Ticks: Tick surveillance
Ticks
Tick surveillance
Authors: Prof Maxime Madder, Prof Ivan Horak, Dr Hein Stoltsz
Licensed under a Creative Commons Attribution license.
TABLE OF CONTENTS
Introduction ....................................................................................................................3
Sampling methods..................................................................................................................3
Detection and identification ....................................................................................................4
Surveillance and analyses ......................................................................................................4
Collection of ticks ..........................................................................................................5
Collection of free-living ticks ...................................................................................................5
Free-living immature ticks: drag sampling (Spickett et al. 1991) .................................................. 5
Free-living adult ticks..................................................................................................................... 6
Drag-sampling ................................................................................................................................................ 6
Vegetation sampling ....................................................................................................................................... 7
Nest/burrow sampling ..................................................................................................................................... 7
Carbon dioxide traps ...................................................................................................................................... 7
Trapping Amblyomma hebraeum adult ticks .................................................................................................. 7
Attraction-Aggregation-Attachment Pheromone (AAAP) trap) (Norval et al. 1989) ........................................ 7
Collection from live hosts ........................................................................................................8
Tick collection sites (Baker & Ducasse 1967) ............................................................................... 9
Tick collection (Londt et al. 1979) ............................................................................................... 10
Tick collection (Matthee et al. 1997) ........................................................................................... 11
Tick collection (Howell et al. 1989) .............................................................................................. 12
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Tick collection (standard females) ............................................................................................... 12
Collection from dead hosts ...................................................................................................13
Tick collection (large mammals) (Horak et al. 1992) ................................................................... 13
Tick collection (large mammals) (Van Dyk & McKenzie 1992) ................................................... 13
Tick collection (small mammals) (Horak et al. 1986) .................................................................. 14
Tick collection (birds) (Horak & Williams 1986) .......................................................................... 14
References ...................................................................................................................15
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INTRODUCTION
The surveillance of vectors and vector-borne diseases is essential for their control. As the livestock
populations grow and with increased trade worldwide, there is the increasing likelihood of vector-borne
disease outbreaks and many are expanding their range into new areas. After the recent import of Brazilian
cattle in Benin and Ivory Coast early in the 21st century, the cattle tick Rhipicephalus (Boophilus) microplus
was found to have been introduced on the animals (Madder et al., 2007, 2011 and 2012). An adequate
surveillance system could have identified the introduction in time allowing successful eradication in an early
stage.
Vector surveillance can be defined as the monitoring of arthropod populations responsible for the
transmission of pathogens. Vector surveillance can be used to:

Better understand vector ecology, for example:
o
Vector population distribution or density
o
Vector species diversity
o
Seasonal variation and population dynamics. This could be important to understand the
transmission dynamics of a pathogen and the resulting epidemiological situation of the
disease.


Detect the presence/absence of a vector population, for example:
o
Detection of an “exotic” vector species in a region not known to be colonized
o
Evaluation of vector control programmes
o
Surveillance of the presence of insecticide resistance genes in a vector population;
Assess the risk of vector-borne pathogen transmission, for example:
o
An early-alert system based on routine pathogen detection in vector populations
o
The evaluation of vector abundance. This information can be used in pathogen transmission
models to estimate the abundance threshold (ratio between vector and host numbers) above
which an epidemic may occur.
Sampling methods
Routine sampling of vector populations is critical in order to understand the estimated levels of both infected
and uninfected arthropods. Arthropods are typically collected, sent to an appropriate laboratory alive, or
preserved in ethanol (70%), and assayed for identification and infection. The methods of collection (e.g.
dragging the ground with a flannel cloth) will vary with the vector as well as the handling and packaging
methods and according to the pathogen and vector involved. For surveillance purposes, arthropods are
trapped, identified, sorted by sex, age, physiological type etc., counted and stored for later assays (Armed
Forces Pest Management Board, 1998).
