LABORATORY COLONIES OF

Michael L. Levin Lauren Schumacher Medical Entomology Laboratory, Rickettsial Zoonoses Branch Centers for Disease Control and Prevention (CDC)

Saravanan Thangamani Insectary Services Division, Galveston National Laboratory University of Texas Medical Branch (UTMB)

Table of Contents

Foreword Introduction Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.1 Equipping and Operating an Insectary 1.2 General Methods for Rearing Ticks 1.3 Culture

Chapter 2: Maintenance of Argasid Ticks in the Laboratory

Appendix I – Recommended reading on Tick life cycles under laboratory conditions.

Appendix II - Vacuum Aspirator Assembly

Foreword

Methods in Tick Research is part of a comprehensive collection of new rearing and handling protocols for vector species of importance to human health was borne out of the Vector Biology Research Resources Workshop held in June 2015 at the National Institutes of Health with the generous support by BEI Resources. This effort was inspired by the BEI manual, Methods in Anopheles Research, started by Mark Benedict and widely expanded by Paul Howell, which has become the gold standard of mosquito rearing and manipulation protocols. It continues to be the go‐to resource for laboratory‐ based scientists conducting basic research and public health entomologists from malaria endemic countries alike.

We would like to thank David Bland, Paul Howell, Michael Levin, Kevin Macaluso, Claudio Meneses, Tobin Rowland, Saravanan Thangamani, and Margaret (Peggy) Wirth, for sharing their techniques and expertise, and for putting together these protocols.

These protocols are intended as living, breathing documents with ample room for improvement based on a specific lab's capacity and infrastructure. They are intended as guidelines only, especially with regards to research involving vertebrate or biohazards, and containment, which require institutional approval tailored to individual laboratories.

We hope that the community can benefit significantly from the generation of this comprehensive set of new protocols and stimulate new work in vector biology and vector‐borne diseases. Kristin Michel ([email protected], Kansas State University) and Lyric Bartholomay ([email protected], University of Wisconsin‐Madison)

To provide feedback on this or any of the vector resources protocols, please send an email to [email protected].

Disclaimer. BEI Resources is funded under contract HHSN272201000027C by the National Institute of Allergy and Infectious Diseases, National Institutes of Health, Department of Health and Human Services. The views expressed in this publication neither imply review nor endorsement by HHS, nor does mention of trade names, commercial practices, or organizations imply endorsement by the U.S. Government. Introduction

Worldwide, ticks are second only to mosquitoes as vectors of human disease.

Many modern studies of the virulence of tick-borne pathogens in animals have been performed using pathogens grown and maintained in artificial culture, cell lines, or serial passages of blood or tissue homogenates with needle inoculation as the primary mode of infection. However, the dynamics, pathogenesis, and symptoms of infection as well as the subsequent immune response to infection and recovery strongly depend on the route of pathogen introduction into a susceptible host. In natural transmission, tickborne pathogens enter the vertebrate host with tick saliva, which assists establishment of infection by modifying the host immunological and cellular responses at the site of tick attachment. Ticks thus provide a modified environment in which bacteria and parasites can differentiate and proliferate and then migrate to other tissues to ensure successful biological transmission to the next invertebrate host. These tick factors can directly influence the infectivity and virulence of tick-borne agents as well as alter host responses. Consequently, conditions of natural transmission are poorly replicated when artificial methods of proliferation, maintenance, and inoculation into animals are used. Results of such laboratory studies may be difficult to extrapolate and apply to natural infections.

The transmission, maintenance, infectivity, virulence, and pathogenicity of tick-borne agents can be studied in the laboratory by using tick bite as the natural mode of infection. Accomplishment of these studies requires maintenance of laboratory colonies of many epidemiologically important hard tick vectors.

General setup of a tick laboratory, tick feeding protocols, and environmental requirements necessary for maintenance of ixodid tick colonies had been previously described by Patrick & Hair (1975)1 and Sonenshine (1999)2. Here we provide step-by- step recommendations for various procedures used in maintenance of ixodid tick colonies based on our 20+ years of experience. A list of publications describing laboratory life cycles of various ticks species and specific environmental conditions required for successful establishment and maintenance of laboratory colonies is included below.

1 Patrick, C. D., and J. A. Hair. 1975. Laboratory rearing procedures and equipment for multi-host ticks (Acarina: ). J. Med. Entomol. 12: 389-390. 2 Sonenshine, D. E. 1999. Maintenance of ticks in the laboratory, pp. 57-82. In K. Maramorosch and F. Mahmood (eds.), Maintenance of Human, Animal, and Plant Pathogen Vectors. Science Publishers, INC, Enfield, NH, USA.

