A Beginner’s Guide to Collecting Questing Hard Ticks (Acari: Ixodidae): A Standardized Tick Dragging Protocol

Copyright © The Author(s) 2020. Published by Oxford University Press on behalf of Entomological Society of America.

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Abstract

Tick-borne diseases are emerging globally, necessitating increased research and coordination of tick surveillance practices. The most widely used technique for active collection of host-seeking, human-biting tick vectors is ‘tick dragging’, by which a cloth is dragged across the top of the vegetation or forest floor and regularly checked for the presence of ticks. Use of variable dragging protocols limits the ability of researchers to combine data sets for comparative analyses or determine patterns and trends across different spatial and temporal scales. Standardization of tick drag collection and reporting methodology will greatly benefit the field of tick-pathogen studies. Based on the recommendations of the Center for Disease Control and Prevention and other ecological considerations, we propose that tick dragging should be conducted to sample at least 750 m 2 along linear transects when habitat allows in a manner that reduces bias in the sampled area, and report density of each tick species and life stage separately. A protocol for constructing a standard drag cloth, establishing linear transects, and drag sampling is presented, along with a downloadable datasheet that can be modified to suit the needs of different projects. Efforts to align tick surveillance according to these standard best practices will help generate robust data on tick population biology.

Tick surveillance in medical and veterinary entomology began in 1902 (Grutzner 1902), when Ixodes ricinus (Linnaeus) were collected from sheep to understand their effect on sheep health. While early efforts focused on collecting ticks directly from their bloodmeal hosts, methods of collecting ticks off-hosts were eventually developed to monitor tick populations during their host-seeking period, especially as their importance as vectors of agricultural and human pathogens grew (Milne 1943; Falco and Fish 1988, 1992; Daniels et al. 2000). Methods such as flagging, dragging, CO₂ baiting, sweeping, and the use of human sentinels have all been employed for surveillance of host-seeking ticks, and specific sampling methodology is highly variable among research groups depending on the research question or surveillance objective (Falco and Fish 1992, Schulze et al. 1997, Gherman et al. 2012, Chong et al. 2013, Eisen and Paddock 2020, Kugeler and Eisen 2020).

Different methods for collecting host-seeking ticks are used because the life history of some tick species necessitate specialized collection techniques. For example, the lonestar tick, Amblyomma americanum (Linnaeus), a vector of Rickettsia rickettsia (Brumpt) (Rickettsiales: Rickettsiaceae), Borrelia lonestari (Swellengrebel) (Spirochaetales: Spirochaetaceae), and Francisella tularensis, (McCoy and Chapin) (Thiotrichales: Franisallaceae) is an aggressively mobile tick and CO₂ traps can effectively lure host-seeking ticks from the local environment (Childs and Paddock 2003, Petry et al. 2010). Nidicolous tick species like Ixodes angustus (Neumann), which is closely associated with rodent hosts, commonly reside in the burrows and nests of their hosts (Easton and Goulding 1974, Furman and Loomis 1984), so their collection necessitates destructive sampling of the host environment or on-host collections (Foley et al. 2011). Other tick species are classified as ambush or ‘sit-and-wait’ predators that quest to the top of vegetation and wait to attach to a host as they pass by (Waladde and Rice 1982). This includes the blacklegged ticks, Ixodes scapularis (Say) and Ixodes pacificus (Cooley and Kohls), both vectors of the Lyme disease pathogen and several other emerging pathogens (Steere and Malawista 1979, Burgdorfer et al. 1985, Spielman et al. 1985, Lane and Burgdorfer 1987, Costero and Grayson 1996, Mun et al. 2006, Teglas and Foley 2006, Barbour et al. 2009). Accordingly, for these ‘sit-and-wait’ ticks, collection methods that provide a large surface area to contact and collect these ticks are useful.