Arthropod sampling data in surveillance involves an estimation of vector density. Vector density in a region is
important to understand because high vector densities have been shown to be associated with (high risk)
outbreaks of certain vector-borne diseases. Many sampling tools are available and the choice of a particular
tool depends on the species and the surveillance question. Because many hard ticks “quest for a host” on
vegetation, they are collected by dragging a large square piece of cloth over the ground. If the vegetation is
too thick, then a square cloth can be made into a flag that can be waved across the vegetation. In addition,
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for ticks that “hunt” for their hosts, CO2 traps are placed on the ground to attract ticks to a “sticky” surface
where they can be collected or they are trapped within a receptacle. Ticks can also be collected from hosts,
but this method may select for tick species that remain on the host for long periods of time or certain stages
such as adult males, and is further complicated by the movement of the host; all of these factors make
sampling design key in determining relative abundance in an area.
Detection and identification
In a vector surveillance program it is essential to collect vectors systematically in time and space and to
determine the species either morphologically or molecularly. In addition, vector surveillance programmes
should include systematic detection and identification of pathogens from a sample of vectors to monitor the
introduction of pathogens transmitted by local tick vectors or newly introduced tick vectors. If the objective is
to isolate the pathogen for identification, then ticks should be collected alive and stored properly for testing.
Alcohol (70%) is used routinely for this purpose. Accurate tick identification is very important especially
because most tick species transmit specific pathogens. After collecting, sorting, identification, labelling, and
placement in a suitable container, the ticks are delivered to an appropriate reference laboratory where they
can be assayed for a pathogen.
Surveillance and analyses
The likelihood of the spread of vector-borne diseases with climatic changes and globalization will lead to
greater use of vector surveillance systems. These systems will benefit from improvements in diagnosis,
knowledge of vector-borne disease ecological systems and reporting (Hitchcock et al., 2007). With the
development and improvement in geological information systems (GIS) that display and analyze
epidemiological data, we have seen an improvement in accuracy, usefulness and timeliness of information
being processed. We can track seasonal and year-to-year trends in animal disease incidence, and by
overlaying climate, vegetation, and other factors, make valuable predictions about potential outbreaks of
vector-borne diseases. There are a variety of satellite derived environmental variables such as temperature,
humidity, and land cover type with vector density that are used to identify and characterize vector habitats.
Remote sensing techniques have been used to map several vector-borne diseases due to mosquitoes, ticks,
black flies, tsetse flies, and sand flies (Kalluri et al., 2007).
Climate changes influence the epidemiology of vector-borne disease and these changes can influence both
vector and pathogen distributions, how pathogens are transmitted, and interactions between vectors and
hosts (Tabachnick, 2009). The challenges today are the development of vector surveillance systems that
continue to collect epidemiological information on vectors, storage of that data, processing of the data, and
analyses that will allow for monitoring of current changes in vector populations and prediction of future
populations changes with our changing global environment. Geographical and seasonal distributions of
vectors are influenced by climatic and land-use changes and thus climate-related environmental factors can
be used as predictive indicators in association with on-going vector surveillance activities. Satellite
measurements and remote sensing techniques cannot identify the vectors themselves, but they can identify
and characterize suitable vector habitats. Remote sensing techniques can aid in the development of
distribution maps and disease risk on a seasonal basis and monitor changes in distributions and disease risk
over time. Maps showing seasonal risks of vector-borne diseases will be critical in monitoring the impacts of
global climate changes on vectors. Remote sensing can be used to determine the influence of environmental
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factors on the spread of vectors or possible increases in distributional boundaries. Remote sensing and other
geospatial technologies are integral to any vector surveillance programme and remains an important tool in
predictive veterinary epidemiology (Martin et al., 2007).
COLLECTION OF TICKS
As mentioned above, ticks can be collected for surveillance purposes. They can also be collected to
determine their geographic distribution. In this case various host species (e.g. cattle, goats and dogs) and
the vegetation can be sampled once at a number of localities within a district or province. To determine the
seasonal abundance of ticks a single or more host species and the vegetation at a fixed locality are sampled
at regular intervals (e.g. monthly).

Certain procedures or requirements may have to be followed or met before the collection of ticks can
commence.

The property/farm owner’s consent must be obtained whether collecting ticks from the vegetation or
live or dead host animals.

If collections are going to be made in national or provincial wildlife reserves it will be necessary to
obtain the required permits from the administrating authorities.

The management of wildlife reserves will usually require a full protocol before collections can start.

If live animals are to be examined an animal ethics clearance as to the wellbeing of the animals
during sampling may be required.

If ticks are to be taken out of a wildlife reserve to a laboratory the necessary permits might have to
be obtained.
