Name: tick artificial membrane feeding for ixodes scapularis
Uploaded: 2022-11-30T14:00:00.000+00:00
Duration: 8 min 53 s
Description: Watch this Scientific Journal Video about Tick Artificial Membrane Feeding for Ixodes scapularis at JoVE.com

1. Preparing the tick membrane chamber
<ol>
	<li>Prepare a flat, non-porous surface such as a plane of glass or ceramic-coated metal base of an arm stand by wiping it down with 70% ethanol and then cover it with a single layer of plastic wrap, making sure that the plastic wrap is flat and without bubbles or wrinkles (see Figure 1A).</li>
	<li>Tape down 100% rayon lens cleaning paper to the prepared surface. Ensure it is flat and slightly taut with tape across all four sides of the paper. Two pieces of 4 in x 6 in lens paper can be made into membranes using the volumes described in step 1.3. 
	NOTE: This is sufficient to make up to 12 feeding chambers (see Figure 1B). If the membrane is for larval ticks, use unryu paper instead of lens cleaning paper.</li>
	<li>Prepare the 00-10 hardness silicone mixture by measuring out 5 mL of each part of the silicone kit into a disposable container, lightly mixing the two liquids before adding 1.5 mL of hexane. Continue mixing until the mixture is homogeneous and thoroughly mixed. 
	NOTE: If making membranes for adult ticks, use 00-50 hardness silicone.</li>
	<li>Use a small squeegee to distribute the silicone mix over the lens paper. Allow to stand for 1 min to ensure that the silicone has soaked into the lens paper. Steadily scrape excess silicone to the side with the squeegee using a small amount of downward pressure. Perform a second pass with the squeegee, exerting no downward pressure, to remove any lines of silicone and produce a smooth layer. 
	NOTE: This step of the procedure may require some practice to produce membranes of optimal thickness for each life stage (adults: 150-200 &#181;m; nymphs: 90-120 &#181;m; larvae: 80-100 &#181;m). A thicker membrane (within the feeding tolerances of the life stage) will usually produce better results by reducing condensation in the chamber and improving the self-healing characteristic of the membrane.</li>
	<li>For larvae only, dip the top of the feeding chamber into Fluon fluoropolymer resin (PTFE-30) several times, allowing it to dry between applications to produce a consistent layer. 
	NOTE: The application of Fluon will form a non-stick layer of fluoropolymer resin similar to a non-stick cooking pan. This will reduce the number of larvae that climb to the top of the chamber21.</li>
	<li>Leave the membrane to cure for at least 24 h in a dust-free environment, such as a closed chemical or biosafety cabinet, before moving on to the next step. See Figure 1C for how the rayon lens paper looks after the silicone has been allowed to dry.</li>
	<li>Prepare the 30-hardness silicone mixture to attach the chambers to the membrane by mixing equal parts of silicone mix A and B.</li>
	<li>Dip the polycarbonate chambers about a quarter of an inch (6 mm) into the silicone mixture, and then place on the membrane sheet, taking care not to overlap any of the tape (Figure 1D).</li>
	<li>Allow the attaching silicone to cure for at least 24 h before moving to the next steps.</li>
	<li>Using a scalpel, carefully cut the membrane around each of the attached chambers, trimming down the edges so that it can fit smoothly into a well of the 6-well plate without much scraping on the sides (Figure 1E). Allow sufficient material outside the chamber to maximize adhesion of the membrane. 
	NOTE: If too much exterior material remains, repeated removal of the chamber from the 6-well plate can cause the membrane to partially detach from the chamber. Repairing detached membranes is described in section 5.</li>
	<li>Cut a small piece of the leftover membrane from each of the corners and the center of the membrane sheet. Remove the plastic wrap from each piece. Measure the pieces with a micrometer to determine the average membrane thickness. 
	NOTE: If the membrane thickness does not fall within the tolerance range for life stage (see step 1.4), the membranes should be discarded and remade. Unryu paper thickness is highly variable due to its thick strands of fiber and thin regions. While this characteristic allows larvae to find regions of optimal thickness, it makes it difficult to determine actual membrane thickness.</li>
	<li>Place the feeding chambers along with one O-ring per chamber into a glass beaker to be autoclaved at 121 &#176;C for at least 20 min to sterilize. Allow the chambers to cool before using them.</li>
</ol>
2. Setting up the tick feed
<ol>
	<li>Prepare a lamp or room so that the lights are on for 16 h and then off for 8 h. If using a lamp, make sure that the water bath in the next step is in the dark for those 8 h.</li>
	<li>Set up a water bath at 34 &#176;C and with supports so that a 6-well plate can sit and float slightly in it. Use small glass beakers sunken in the water bath as supports. Use a transparent cover for the water bath to maintain a light-dark cycle for the ticks; if need be, use a T-stand and plastic wrap to make a cover. To reduce the risk of contamination, add 0.02% benzalkonium chloride to the water bath and set another water bath to 36 &#176;C for warming chilled blood.</li>
	<li>Take out the preprepared, autoclaved membrane chambers, placing the O-ring around the chamber (Figure 1F), and then fill each chamber with enough 70% ethanol to cover the membrane. Let sit in a 6-well plate well for 5 min and then look for any leaks between the chamber and membrane and in the membrane itself. 
	NOTE: If the membrane is leaking, ethanol will accumulate in the base of the well. Leaky membranes should be discarded.</li>
	<li>Empty out the ethanol from the chambers and then let it air dry inside a biosafety cabinet or laminar flow hood. 
	NOTE: This may take several minutes depending on ambient humidity.</li>
	<li>While waiting, heat-inactivate the complement in the blood by heating at 56 &#176;C for 40 min. Supplement mechanically defibrinated bovine blood with 2 g/L glucose.</li>