Host-seeking within the Ixodidae family is facilitated by the tick’s Haller’s organs, a pair of sensory organs located on the first segment of their first pair of legs (Nuttall et al. 1908, Waladde and Rice 1982, Steullet and Guerin 1992). When host-seeking, these ticks crawl to the top of vegetation and maneuver their first two legs in anticipation of a passing bloodmeal host. Typically, this behavior is shown in generalist hard tick species, such as I. scapularis, I. pacificus, Dermacentor occidentalis (Say), Dermacentor andersoni (Stiles), A. americanum, and Amblyomma maculatum (Koch) (Eisen and Paddock 2020). One-host ticks, or ticks that spend their complete life cycle on a single host such as Rhipicephalus annulatus (Say), and nidicolous ticks do not typically exhibit questing behavior (Furman and Loomis 1984) due to a strong association with single or limited set of hosts. Host specificity and lack of questing behavior minimize the threat of one-host and nidicolous ticks in the transmission of pathogens to humans and other domestic or agricultural hosts. In contrast, questing tick species are often generalists in their host associations, and therefore pose a greater risk of pathogen transmission to susceptible hosts. However, even within the same tick species complex, questing behavior, and therefore human risk, can vary greatly across different geographic regions (Diuk-Wasser et al. 2006, Teel et al. 2010, Hamer et al. 2012, Salkeld et al. 2014, Arsnoe et al. 2019). For example, I. scapularis tends to quest more frequently and to higher heights in the northern United States relative to the southern United States (Arsnoe et al. 2019), a behavioral difference that correlates with the high incidence of human Lyme disease in the north relative to the south. Differences in tick questing behavior are likely attributed to variation in climate, host feeding patterns, and tick genetics.

Two of the most commonly used sampling techniques for host-seeking hard ticks are dragging and flagging. These methods are used for the active collection of ticks, and although they remove ticks from the environment, they are not recognized to have an effect on diminishing or controlling tick populations (Daniels et al. 2000, Tälleklint-Eisen and Lane 2000). Both methods exploit the questing behavior by dragging or sweeping a heavy cloth across leaf litter or vegetation to pick up host-seeking ticks (Waladde and Rice 1982, Furman and Loomis 1984, Steullet and Guerin 1992). These two methods are often used in the literature interchangeably but are actually distinct sampling techniques, usually motivated by different objectives (Carroll and Schmidtmann 1992, Dantas-Torres et al. 2013, Rulison et al. 2013, Newman et al. 2019). In drag sampling, the collector moves along an established transects with the drag cloth trailing behind them and in contact with the vegetation, resulting in a known sampling area that can therefore generate estimates of tick density. Accordingly, dragging is the method of choice when one wishes to calculate epidemiologically important metrics including the density of nymphs (DONs) and density of infected nymphs (DINs) (Ostfeld et al. 2006, Gatewood et al. 2009, Diuk-Wasser et al. 2012, Pepin et al. 2012, Eisen and Paddock 2020). These metrics can then be compared between sites and over time (Guerra et al. 2002, Brownstein et al. 2005). In contrast, the flagging technique uses a similar device as the previously described drag cloth, but more closely resembles a flag attached to a pole. The collector uses this adapted flagging device by waving it over vegetation, representing the motion of waving a flag (Dantas-Torres et al. 2013, Rulison et al. 2013). The benefit of flagging is that the collector has more control over the cloth and dowel so the technique can be used in a more targeted manner to collect a certain species or life stage of tick, especially adult ticks, and usually aims to collect a minimum sample size of ticks. Flagging is generally used to determine the presence or lack of detection of ticks (Burgdorfer et al. 1985, Liz et al. 2001) and tick density is not typically evaluated using this method. Further, flagging is also more difficult to standardize between collectors, as sampling effort is not standardized, adding challenges to comparisons of results of different flagging studies (Kugeler and Eisen 2020).

The Centers for Disease Control and Prevention recently issued suggested procedures for Ixodes surveillance (Eisen et al. 2019). Building upon the CDC guidelines, we propose a standardized tick dragging protocol with specific attention to sampling area, standardizing the sampling interval distance, and uniform reporting of tick density. The goal of this paper is to provide a standardized tick dragging protocol, which could be useful to new tick researchers or those with an interest in ensuring that relevant data are collected in a standardized way to facilitate comparisons to other studies. We also provide detailed instructions on how to construct a standard tick drag cloth, include a step-by-step protocol for how to set up a linear transect, and provide a template for a field data sheet that can be adapted to suit the needs of individual research projects.

Protocol

Overall, the method of tick dragging involves walking through the designated area to be sampled with a 1-m by 1-m 2 white cloth dragging behind the collector ( Fig. 1A ). The size of the cloth allows for determination of the area sampled; the distance of meters dragged is multiplied by the size of the cloth in order to determine the total area dragged, which is needed to determine ticks per unit area. After dragging for 15 m, the collector stops to check the drag cloth and remove any ticks which can be killed and preserved by placing into 70–95% ethanol, or kept alive by placing into small chambers that allow for humidity and air exchange. To drag sample a predetermined area of at least 750 m 2 , a grid or linear transects can be established. Drag sampling along the same grid or transect can be done repeatedly for longitudinal studies to define host-seeking phenology or compare year-to-year density, for example.

