Tick paralysis (toxicity) is an acute, progressive, symmetrical, ascending motor paralysis caused by salivary neurotoxin(s) produced by certain species of ticks. With some species, other signs of systemic "single organ" toxicity (eg, cardiac, airway, bladder, lung, esophagus, etc) may be seen separate to or within the classic paretic-paralysis presentation. People (usually children) and a wide variety of other mammals, birds, and reptiles may be affected. Human cases of tick paralysis caused by the genera Ixodes, Dermacentor, and Amblyomma have been reported from Australia, North America, Europe, and South Africa; these three plus Rhipicephalus, Haemaphysalis, Otobius, and Argas have been associated with paralysis to varying degrees in animals.
This toxicity is unique, because it is a pulsed toxin flow associated with repeat tick feeding over a set period of time. Animals are generally affected by paralysis, but there are also very odd presentations of associated toxicity. Deterioration can be unpredictable and rapid in some cases, and some animals may have prolonged and unexplainable recovery. Very severe cases require access to ICU facilities for artificial ventilation to maximize recovery rates, and generally, only tick removal and use of TAS and antibiotics have a major effect on overall mortality.
Etiology, Epidemiology, and Pathogenesis
The potential to induce paralysis has been demonstrated, described, or suspected in 64 species of ticks belonging to 7 ixodid and 8 argasid genera. On the eastern coast of Australia, the paralysis tick I holocyclus (and to a lesser extent I cornuatus and I hirstii, in which morphologic classification has been shown to be unreliable) causes the most severe form of tick paralysis, with a mortality of as much as 10% in dogs (usually 4%–5%), irrespective of therapy.
In North America, D andersoni (the Rocky Mountain wood tick) and D variabilis (the American dog tick) are the most common causes. Sheep, cattle, and people may be affected, as well as dogs. D albipictus, I scapularis, Amblyomma americanum, A maculatum, R sanguineus, and O megnini may cause paralysis. In fowl, Argas radiatus and A persicus have caused paralysis. In Africa, I rubicundus (Karoo tick paralysis) and R punctatus in South Africa, R evertsi evertsi and Argas walkerae in sub-Saharan Africa, and R evertsi mimeticus in Namibia can cause the disease. Cats appear to be resistant to the disease caused by these ticks but are affected by I holocyclus. Toxicity is usually less severe (than in dogs), does not include chest complications, and has a better prognosis.
I holocyclus in Australia causes a much more severe disease than that seen in North America and elsewhere. Dogs and cats are affected, as well as sheep, goats, calves, foals, horses, pigs, flying foxes, poultry, birds (ostrich), reptiles (snakes and lizards), and people. Both local (less common) and systemic paresis and paralysis are seen. The natural hosts (bandicoots) are rarely affected, presumably acquiring immunity at an early age. However, without exposure to toxin, they too become susceptible.
Host factors influencing epidemiology include species, sensitivity to toxin, age, acquired immunity, field behavior, concurrent work demands, reaction to environmental factors, skin reactivity, and population density. Antitoxin immunity, starting at least 2 wk after primary tick exposure and lasting a few weeks, can be boosted by further infestations, but chronic tick exposure eventually is associated with a decline in immunity, possibly due to toxin-neutralizing effects by the host. Tick factors include age, toxin absorption and circulation dynamics, paralysis-inducing capability, other toxic effects, sexual activity, rate and volume of secretions, and the frequency of the sucking phase.
The maximal prevalence of tick paralysis is associated with seasonal activity of female ticks, mainly in spring and early summer, but in some areas ticks are active throughout the year. Environmental factors such as temperature and humidity also play a major role in tick morbidity and mortality (ie, ticks are easily killed by both hot/dry and wet conditions). Modern rapid transport of ticks attached to people, animals, or plant material can give rise to isolated cases of tick paralysis, far removed from the particular geographic area (or country) where the ticks are naturally found. The diagnosis may be delayed when such infested animals travel to areas where such paralysis associated with reduced levels of acetylcholine is not typically seen.
Toxicity does not relate directly to tick size, number, or duration of attachment. The clinical signs produced in various hosts depend on several variables, including toxin secretion rate, local site responsiveness, host immunity and susceptibility, and specific organ susceptibility.