	<li>After the membrane is dry, apply ~20 &#181;L of phagostimulant to the inside of the chamber and then spread it around the surface by tilting the chamber. See section 6 for instructions regarding preparing a phagostimulant from tick frass.</li>
	<li>Wait until the phagostimulant is dry before adding the ticks to the chamber with either a brush or forceps. Work as quickly as possible when transferring the ticks to the chamber to prevent escape. Put the ticks in a tube on ice for 1-2 min before transferring them to slow down their ability to escape. After the ticks have been transferred, seal the top with parafilm; do not overstretch the parafilm, as the heat from the water bath can cause it to rip. 
	NOTE: Forceps work best with nymphal and adult life stages and brushes work best with larval life stages.</li>
	<li>Add 5 mL of the heat-inactivated blood per well/chamber into a conical tube and place in the water bath set at 36 &#176;C. Bring chilled blood up to at least room temperature before exposing the ticks to it. Set aside four chambers per 6-well plate for easy handling of chambers. More chambers per plate should be avoided, however, depending on the size of the water bath, more plates can be used.</li>
	<li>Add 5 &#181;L of 3 mM ATP and 50 &#181;L of 100x stock of penicillin/streptomycin/fungizone per 5 mL of blood to the tube.</li>
	<li>Transfer 4.5 mL of the blood into a well of a 6-well plate, and then gently place the chamber into the well, adjusting the O-ring height on the chamber so that the membrane sits in the blood but the blood level around the side does not overtop the well. Place the well into the blood at an angle to avoid air bubble formation between the blood and the membrane. 
	NOTE: A completed plate is shown in Figure 2.</li>
	<li>Place the lid of the 6-well plate on top of the feeding chambers; then, put the 6-well plate with chambers into the prepared 34 &#176;C water bath and close the lid. 
	&#8203;NOTE: The lid on top will help stop condensed water from contacting the parafilm and diluting the blood in the wells.</li>
</ol>
3. Maintaining the feeding ticks by changing the blood every 12 h
<ol>
	<li>Aliquot 5 mL of the heat-inactivated blood per well into a tube and warm it up in a 36 &#176;C water bath. Thaw the ATP and penicillin/streptomycin/fungizone in the water bath at the same time.</li>
	<li>Add 5 &#181;L of 3 mM ATP and 50 &#181;L of 100x stock of penicillin/streptomycin/fungizone per well to the blood tube. Transfer 4.5 mL of blood into a well of a fresh 6-well plate.</li>
	<li>Take out the 6-well plate with the tick chamber from the water bath. Remove the chamber from the well and rinse the outside of the chamber and membrane with 10 mL of sterile 1x PBS to remove the blood.</li>
	<li>Using autoclaved filter paper, gently dab dry the membrane and chamber to remove the excess PBS. Replace the parafilm on the top of the chamber. If there is a lot of condensation inside the chamber, dab dry the wet spots with autoclaved filter paper before sealing with fresh parafilm.</li>
	<li>Place the tick chamber into the new 6-well plate with fresh blood, adjusting the O-ring height if needed. Repeat steps 3.3-3.5 for each chamber on the plate. Place the lid of the 6-well plate over the tops of the chambers. Move the new 6-well plate back into the 34 &#176;C water bath.</li>
	<li>Repeat steps 3.1-3.5 every 12 h until the conclusion of the tick feed. Wait for the ticks to detach from the membrane by themselves when engorged or gently remove them from the membrane with soft-touch tweezers while still attached to obtain partially engorged ticks.</li>
</ol>
4. Antifungal treatment
NOTE: Perform only when fungal growth is seen on the membrane. Fungus is likely to form on the blood side of the membrane if the feeding is of sufficient duration. The first indication of fungal contamination is small (1-3 mm) flakes of coagulated blood visible on the membrane. When fungal contamination is noted, antifungal treatments can prolong the duration of the experiment and improve engorgement success.
<ol>
	<li>Dissolve 10,000 U nystatin per mL in distilled water, 5 mL per feeding chamber. Filter-sterilize with a 0.1 &#181;m syringe filter.</li>
	<li>Add 5 mL of nystatin solution per well of a new 6-well plate, one well for each feeding chamber.</li>
	<li>Place PBS-rinsed chambers into the solution. Let sit for 10 min.</li>
	<li>Rinse the treated membranes with PBS before putting back into fresh blood.</li>
	<li>Repeat the antifungal treatment every 2-3 days (depending on the extent of fungal contamination) until the completion of the experiment.</li>
</ol>
5. Re-adhering detached membranes with cyanoacrylate glue
NOTE: Perform only when membrane detachment is noticed.
<ol>
	<li>Membranes may partially detach from the feeding chambers, especially during removal from the 6-well plate. If this happens, rinse the membrane of blood with PBS.</li>
	<li>Gently dry the affected area. It does not need to be perfectly dry, but the presence of moisture causes the glue to set faster. 
	NOTE: Re-adhering the membrane limits the working time, however, depending on detachment size, a faster setting time may be desirable.</li>
	<li>Squeeze a small amount of cyanoacrylate glue into the gap where the membrane has pulled away from the chamber. Hold in place for ~2 min to allow the glue to set.</li>
	<li>Place the chamber back into the blood in the 6-well plate.</li>
</ol>
6. Making phagostimulant
NOTE: Perform at the end of a feed or before a membrane feed has started.
<ol>
	<li>Collect tick feces from either a prior membrane feed or another source of feeding ticks. Store the feces at -20 &#176;C prior to making the phagostimulant. Crush the pellets of tick feces and mix 0.1 g per 1 mL of water. Add 1 mM tripeptide reduced glutathione, dissolve, and mix thoroughly by vortexing.</li>
	<li>Filter-sterilize with a 0.1 &#181;m syringe filter.</li>
	<li>Freeze at -20 &#176;C until needed.</li>
</ol>