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.1 Equipping and Operating an Insectary Page 1 of 3

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.1 Equipping and Operating an Insectary Michael L. Levin and Lauren Schumacher Medical Entomology Laboratory, Rickettsial Zoonoses Branch Centers for Disease Control and Prevention Use of laboratory animals as hosts for blood-sucking remains a time-proven and the most efficient method for establishment and propagation of slowly feeding ixodid ticks, despite introduction of techniques involving artificial feeding on either animal skins or synthetic membranes. New Zealand White rabbits are usually most accessible and most suitable hosts routinely used for establishment and maintenance of a large variety of multi-host tick species. Here we describe Standard procedures of maintaining colonies of multi-host ixodid ticks by feeding all developmental stages (larvae, nymphs, and adults) upon NZW rabbits. When needed, the same procedures here can be easily adapted to other species of laboratory or domestic animals from mice to dogs and goats. A summary of our experience in maintaining laboratory colonies of I. scapularis, I. pacificus, A. americanum, D. variabilis, D. occidentalis, H. leporispalustris, and R. sanguineus with descriptions of the complete laboratory life cycles and reliable production of uninfected ticks under standardized conditions had been published by Troughton and Levin (2007). Biosafety Establishment and maintenance of live colonies of ticks carry substantial biological risk for the personnel both inside and outside the tick-handling facility. This risk is associated with ticks’ obligatory hematophagy, ability to crawl under the protective equipment (PPE) and personal clothing, remain hidden and/or attached to the host as well as survive on or under furniture (e.g. on a counter, in an elevator, on a door handle, on a telephone receiver) for long periods of time. Unlike flying insects, whose escape and dispersal may be impeded by airlocks, ticks can and will crawl through doorways, or hitch a ride on clothing and packaging. In addition, ticks are competent and efficient vectors of multiple pathogenic and obligate or facultative symbiotic microorganisms. While it is possible to test representative samples of ticks in the colony for the presence or absence of known pathogens, there are no guarantees that colonized ticks do not carry some yet unknown pathogenic organisms (e.g. only recently discovered Panola Mountain Ehrlichia, Heartland virus), or that bacteria previously known as

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.1 Equipping and Operating an Insectary Page 2 of 3 endosymbionts of ticks are not indeed horizontally transmissible pathogens (e.g. Rickettsia slovaca, R. amblyommii). Therefore, it is imperative that live ticks are always handled in a specially designed and designated laboratory with implemented, strict biosafety procedures – at least ACL level 2 precautions – preventing an escape of hematophagous arthropods. All live ticks, regardless of their origin and/or the goal of a particular study, should be treated and handled as potentially infected. Appropriate PPE must be worn at all times by all personnel entering the designated facility regardless of task(s) performed or its duration. PPE must be inspected for ticks and appropriately removed when exiting the facility – Tick Laboratory PPE must not be allowed outside of the tick-designated area. Personal Protective Equipment Materials Needed:  Disposable White Gown or Coveralls  Hair cover  Disposable Gloves  Shoe covers (as needed for animal work)  Respirator (as needed for animal work)  Face shield or safety glasses (as needed for animal work) Proper personal protective equipment (PPE) should be worn at all times when working in the laboratory. Personal protective equipment required for the lab includes a white gown/ coveralls, a hairnet or a cap (hair must not be touching the gown), and properly fitting gloves. When putting on the gloves it is important to wear them pulled over the sleeves of a gown/coveralls to ensure that your wrists are covered at all times, and ticks could not crawl under the sleeves. The easiest way to ensure that the sleeve stays tacked into the glove at all times is to poke a hole through the end of the sleeve just beyond the hem, fit thumb through the hole, and then put on a glove, sliding it over the sleeve. When working with animals the personal protective equipment required consists of either coveralls or a white gown and shoe covers, hair cover, properly fitted respirator, face shield, and gloves.

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.1 Equipping and Operating an Insectary Page 3 of 3 Workstation Set-Up Materials Needed:  White work surface  Waste bucket (~ 1 gal) with 10% bleach solution and a lid.  Spray bottle with 10% bleach or Lysol spray bottle.  12 x 75mm Polystyrene tubes or similar (housing of engorged larvae, flat and engorged nymphs, flat adult ticks)  Flanged 12mm plug tops (stoppers for 12 x 75mm tubes)  Flanged 12mm plug tops with the tip cut off (opened)  11.1 ml snap cap Polystyrene containers or similar (housing of engorged female ticks – “oviposition containers”)  Snap cap lids (lids for containers)  Snap cap lids with 4-8 mm hole (open)  Nylon mesh (e.g. Voile White Decorator fabric – Hancock Fabrics) precut ~1x1”  Ruler  Scissors  Loew-Cornell 795 Paint brushes (size 2 and 6) or similar small artist paint brushes  Paper towels  Permanent marker  Fine point permanent marker  Vacuum with aspirator (desirable though not absolutely necessary)  White tray for sorting ticks (approx. 2.5” deep)  Gloves  Masking tape for catching and disposing of escaped ticks – “spill cleanup”.  Petroleum jelly  Container with 10% solution of bleach for disposal of waste materials including used vials  Lysol spray for decontamination of the sorting trays