Systemic toxicity follows injection of toxin(s) into the host, especially during periods of rapid engorgement, although large numbers of larval or nymphal ticks may also cause the same paralysis of the neuromuscular junction. The toxin is presumed to travel from the attachment site via the lymph to the systemic circulation and thus to all areas of the body, where it has a direct effect on cellular potassium channels and thus on intracellular calcium levels. However, primary hypoventilation is the main cause of death in most severe cases, in which alveolar disease may also be present.
In tick paralysis other than that caused by I holocyclus, clinical signs are generally seen ~5–9 days after tick attachment and progress over the next 24–72 hr. When I holocyclus is involved, clinical signs usually appear in 3–5 days (rarely longer, eg, up to 18 days, possibly with virginal ticks) after attachment and usually progress rapidly throughout the next 24–48 hr. Time periods can vary with I holocyclus because of tick factors, environmental humidity, temperature (microclimate), and host factors. Both "shorter onset to severe" signs and delayed "quiet" attachments with minimal signs may be seen. Removal of I holocyclus ticks does not immediately halt progression of disease. In severe cases, death from respiratory muscle failure and other respiratory complications can occur within 1–2 days of the onset of signs.
Early signs may include change or loss of voice (due to laryngeal paresis); hindlimb incoordination (presumed to be due to weakness and not central CNS ataxia); change in breathing rhythm, rate, depth, and effort; gagging, grunting, or coughing; regurgitation or vomiting; and pupillary dilation. Dogs with a grunt are believed to have increased airway resistance.
Hindlimb paralysis begins as slight to pronounced incoordination and weakness, which is best observed with the animal turning or walking away from the observer (or when climbing stairs or jumping up). As paralysis progresses, the animal becomes unable to move its hindlimbs and forelimbs, to stand, to sit, to right, and finally to lift its head.
A four-stage classification system based on systemic limb activity may enable clinical predictability. In stage 1, the dog's voice is changed (usually noticed retrospectively), and the dog is weakened but can still walk and stand. In stage 2, the dog cannot walk but can stand. In stage 3, the dog cannot stand but can right. In stage 4, the dog cannot right. Stages 3 and 4 (~30% of cases) indicate a poor prognosis. However, some dogs show few signs because of low levels of toxin or high levels of protective skin or systemic immunology, and some show signs in only one organ (eg, esophageal paralysis). Sensation is usually preserved, but it is increasingly harder to detect the clinical responses to stimuli due to lower motor neuron paralysis. (Visual analog scale scoring is also performed for the neuromuscular junction, overall toxicity, and dyspnea, with highly predictive results).
Breathing abnormalities include choke, upper respiratory tract obstruction, bronchoconstriction (especially seen early in cats), progressive fatigue of respiratory muscles, and aspiration of esophageal and/or gastric contents (due to loss of pharyngeal and laryngeal function), leading to aspiration pneumonia. Aspiration can be significant, and the lung severely affected before any obvious signs. It is possible to have a silent (no crackles), severely pneumonic lung if there is poor airflow into the affected lobe. Some dogs have profound dyspnea, no crackles, and extensive pulmonary radiographic opacity (due to aspiration pneumonia); such cases are usually terminal. Dogs with upper respiratory tract obstruction have a marked expiratory stridor (not the classic inspiratory stridor of primary laryngeal paralysis of large breeds), often with the head and forelegs extended to maximize air flow and exchange. If there is chest disease as well, the animal is usually very dyspneic. A thrill can be felt at or just below the larynx in association with the obstructed expiratory effort and stridor. The upper respiratory tract lesion can be easily missed, especially if the dog is paralyzed. Often the respiratory rate is high and forced. In cats, the doll test can be used to assess upper respiratory tract function. If finger/thumb compression of the chest induces a stridor, then this supports paresis or paralysis, irrespective of other respiratory tract defects. It is essential that any upper respiratory tract obstruction be diagnosed, because the associated workload, anxiety, and resultant fatigue can quickly become terminal.
Paralysis of esophageal muscles develops in most dogs (but not cats), with or without obvious esophageal dilation. Saliva and ingested food or fluid pool in the esophagus and may be regurgitated into the pharynx and mouth. Loss of pharyngeal function makes it difficult for the animal to clear material from the upper respiratory tract, which may then lead to aspiration pneumonia.