<ol>
	<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> Morbidity and Mortality Weekly Report. 67 (17), 496-501 (2018).</li><li>Busch, J. D., 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=Widespread+movement+of+invasive+cattle+fever+ticks+(Rhipicephalus+microplus)+in+southern+Texas+leads+to+shared+local+infestations+on+cattle+and+deer.">Widespread movement of invasive cattle fever ticks (Rhipicephalus microplus) in southern Texas leads to shared local infestations on cattle and deer.</a> Parasites &amp; Vectors. 7, 188 (2014).</li><li>S&#252;ss, J., Klaus, C., Gerstengarbe, F. W., Werner, P. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=What+makes+ticks+tick?+Climate+change,+ticks,+and+tick-borne+diseases.">What makes ticks tick? Climate change, ticks, and tick-borne diseases.</a> Journal of Travel Medicine. 15 (1), 39-45 (2008).</li><li>Gray, J. S., Dautel, H., Estrada-Pe&#241;a, A., Kahl, O., Lindgren, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+climate+change+on+ticks+and+tick-borne+diseases+in+Europe.">Effects of climate change on ticks and tick-borne diseases in Europe.</a> Interdisciplinary Perspectives on Infectious Diseases. 2009, 593232 (2009).</li><li>Sonenshine, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Biology of Ticks. , (1991).</li><li>Schwan, T. G., Piesman, J., Golde, W. T., Dolan, M. C., Rosa, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Induction+of+an+outer+surface+protein+on+Borrelia+burgdorferi+during+tick+feeding.">Induction of an outer surface protein on Borrelia burgdorferi during tick feeding.</a> Proceedings of the National Academy of Sciences. 92 (7), 2909-2913 (1995).</li><li>Massung, R. F., Priestley, R. A., Miller, N. J., Mather, T. N., Levin, M. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inability+of+a+variant+strain+of+Anaplasma+phagocytophilum+to+infect+mice.">Inability of a variant strain of Anaplasma phagocytophilum to infect mice.</a> The Journal of Infectious Diseases. 188 (11), 1757-1763 (2003).</li><li>Koci, J., Bernard, Q., Yang, X., Pal, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Borrelia+burgdorferi+surface+protein+Lmp1+facilitates+pathogen+dissemination+through+ticks+as+studied+by+an+artificial+membrane+feeding+system.">Borrelia burgdorferi surface protein Lmp1 facilitates pathogen dissemination through ticks as studied by an artificial membrane feeding system.</a> Scientific Reports. 8 (1), 1910 (2018).</li><li>Levin, M. L., Schumacher, L. B. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Manual+for+maintenance+of+multi-host+ixodid+ticks+in+the+laboratory.">Manual for maintenance of multi-host ixodid ticks in the laboratory.</a> Experimental and Applied Acarology. 70 (3), 343-367 (2016).</li><li>Hart, T., Yang, X., Pal, U., Lin, Y. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+Lyme+borreliae+proteins+promoting+vertebrate+host+blood-specific+spirochete+survival+in+Ixodes+scapularis+nymphs+using+artificial+feeding+chambers.">Identification of Lyme borreliae proteins promoting vertebrate host blood-specific spirochete survival in Ixodes scapularis nymphs using artificial feeding chambers.</a> Ticks and Tick-Borne Diseases. 9 (5), 1057-1063 (2018).</li><li>Hart, T. 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=Host+tropism+determination+by+convergent+evolution+of+immunological+evasion+in+the+Lyme+disease+system.">Host tropism determination by convergent evolution of immunological evasion in the Lyme disease system.</a> PLoS Pathogens. 17 (7), (2021).</li><li>Bernard, Q., 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=Plasticity+in+early+immune+evasion+strategies+of+a+bacterial+pathogen.">Plasticity in early immune evasion strategies of a bacterial pathogen.</a> Proceedings of the National Academy of Sciences. 115 (16), 3788-3797 (2018).</li><li>Pierce, A. E., Pierce, M. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+note+on+the+cultivation+of+Boophilus+microplus+(Canestrini,+1887)+(Ixodidae:+Acarina)+on+the+embyonated+hen+egg.">A note on the cultivation of Boophilus microplus (Canestrini, 1887) (Ixodidae: Acarina) on the embyonated hen egg.</a> Australian Veterinary Journal. 32 (6), 144-146 (1956).