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.2 General Containment and Handling Page 1 of 5 1.2 General Containment and Handling It is recommended that the doorframes to laboratories holding ticks are surrounded with petroleum jelly, to prevent ticks from escaping the designated facility. Petroleum jelly should be replaced once a month. If petroleum jelly is not used a similar method should be in place to prevent ticks from leaving the laboratory, in the event that they escape. The workstation should be set up on a white surface (Figure 1). This makes it easier to spot ticks easily in case they climb out of the tray. Vial/tube stoppers with holes in them (open leads) fitting the tubes and containers that house ticks, pre-cut nylon mesh, the waste bucket with 10% bleach (and an aspirator for sorting/counting ticks – Appendix II) must be prepared beforehand. To make the open flanged plug tops, place the stopper on a hard surface with the tip (bottom) of the lid facing up and make a hole using a 4 mm biopsy punch. Alternatively, tip can be cut off using a knife or a box cutter (Figure 2). Snap cap lids can be cut either similarly using a 4 mm biopsy punch or an office desktop hole-punch. Nylon mesh allows sufficient ventilation of tubes (through holes in the leads) while preventing larvae from escaping from the containers. It should be pre-cut into small pieces approximately 1x1”. Used tubes, tube stoppers, gauze, and dead ticks should be discarded into a large (gallon sized) vessel ~3/4 filled with 10% bleach. Paper towels used for cleaning and drying engorged ticks may be placed into a sealed Ziploc bag to ensure that any ticks that were thought to have been dead

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.2 General Containment and Handling Page 2 of 5 are securely sealed inside. Sorting tray(s), tweezers, brushes and any other reusable instruments must be cleaned with Lysol spray before and after working with each cohort of ticks. This will minimize risk of inadvertent contamination of tick colonies with enthomopathogenic fungi and mites, especially when working with arthropods recently derived from the field. Cotton fabrics, filter paper, paper labels, and so on should be avoided as they prone to getting moldy in the high humidity environment, which will result in tick die-offs both during and after molting. Masking tape is useful for capturing stray ticks. It, however, is not efficient as an escape barrier because most of ticks are quite capable of walking on it. Ticks placed in the sorting tray must not be left unattended however briefly. All tools and supplies including extra disposable gloves must be arranged around the work station in such a manner that they are easily accessible (Fig. 1) and can be reached without diverting one’s attention from the sorting tray while it contains ticks (either live or seemingly dead). Housing Ticks between feedings

Materials Needed:

 Incubator(s)  High humidity chamber(s) - Dry-Keep Desiccator Cabinets or any other Desiccator where water (or salt solution) can be placed on the bottom to maintain desired high levels of relative humidity.  Water (or salt solution)  Fine point permanent marker  Felt tip permanent marker  12 x 75 mm polystyrene tubes or similar tubes for storing ticks  11.1 ml polystyrene containers or similar container for storing engorged females  Open flanged plug tops (lids that fit tubes)  Open top snap cap lids (lids that fit containers)  Precut Nylon mesh  Plastic containers for storing tick tubes (empty boxes from pipette tips or Tupperware containers work well)

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.3 General Containment and Handling Page 3 of 5 Placement of Ticks in Polystyrene Containers All tubes in which the ticks are stored in should be capped with a piece of nylon mesh fully covering the tube and a coordinating open lid (Figure 2). This will ensure that the ticks are secure inside the tube, and that there is sufficient airflow/humidity reaching the ticks. Limiting numbers of ticks placed in tubes prevents overcrowding, which may result in decreased survival and longevity. It is not advisable to place any filter paper inside tick tubes or containers. Although such paper may minimize condensation inside the tubes and somewhat control overcrowding, it also provides a substrate for fungi growth, which should be prevented by all means. In addition, polystyrene tubes are much less susceptible to water vapor condensation than traditionally used glass containers. Flat nymphs are stored in 12x75mm polystyrene tubes capped with nylon mesh and an open top. Flat nymphs can be stored up to 50 nymphs per tube, with the exception of spp. No more than 25 Ixodes spp. nymphs should be stored per tube as these nymphs are more susceptible to overcrowding. Adults are stored up to 10 per 12x75mm polystyrene tube capped with nylon mesh and an open top. It is better to keep males and females in separate tubes. For Ixodes spp., separation of adult ticks by sex immediately after nymphal molting is required to prevent inbreeding and a die-off in males as these ticks are capable of mating off host – almost as soon as their cuticle hardens after the ecdysis. Engorged females are stored individually; this allows to better control numbers of larvae per container as well as correlate specific progenies to the PCR results in individual spent females. The size of the tube depends on the size of the engorged ticks. Tubes should be sealed with a piece of nylon mesh and the coordinating open lid. Subsequent eggs and larvae are kept in the same tube as manipulations of eggs tend to decrease their hatchability and transfers of hatched larvae from one container to another usually results in decreased longevity. When sorting and storing engorged larvae and nymphs for molting, it is especially important to limit numbers of ticks per tube as overcrowding results in quick accumulation of waist, growth of fungi, and subsequent die-offs of both pre-molt and freshly molted ticks. For that reason, we recommend housing only up to 25 engorged larvae per 12x75mm polystyrene tube, or up to 100 engorged larvae per 11.1 ml container. Molting success of Ixodes spp. larvae drops significantly in large groups, therefore we recommend always storing Ixodes larvae in 12x75mm tubes, up to 25/tube. Engorged larvae of other genera may be stored in either 12x75mm tubes or

