association of wolbachia with heartworm disease in

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ASSOCIATION OF WOLBACHIA WITH HEARTWORM DISEASE IN CATS AND DOGS

By

PATRICIA ANN DINGMAN

A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT

OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE

UNIVERSITY OF FLORIDA

2009

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© 2009 Patricia Ann Dingman

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To my cats, Sara and CC and my dog, Ellington Foxworthy

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ACKNOWLEDGMENTS

I thank Nicholas Ronnquist for being patient and understanding. I thank my mentor

and inspiration, Dr. Julie Levy. She has given me so many opportunities to succeed and

has greatly contributed to my personal and professional growth. I would like to thank Dr.

Calvin Johnson, Dr. Ellis Greiner, and Dr. Charles Courtney for their patience and

support.

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TABLE OF CONTENTS page

ACKNOWLEDGMENTS ...............................................................................................................4

LIST OF TABLES...........................................................................................................................7

LIST OF FIGURES .........................................................................................................................8

ABSTRACT ....................................................................................................................................9

CHAPTER

1 DIROFILARIA IMMITIS ........................................................................................................11

Identification...........................................................................................................................11 Life Cycle/Pathogenesis .........................................................................................................11 Epidemiology..........................................................................................................................13 Heartworm Disease.................................................................................................................13 Heartworm-Associated Lung Pathology.................................................................................14 Diagnosis of Heartworm Infection .........................................................................................15 Heartworm Treatment.............................................................................................................15

2 WOLBACHIA..........................................................................................................................17

Biology ...................................................................................................................................17 Endosymbiotic Relationship with D. immitis .........................................................................17

3 EXPERIMENTAL DESIGN..................................................................................................20

Materials and Methods ...........................................................................................................20 Animals............................................................................................................................20 Sample Collection ...........................................................................................................21 Serology...........................................................................................................................21 Parasitology .....................................................................................................................22 Pulmonary Histologic Morphometry...............................................................................22 Wolbachia DNA Detection..............................................................................................23 Wolbachia Immunohistochemistry..................................................................................24 Statistical Analyses..........................................................................................................24

4 RESULTS...............................................................................................................................32

Statistical Analyses.................................................................................................................32 Serology...........................................................................................................................32 Parasitology .....................................................................................................................32 Detection of Wolbachia ...................................................................................................33 Pulmonary Histologic Morphometry...............................................................................33

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5 DISCUSSION.........................................................................................................................44

6 CONCLUSIONS ....................................................................................................................46

LIST OF REFERENCES...............................................................................................................47

BIOGRAPHICAL SKETCH .........................................................................................................51

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LIST OF TABLES

Table page 3-1 Pulmonary arteriolar lesion scoring system.......................................................................25

3-2 Bronchiolar lesion scoring system. ....................................................................................26

3-3 Pulmonary interstitial and alveolar smooth muscle scoring system. .................................27

4-2 The prevalence of Ancylostoma spp. and ascarids in cats and dogs with pulmonary lesions ..............................................................................................................36

4-3 Frequency of pulmonary lesions in cats and dogs with and without Wolbachia...............37

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LIST OF FIGURES

Figure page 3-1 Representative feline pulmonary tissues demonstrating lesion scores in

arterioles.............................................................................................................................28

3-2 Representative feline pulmonary tissues demonstrating lesion scores in bronchioles.........................................................................................................................29

3-3 Representative feline pulmonary tissues demonstrating lesion scores in alveolar smooth muscle......................................................................................................30

3-4 Representative feline pulmonary tissues demonstrating lesion scores in interstitium .........................................................................................................................31

4-1 Feline D. immitis antibody titer versus Wolbachia surface protein (WSP) antibody titer in all cats......................................................................................................38

4-2 Canine D. immitis antibody titer versus Wolbachia surface protein (WSP) antibody titer in all dogs ....................................................................................................39