</li><li>Doube, B. M., Kemp, D. 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+influence+of+temperature,+relative+humidity+and+host+factors+on+the+attachment+and+survival+of+Boophilus+microplus+(Canestrini)+larvae+to+skin+slices.">The influence of temperature, relative humidity and host factors on the attachment and survival of Boophilus microplus (Canestrini) larvae to skin slices.</a> International Journal for Parasitology. 9 (5), 449-454 (1979).</li><li>Kr&#246;ber, T., Guerin, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+in+vitro+feeding+assay+to+test+acaricides+for+control+of+hard+ticks.">An in vitro feeding assay to test acaricides for control of hard ticks.</a> Pest Management Science. 63 (1), 17-22 (2007).</li><li>Kuhnert, F., Diehl, P. A., Guerin, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+life-cycle+of+the+bont+tick+Amblyomma+hebraeum+in+vitro.">The life-cycle of the bont tick Amblyomma hebraeum in vitro.</a> International Journal for Parasitology. 25 (8), 887-896 (1995).</li><li>Kr&#246;ber, T., Guerin, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+feeding+assays+for+hard+ticks.">In vitro feeding assays for hard ticks.</a> Trends in Parasitology. 23 (9), 445-449 (2007).</li><li>Andrade, J. J., Xu, G., Rich, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+silicone+membrane+for+in+vitro+feeding+of+Ixodes+scapularis+(Ixodida:+Ixodidae).">A silicone membrane for in vitro feeding of Ixodes scapularis (Ixodida: Ixodidae).</a> Journal of Medical Entomology. 51 (4), 878-879 (2014).</li><li>Oliver, J. D., 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=Infection+of+immature+Ixodes+scapularis+(Acari:+Ixodidae)+by+membrane+feeding.">Infection of immature Ixodes scapularis (Acari: Ixodidae) by membrane feeding.</a> Journal of Medical Entomology. 53 (2), 409-415 (2016).</li><li>Oliver, J. D., 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=Growth+dynamics+and+antibiotic+elimination+of+symbiotic+Rickettsia+buchneri+in+the+tick+Ixodes+scapularis+(Acari:+Ixodidae).">Growth dynamics and antibiotic elimination of symbiotic Rickettsia buchneri in the tick Ixodes scapularis (Acari: Ixodidae).</a> Applied and Environmental Microbiology. 87 (3), (2021).</li><li>Graham, E. E., Poland, T. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficacy+of+Fluon+conditioning+for+capturing+cerambycid+beetles+in+different+trap+designs+and+persistence+on+panel+traps+over+time.">Efficacy of Fluon conditioning for capturing cerambycid beetles in different trap designs and persistence on panel traps over time.</a> Journal of Economic Entomology. 105 (2), 395-401 (2012).</li><li>Munderloh, U. G., Liu, Y., Wang, M., Chen, C., Kurtti, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Establishment,+maintenance+and+description+of+cell+lines+from+the+tick+Ixodes+scapularis.">Establishment, maintenance and description of cell lines from the tick Ixodes scapularis.</a> Journal of Parasitology. 80 (4), 533-543 (1994).</li><li>Lehane, 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=Prevalence+of+single+and+coinfections+of+human+pathogens+in+Ixodes+ticks+from+five+geographical+regions+in+the+United+States,+2013-2019.">Prevalence of single and coinfections of human pathogens in Ixodes ticks from five geographical regions in the United States, 2013-2019.</a> Ticks and Tick-Borne Diseases. 12 (2), (2021).</li><li>Gonz&#225;lez, J., Bickerton, M., Toledo, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Applications+of+artificial+membrane+feeding+for+ixodid+ticks.">Applications of artificial membrane feeding for ixodid ticks.</a> Acta Tropica. 215, (2021).</li><li>Kr&#243;l, N., 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=Evaluating+transmission+paths+for+three+different+Bartonella+spp.+in+Ixodes+ricinus+ticks+using+artificial+feeding.">Evaluating transmission paths for three different Bartonella spp. in Ixodes ricinus ticks using artificial feeding.</a> Microorganisms. 9 (5), 901 (2021).</li><li>Anderson, J. F., Magnarelli, L. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biology+of+ticks.">Biology of ticks.</a> Infectious Disease Clinics of North America. 22 (2), 195-215 (2008).</li></ol>

