Tick paralysis in Australia caused by Ixodes holocyclus Neumann

Abstract

Ticks are obligate haematophagous ectoparasites of various animals, including humans, and are abundant in temperate and tropical zones around the world. They are the most important vectors for the pathogens causing disease in livestock and second only to mosquitoes as vectors of pathogens causing human disease. Ticks are formidable arachnids, capable of not only transmitting the pathogens involved in some infectious diseases but also of inducing allergies and causing toxicoses and paralysis, with possible fatal outcomes for the host. This review focuses on tick paralysis, the role of the Australian paralysis tick Ixodes holocyclus, and the role of toxin molecules from this species in causing paralysis in the host.

Many forms of tick toxicosis affect humans and other animals (Gothe and Neitz, 1991; Mans et al., 2004). According to Gothe (1984), a tick toxicosis is defined as a ‘generalized, experimentally standardizable, reproducible disease syndrome induced by one or a few potent ticks, even on first infestation of a physiological susceptible vertebrate species without participation of an immunopathological reaction during or following the tick feeding’. Such toxicoses are mostly caused by ixodid or hard ticks (Stone and Wright, 1981) and, in their most severe form, result in paralysis of the infested host. About 69 species of ticks from around the world are capable of inducing paralysis (Gothe and Neitz, 1991), the most important being Ixodes holocyclus in Australia, Dermacentor andersoni, De. variabilis and Argas (Persicargas) radiatus in North America, Ix. rubicundus in South Africa, Rhipicephalus evertsi evertsi and Ar. (Pers.) walkerae in Ethiopia, and Ar. (Pers.) radiatus in the Nearctic region of North America (Stone, 1986). Some important paralysing ticks and their distributions are summarized in Table 1. In Australia, Ix. holocyclus can cause paralysis in humans, dogs, cats, sheep, cattle, goats, pigs and horses but predominantly infests dogs, cats and humans (Stone, 1986). It appears to be the most potently toxic tick species, with a single tick capable of killing a large dog (Stone and Wright, 1981) or sheep (Sloan, 1968). Although the Tasmanian paralysis tick Ix. cornuatus has been reported to cause bulbar paresis and respiratory failure in humans (Tibballs and Cooper, 1986) and dogs (Beveridge et al., 2004), its habitat is more restricted than that of Ix. holocyclus and very few cases of paralysis have been associated with this tick. In Australia, tick paralysis has mainly been seen as a problem in veterinary medicine, affecting approximately 10,000 companion animals/year (Stone and Aylward, 1987). Occasionally, however, the climatic conditions become particularly favorable to tick survival, tick densities reach very high levels, and the number of humans being bitten by Ix. holocyclus increases. Although rarely severe, tick paralysis caused by Ix. holocyclus can be fatal.

Table 1. Distribution of the main tick species that cause paralysis (Gregson, 1973; Murnaghan and O’Rourke, 1978; Gothe et al., 1979).

Tick species	Country/region	Host(s)
ixodid
Amblyomma maculatum	Uruguay	Dogs
	U.S.A.	Humans
Dermacentor andersoni	Northern U.S.A. and Canada	Livestock, humans
Dermacentor auratus	India	Domestic animals
Dermacentor occidentalis	U.S.A.	Cattle, horses, deer
Dermacentor sp.	Italy	Cattle, goats, sheep
Dermacentor variabilis	Eastern U.S.A.	Dogs, humans
Haemaphysalis cinnabarina	U.S.A.	Humans (one fatal case)
Haemaphysalis kutchensis	India	Rabbits
Haemaphysalis punctata	Crete	Sheep, dogs, cats
	Macedonia, Bulgaria	Sheep, goats, calves, chickens
Hyalomma truncatum*	South Africa	Humans (rarely), sheep, goats, pigs, dogs
Ixodes brunneus	U.S.A.	Birds
Ixodes crenulatus	Ukraine	Sheep
Ixodes hexagonous	England and France	Humans
Ixodes holocyclus	Australia	Cats, pigs, cattle, horses, sheep, goats, humans
Ixodes ricinus	Crete	Livestock, dogs, cats
	Former Yugoslavia	Cattle, sheep, goats
	Turkey	Sheep, goats, calves
	Azerbaijan	Foals
Ixodes rubicundus	Africa	Humans (rarely), sheep, dogs, cattle, jackals
Ixodes scapularis	U.S.A.	Dogs
Rhipicentor nuttalli	South Africa	Dogs
Rhipicephalus evertsi†	Africa	Lambs, calves, dogs
Rhipicephalus sanguineus	Venezuela	Dogs
Rhipicephalus simus	Africa	Humans (rarely)
argasid
Argas persicus	Africa	Chickens
Argas walkerae	Africa	Chickens
Ornithodoros lahorensis	Kazakhstan	Sheep
Ornithodoros savignyi	Nigeria	Cattle
Otobius megnini	Africa	Cattle