Vomiting (with evidence of bile) may occur in I holocyclus paralysis; a central action of toxin on the vomiting center has been suggested. Most cases of vomiting reported by clients are probably regurgitation, although drug-induced vomiting can be a complication. Dogs will gag and retch in an attempt to clear secretions and move their head and jaw in an odd way, associated with a characteristic groan, to further attempt clearance of materials.
Body temperature may be normal in the early stages; however, due to the toxin's effect on arteriovenous anastomoses (shunts), normal thermoregulation is lost. This can cause hyper- and hypothermia as animals are affected by local environmental factors. Shivering is also lost in severe cases. Profound hypo- and hyperthermia can occur suddenly and can be easily misdiagnosed; hypothermia clinically resembles tick paralysis in several ways. When body temperature is restored, the level of tick paralysis in some cases can be mild.
Rarely in dogs, acute congestive heart failure can present with extensive pulmonary edema due to diastolic myocardial dysfunction (the myocardium is unable to correctly relax, reducing efficient chamber filling and therefore systolic cardiac output). Venous return may also be reduced, and systemic venous pressure is increased.
Some dogs have a prolonged QT interval on ECG, which can result in a lethal ventricular arrhythmia. The frequency of these unexplained deaths, which follow complete gross clinical recovery, is not known, but most veterinarians who treat many cases report such events.
Cats with moderate to severe toxicity can be anxious. It is essential not to interfere with these animals until they have settled in their cage. If procedures are forced on them, these animals can die from obstructive dyspnea and the (presumed) associated hypoxemia, acidosis, and hypercapnea. Animals can deteriorate if compromised by excessive hospital stress (eg, nursing attention, noise, smell).
Cats may present with an "asthma-like" airway constriction, usually when they are mildly paretic; expiratory wheeze on auscultation, forced abdominal expiratory effort, and very easily induced exercise intolerance are classic signs at this time. These cats often have a positive doll test and will, after a few steps, sit on their hindquarters with the chest in a more upright vertical position than normal, often with an increased respiratory focus or effort. Feline "asthma" can be easily misdiagnosed at this stage if a tick is not found or suspected.
The presence of a tick in conjunction with the sudden appearance of limb weakness and/or respiratory impairment is diagnostic. The offending tick may no longer be attached, but a “skin crater” (a hole 1–2 mm deep and 1–3 mm wide, surrounded by a variably raised and inflamed area) confirms the diagnosis. Sometimes neither tick nor crater can be found (ticks attached deep in the ear, between toes, or in the mouth or anus may be missed). However, with the appropriate clinical signs, in a known tick area without another obvious cause of lower motor neuron or neuromuscular disease, tick antitoxin serum (TAS) treatment is still indicated. Recovery after treatment subjectively confirms the provisional diagnosis.
Specific laboratory diagnostic techniques are not available, but procedures that may be generally helpful include a PCV, serum protein and radiography, to assess presence and degree of pulmonary edema, megaesophagus, and pneumonia due to aspiration. Specific signs (eg, congestive heart failure, urethral obstruction) require routine evaluation and treatment of that body area or system.
Botulism, polyradiculoneuritis, acute peripheral neuropathies, snakebite, hypokalemia, and toadfish and ciguatera toxicity are some differential diagnoses. In regions where ticks are endemic, tick paralysis is usually high on the list of differential diagnoses for any flaccid, clinically ascending motor paralysis. It should also be considered in the differential diagnosis of megaesophagus, unexplained vomiting, acute left congestive heart failure (dogs), or "asthma" (cats). The tick season is usually well known for various areas (eg, a local creek) within the environment of a particular practice, and often most tick paralysis cases come from a few, well-defined, highly endemic areas.
Blood and serum values are unchanged in the early stages. Increased PCV (with normal serum protein) indicates a fluid shift into the lungs and a more guarded prognosis. Other changes may include increased levels of blood glucose, cholesterol, phosphate, and CK, and a decrease in blood potassium levels, but none of these changes are specific for tick paralysis or indicate severity or prognosis.