Artificial membrane feeding of ticks provides a useful tool for a variety of experimental procedures, but is not likely to replace animal feeding for all applications. Maintaining large colonies of ticks at all life stages without animal feeding is generally untenable. Instead, the artificial feeding system is valuable for other purposes such as infecting ticks with pathogens not supported by model hosts, evaluating the impacts of controlled dosages of compounds or microorganisms on the ticks in a simplified feeding envi...

Tick-borne diseases strongly impact the health of humans and animals across the world, being responsible for more than two-thirds of all vector-associated illness in the USA from 2004 to 20161. Additionally, case numbers have been growing in recent years, with more people and livestock being affected by ticks and their associated diseases2,3. While there are likely numerous causes for the upward trend in case numbers, the changing climate is an important factor3,4. The predicted ongoing increase in the number of tick-borne disease cases underlines the need to develop new tools to investigate the relationships between ticks and the pathogens they transmit.
It is known that ticks undergo changes in physiology and gene expression during feeding and that these changes play a role in pathogen transmission5,6. It may be difficult to perform studies that examine the effects of full and partial feeding on pathogen transmission and acquisition using animal models, particularly in situations where rodent models are not susceptible to infection by a particular pathogen. For example, Anaplasma phagocytophilum Variant-1 strain is naturally transmitted between Ixodes scapularis and deer but is unable to infect mice, complicating tick infection in the lab7. Artificial feeding systems can also be applied to help study pathogens such as Borrelia burgdorferi via the use of transgenic mutants that have gene deletions that inhibit transmission or infection8. Using an artificial feeding system helps researchers isolate the genes&#39; role by allowing infection or transmission to only occur on the tick&#39;s side, thereby isolating any host response that may confound such studies.
Similarly, some life stages of ticks involved in disease and animal transmission may not be induced to feed on common laboratory model species. Ixodes scapularis females, for example, must be fed on larger animals, typically rabbits9. While often accessible for laboratory experimentation, the administrative and husbandry requirements of using rabbits exceeds those of small rodents and may be prohibitive for some laboratories. Other tick species, particularly those of veterinary concern, must be fed on cattle or other large animals that are not practical to use in most laboratories. In vitro feeding and infection methods, such as artificial membrane feeding, provide alternatives to using large or exotic host animals.
Additionally, the use of an artificial feeding system allows certain analyses that may not be possible with traditional animal feeding methods. One such example is that, by separating the blood source from the feeding mechanism, examination of the role that different hosts&#39; blood may have in B. burgdorferi transmission becomes possible10. This examination of host blood and the role blood itself plays in the absence of the host immune response is an important factor in being able to understand pathogen transmission cycles and one that artificial feeding systems are able to help answer11. It also becomes possible to quantify the exact transmission numbers of a pathogen during a feed rather than just examining transmission success and establishment in a host8,12.
Some of the first artificial feeding membranes made for hard ticks were made out of animal skins or animal-derived membranes in the 1950s and 1960s13,14. Due to the biological nature of these membranes, there were problems with both the production of new membranes and shelf life. In the 1990s, fully artificial membranes were developed that utilized a backing of netting, paper, or fabric with silicone impregnation15,16. Silicone was ideal as its physical properties mimic skin&#39;s stretchiness and slight tackiness, along with its bio-inherent nature. Building on this, Krober and Guerin, whose work this technique was based on, described a silicone-impregnated rayon membrane feeding technique for the artificial feeding of I. ricinus17.
Refinement of the methods for I. scapularis, a closely related species, has led to notable differences in the hardness of silicone used in membrane impregnation, the recipe for membrane production, dimensions of the chamber, and the attachment stimulant. While the refinements reported in this study have resulted in similar membrane characteristics as those reported by Andrade et al., who also developed a silicone-based membrane based on Krober and Guerin for use in I. scapularis, there is a difference in the silicone impregnation steps, which allows the flexibility to utilize this protocol for immature life stages of I. scapularis15,18. This study also describes additions and technical alterations based on repeated use of this method, best practices that result in a successful feed, and troubleshooting of problems that may arise. This method has been used to feed all active life stages, infect ticks with pathogenic bacteria, and expose ticks to multiple dosages of antibiotics19,20. While the artificial membrane feeding method shown is for I. scapularis, this method is readily adaptable to other species of ticks with minor modifications in membrane thickness.