^*Cause of ‘sweating sickness’.

^†Cause of ‘spring lamb paralysis’.

Ixodes holocyclus is found along the eastern coastal strip of Australia and is most abundant from September to about March. Paralysis is induced by a toxin that is transmitted to the host in the saliva of a female Ix. holocyclus when the tick takes a bloodmeal. During feeding, toxicity in the salivary glands increases, peaking after 4–5 days of engorgement (Goodrich and Murray, 1978). The toxin produced by Ix. holocyclus inhibits acetylcholine release at the neuromuscular junction and tends to cause more severe neurological impairment than that associated with De. andersoni or De. variabilis in North America (Grattan-Smith et al., 1997).

Historically, the Australian explorers Hamilton Hume and William Hovell described ticks biting humans as early as 1824, during their journey between Lake George (Canberra) and Port Phillip (Victoria) (Gregson, 1973). Bancroft (1884) was the first to report on tick toxicosis in humans in Australia, describing two cases with weakness and blurred vision. A human fatality caused by the bite of an (apparently uninfected) tick was first described by Cleland (1912). Between 1914 and 1942, 20 human fatalities in Australia were attributed to tick bite, all but three of the victims being children (Murray and Koch, 1969).

CLINICAL PRESENTATION

Although tick paralysis in humans is most commonly reported in children aged 1–5 years, the problem does also occur in older children and adults (Grattan-Smith et al., 1997; Inokuma et al., 2003). The clinical presentation usually involves a combination of lethargy, weakness and unsteadiness of gait, often with ataxia, loss of appetite, dilated pupils, ascending symmetrical paralysis, slurred speech and depressed deep-tendon and gag reflexes (Grattan-Smith et al., 1997; Edlow and McGillicuddy, 2008). Laboured breathing, bradycardia, decreased oxygen saturation and asystole have also been reported (Grattan-Smith et al., 1997). In addition to neuromuscular paralysis, the toxin from Ix. holocyclus can produce myocarditis in children (Pearn, 1966). In older children or adults, double or blurred vision and photophobia may develop (Grattan-Smith et al., 1997; Edlow and McGillicuddy, 2008). Facial weakness or paralysis may be present and ascending flaccid paralysis can also occur if the tick is not removed quickly (Pearn, 1977). It is also worth noting that the removal of Ix. holocyclus from a patient can worsen the patient’s condition. In one case described by Miller (2002), for example, facial-nerve paralysis occurred hours after the removal of Ix. holocyclus ticks from around the patient’s face. Similarly, tick removal apparently led to deterioration in four of the six cases described by Grattan-Smith et al. (1997). Stone et al. (1989) hypothesised that such delayed toxicity was caused by the disruption of the feeding lesion during the removal of the tick, which caused the release of toxin that had been bound to cells or tissues. Surprisingly, despite the continued deterioration and then slow recovery seen in many Australian patients following the removal of Ix. holocyclus, removal of De. andersoni and De. variabilis from North American patients results in recovery within a day (Garrettson, 1984).