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.2 General Containment and Handling Page 4 of 5 11.1 ml polystyrene containers. For the same reasons, we recommend storing engorged nymphs in small groups of 10/tube (see Appendix I for more detales). Each tube or vial should be clearly labeled with identifying information including (a) species of ticks, (b) date of engorgement, (c) number of ticks in the tube and the life stage, (d) ID of the animal host (Figure 3). This information can be either handwritten using a fine-point permanent marker or printed on plastic adhesive labels. As stated above, paper labels should be avoided. All tubes holding ticks from the same cohort should be stored together in a plastic box clearly labeled with the same identifying information as on the tubes (Figure 4). These boxes are stored in desiccators within incubators. The type of plastic box is not important, but the use of cardboard boxes for tick storage should be avoided since the cardboard will begin to mold inside the desiccators which could affect the longevity of the ticks.

Desiccators and Incubators Ixodid ticks (unlike many argasids) in general require high relative humidity (rH) for their survival. We routinely keep rH in our humidity chambers at or above 90%. To achieve this, a tray with water (or salt solution if desired) is placed on the bottom of each desiccator (Fig. 4). There only needs to be enough liquid (~1/4”) to cover the surface of the trays in order to keep the desiccators at the correct humidity level. If using pure water, the trays need to be cleaned regularly (every 1-2 weeks) to prevent mold growth. Use of saturated salt solutions may delay the mold growth. However, one must keep in

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.2 General Containment and Handling Page 5 of 5 mind that the water vapor concentration and therefore the relative humidity over a salt solution is less than that over pure water. Ticks are sensitive to both the ambient temperature and photoperiod. Increase in ambient temperature in general results in expedited development of ticks (molting, gestation, oviposition, hatching) but also in significantly shortened life span of unengorged ticks. We found that keeping ticks in environmental chambers (incubators) sat at 22°C ± 1°C and long photoperiod of 16:8 (L:D) provides reasonable balance between the rate of development and longevity [Troughton, 2007 #8032]. Long photophase also helps to prevent occurrence of the diapause in (most) Ixodes spp.

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.3 Tick Culture Page 1 of 10 Chapter 1.3 Tick Culture Feeding ticks on animals New Zealand White rabbits are usually most accessible and most suitable hosts routinely used for establishment and maintenance of a large variety of tick species. Here we describe Standard procedures of maintaining colonies of ixodid ticks by feeding all developmental stages (larvae, nymphs, and adults) upon NZW rabbits. When needed, the same procedures here can be easily adapted to other species of laboratory or domestic animals from mice to dogs and goats. Preparing Rabbits for Materials Needed:  Water moats (metal or plastic trays filled with 1/5-1” of water) placed under the individual animal cages.  Petroleum jelly.  Drugs and equipment necessary for anesthesia/sedation of animals (e.g. mixture of Ketamine | Xylazine 25-35 | 5 mg/kg, IM).  Hair clipper with #40 or #50 clipper blade.  Cotton Stockinette (2-3” diameter) pre-cut to 4-5” segments.  Nylon stocking (e.g. nylon pantyhose) pre-cut to 4-5” segments  Medical or veterinary grade skin adhesive - for attachment of feeding bags to the skin (technical or household glues and adhesives can cause skin irritation and flake off before ticks have time to complete their feeding)  Plastic bottles 2.5-3.0” diameter  Rubber bands  Elizabethan collars of appropriate sizes  Waterproof tape (e.g. “Johnson and Johnson” medical bandage tape).  Permanent marker (e.g. Sharpie)  Large capacity vacuum (e.g. Shop Vacuum) to collect clipped hair.