<table><tbody><tr><td>00-10 Hardness Silicone</td><td>Smooth-On</td><td>Ecoflex 00-10</td><td>Trial size from Smooth-On Store</td></tr><tr><td>00-50 Hardness Silicone</td><td>Smooth-On</td><td>Ecoflex 00-50</td><td>Trial size from Smooth-On Store</td></tr><tr><td>30 Hardness Silicone</td><td>Smooth-On</td><td>Mold Star 30</td><td>Trial size from Smooth-On Store</td></tr><tr><td>6-well cell culture plates</td><td>Corning Incorporated</td><td>3516</td><td></td></tr><tr><td>Adenosine triphosphate (ATP)</td><td>Millipore Sigma</td><td>A1852-1VL</td><td>Used to make an aqueous solution of 3 mM ATP that has been filter sterlized via 0.2 micometer filter</td></tr><tr><td>Bovine blood</td><td>HemoStat</td><td>DBB500</td><td>Mechanically defibrinated; 500 mL is usually sufficient for one experiment</td></tr><tr><td>Clingwrap</td><td>Fisherbrand</td><td>22-305654&amp;nbsp;</td><td></td></tr><tr><td>Filter Paper</td><td>Fisherbrand</td><td>09-790-2C</td><td>Autoclave and let cool before using. Can use Fine quality instead of medium too</td></tr><tr><td>Fluon (aqueous polytetrafluoroethylene)</td><td>Bioquip</td><td>2871</td><td>Available from other sources such as https://canada-ant-colony.com/products/fluon-ptfe-10ml</td></tr><tr><td>Glucose</td><td>Millipore Sigma</td><td>G8270-100G</td><td></td></tr><tr><td>Hexane</td><td>Millipore Sigma</td><td>139386-100ML</td><td></td></tr><tr><td>Lens paper</td><td>Fisherbrand</td><td>11-995</td><td>100% rayon</td></tr><tr><td>Nystatin &amp;nbsp;</td><td>Gold Biotechnology</td><td>N-750-10</td><td></td></tr><tr><td>Parafilm</td><td>Fisherbrand</td><td>S37440&amp;nbsp;</td><td></td></tr><tr><td>Penicillin/streptomycin/fungizone</td><td>Gibco</td><td>15240-096</td><td>Or equivalent generic with concentration as follows (10,000 units/mL of penicillin, 10,000 &amp;micro;g/mL of streptomycin, and 25 &amp;micro;g/mL of Amphotericin B)</td></tr><tr><td>Phagostimulant</td><td>Made in House</td><td></td><td>Collected from prior tick feeds</td></tr><tr><td>Polycarbonate Pipe</td><td>McMaster-Carr</td><td>8585K204&amp;nbsp;</td><td>Cut to 45 mm length, 1.25 inch outer diameter, 1 inch inner diameter. Cutting requires a chop saw grinding wheel.</td></tr><tr><td>Rubber O-rings</td><td>McMaster-Carr</td><td>9452K38&amp;nbsp;</td><td>5 mm thick, 1.25 inch inner diameter</td></tr><tr><td>Soft touch forceps</td><td>VWR</td><td>470315-238&amp;nbsp;</td><td></td></tr><tr><td>Super glue</td><td></td><td></td><td>cyanoacrylate glue</td></tr><tr><td>Unryu paper&amp;nbsp;</td><td>Art supply stores</td><td></td><td>mulberry fiber 10 g/m&lt;sup&gt;2&lt;/sup&gt;. Purchased at Wet Paint art supply store, St. Paul, MN, USA</td></tr></tbody></table>