DIFFERENTIAL DIAGNOSIS

Edlow and McGillicuddy (2008) suggested that ‘the first and most important factor in diagnosing tick paralysis is thinking of it’ and that Guillain–Barré syndrome (GBS) may be commonly confused with tick paralysis. In addition to GBS, viral encephalomyelitis, botulism, myasthenia gravis, spinal-cord compression, transverse myelitis, organophosphate poisoning, porphyria, heavy-metal poisoning and diphtheria may also need to be considered in a differential diagnosis. Edlow and McGillicuddy (2008) presented an informative guide to some clinical markers that may be of use in differential diagnosis (see Table 2).

Table 2. Factors to be considered in the differential diagnosis of tick paralysis [an adaptation of the factors described by Edlow and McGillicuddy (2008)]^*.

	Condition or disease
Sign or symptom	Tick paralysis	Guillain–Barré syndrome	Transverse myelitis	Spinal-cord compression	Botulism	Viral encephalomyelitis or poliomyelitis
Fever	Absent	Rare	Variable (depending on aetiology)	Absent (except in some cases of epidural abscess)	Absent	Present
Pain	Rare	Rare	Frequent	Frequent	Absent	Variable
Sensory abnormalities	Generally absent	Frequent prodromally	Frequent	Frequent	Absent	Not prominent
Dilated pupils	Can be present (Australia) but rare in North America	Rare	Absent	Absent	Present	Absent
Protein in cerebrospinal fluid	Normal	Elevated	Often elevated	Variable (depending on aetiology)	Normal	Elevated
Leucocytes in cerebrospinal fluid	Normal	Normal	Elevated	Variable (depending on aetiology)	Normal	Elevated
Results of MRI scan	Normal	Usually normal	Abnormal	Abnormal	Normal	Abnormal

^*This is a partial list that is not meant to be encyclopaedic.MRI, Magnetic-resonance imaging.

TREATMENT

In humans, untreated tick paralysis can be fatal. Despite the risk of a short-term worsening in the patient’s symptoms, locating and removing all ticks from the patient’s body is an important part of therapy (Edlow and McGillicuddy, 2008). Ticks are most commonly found on the scalp behind an ear but, as multiple ticks may be attached, a thorough investigation of the patient’s entire body may be warranted. Although tick anti-toxin is available, the heterologous nature of the antiserum can induce serum sickness and anaphylactic shock in humans (Stone et al., 1982). Progression of paralysis may require mechanical respiratory ventilation in an intensive-care unit (Grattan-Smith et al., 1997). The administration of antibiotics, to protect against tick-borne pathogens such as Rickettsia and Orientia, may also be justified (Inokuma et al., 2003).

NATURAL HOSTS OF Ixodes holocyclus

The most common native hosts of Ix. holocyclus are the long-nosed bandicoot (Perameles nasuta), the short-nosed or northern brown bandicoot (Isoodon macrourus) and the southern brown bandicoot (Is. obesulus) (Doube, 1975) but this tick has has been found on a wide variety of native animals (Table 3). Ixodes holocyclus also parasitises livestock (sheep, goats, horses, pigs, chickens and ducks) and companion animals (cats and dogs) and is frequently associated with humans.

Table 3. The many native hosts of Ixodes holocyclus in Australia (Roberts, 1960; Domrow and Derrick, 1965; Marks and Cribb, 1966; Doube, 1975; Jackson et al, 2000; Campbell et al., 2003).