Echocardiography reveals both diastolic and secondary systolic myocardial dysfunction associated with reduced ventricular filling, possibly due to both peripheral venous pooling and poor diastolic myocardial relaxation. Nonstressful radiography gives the best available prognostic support, and pulse oximetry the best continuous assessment of oxygenation. Capnography helps assess the functional level of ventilation. Arterial blood gas analysis (although invasive) gives the best overall assessment of cardiopulmonary function. However, the stress of any such testing should be considered; positioning for chest radiographs (eg, dorsoventral to lateral) can tip animals into a terminal hypoventilatory decline associated with acute respiratory or cardiac arrest.
In most infestations (except I holocyclus), removal of all ticks usually results in improvement within 24 hr and complete recovery within 72 hr. If ticks are not removed, death may occur from respiratory paralysis in 1–5 days. Removal of I holocyclus ticks does not immediately halt progression of disease. Clinical signs can deteriorate for ~24 hr and longer, but most dogs start to improve in 6–12 hr after TAS therapy. In any infestation, removal of all ticks is absolutely necessary. The entire integument should be searched, diligently and repeatedly, especially on long-haired animals or those with thick coats. Most ticks (80%) are located around the head or neck, but they can be found anywhere on the body. Plucking the tick(s) yields the best result (in dogs) and does not induce anaphylaxis.
Therapy for tick toxicity must address primary tick toxemia and paralysis, secondary issues (eg, esophageal reflux, aspiration pneumonia), and potential tertiary factors (eg, chronic weakness, esophageal stricture).
TAS is an immune serum against the toxin (similar to tetanus antitoxin) and is the product of choice. It should be given as early in the disease as possible; subsequent “top up” doses are not effective, because they are too late. For dogs, a minimal dosage of 0.5–1 mL/kg, should be given slowly IV throughout at least 20 min to avoid any shock reaction. Rapid IV use can induce clinical reactions in >80% of dogs. Anaphylaxis can occur unpredictably (as with all products), necessitating the use of high-dose, soluble cortisol and rapid fluid loading, etc. Based on retrospective case studies, cats are believed to be somewhat more susceptible than dogs, presumably with a second dose, a few weeks (not days) after the first dose.
Animals with multiple ticks or in the early stages of paralysis should receive a higher dose, because these cases have the most unbound toxin to neutralize. However, there are no data for the exact dosage rates required, and batch and brand levels of protective immunoglobulin may vary. Severely affected dogs may have less remaining unbound circulating toxin for TAS to neutralize. However, debate persists about the required (possibly less) dose of TAS. It has been suggested that a standard dose should be given, based on the amount needed to neutralize the pulsed toxin flow from one tick rather than on a weight basis; a minimal dose of 10–20 mL is recommended for dogs (and 5–10 mL for cats). However, until the level of remaining unbound toxin in the affected animal and the level of specific protective immunoglobulin can be better assessed, a set dose rate cannot be established (the level of disease is reflective of the level of bound toxin, which is not affected by TAS).
TAS given IP is the best alternative in cats for which the IV route is an issue (eg, respiratory distress, restraint dangers, dyspnea). However, its clinically effective half-life is believed to be short (days not weeks), and it will have no effect if the toxin is already tissue bound and the animal is severely ill or about to become so (with toxin in the perivascular space).
Minimization of stress and anxiety is essential. Acepromazine (0.03 mg/kg) may be given SC before any other medication or handling that may upset the animal. However, high doses should be avoided, especially if the animal is depressed, hypotensive, or hypothermic. (Overdosage may induce hypotension and hypothermia.) Opiates are an alternative (eg, methadone, 0.3–0.5 mg/kg, SC, IM). Oxygen therapy (nonstressful, usually nasal) is implemented (as indicated), but progressive disease requires more intensive therapy.
General anesthesia is indicated in animals that are severely fatigued and dyspneic, to allow for better administration of oxygen, esophageal drainage, and upper respiratory tract suction. Pentobarbitone can be used as a constant-rate infusion or given periodically IV to induce light anesthesia, with repeat doses as needed. Another potential benefit of pentobarbitone may be control of long QT syndrome. The chief benefits of some form of anesthesia (eg, profofol) are to reduce dyspnea, enable muscle rest, and help overcome primary muscle fatigue and general exhaustion. Periods of 6–8 hr of light anesthesia are best, with reassessment of clinical status after each period.