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.

A successful feeding depends on whether a partial or full engorgement is desired. Successfully fed I. scapularis turn a shade of gunmetal grey for adults and detach on their own from the membrane. However, if they are at least pea-sized, they may be detached from the membrane when finishing up the feeding. For immature stages of I. scapularis, the size for fully engorged ticks varies, and because, unlike adults, they do not exhibit a color change, detachment is the best way to determine if a tick is ful...

Watch this Scientific Journal Video about Tick Artificial Membrane Feeding for Ixodes scapularis at JoVE.com

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.

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 ...

tick-artificial-membrane-feeding-for-ixodes-scapularis

Research

JoVE Journal

Biology

3.1K Views.  University of Minnesota. 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.

Video: Tick Artificial Membrane Feeding for Ixodes scapularis

Testing Nicotine Tolerance in Aphids Using an Artificial Diet Experiment

Plants may upregulate the production of many different seconday metabolites in response to insect feeding. One of these metabolites, nicotine, is well know to have insecticidal properties. One response of tobacco plants to herbivory, or being gnawed upon by insects, is to increase the production of this neurotoxic alkaloid. Here, we will demonstrate how to set up an experiment to address this question of whether a tobacco-adapted strain of the green peach aphid, Myzus persicae, can tolerate higher levels of nicotine than the a strain of this insect that does not infest tobacco in the field.

In this video and assay is demonstrated that tests the tolerance of nicotine to two types of aphids one that infests tobacco plants in the field and one that does not.

Plants may upregulate the production of many different seconday metabolites in response to insect feeding. One of these metabolites, nicotine, is well ...

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 ...

Immunology and Infection

Preparation of Artificial Bilayers for Electrophysiology Experiments

Planar lipid bilayers, also called artificial lipid bilayers, allow you to study ion-conducting channels in a well-defined environment. These bilayers can be used for many different studies, such as the characterization of membrane-active peptides, the reconstitution of ion channels or investigations on how changes in lipid bilayer properties alter the function of bilayer-spanning channels.  Here, we show how to form a planar bilayer and how to isolate small patches from the bilayer, and in a second video  will also demonstrate a procedure for using gramicidin channels to determine changes in lipid bilayer elastic properties.   We also demonstrate the individual steps needed to prepare the bilayer chamber, the electrodes and how to test that the bilayer is suitable for single-channel measurements.