Collection of free-living ticks
Free-living immature ticks: drag sampling (Spickett et al. 1991)
Ten 1 000 mm x 100 mm flannel strips are attached adjacent to one another by means of Velcro
tape on a 1 200 mm-long wooden spar. Each collection is made by an operator pulling this spar, with
the flannel strips attached, by means of a string or twine harness attached to its ends, for a distance
of 250 m over the vegetation. At the end of each drag the flannel strips are individually detached
from the spar and the ticks are removed with fine-point forceps and placed in vials containing 70 %
ethyl alcohol. Each cleaned flannel strip is then re-attached to the wooden spar.
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Video: Drag sampling (http://youtu.be/5eLT1C4s4Ks)
Ideally three drags are performed in each of grassland, woodland and gully sub-habitats within larger
habitats. Drags are not done over dew-laden grass early in the morning or over grass after rain, as
this wets the flannel strips and decreases their efficacy. All ticks collected are identified and counted
under a stereoscopic microscope.
Free-living adult ticks
Drag-sampling
Many adult ixodid ticks can be collected whilst questing for hosts from the vegetation. This is can be
done by the dragging-sampling technique already described.
Video of a questing adult Rhipicephalus appendiculatus tick
(http://youtu.be/gGa6AkriVFw)
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Vegetation sampling
Adult ticks can be collected by hand from the tips and stems of grass within a specified measured
area or alongside a measured length of road or a path. This and the previous method are only
applied to “exophilic” ticks which quest for a host from the vegetation.
Nest/burrow sampling
This involves sampling the nest or the burrow of a host. A number of quite elaborate vacuum
systems have been described, which collect and separate ticks from their host's burrows (vacuum
extraction). Multiple sampling of these burrows over a 3 to 4 day period may extract large numbers
of ticks. This type of capture technique may be used in conjunction with CO2. It is particularly useful
for sampling ticks that are difficult to reach, such as Ornithodoros in warthog burrows.
Carbon dioxide traps
Carbon dioxide traps, which simulate the production of CO 2 by the host, have been developed to
attract both ixodid and argasid ticks. The source of CO2 can vary from very sophisticated CO2
producing traps (chemical traps), to simple, easily available sources such as animals, dry ice or
carbon dioxide cylinders.
Trapping Amblyomma hebraeum adult ticks
Adult A. hebraeum do not quest from the vegetation and are seldom seen as they are usually hidden
beneath the leaf litter under bushes. Only when an animal passes or there is a strong source of CO 2
nearby will these ticks be enticed to leave their microhabitat in search of a host. Once on the soil
surface the ticks are non-directionally active and can be caught moving around on the ground.
To elicit a directional response from adult A. hebraeum a volatile pheromone mixture obtained from
feeding male ticks needs to be added to the CO2. The adult ticks (males and females) as well as
nymphs can detect the pheromone source and they "home in" on it from several metres away.
Attraction-Aggregation-Attachment Pheromone (AAAP) trap) (Norval et al. 1989)
Locating suitable sites: This trap can be used to locate the presence of adult A. hebraeum in a
particular site or area. Before this can be done, however, areas ecologically suitable for the survival
of these ticks (shaded area/thick leaf litter) need to be identified.
CO2 source: The ticks are attracted from the leaf litter by means of a 0,5 kg block of dry ice placed
centrally. The leaf litter around the dry ice should be removed so that the ticks, which have been
attracted, are more visible to the collector.
Pheromone source: Feeding adult male ticks are removed from an un-dipped cow and used as the
source of pheromone. About 100 ticks are removed and immersed in a brown glass bottle containing
± 100 mℓ of diethyl ether. These ticks are left in the bottle overnight so that the pheromone can be
absorbed into the ether. The ether is then decanted and is stored in a brown glass bottle in a cool,
dark place.
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To use the pheromone place a few drops of the liquid on a filter paper and allow the ether to
evaporate (few minutes). The impregnated filter paper is then attached to a metal tent peg, which is
knocked into the ground near the CO2 source.
At sites in which adults of A. hebraeum are present their presence will soon be noticed because of
their activity in the vicinity of the trap. The adult ticks are large and ornate and move rapidly and
hence are reasonably easy to see and collect. Use forceps to collect the ticks and place them in
individual containers before transporting them to the laboratory for further identification and
processing.