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.3 Tick Culture Page 2 of 10 New Zealand White Rabbits being used for tick feedings should be housed in cages held above water moats (Figure 5). The edges of the water moats should also be lined with petroleum jelly. This setup ensures that ticks cannot escape from the cage if feeding bags become damaged or detached. In addition, it is strongly recommended that the door frame in the room housing tick-infested animals is covered with petroleum jelly to ensure that escaped ticks would not crawl out in to hallways. 1. First, determine how many feeding bags will be placed on rabbits, and cut the needed number of stockinette and nylon for the bags. -Each feeding bag holding adult ticks will require two (2) 4-5” segments of cotton stockinette -Each feeding bag holding larval or nymphal ticks will require one (1) 4-5” segment of cotton stockinette for the outer layer AND one (1) 4-5” segment of nylon stocking (pantyhose) for the inner layer. 2. Anesthetize/sedate a rabbit. 3. While waiting for anesthesia to take effect, prepare double-layered feeding bags for ticks. Bags for larval and nymphal ticks have an inner nylon bag and outer stockinette. This is necessary to ensure that small immature ticks do not escape from the bags through hales in the stretchable cotton webbing. Bags for adult ticks can be made out of 2 layers of stockinette.

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.3 Tick Culture Page 3 of 10 To prepare feeding bags, stretch the inner layer (cotton stockinette for adults or nylon stocking for immature ticks) over the bottom of a plastic bottle. Stretch the outer layer - a piece of stockinette over the inner layer. Turn the top edge of the outer layer stockinette down to create a “flange”, then turn the inner stockinette/nylon down in the same manner making sure that the inner “flange” is shorter than the outer one (Figure 6). 4. Once the rabbit is sedated, clip hair on rabbit’ dorsum as close to the skin as possible. Use surgical clipper blades #40 or #50. 5. Hobble the rabbit’s back legs with waterproof medical tape (Figure 7). Leave ~ 2 inches of space between the rabbits’ feet so the animals can move around the cage and sit normally; and ensure that the tape is not wrapped too tightly around the foot itself, cutting off the blood circulation. Gently wrap the medical tape twice around the tibia just above tarsal joint back leg above the foot. Without tearing the tape, bring the spool of tape to the other leg and wrap it twice around the other leg in the same position, making sure to leave about 2” of tape between the two legs. Without tearing the tape, bring the spool of tape back to the first leg, pressing the tape to the sticky side of the 2” space of tape left between the feet, to strengthen the hobble and keep the tape from sticking to the rabbit’s fur. Pull about 4.5” of extra tape and tear the tape from the spool. Bring the tape around the leg one last time, and wrap the extra 4” of tape around the two inch hobble between the two feet to further reinforce the hobble. If hobbles break, the rabbits are prone to scratch and damage the bags.

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.3 Tick Culture Page 4 of 10 6. Apply an adhesive on the “flanges” of both layers of the feeding bag (Figure 8). Make sure that glue covers both flanges all the way around without gaps. Place the bottle with the bags on the shaven area of the skin and unfold flanges the bag with glue down onto the rabbit’s skin (Figure 9). Press the bags/glue firmly onto the back of the rabbit. While holding the base of the bag the rabbit’s back with one hand, very gently remove the bottle from inside the bags with the other. Secure the open end of a bag with a rubber band, being careful not to disrupt the bags. 7. Several feeding bags may be placed on each rabbit if needed (Figure 10). 8. Once all bags are in place, put an e- collar around the rabbit’s neck, leaving space (you can fit two fingers) between the collar and the neck.

9. Being mindful of freshly glued bags, place the rabbit in the tick-feeding cage for recovery. Anesthesia/sedation need to last long enough for the adhesive to harden (even if not to dry completely) 10. Take rabbit ID card from holding cage and write rabbit number with the sharpie. Place cage card in holder on tick feeding cages 11. Repeat for all rabbits according to the feeding schedule.