Latin name of host	Common English name of host
Aepyprymnus rufescens	Australian marsupial bettong (rat-kangaroo)
Ailuroedus crassirostris	Green cat bird
Antechinus flavipes	Marsupial mouse
Corvus coronoides	Australian raven
Corvus orru	Australian crow
Cracticus nigrogularis	Pied butcher bird
Dasyurus maculatus	Tiger quoll
Dendrolagus lumholtzii	Tree kangaroo
Gymnorhina tibicen	Australian magpie
Isoodon obesulus	Southern brown bandicoot
Isoodon macrourus	Short-nosed bandicoot
Hydromys chrysogaster	Eastern water rat
Macropus sp.	Kangaroos
Manorina melanocephala	Noisy miner
Melomys cervinipes	Mosaic-tailed rat
Melomys lutillus littoralis	Brown rat
Perameles nasuta	Long-nosed bandicoot
Phascogale tapoatafa	Red-tailed phascogale
Phaseolarctos cinereus	Koala
Pitta sp.	Pitta bird
Platycercus elegans	Crimson rosella
Pteropus conspicillatus	Spectacled flying fox
Ptilonorhynchus violaceus	Satin bower bird
Rattus conatus	Native rat
Ra. culmorum	Pale field rat
Ra. fuscipes	Australian bush rat
Ra. rattus	Black rat
Ra. sordidus	Cane field rat
Strepera graculina	Pied currawong
Tachyglossus aculeatus	Echidna
Thylogale stigmatica	Red-legged pademelon
Trichosurus caninus	Mountain brush-tail possum
Tr. vulpecula	Coppery brush-tail possum
Uromys caudimaculatus	White-tailed native rat
Vombatus ursinus	Wombat
Wallabia dorsalis	Black-striped wallaby
W. rufogrisens	Red-necked wallaby

PRODUCTION OF Ix. holocyclus TOXIN IN THE TICK’S SALIVARY GLANDS

The toxin from Ix. holocyclus is produced in the salivary glands (SG) of feeding female ticks and transferred to the host via the tick’s saliva (Stone et al., 1983a). The SG of ticks consist of numerous alveoli, called acini, classified into four types known as I, II, III and IV. Both genders have acini of types I, II and III but only males have type IV. The acini produce and release biologically active compounds. Each acinus consists of one or two layers of cells around a lumen. Agranular cells characterise the type-I acinus whereas the cells in type-II and type-III acini are granular and are categorised, according to their histology, into cell types A–F, with cell types A–C and D–F occurring in type-II and type-III acini, respectively (Kaufman, 1983). Although Binnington and Stone (1981) interpreted the results of histological investigations as indicating that cells of type B (from the type-II acini of Ix. holocyclus) might contain the toxin responsible for tick paralysis, Stone et al. (1989) subsequently identified cells of type E (from type-III acini) as the probable source.

A tick’s saliva contains a plethora of biologically active molecules that modulate the host’s immune response to tick infestation, facilitating the formation of a feeding pool in the host’s tissue (Ribeiro, 1995; Wikel, 1996, 1999; Schoeler and Wikel, 2001). To allow successful feeding over several days, ticks need to overcome the host’s haemostatic, inflammatory and immune mechanisms (Ribeiro and Francischetti, 2003), and they produce many compounds, including anti-coagulants (such as gelatinases), fibrinolytic enzymes and anti-platelet and vasodilator substances (Kemp et al., 1983), to achieve this goal. Anti-coagulant has been reported in the saliva of Ar. persicus (Hoeppli and Feng, 1933), De. sinicus (Hoeppli and Feng, 1933), Ornithodoros moubata (Hawkins and Hellmann, 1966), Ix. ricinus (Ross, 1926) and Ix. holocyclus (Kaire, 1966). These substances generally interfere with coagulation via activated factor X (fXa) or thrombin. The anti-platelet substance apyrase has been detected in the saliva of Ix. dammini (Ribeiro et al., 1985) and Orn. moubata (Ribeiro et al., 1991) and various forms of prostaglandins are also produced in tick saliva to hinder platelet formation (Bowman et al., 1995). Prostaglandins act as vasodilators as well as having anti-inflammatory, immunosuppressive and antihaemostatic activities (Bowman et al., 1996). Usually, tick saliva contains at least one anti-clotting agent, one anti-platelet agent and a vasodilator, which together prevent coagulation (Ribeiro and Francischetti, 2003). Although the saliva of some tick species (which have hypostomes that only penetrate a small way into the host dermis) also contains cement to anchor the tick to the host, the hypostome of Ix. holocyclus reaches up to 980 μm into a host’s skin (Allen et al., 1977) and does not need the support of any such cement.