Collection from live hosts
To determine the attachment sites of ticks, on host collection is by far the most ideal method. Several
authors have described predilection sites of tick attachment on cattle. Normally, each animal requires several
hours for complete de-ticking but this largely depends on the number of parasitizing ticks. A pair of forceps is
used to remove ticks: the ticks are stored in 70% alcohol until identification.
Collection of ticks from live cattle.
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Tick collection sites (Baker & Ducasse 1967)
To determine the exact predilection sites of cattle ticks, the animal's body can be divided into 18 areas, not
necessarily corresponding to the recognised anatomical regions: (1) Muzzle; (2) Peri-orbital zones; (3) Head
(delimited by a vertical line drawn from the base of the ears ventralward over the throat latch but excluding
(1&2); (4) Pinnae (both surfaces); (5) Ear passages; (6) Poll (including mane and upper neck border to withers);
(7) Neck (lateral surfaces); (8) Dewlap; (9) Axilla (delimited by a line joining the points of the two shoulders
cranially and by one running from one olecranon to the other caudally); (10) Sternum (caudal sternal and
xiphoid regions up to the umbilicus); (11) Belly and groin (post-umbilical and inguinal regions including
udder/scrotum); (12) Lower perineum (ventral to vulva in the female or anus in the male to base of
udder/scrotum); (13) Upper perineum (from base of tail, around anus, including vulva in the female); (14) Tail;
(15) Tail brush; (16) Feet (below fetlocks); (17) Legs (from fetlocks to elbows/stifles); (18) Rest of body (lateral
thoracic, abdominal, gluteal and femoral regions).
Each of the sites is handled separately for the purpose of tick collection. The ear passage is
carefully de-ticked by means of a fine spoon-curette. In the other sites, tick removal is accomplished
by means of forceps for large ticks and a fine nit-comb for smaller ticks and the immature stages.
Each site is diligently combed, all hair, debris and ticks being collected into a specially adapted
plastic funnel and thence transferred into separate and permanently marked plastic bottles. Each
animal requires some 3,5 hours for complete de-ticking. The field collection bottles are then packed
in specially constructed boxes and dispatched to the laboratory for further attention.
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Considerable difficulty may be experienced in sorting the ticks owing to the large amounts of hair,
debris and wax contained in the collection bottles. By placing each collection into a specially
constructed stainless steel sieve with 150 micron apertures, and immersing the contents in boiling
10 % NaOH solution for varying lengths of time, depending upon the amount of extraneous matter
present, the hair can be digested, and sorting and examination can be greatly facilitated. Great care
must be exercised to ensure that the ticks are not over-boiled, as this causes them to burst and
makes the task of identification very difficult. After boiling, the sieve contents are collected and
preserved for later examination.
Tick collection (Londt et al. 1979)
Total tick counts of the sort undertaken by Baker & Ducasse (1967) are not always possible for
practical reasons. Instead, six clearly defined sites on each host animal can be selected for study
because of their importance as feeding sites for the different stages of the commoner cattle ticks. All
the ticks on these sites are removed, either by hand or with forceps, and taken to the laboratory for
study. The ear pinnae and lower border of the dewlap are scraped with a sharp knife for the removal
of the immature stages of R. appendiculatus, but this sampling method is not used on the other
predilection sites.
The sites are as follows:
Pinna: (Site 4 of Baker & Ducasse 1967). Both surfaces of a single ear of each bovine are sampled.
This site is important for all stages of R. appendiculatus and immature R. (B) decoloratus. The actual
ear passage can also be included but care must be taken with collections not to damage the ear
canal but also not to damage the ticks. This site is important for the immature stages of R. evertsi
evertsi.
Neck: (Sites 7, 8 and part of 6 of Baker & Ducasse 1967). This site includes the lateral surfaces of
the neck, the dewlap and the mane. Only one side of each bovine is sampled. This is an important
site for all stages of R. (B) decoloratus and for R. appendiculatus larvae.
Leg: (Sites 9, 16 and 17 of Baker & Ducasse 1967). This site includes the axilla, leg (from elbow to
fetlock) and foot (below fetlock). Only one foreleg of each survey animal is sampled. This site is
important for the feeding of all stages of R (B) decoloratus, nymphs and larvae of A. hebraeum and
larvae of R. appendiculatus and adult Hyalomma spp.