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.3 Tick Culture Page 5 of 10 12. Vacuum all of the rabbit fur from the work area and equipment including clippers, plastic bottles, plastic carrier, etc. Spray the counter with 10% bleach and wipe clean. 13. Wait for at least 3 hours before placing tick into the feeding bags to ensure that the glue is completely dry. Placing Ticks on Rabbits Note: Always keep one hand on an open bag while placing ticks inside. This will allow you to quickly pinch the bag closed if the rabbit were to begin to move to avoid any chance of ticks coming out. 1. Count and combine ticks for each feeding bag. To avoid overcrowding, place 1 batch of larvae per larval bag, 200-300 nymphs per nymphal bag, and 20-30 pairs of adults per adult bag. Placement of too many ticks within the limited space of a bag may cause an immediate hypersensitivity reaction and rejection of attached ticks resulting in low feeding success and further decreased survival during the molt. Whenever possible, make note of exactly how many ticks you placed into each bag to ensure that all ticks are recovered after feeding (male ticks may stay attached and hidden in the growing hair). It is easiest and safest to place all of the ticks destined for the same feeding bag into a single tube or vial. 2. Prepare a 1 L waste beaker filled approximately ¾ with 10% bleach solution to discard emptied tick tubes. Add a couple of drops of soap or detergent to the beaker and stir avoiding making any bubbles. Extra detergent in the solution will ensure that any live ticks remaining in the tubes would sink and be killed. 3. Remove rabbit from cage. Open the first bag and fold the open ends of the outer bag down, then fold the open ends of the inner bag down, over the outer bag. This will allow you to be able to see inside the bag. 4. Grab the tube of ticks that will go into this bag. Place your index finger over the lid of the tube, and firmly tap the tube on the table to knock all of the ticks to the bottom of the tube. Make sure there are not any ticks on the lid. Open the tube (if working with larvae immediately place the lid into the bleach solution) and quickly, but carefully, dump the ticks into the bag. Keep your less dominant hand around the bag as you turn the tube into the bag. Use your index finger and thumb of the hand on the bag to grab the upside down tube (still in the bag) leaving your other fingers around the bag. Keeping your hand on the bag will allow you to be able to quickly hold the bag closed in the event that the rabbit begins to move, or ticks begin to crawl up. Making sure the open end of the

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.3 Tick Culture Page 6 of 10 tube is well within the bag, use your index finger on your dominant hand to firmly flick the tube to knock all of the ticks into the bag. For larvae, limit yourself to one or two firm flicks. You will not be able to safely empty all of the larvae into the bag. They will begin to crawl up the bag if you try. Get the most you can. 5. For larvae: Keeping your hand on the bag in the same position as described, use your dominant hand to grab the tube and submerge it into the Clorox solution. For nymphs and adults: Keeping your hand on the bag in the same position as described, use your dominant hand to flip the empty tube back over, still holding it with your index finger and thumb of the hand on the bag. Grab the lid to the tube and mesh, and reseal the tube, just in case there are still ticks inside that you may have overlooked without realizing it, place the tube aside. Work very carefully, but quickly to ensure that ticks do not begin to crawl up and out of the bag. 1. Seal the inner bag. Unfold the inner bag from the outer stockinette. If the inner bag is nylon, pinch the bag above where any ticks have crawled up to with your non- dominant hand, and twist the material above with your dominant hand. Fold only the twisted part of the material over itself, and seal it tightly with a rubber band (Figure 11). If the inner bag is made out of cotton stockinette, fold the open end down approximately 1/3 of the length of the bag and then fold the crease in an accordion manner. Fasten it tightly with a rubber band. Fold and fasten the outer stockinette bag in the same accordion fashion. Both bags should be securely sealed with rubber bands.

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.3 Tick Culture Page 7 of 10 Collecting Ticks from Rabbits Note: Animals infested with ticks must be checked regularly (daily) to ensure integrity and security of feeding bags as well as to collect detached ticks. If left inside the bags, engorged ticks will crawl around irritating the animal, which can damage them in its efforts to alleviate irritation. In addition, engorged larvae and even nymphs of many tick species are prone to desiccation and can die if left in the bags for more than 24 hours. Materials Needed:  Bench-top vacuum pump. Laboratory pumps with HEPA filters are recommended to prevent tick feces from being aerosolized. Alternatively, a vacuum trap flask with Lysol solution or mineral oil can be used to capture any aerosolable materials (Figure 12).  50-100 ml side-arm flasks – for collection of engorged larvae and nymphs  Rubber stopper with a glass tube bent at an obtuse angle.  Cotton ball  Polystyrene vials – for collection of engorged adult ticks.  Forceps

1. Place a small wad of cotton into the opening of the side-arm on the flask intended for collection of engorged larvae or nymphs (Figure 13). Cotton will prevent ticks from being sucked into the pump, but do not “overstuff” the side-arm as it will impede the airflow and decrease suction. 2. Plug the flask using a rubber stopper with an angled glass tube. The lower portion of the tube should extend below the stopper to approximately ½” – ¾” from bottom of the flask. Attach the side-arm of