The diversity of the compounds used by haematophagous arthropods is astounding (Ribeiro, 1995) and the biological functions of many of the salivary proteins secreted by such arthropods remain unknown.

ISOLATION AND CHARACTERISATION OF THE Ix. holocyclus TOXIN

Kaire (1966) was the first to isolate the toxin from Ix. holocyclus and found it was resistant to digestion with pepsin, trypsin and papain. In a series of elegant experiments, Stone et al. (1983a) then proved that the toxin was injected into the tick’s hosts with the tick’s saliva. In these investigations, Ix. holocyclus were allowed to feed on mice for 4 days prior to their exposure to an in-vitro feeding system in which they fed on tissue-culture medium through an artificial membrane. The medium that contained saliva from the pre-fed ticks was then concentrated and injected into mice, where it produced classic tick paralysis, similar to that induced by inoculation of Ix. holocyclus salivary-gland extracts (SGE). The toxin found in Ix. holocyclus was called holocyclotoxin. Separation of Ix. holocyclus SGE by gel filtration and high-performance liquid chromatography (HPLC) subsequently yielded proteins of 50–80 kDa (Stone et al., 1979, 1983b; Stone and Aylward 1987; Stone, 1988). Holocyclotoxin was found to be a proline-rich glycoprotein that was resistant to digestion with pronase (B. F. Stone, unpubl. obs.). While it was initially thought that apparent inconsistencies seen in the molecular sizes of fractions isolated from Ix. holocyclus SGE were the result of variable resolution in the electrophoresis protocols that had been used, Malik (1991) suggested that several different toxic components were present in the SGE, including the holocyclotoxin (Stone and Wright, 1981) that causes paralysis and another toxin that was lethal but did not induce flaccid paralysis.

In an attempt to produce an immunogenic but less toxic antigen from the SGE of Ix. holocyclus, as a first step in the production of a vaccine against tick paralysis in Australia, Stone et al. (1986) treated SGE with glutaraldehyde. Although this led to a better antitoxin response, it was decided that the number of engorged ticks (and hence the volume of SGE) that could be produced was too low to meet the demands for the production of a vaccine based on glutaraldehyde-treated SGE.

Thurn et al. (1992) isolated a neurotoxin from Ix. holocyclus by binding SGE to pinched-off nerve terminals in rat-brain synaptosomes. Three polypeptides, known as holocyclotoxin 1, 2 and 3 (HT-1, HT-2 and HT-3) were isolated. Each had a molecular mass of about 5 kDa. Subsequently, HT-1 was isolated and defined by N-terminal amino-acid sequencing (Thurn, 1994). The corresponding gene, which was isolated, sequenced and characterised by Masina and Broady (1999), showed a high homology with the gene coding scorpion neurotoxin.

Antibodies directed against an immunogenic recombinant form of HT-1 have been found to be protective against holocyclotoxin (Thurn et al., 1992) and the recombinant antigen shows great promise as a vaccine candidate. HT-1, HT-2 and HT-3 seem to be part of the same gene complex (Nicholson et al., 2006).

The 68-kDa toxin in Rhipicep. evertsi SGE and the 60-kDa toxin in Ar. walkerae SGE also cause ascending flaccid paralysis but perhaps via a different mechanism to that occurring with Ix. holocyclus, since these toxins impair the conduction of impulses along peripheral nerve fibres (Gothe et al., 1979; Gothe and Laemmler, 1982; Viljoen et al., 1986, 1990; Crause et al., 1994).