4-3 Severity of pulmonary lesions in cats with or without detectable Wolbachia DNA and/or Wolbachia surface protein in lung tissues.....................................................40

4-4 Severity of pulmonary lesions in dogs with or without detectable Wolbachia DNA and/or Wolbachia surface protein in lung tissues.....................................................41

4-5 Severity of pulmonary lesions in cats with or without detectable circulating Wolbachia surface protein antibodies (WSP Ab) ..............................................................42

4-6 Severity of pulmonary lesions in dogs with or without detectable circulating Wolbachia surface protein antibodies (WSP Ab) ..............................................................43

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Abstract of Thesis Presented to the Graduate School of the University of Florida in Partial Fulfillment of the

Requirements for the Degree of Master of Science

ASSOCIATION OF WOLBACHIA WITH HEARTWORM DISEASE IN CATS AND DOGS

By

Patricia Ann Dingman

May 2009 Chair: Julie Levy Major: Veterinary Medical Sciences

Although the presence of adult Dirofilaria immitis in the pulmonary arteries and its

associated arteritis and thromboembolic disease can explain some of the manifestations

of canine and feline heartworm disease, the cause of other findings remains unclear. For

example, cats with D. immitis antibodies but lacking adult parasites in the pulmonary

arteries frequent develop histological lesions of Heartworm-Associated Respiratory

Disease. All D. immitis parasites harbor Wolbachia and D. immitis-infected animals can

have circulating Wolbachia antibodies and pro-inflammatory Wolbachia antigens

deposited in tissues. Little is known about the role Wolbachia plays in the pulmonary

pathology of animals naturally infected with D. immitis. The purpose of this study was to

determine the contribution of Wolbachia to the pathogenesis of natural heartworm

disease in cats and dogs. We hypothesized that animals having sufficient Wolbachia

burden to be detected in pulmonary tissue by immunohistochemistry and/or by PCR

would have more severe pulmonary disease than those with bacteria below the limits of

detection. We further hypothesized that animals that were immunoreactive to pro-

inflammatory WSP would have more severe pulmonary lesions than those that were

seronegative for WSP antibodies. Blood and pulmonary tissue samples were collected

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from cats and dogs representing three different D. immitis infection statuses: heartworm-

free, heartworm-exposed, heartworm-infected. There was a positive but weak correlation

between the magnitude of D. immitis antibody titers and WSP antibody titers in cats

(r=0.57, p<0.001) and in dogs (r=0.39, p<0.001). Pulmonary lesions were more common

in HW-infected animals than in HW-free animals. Pulmonary arteriolar occlusion was

more common in HW-infected cats (57%) (p=0.003) than in HW-infected dogs (17%).

Although pulmonary lesions were most common in HW-infected animals, there was no

clear additive effect when either Wolbachia DNA/WSP was detected in pulmonary tissue

or when circulating WSP antibodies were detected. Similarly, there were no significant

differences in the magnitude of pulmonary lesion scores within each HW-infection status

group regardless of whether Wolbachia DNA/WSP or WSP antibodies were detected.

The relationship between Wolbachia and pulmonary pathology in heartworm-infected

animals remains to be determined. The lack of clear evidence for a role of Wolbachia in

heartworm disease creates a dilemma for veterinarians treating cats and dogs in D.

immitis-endemic areas. Although the indiscriminant use of antibiotics should be avoided,

many clinicians prescribe doxycycline based on the favorable responses observed in

human filarial diseases and on promising results from the first published studies of

doxycycline use in D. immitis-infected dogs.

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CHAPTER 1 DIROFILARIA IMMITIS

Identification

Dirofilaria immitis (heartworms) are filarial helminths that belong to the phylum,

Nematoda (roundworms) and superfamily Filaroidea. Other filarial nematodes, including

Wuchereria bancrofti and Brugia malayi, are known to cause human lymphatic filariasis

(elephantiasis), and Onchocerca volvulus is a causative agent for human river blindness disease.