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.3 Tick Culture Page 8 of 10 the flask to the inlet of a vacuum pump (or a filtering flask) the using rubber or Tygon tubing. 3. Remove a rabbit from the cage and place it on the counter (examination table). Make sure not to pull on the feeding bags or to crush engorged ticks inside. 4. Examine the rabbit to make sure that all bags are still securely attached to the rabbits back and that none of them have any holes (if they do have holes or have become unattached, glue another layer of stockinette around the damaged bag). 5. Open the feeding bag by removing rubber bands and unfolding the material. Remember to keep your hand on the bag at all times while the bag is open. Collect (vacuum) the engorged larval or nymphal ticks crawling inside the bag using a side-arm flask with angled glass tube (Figure 14). Collect engorged female ticks by (gloved) hand into large polystyrene vials. Male ticks will be removed with forceps when all females in the bag have finished feeding - use forceps only to remove males as fully engorged females are easily damaged by hard instruments. 6. Engorged ticks must be removed at least once per day. Larvae of Ixodes spp. and Rhipicephalus spp. need to be collected twice a day to avoid desiccation. 7. Reseal the inner and outer layers of the feeding bag as described previously immediately after collecting ticks. 8. Bags, E-collar, and hobbles can be removed after all ticks on a particular rabbit have completed their engorgement and/or have been removed. Cleaning and Storing Engorged Ticks Engorged ticks collected from animals are usually externally contaminated with tick feces and animal blood. This contamination may result in ticks becoming moldy and dying when stored in high humidity chambers. It is therefore advisable to clean ticks before placing them into appropriate holding containers.

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.3 Tick Culture Page 9 of 10 1. Easiest way to clean these is by placing them in a strainer and using running water to dislodge all external contaminants. 2. Surface-dry the washed ticks by dumping them onto a paper towel. 3. Sort and count live engorged ticks using a paintbrush (Figure 15) or an aspirator (Appendix II). 4. Cleaned ticks are stored in appropriate labeled containers as described above in #IV “Housing Ticks between feedings”. 5. If you find a tick outside of the tray, use masking tape to immediately pick it up, and put the tape inside a Ziploc bag. 6. Used paper towels with any remaining dead ticks should be folded and placed into a sealed Ziploc bag. Cleaning Molted Ticks

Freshly molted ticks should be moved into clean containers. Waste accumulating in the vials during the molting process (tick feces, dead skins) are prone to becoming moldy in the high humidity environment; and growing fungi can entrap and kill ticks. 1. Prepare washing solution by adding a drop of liquid dish soap to 1-2 L of water. Pour a small amount of solution (about 50ml) into a small beaker. 2. Take a vial with molted ticks and firmly tap the bottom of the vial on a hard surface to knock all ticks to the bottom of the vial. Hold the lid of the vial with your index finger while knocking to ensure that it does not come off inadvertently. 3. Make sure there are no ticks clinging to the lid and open the tube. Place the lid and mesh into the white tray. Knock the flat ticks out of their tubes into the beaker with washing solution. Ticks remaining in a tube can be moved using a small paint brush or tweezers. Ensure that there are no ticks remaining on the lead and nylon mesh, or in the vial and discard all into waste bucket with 10% bleach solution. Several vials containing ticks from the same cohort can be emptied into the beaker – one after another.

Chapter 1: Maintenance of Multi-Host Ixodid Ticks in the Laboratory 1.3 Tick Culture Page 10 of 10 4. Swirl the ticks around in the cleaning solution either by gently rocking the beaker or using a brush. 5. Pour the cleaning solution and ticks into a strainer over an empty container. Make sure that no ticks remain in the beaker. Allow the water to flow through the strainer. Very gently tap out any excess water from the strainer. Take the strainer and push it gently on the paper towel to absorb the water. Flip the strainer over and flick the ticks out of the strainer onto a dry paper towel. Quickly but carefully inspect the strainer for ticks that may still be caught in the strainer and use a paint brush to move them to the tray. Note: Occasionally glance at your strainer while cleaning the ticks to make sure there are not any ticks crawling on it that you may have missed. 6. Dry the ticks by folding the paper towel over the ticks. Unfold the paper towel. 7. Use a paint brush to gently pick up each thoroughly dried, living tick and put them into tubes. 25 flat nymphs per tube for Ixodes, 50 flat nymphs per tube for other species, and 10 flat adults per tube. Collect all of the ticks in the tray, being careful not to allow ticks to crawl up the sides of the tray. Do not allow your PPE to touch the sides of the tray, to make sure ticks do not get the chance to crawl onto your PPE. 8. If for some reason you find ticks outside of your tray immediately pick them up with masking tape. This is to make sure you do not inadvertently mix a tick in from a different cohort. 9. Fold the paper towel over several times to seal dead ticks inside. Put paper towel inside Ziploc bag when finished to make sure that ticks don’t escape from the garbage. Inspect the tray to make sure it is free of ticks. 10. Label all clean tubes with sharpie: species, number of ticks, host, and date written on the cohort box. Put vials with clean ticks in their plastic box and place them back in the designated desiccator. 11. Spray the inside of the tray with Lysol solution and wipe clean. Ziploc bags holding the used paper towels can be placed into a larger Ziploc or autoclave bag (double-bag) and deposited into the biohazard waste. Clean beakers with 10% bleach and rinse it out. Clean the paint brushes with 10% bleach and allow to air dry. Clean the strainer with Lysol solution or 10% Clorox. Rinse the strainer, shaking it out the remove excess water, and spray it with 70% ethanol. Allow it to dry before use.