TOXICITY OF TICKS

An interesting clinical phenomenon is the fact that Ix. holocyclus need several days of engorgement on their host before signs of paralysis manifest. Ross (1935) and Goodrich and Murray (1978) observed that the SG from Ix. holocyclus only produced marked toxicity, when injected into mice, if they came from ticks that had previously fed on dogs for 5 days. The SG collected from ticks that had fed for only 3 or 4 days on dogs produced much less toxicity when injected into mice (Kaire, 1966; Goodrich and Murray, 1978). Binnington and Stone (1981) subsequently confirmed that the toxicity of homogenates of Ix. holocyclus SG depended on the duration of tick feeding. Thus, although SG collected 1 or 2 days after the onset of feeding showed very little toxicity, the toxicity of SG increased with progressive feeding, peaking 4–5 days post-infestation and then decreasing until complete repletion. In both Ix. holocyclus (Binnington and Stone, 1981) and Rhipicep. evertsi (Neitz and Gothe, 1986), the timing of the peak production of toxin in the SG was found to coincide with the rapid engorgement phase of feeding (on days 4–5 post-infestation). The paralysis caused by Ix. rubicundus or De. andersoni is easily reversed, at least in its early stages, by removing the attached tick(s) (Stone, 1988). Paralysis in chickens induced by larvae of the genus Argas disappears when the replete larvae drop off the host (Mans et al., 2004). Paralysis caused by Ix. holocyclus, however, cannot be easily reversed and affected animals may still die despite removal of the ticks.

Potency levels within one tick species, such as De. andersoni (Gregson, 1973), can vary considerably; it is suspected that De. andersoni has geographical variants with different levels of toxicity. Although Ix. holocyclus ticks generally appear to be consistently toxic, with one tick potent enough to kill a dog, there is also some evidence of variation in toxin potency in this species (Alexander, 1986; Curtin, 1986; Jones, 1986; Pursell, 1986; Atwell and Fitzgerald, 1994).

Toxicity of the Immature Stages of Ticks

Riek (1957) homogenized the eggs from each of 22 species of tick [17 ixodids (including Ix. holocyclus) and five argasids] and injected the homogenates into laboratory animals to test for toxic reactions. Toxaemia in the animals injected with ixodid-egg homogenates, leading to death without paralysis, was observed. Similar egg toxicity (attributed to an ‘ixovotoxin’) has since been reported for additional ixodid species (Gregson, 1973). The relationship between ixovotoxin and the toxins found in SGE is not known (Sonenshine, 1993).

When injected into guinea pigs, extracts from larval Ix. holocyclus caused paralysis (Oxer and Ricardo, 1942) whereas extracts from the larvae of Rhipicep. (B.) microplus, Ix. ricinus, Hyalomma dromedarii and Haemaphysalis bispinosa proved to be lethal — with clinical signs similar to those induced by ixovotoxin — but produced no paralysis (Riek, 1957).

Extracts from the nymphal stages of Ix. holocyclus and the Karoo paralysis tick, Ix. rubicundis, also induced paralysis, and were fatal, when injected into guinea pigs (Oxer and Ricardo, 1942) and rabbits (Spickett et al., 1989), respectively. In contrast, extracts of the nymphs of Rhipicep. (B.) microplus or Ha. bispinosa were non-fatal when injected into guinea pigs (Riek, 1957).

PHYSIOLOGICAL REACTION OF THE HOST TO TICK TOXIN

Toxin from Ix. holocyclus is now believed to induce motor paralysis as the result of action at the host’s neuromuscular junctions (Cooper and Spence, 1976). In dogs paralysed by Ix. holocyclus, Cooper et al. (1976) found nerve action potentials and maximum nerve conduction velocities to be normal but muscle action potentials to be reduced. In humans and other animals affected by holocyclotoxin, the cortex of the central nervous system (CNS) seems generally unaffected, since the cases remain alert and conscious and sensations are retained (Stone, 1986).

Although Ross (1926) concluded that the toxin of Ix. holocyclus acts on motor neurones in the CNS (because humans paralysed by the toxin showed loss or modification of reflexes and normal contractions in their paralysed muscles), this conclusion was not supported by the results of subsequent research. Using phrenic-nerve–hemidiaphragm preparations, Cooper and Spence (1976), for example, demonstrated a temperature-dependent inhibition of transmitter release by holocyclotoxin at neuromuscular junctions; at relatively low temperatures, muscle contraction following nerve stimulation occurred more readily than at higher temperatures. Cooper and Spence (1976) concluded that holocyclotoxin inhibits transmitter release at neuromuscular junctions via an intermediate step between the terminal membrane and the release of acetylcholine.