Planar lipid bilayers, also called artificial lipid bilayers, allow you to study ion-conducting channels in a well-defined environment. Here, we demonstrate the individual steps needed to prepare the bilayer chamber, the electrodes and how to test that the bilayer is suitable for single-channel measurements.

Planar lipid bilayers, also called artificial lipid bilayers, allow you to study ion-conducting channels in a well-defined environment. These bilayers can ...

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 ...

Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the lipidic cubic phase or in meso method. The method has been shown to be quite versatile in that it has been used to solve X-ray crystallographic structures of prokaryotic and eukaryotic proteins, proteins that are monomeric, homo- and hetero-multimeric, chromophore-containing and chromophore-free, and alpha-helical and beta-barrel proteins. Recent successes using in meso crystallization are the human engineered beta2-adrenergic and adenosine A2a G protein-coupled receptors. Protocols are presented for reconstituting the membrane protein into the monoolein-based mesophase, and for setting up crystallizations in the manual mode. Additional steps in the overall process, such as crystal harvesting, are to be addressed in future video articles. The time required to prepare the protein-loaded mesophase and to set up a crystallization plate manually is about one hour.

Herein is described the procedure implemented in the Caffrey Membrane Structural and Functional Biology Group to set up manually crystallization trials of membrane proteins in lipidic mesophases.

A detailed protocol for crystallizing membrane proteins by using lipidic mesophases is described. This method has variously been referred to as the ...

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 ...

Feeding and Quantifying Animal-Derived Blood and Artificial Meals in Aedes aegypti Mosquitoes

Females of certain mosquito species can spread diseases while biting vertebrate hosts to obtain protein-rich blood meals required for egg development. In the laboratory, researchers can deliver animal-derived and artificial blood meals to mosquitoes via membrane feeders, which allow for manipulation of meal composition. Here, we present methods for feeding blood and artificial blood meals to Aedes aegypti mosquitoes and quantifying the volume consumed by individual females.
Targeted feeding and quantification of artificial/blood meals have broad uses, including testing the effects of meal components on mosquito behavior and physiology, delivering pharmacological compounds without injection, and infecting mosquitoes with specific pathogens. Adding fluorescein dye to the meal prior to feeding allows for subsequent meal size quantification. The meal volume consumed by mosquitoes can be measured either by weight, if the females are to be used later for behavioral experiments, or by homogenizing individual females in 96-well plates and measuring fluorescence levels using a plate reader as an endpoint assay. Meal size quantification can be used to determine whether changing the meal components alters the meal volume ingested or if meal consumption differs between mosquito strains. Precise meal size quantification is also critical for downstream assays, such as those measuring effects on host attraction or fecundity. The methods presented here can be further adapted to track meal digestion over the course of days or to include multiple distinguishable markers added to different meals (like nectar and blood) to quantify the consumption of each meal by a single mosquito.
These methods allow researchers to singlehandedly perform high-throughput measurements to compare the meal volume consumed by hundreds of individual mosquitoes. These tools will therefore be broadly useful to the community of mosquito researchers for answering diverse biological questions.

Feeding and Quantifying Animal-Derived Blood and Artificial Meals in Aedes aegypti Mosquitoes

The goal of this protocol is to deliver animal-derived and artificial blood meals to Aedes aegypti mosquitoes through an artificial membrane feeder and precisely quantify the volume of meal ingested.

Females of certain mosquito species can spread diseases while biting vertebrate hosts to obtain protein-rich blood meals required for egg development. In ...

Human Exposure to Larval Ticks for Tickborne Disease Research

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.

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.

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

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 ...

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 ...

Tick Artificial Membrane Feeding for Ixodes scapularis

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Rozdziały w tym wideo

Testing Nicotine Tolerance in Aphids Using an Artificial Diet Experiment

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

Preparation of Artificial Bilayers for Electrophysiology Experiments

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

Crystallizing Membrane Proteins for Structure Determination using Lipidic Mesophases

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

Feeding and Quantifying Animal-Derived Blood and Artificial Meals in Aedes aegypti Mosquitoes

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

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