularis, is described here.&#160;This technique can be used to deliver components, such as CRISPR/Cas9, for stable germline transformations.

Embryo Injection Technique for Gene Editing in the Black-Legged Tick, Ixodes scapularis

The present protocol describes a method for injecting tick embryos. Embryo injection is the preferred technique for genetic manipulation to generate transgenic lines.

Ticks can transmit various viral, bacterial, and protozoan pathogens and are therefore considered vectors of medical and veterinary importance. Despite ...

Tick Artificial Membrane Feeding for Ixodes scapularis

Ticks and their associated diseases are an important topic of study due to their public health and veterinary burden. However, the feeding requirements of ticks during both study and rearing can limit experimental questions or the ability of labs to research ticks and their associated pathogens. An artificial membrane feeding system can reduce these problems and open up new avenues of research that may not have been possible with traditional animal feeding systems. This study describes an artificial membrane feeding system that has been refined for feeding and engorgement success for all Ixodes scapularis life stages. Moreover, the artificial membrane feeding system described in this study can be modified for use with other tick species through simple refinement of the desired membrane thickness. The benefits of an artificial membrane feeding system are counterbalanced by the labor intensiveness of the system, the additional environmental factors that may impact feeding success, and the need to refine the technique for each new species and life stage of ticks.

Tick Artificial Membrane Feeding for Ixodes scapularis

Presented here is a method to blood feed ticks in vitro via an artificial membrane system to allow for partial or full engorgement of a variety of tick life stages.

Ticks and their associated diseases are an important topic of study due to their public health and veterinary burden. However, the feeding requirements of ...

A Model for Experimental Exposure of Humans to Larval Ixodes scapularis Ticks

Summary

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Chapters in this video

RNA Interference in Ticks

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

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

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

Rearing Ixodes scapularis, the Black-legged Tick: Feeding Immature Stages on Mice

Disclosing Hemolymph Collection and Inoculation of Metarhizium Blastospores into Rhipicephalus Microplus Ticks Towards Invertebrate Pathology Studies

A Capsule-Based Model for Immature Hard Tick Stages Infestation on Laboratory Mice

Isolation of microRNAs from Tick Ex Vivo Salivary Gland Cultures and Extracellular Vesicles

Embryo Injection Technique for Gene Editing in the Black-Legged Tick, Ixodes scapularis

Tick Artificial Membrane Feeding for Ixodes scapularis