Microfilariae (mff) are mobile vermiform embryos and are the microscopic immature stage

of D. immitis circulating in the blood of the vertebrate host. They are not sheathed and have a

straight tapered tail. Their size ranges from 286-340 µm in length and 6-7 µm in diameter. The

physical characteristics of D. immitis microfilariae distinguish them from other filarial species

such as Dipetalonema reconditum (Yabsley et al., 2004).

Dirofilaria immitis show sexual dimorphism; adult males range between 12 and 20 cm and

are much shorter in length than the female worms, which can grow as long as 30 cm in length.

Both sexes are thin, measuring less than 5 mm wide and white in color. They are cylindrical in

shape and a psuedocoelon is present. Males have coiled (corkscrew) tails, whereas the females’

tails are straight.

Life Cycle/Pathogenesis

There are three factors involving D. immitis transmission. First there needs to be an

infected vertebrate with circulating microfilariae, most commonly a domestic dog, which is a

definitive host for D. immitis. Heartworm-infected canids have circulating microfilariae and are

considered the natural reservoir for these parasites. Although cats are naturally more resistant to

D. immitis, feline infection is likely to occur anywhere the parasite is found in canids.

Microfilaremia is uncommon in cats and therefore for a cat to become infected with D. immitis

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there needs to be an infected canid nearby (Genchi et al., 2008). Environmental conditions must

be appropriate for D. immitis larval development because environmental temperature affects

growth within the mosquito and microfilariae. Finally, vector competent mosquitoes such as the

Culex spp. must feed on both infected canids and susceptible hosts to effect transmission (Nelson

et al., 2005a; Ralston et al., 1998). Almost 70 species belonging to the family Culicidae are

considered potential vectors (Aedes spp., Anopheles spp., Culex spp., and Mansonia spp.).

To begin the cycle, a female mosquito feeds on a microfilaremic dog and the microfilariae

are ingested during the blood meal. The microfilariae pass through the hemocoel and into the

malpighian tubules of the mosquito. There they mature into first stage larvae (L1s) and molt to

second stage larvae (L2s) within 10 days. Ten to fourteen days later, they become third stage

larvae (L3s), which is the infective stage. The L3s migrate to the salivary glands and proboscis

and when the mosquito feeds, the L3s are deposited onto the skin and then enter the wound made

by the mosquito. The larvae begin to develop and molt into the 4th stage larva in 3-4 days.

Fourth-stage larvae migrate to the muscle fascia of the thorax and abdomen, molt, and then L5s

enter the circulatory system over a period of 3 to 4 months. They are carried to the pulmonary

arteries, heart, and lungs via blood circulation 4 to 7 months post infection. Once in the heart and

lungs they mature into adult worms approximately 6 months after infection if both male and

female worms are present. Adult female worms begin to produce microfilariae 6 to 9 months

(180 to 190 days) after the initial infection, which is the infectious period when D. immitis can be

acquired by mosquitoes and then transmitted to other cats or dogs. In cats, arrested development

of the worm is common because cats are not the natural host. Fewer worms reach maturity in

cats compared to dogs. Only about 20% of D. immitis larvae become adult worms in cats,

whereas up to 75% of D. immitis larvae in dogs become adults. Microfilariae were shown to

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remain in blood circulation for up to two years in a dog that was experimentally inoculated with

microfilaremic blood (Underwood and Harwood, 1939). Adult worms in dogs can live up to 5 to

7 years in dogs (Newton, 1968) and 2 to 3 years in cats. Aberrant migration of the parasite has

been documented, resulting in disease associated with dysfunction of the target tissue (Nogami

and Sato, 1997).

Epidemiology

In areas endemic for D. immitis, prevalence in dogs that are not on heartworm preventive

can exceed 50%. Feline heartworm prevalence is often 5-20% of the positive dog population

(Atkins et al., 1998; Bowman et al., 2007; Carleton and Tolbert, 2004; Hermesmeyer et al., 2000;

Miller, 1998; Nogami and Sato, 1997; Patton and McCracken, 1991; Ryan and Newcomb, 1995).