Tail: (Sites 14 and 15 of Baker & Ducasse 1967). This site includes the tail and tail brush, and is
important for the feeding of R. simus, H. truncatum and A. hebraeum adults.
Upper perineum: (Site 13 of Baker & Ducasse 1967). This site, extending from the base of the tail
to about 10 cm below the anus, is very important for the feeding of R. evertsi evertsi, A. hebraeum
and H. marginatum rufipes adults.
Lower perineum: (Site 12 of Baker & Ducasse, 1967). This site, extending from below the upper
perineum to the base of the scrotum, is an important feeding site for A. hebraeum adults.
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Tick collection (Matthee et al. 1997)
Sketch outline of the individual sites of collection on the impala. (1) muzzle, (2) head, (3) pinna, (4) neck, (5)
fromt leg, (6) hind leg, (7) sternum, (8) abdomen, (9) tail and (10) rest of the body. Matthee et al. (1997)
Ten defined sites of tick attachment from the predilection sites described by Baker & Ducasse (1967)
and Londt et al. (1979) are chosen, namely: (1) Muzzle. (2) Head. Excluding the muzzle.
Delimited by drawing a line from the base of the pinna downward over the throat-latch. (3) Pinna:
both surfaces. (4) Neck. (5) Foreleg. (6) Hindleg. (7) Sternum. (8) Abdomen. (9) Tail: half the
tail, the tail brush, upper and lower perimeum. (10) Body.
Each of the sites is treated separately for the purpose of ectoparasite collection. The ten sites are
delineated with a black permanent marker pen on the one half of the skin. The area of skin to be
shaved within each predilection site is delineated using one of three plastic templates of different
sizes: 50 mm x 50 mm; 50 mm x 75 mm; and 75 mm x 120 mm. The size of the template used is
determined by the area of the predilection site, for example the small 50 mm x 50 mm template is
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used on the head. The templates are placed on the skin in the predilection sites and the outline
marked with a marker pen.
The placement of the templates on six of the sites is determined in the following way:
Head: A 50 mm x 50 mm template is placed halfway between the eye and the lower jaw.
Neck: A 50 mm x 75 mm template is placed halfway between the lower edge of the jaw and the
shoulder.
Legs: A 50 mm x 75 mm template is placed at the elbow of the front leg and the stifle of the hind leg,
and the entire surface of the front and hind fetlocks, is shaved.
Sternum: A 50 mm x 75 mm template is placed in the centre of the site.
Abdomen: A 50 mm x 75 mm template is placed in the centre of the site.
Body: A 75 mm x 120 mm template is placed on the body at the shoulder and another at the hip.
The hair inside the marked area is closely shaved and the surfaces of the pinna, fetlocks and feet
and the tail are completely shaved with a Minora razor. The hair and parasites collected from the
respective sites are placed separately in pre-marked bottles containing 10 % formalin. The formalin
is decanted and the material that has been collected in the bottles is immersed in 10 % NaOH
solution until the hair dissolves. Thereafter the solution is sieved through a sieve with 150 m
apertures and the parasites retained on the sieve are collected and preserved.
Tick collection (Howell et al. 1989)
This method is particularly applicable to small rodent hosts, but could also be used on other small
mammals. After the animals have been trapped they are transferred to holding cages over water.
Ticks detaching from the animals are collected from the water each morning and evening. This can
be done by sieving or by decanting the water. Any adult ticks that may have dropped from the
animals are immediately placed in 70% ethyl alcohol for later identification and counting. The
engorged immature ticks that have dropped are gently dried and placed in glass vials in an
acaridarium to allow them to moult to the next developmental stage as this facilitates identification
and counting. Any ticks that die during this process are also identified and counted. Two days after
no further ticks have dropped from the caged animals the animals can be released at the sites at
which they were captured.
Instead of water the caged animals can be kept over moist paper towelling. In this case the edges of
the container in which the moist paper is placed must be treated or fitted with non-escape material to
prevent the ticks that have dropped from escaping.