Chapter 2: Maintenance of Argasid Ticks in the Laboratory Page 1 of 2

Chapter 2: Maintenance of Argasid Ticks in the Laboratory- SOP

Saravanan Thangamani Insectary Services Division, Galveston National Laboratory University of Texas Medical Branch (UTMB)

Purpose The purpose of this standard operating protocol (SOP) is to describe procedures to rear and maintain argasid (soft) ticks.

In-Vivo Tick Feeding Procedures

1. Conduct the following procedures on a white tray with masking tape around the periphery of the tray. The tape would serve as a containment barrier if a tick would like to escape.

2. Anesthetize the mice as per the approved IACUC protocol. Alternatively, mice could be restrained in a small mesh cage (without anesthesia) during tick feeding.

3. Chill the tick vials prior to handling them.

4. Larval ticks are painted onto the neck and upper-back of the mouse using fine brush. Nymphal and adult ticks may be placed on the neck and upper-back of the mouse using blunt end forceps. Between 30-60 mins, ticks fed to repletion will drop-off the mouse. Collect the fed ticks with blunt-end forceps (without puncturing) and transfer to a fresh glass vial for housing inside the incubator.

Tick Housing

1. House ticks in a sterile clear-glass vials (Wheaton Inc) with a plastic snap cap. For sufficient air exchange, cut 2-4 2mm holes in the snap cap and secure the lid sterile fine mesh.

Chapter 2: Maintenance of Argasid Ticks in the Laboratory Page 2 of 2

2. Sterile mouse bedding material may be added to the vial. This will allow the ticks to burrow in between the bedding material (simulating the natural environment).

3. Keep the Storage vials inside a glass desiccator containing saturated salt solution in the desiccator basin (saturated potassium nitrate and potassium sulfate will provide 94% and 97% relative humidity). Alternatively, plaster of Paris and charcoal could be used at the vial base to maintain humidity.

4. Apply Vaseline grease around the rim of the desiccator to maintain humidity.

5. Storage vials must be labeled with the species, sex in the case of adult ticks, number of individuals for each life cycle stage, date of collection and name of the PI.

6. Store the desiccators in an environmental chamber maintaining 23°C at 95% relative humidity with 14:10 hour dark/ light cycle (this information is specific for . Some species might require different housing conditions).

Appendix I.

List of references for tick life cycle under laboratory conditions:

Tick References Troughton and Levin, 2007 Amblyomma triste Labruna et al, 2003 Amblyomma auricularium Faccini et al 2010 Amblyomma incisum Szabó et al 2009. Amblyomma brasiliense Sanches et al, 2008 Amblyomma marmoreum Fielden et al 1992 Amblyomma ovale Martins et al 2012 Dermacentor silvarum Liu et al 2005, Yu et al, 2010 Dermacentor occidentalis Troughton and Levin, 2007 Troughton and Levin, 2007 Hyalomma asiaticum Chen et al 2009 Hyalomma rufipes Chen et al 2012 Haemaphysalis qinghaiensis Ma et al 2013 Haemaphysalis leporispalustris Troughton and Levin, 2007 Hyalomma marginatum Knight et al 1978 Haemaphysalis leachi Jacobs et al 2004 Ixodes minor Banks et al, 1998 Rhicephalus sanguineus Jacobs et al 1992 Ixodes luciae Labruna et al 2009 Dotson and Oliver, 1995 Ixodes woodi Banks et al 1998 Ixodes scapularis Ogden et al 2004 Padgett and Lane 2001 Ixodes loricatus Schumaker et al 2000 Konnai et al 2008 Ixodes scapularis Troughton and Levin, 2007 Ixodes pacificus Troughton and Levin, 2007 Jacobs et al 1992, Troughton and Levin, 2007 Rhipicephalus bursa Yeruham et al 2000

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(: Ixodidae) in the laboratory. J Med Entomol. 1998 Jul;35(4):496‐9. PubMed

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Banks CW, Oliver JH Jr, Hopla CE, Dotson EM. Laboratory life cycle of Ixodes woodi (Acari: Ixodidae). J Med Entomol. 1998 Mar;35(2):177‐9. PubMed PMID:

9538581.

Chen Z, Yu Z, Yang X, Zheng H, Liu J. The life cycle of Hyalomma asiaticum kozlovi Olenev, 1931 (Acari: Ixodidae) under laboratory conditions. Vet

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