Demonstration of tick dragging in the field. The series of panels show (A) walking to the pace of the ‘wedding march’ (approximately 50 m/min, a 30-min mile, or 1.8 miles/h) with the drag cloth trailing behind the collector, (B) pausing after 15 m to remove any collected ticks, and (C) a size comparison of Ixodes pacificus at each life stage to demonstrate the differences in size of the three tick life stages. Photographs by Andrea Swei and Sam McDuff.

Building a 1-m 2 Drag Cloth

Materials

Heavy cotton flannel fabric that is white or another light color to facilitate visibility of ticks. The dimensions should be a minimum of 1.25 yards by 1.25 yards (114 cm by 114 cm). Wash the material with fragrance-free soap before cutting and sewing, to allow for material shrinkage.

Wooden dowel 1″ diameter (2.4 cm). In the United States, dowels are typically sold in 45″ lengths (1.143 m).

Nylon rope with a thickness of 0.5″ (1.27 cm) for attachment to the dowel. The length at which the rope is cut depends on the height of the individual, but generally we recommend about 3 m of rope per drag cloth, in order to fasten the rope to the dowel and provide length for the entire cloth to comfortably drag on the ground, about a half a meter behind the tick collector.

An electric drill with a 9/16″ (14.5 mm) drill bit to accommodate for the width of the rope used. The hole for the rope can also be screwed by hand.

Metal fishing weights (‘sinkers’) or other small weights to add approximately 8–12 ounces of total weight to the bottom of the drag cloth. Another option alternative to the individual weights is to insert a 90-cm metal chain. The chain method over the free weights may prevent the weights from moving inside the hem.

Instructions

1) Orient the fabric with a top, bottom, front, and back ( Fig. 2 ). Measure and cut the cloth to a width of 102.4 cm and length of 114 cm.

Diagram illustrating how to construct a drag cloth including (A) the final drag cloth design, (B) hemming the sides by 1.2 cm, (C) building the enlarged hems for the dowel slot and the weights pocket, and (D) attaching the rope to the dowel. The front is the ‘clean’ side with no hem fringe and comes in contact with the leaf litter or vegetation ( Fig. 1A ). The back is the side that will face upwards when being dragged. By maintaining the distinction of each side of the drag cloth, the hems will remain intact longer and will reduce the risk of overlooking larvae lost inside the fringes of the hem.

2) Create the hem by folding in 1.2 cm from the sides and pin down with sewing pins ( Fig. 2B ). The fringe of the hem is the back side. Once sewing is complete, remove the pins.

3) To create the hemmed top loop and the bottom pocket, for both dowel and weights, measure and fold in 7 cm from the top and bottom edge of the drag cloth and pin. The fringe of the top and bottom hem should also be on the back side. The hemmed top loop and bottom pocket will both be 7 cm, resulting in a finished area of 1 m by 1 m.

4) Insert the chosen material of weight at the bottom pocket and sew the edges closed. For ease of laundering, the bottom pocket can be fastened with buttons or Velcro, rather than sewn shut, to allow for the weights to be removed before cleaning. To better secure the free individual weights, a few vertical lines can be sewn in the hem to prevent the free weights from moving around.

5) Insert the dowel in the top hem loop. The ends of the dowel should be slightly longer than the drag cloth on both sides to attach the rope. By leaving the top hem loop open, this allows for easy removal of the cloth from the dowel before washing.

6) Drill a hole through the dowel 3.8 cm from each end. The hole should be big enough to accommodate the rope.

7) Cut the rope to a length of 3 m and fasten it through the holes of the dowel ( Fig. 2A ). This rope length can be adjusted according to the height of the collector to maintain a 45° angle between the drag cloth and the surface being dragged by tying a knot to reduce the length. Tie the rope in a sturdy knot that can be untied in order to easily remove the cloth for cleaning ( Fig. 2D ).

Considerations for Drag Cloth Material

Corduroy is also commonly used in drag cloths. The fabric wales (ridges) increase surface area such that the sampling cloth size may be greater than 1 m 2 which may be a consideration in density estimates. Additionally, depending on wale size, larvae may be overlooked as they are quite small and can hide in the crevices of the corduroy ( Fig. 1C ). Fabrics such as muslin have had better success in collecting A. americanum (Newman et al. 2019), but this material may need to be replaced more often due to its relative thinness. Denim may also be considered, but make sure that it is light enough in order to see the ticks (Eisen et al. 2019). In congruency with the CDC, we recommend using a thick white flannel material, as it is easy to wash, durable, less expensive, and easier to find at a local fabric store. When soiled, the drag cloth can be laundered (with dowel and weights removed) with non-fragrance laundry soap in a regular washing machine system.