Mechanical or manual ventilation may be required but should be carefully assessed because recovery can be delayed, especially in brachycephalic animals. Longer-term ventilation cases can have a 70% recovery rate. It is essential to assess pulmonary (expired CO2 levels) and alveolar (pulse oximetry) ventilatory capacity and to be aware of profound respiratory muscle fatigue. Alveolar disease (edema and/or pneumonia) has a poor prognosis in such cases.
Atropine (repeat every 6 hr, lowest dose) can be used if GI and respiratory secretions are excessive, but its effect on tear secretion (and the host's potential for eyelid paralysis, reduced blink reflex, and corneal drying) and cardiac rate and rhythm changes should be considered.
Antiemetic therapy should be used in animals that are vomiting, which is usually a poor prognostic sign. If the animal is regurgitating, the esophagus should be aspirated along with the upper respiratory tract. Correct drainage positioning then becomes a vital factor in helping to avoid aspiration. Care is needed with gastroesophageal reflux cases regarding their chronicity and tissue damage.
Broad-spectrum bactericidal antibiotics are indicated (especially in severe cases) to help avoid development of aspiration pneumonia, but they must be given as soon as possible. Dogs with upper respiratory tract obstruction require either tracheotomy or anesthesia and intubation to overcome the potentially lethal effects of such obstruction.
Diuretics (eg, furosemide) with maximally appropriate oxygen treatment are indicated to treat congestive heart failure. Verapamil (0.1 mg/kg, IV bolus) has been used to help relieve the basic toxic myocardial effect of a failure to relax. The toxin does unbind, so if the animal can be kept free of terminal pulmonary edema (or arrhythmia), the cardiac failure will reverse over a few days, provided routine support is given. Esmolol has been used to treat affected animals that have a long QT interval and the potential for a lethal, unpredictable ventricular arrhythmia.
Fluid therapy should be used with great care, because pulmonary edema can be induced easily. Staying below maintenance levels and ensuring the animal is assessed for edema, both before and during IV fluid therapy, should be routine. Dehydration can occur in tick paralysis but not usually in routine cases until the second day of hospitalization, when increased PCV and protein values may be evident. In small patients, SC or IP fluids can be given if lung status is a concern. Exceptional cases may require extensive rehydration (eg, paralyzed in the sun with high humidity and temperature for a day before presentation), but the extent of the underlying organ dysfunction should be assessed before intensive fluids are given.
The asthma-like disease in cats is hard to reverse, because routine bronchodilators do not seem to be effective.
Muscle fatigue can be reduced (with recovery of some muscle strength) by short periods (6–8 hr) of anesthesia. The animals remain hypercapneic but, with endotracheal intubation and O2 therapy, can establish reasonable hemoglobin saturation levels (>95%), provided there is no significant alveolar disease.
Intoxicated animals lose their ability to regulate body temperature. Animals that have fallen below 32°C (90°F) for a long period may be hard to resuscitate. Various heating mechanisms are used (hot water bottles, blankets, hot air flow blankets), but peripheral heat absorption cannot occur if arteriovenous anastomoses (shunts) are shut due to the effect of the toxin and the host's vasoconstrictive reaction to hypothermia. Warmth applied at the lower limbs (especially the hindlimbs) will be of maximal benefit; direct application to the groin area may also potentially be useful. Some animals may need warmed fluids, IV or rectally, to reverse a very cold presentation (eg, ≤32°C). Sudden hyperthermia (>42°C) can be seen in hospitalized dogs. They usually show exaggerated head and possibly foreleg movements and signs of anxiousness. With cooling (eg, wet towels, direct fan flow, high rate of air changes), these signs abate.
Because the animal's condition is expected to deteriorate after ticks are removed and TAS is given, hospitalization with minimally invasive monitoring and good nursing care is necessary. The animal should be kept in a quiet, dark, comfortable area of the hospital where it can be easily seen. It should be placed on the sternum to maximize lung function. Lateral recumbency, left side down with the shoulder (not the pharynx or neck) as the highest point, is the best position for drainage. If possible, slight “head down” is also advised. Animals should never be rotated unless it can be done frequently (every 1–2 hr), day and night.