When Campbell and Atwell (2002) investigated cardiovascular functions in dogs with tick paralysis, they found that the dogs were suffering from acute left-sided congestive heart failure. Prolonged QT intervals (which represent the time needed for ventricular depolarization and repolarization) were seen in electrocardiographic investigations of dogs (Campbell and Atwell, 2002), bats (Campbell et al., 2003) and rats (Campbell et al., 2004) affected by tick paralysis caused by Ix. holocyclus. The salivary toxin of Ix. holocyclus directly affects vascular and cardiac potassium channels by blockade, and this action differs from the respiratory distress caused by progressive muscle paralysis (Atwell et al., 2001).

It is not known if the limb paralysis and cardiovascular changes seen in mammals bitten by Ix. holocyclus are elicited by one or two separate toxins. In animals paralysed by the toxin produced by De. andersoni, paralysis was found to be the result of presynaptic blockage of conduction at neuromuscular junctions (Murnaghan and O’Rourke, 1978; Gothe et al., 1979; Stone, 1988). When acetylcholine was injected into paralysed muscle preparations, the muscles responded normally, indicating that acetylcholine is either not produced or not released in the paralysed muscle (Murnaghan and O’Rourke, 1978).

VACCINE

Over the last three decades there has been some research into a vaccine that will protect against tick paralysis. As already mentioned, Stone et al. (1986) treated a partially purified toxin from Ix. holocyclus SG with glutaraldehyde to produce an inactivated and stabilized antigen (toxoid). In rabbits, compared with untreated SGE, this preparation showed enhanced immunogenicity, with relatively high toxin neutralization titres that were achieved in a relatively short time (Stone and Neish, 1984). In dogs, the toxoid showed promising results (Stone et al., 1986). Based on the results of limited terminal amino-acid sequencing of toxic proteins isolated from Ix. holocyclus SG, a putative toxin-coding gene was cloned and expressed recombinantly in bacteria, to produce large quantities of toxin for vaccine research and development (Beckmann, 1989). Unfortunately, the monoclonal antibodies that were generated for toxin isolation were later shown to be unsuccessful in neutralizing the Ix. holocyclus paralysis toxin (B. F. Stone, unpubl. obs.). In another study (Crause et al., 1994), neutralizing monoclonal antibodies were produced to the paralysing toxin of Rhipicep. evertsi and these antibodies were shown to cross-react with the paralysis toxin from Ar. walkerae.

SUMMARY AND CONCLUSIONS

In Australia, tick paralysis is a potentially fatal illness of importance to both human and veterinary medicine, largely attributable to holocyclotoxin from Ix. holocyclus. This toxin is produced in the salivary glands of feeding ticks and transferred to the host via the ticks’ saliva. In the host, the toxin results in a presynaptic decrease in acetylcholine release at the neuromuscular junction. Clinical presentation usually involves a combination of lethargy, weakness, unsteadiness of gait, dilated pupils, ascending symmetrical paralysis, slurred speech and depressed deep-tendon and gag reflexes. Laboured breathing, bradycardia, decreased oxygen saturation and asystole may also be observed. Treatment involves removing the tick, although the removal of the tick may trigger further deterioration in the patient’s clinical condition, and respiratory support may be required in an intensive-care unit. Tick anti-serum is available but may cause acute allergic reactions.

Monoclonal antibodies to Ix. holocyclus toxin are required for the identification and characterisation of tick-toxin components and purification of toxin-specific antigens. The production of potent neutralizing antibodies may provide the basis for a more standardized and specific immunotherapy against tick paralysis.

An immuno-assay to detect and quantify antibodies to tick toxin would be extremely useful, not only for evaluating the immune status of patients but also for monitoring sera following immunization for monoclonal-antibody production, and for screening the resultant hybridomas.