Tick collection (standard females)
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Standard female ticks are those engorging female ticks of various species of which the length of the
idiosoma has reached a specific measurement that indicates that the tick is likely to detach and drop
from the host animal within the next 24 h. These measurements are ± 9,5 mm for the Amblyomma
spp., 4,0 mm to 4,5 mm for the Boophilus spp., ± 7,5 mm for the Hyalomma spp., and 5,0 mm to 6,0
mm for the Rhipicephalus spp. These female ticks, which are usually easily visible, can be collected
from one whole side of the host animal, or patch-sampling of the various predilection attachment
sites of the different tick species can be done. The female ticks collected in this way can then be
preserved in 70 % ethyl alcohol, identified and counted at the laboratory.
For small rodent hosts or other small mammals, ticks can be recovered by placing the hosts in cages
over water after they have been caught in live traps.
Live trapping of rodents
Ticks collected from hosts are normally not used to determine vector competence for pathogens as
false positives may arise resulting from infected blood ingested by the tick.
Collection from dead hosts
Tick collection (large mammals) (Horak et al. 1992)
After the animals have been killed they are transported to the laboratory. There the carcass of each
animal is skinned and half the skins of the head, half the skin of the body and upper legs, the whole
skin of the tail as well as one lower front leg and one lower back leg with skin attached are placed
separately in plastic bags. A tick-detaching agent is added to the skins in the bags that are tightly
secured and stored overnight. The following morning the skins are thoroughly scrubbed with brushes
with steel bristles and washed. The tick-detaching agent remaining in the plastic bags and the
material obtained from scrubbing and washing the skins are sieved on sieves with 150 m apertures.
The residues in the sieves are collected, preserved in 10 % formalin and stored.
Tick collection (large mammals) (Van Dyk & McKenzie 1992)
In this method the skin is subjected to NaOH digestion, and the operator must wear protective
clothing. The respective portions of skin of the animal are immersed in separate baths of 10 %
NaOH for 3 to 7 days until the hair is sufficiently loosened from the skin. he skin portions are then
scraped to remove the hair and also the sludge that has accumulated on the skin. Thereafter the
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skin portions are discarded. The sludge containing the ticks is left in the NaOH solution until all the
hair is dissolved, thereafter the solution is sieved and the residue on the sieve is collected and
preserved with 10 % formalin. Great care must be taken to monitor the digestion process in case the
ticks themselves are digested beyond identification.
Tick collection (small mammals) (Horak et al. 1986)
Ectoparasites are recovered from these animals by placing the whole small animal, immediately
after it has been killed, in a sturdy plastic bag and transporting it to the laboratory. There the animal
can be eviscerated if necessary. Thereafter the whole animal, with its skin intact, is returned to the
plastic bag and sufficient tick-detaching agent added to immerse the animal. It is left in the bag until
the following morning, and then thoroughly scrubbed with a brush with 20 mm long steel bristles and
washed, particular attention being paid to the external ear canals, neck and the feet. The scrubbings
and washings plus the contents of the plastic bag are poured on to a sieve with 150 m apertures
and sieved. The material that is retained on the sieve is collected, preserved with formalin and
stored.
Really small mammals such as rats and mice can be treated in the same way and thereafter the
whole animal can also be examined under the stereoscopic microscope.
Tick collection (birds) (Horak & Williams 1986)
Once the guineafowl or birds of similar size have been shot the whole carcass is immediately placed
in a sturdy plastic bag that is securely closed and transported to the laboratory. At the laboratory the
bird is decapitated just caudal to the level where the bare neck joins the feathered neck and the
feathered portion of the carcass is skinned. The wing-tips are not skinned, but the bone is severed
and the whole wing-tip included with the skin. The head, the skin (including wing-tips) and the legs
are returned to the plastic bag and sufficient diluted tick detaching agent is added to cover all this
collected material. The following morning the bag is opened and its contents are poured out and
thoroughly washed with a strong jet of water over a sieve with 150 m apertures. The fine material
that is retained by the sieve is collected for microscopic examination, and the head, skin, wings and
legs, which have been washed, are also retained for examination.
The whole head and neck and the whole back and tail, one wing, one side and one leg and all the
material that has been retained by the sieve are examined separately under a stereoscopic
microscope. Ticks present on or in this material are collected, counted and identified. The number of
ticks recovered from the wing, the side and the leg are doubled and added to the numbers recovered
from the remainder of the material in order to calculate the total tick burden of each bird.
In the case of small birds the whole bird can be examined under the stereoscopic microscope.
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at 5 different ecological sites in heartwater endemic regions of South Africa. Experimental and
Applied Acarology, 24: 971-982.
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