Because the animal cannot void, catheterization is necessary, with the bladder expressed at least twice daily to avoid infection. As with other localized tick toxicity effects, this may persist beyond the period when the animal has generally recovered. Eye protectants should be used to prevent corneal ulceration or dryness (lid closure, artificial tears, contacts). Suction of the pharynx, larynx, and proximal esophagus minimizes upper respiratory tract distress caused by saliva pooling and regurgitation. An esophageal tube may be slowly inserted to remove any pooled material; in some cases, this is voluminous and possibly prevents choke (seen mostly in brachycephalic breeds with laryngeal blockage by foreign material). Fluid and oxygen therapy should be monitored to avoid overhydration or under-supply, respectively. Nutritional support should be performed carefully to ensure that GI and respiratory function can cope with any offered food and water.
Repeated tick searches should be performed during hospitalization, especially if the animal deteriorates unexpectedly or is slow to recover. Long or matted hair should be clipped, especially about the head and neck. Application of an acaricide may kill any ticks missed in searching. However, the stress of searching, clipping, or bathing can be detrimental in severely affected or nervous animals, in which sedation is recommended.
Appropriate and timely treatment (TAS and antibiotics, especially in severe cases) saves ~95% of affected animals, but ~5% of animals are likely to die despite all treatment efforts, especially those with aspiration, and advanced respiratory paralysis and dyspnea. Most animals (>80%) have only one tick and a large attachment crater. Prolonged recovery and weight loss can be seen with various complications, and death can also occur due to choke, respiratory muscle fatigue, cardiac arrhythmias, congestive heart failure, and cardiopulmonary arrest. Older animals are at greater risk, as are very young pups. Proportionally more severe cases are seen at the start of the season, and a second (close to the first) infestation will be more severe. Dyspnea, crackles, and wheezes are poor prognostic signs, as are high neuromuscular junction scores (3 and 4) or high visual analog scale scores (≥75%) for toxicity or respiratory distress.
Before discharge, the drop test can be used in cats to assess neuromuscular function and three-dimensional gravitational control. Cats should be able to correct a fall from 10–20 cm above the top of the table. Still-affected cats will not correct in time and land more heavily, with the chin hitting the padded table top. Recovered cats land lightly with good head control. Jumping up to and down from the cage can also be used to assess muscle strength in cats. In dogs, jumping down from a cage can induce stridor, indicating unresolved respiratory paresis with forced expiratory air flow, because the unsupported abdominal momentum affects the diaphragm and lung air flow, producing a high-volume expiration. Lifting a dog (with the holder's arms wrapped outside the fore- and hindlegs) with unresolved tick paralysis often produces stridor, indicating abnormal laryngeal function. Animals should be able to eat, drink, and walk normally without any stridor before discharge.
Owners should be advised to continue searching recovered animals for ticks; use appropriate preventive methods to avoid reattachment of ticks; and avoid high temperatures, stress, or strenuous exercise for at least the first month. Smaller, more frequent meals may also be indicated if there was esophageal dysfunction. This rest period especially applies to working farm dogs, in which early overexercise may lead to permanent muscle damage.
Prevention and Control
Owners should not rely solely on chemical control to prevent tick infestation, because no product is totally effective, and a single attached tick can cause the disease. They should be advised about when and where their pets will be at risk; encouraged to thoroughly search the coat daily; keep the coat as short as possible (to aid searching); and understand the efficacy, appropriateness, safety, and limitations of available preventive products (sprays, topical spot-ons, tablets, and collars). Combination therapy (eg, spray and collar) may give better results by using two modes of action, but there are no published data to support this concept.
Attempts to produce an effective vaccine against the I holocyclus toxin have so far been unsuccessful, as have been attempts at “in-field” tick control. Specific RNA studies show that ticks vary geographically, and such genetic differences may explain why clinical signs of tick paralysis and visual analog scale toxicity scores can vary between different areas at the same time of the year in the same season.
Last full review/revision January 2014 by Rick Atwell, BVSc, PhD, FACVSc