Recently a nationwide survey estimated the feline heartworm prevalence rate, determined by

antigen testing, to be 0.6 times the canine prevalence (Lorentzen and Caola, 2008). Northern

Florida is a high endemic area and many dogs are not on preventive. The prevalence in

unprotected dogs is 25-50%. In Northern Florida, feline heartworms are found in 5% of the cat

population in an animal shelter (Levy et al., 2003; Snyder et al., 2000). Studies have

demonstrated conflicting results when examining the cats’ gender as a predisposing risk factor

for D. immitis. Some researchers have found male cats to be at a higher risk for heartworm

infection (Kramer and Genchi, 2002; Levy et al., 2003) whereas others reported gender not being

a determining factor for infection (Atkins et al., 2000; Genchi et al., 2008; Liu et al., 2005).

Heartworm Disease

Heartworm infection can have different effects on cats and dogs. Dogs have higher parasite

intensities, often reaching greater than 30 adult worms compared to cats that have only 1-3 adult

worms (Nogami and Sato, 1997; Ryan and Newcomb, 1995). Dogs are often microfilaremic

versus cats are not usually microfilaremic and it is much easier to diagnose infection in dogs than

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in cats. Clinical signs in dogs include coughing, exercise intolerance, dyspnea, hepatomegaly,

syncope, and ascites. Clinical signs in cats include lethargy, coughing, anorexia, chylothorax,

and vomiting. Commonly cats have dyspnea or respiratory distress similar to feline bronchial

disease (asthma) and dogs experience right-heart failure (Nelson et al., 2005a, b). Often times,

there are no signs. When signs do occur, they are often the result of the arrival of L5s in the

pulmonary arteries which leads to acute pneumonitis. Signs may also occur when the deaths of

adult worms cause thromboembolism and anaphylaxis and may result in sudden death (Dillon,

1998).

Heartworm-Associated Pulmonary Pathology

Although the presence of adult D. immitis in the pulmonary arteries and its associated

arteritis and thromboembolic disease can explain some of the manifestations of canine and feline

heartworm disease (HWD), the cause of other findings remains unclear. This is particularly true

for cats, which frequently develop generalized severe bronchointerstitial disease and

extrapulmonary signs in response to very low worm intensitites (Atkins et al., 2000). In cats,

even a single worm can produce fatal disease.

Browne et al., (2005) examined 630 cats from an animal control shelter and assigned them

to three groups (HW-infected, HW-exposed, and HW-free) based on serological tests and

necropsy findings. Pulmonary lesions characterized by pulmonary arterial occlusive hypertrophy

were common in cats with adult worms and in cats that were free of adult worms but with D.

immitis antibodies, suggesting that even transient infection leave cats with long-lasting

pulmonary pathology. Occlusive hypertrophy is a characteristic of feline heartworm disease and

is defined as >95% occlusion of the arteriolar lumen by an increase in the thickness of the tunica

media. Almost 80% of the HW-infected cats had hypertrophy, followed with the HW-exposed

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(50%) and HW-free (13%) and there was a significant association between the number of

occluded vessels and HW status.

Diagnosis of Heartworm Infection

There are many blood tests for diagnosis of heartworm infection. A blood concentration

test such as the Knotts method or membrane filtration can detect circulating microfilariae in the

blood stream. Dirofilaria immitis antigen tests detect proteins shed from female adult worms.

False-negatives occur if the dog or cat has an all male, a low worm burden, or juvenile worms.

Dirofilaria immitis antibody tests indicate exposure to heartworms, but not necessarily current

infection. Enzyme-linked immunosorbent assays (ELISA) or non-ELISA lateral flow tests can be

used to detect D. immitis antigen or antibodies. Additional support for a diagnosis of D. immitis

infection includes thoracic radiography and echocardiography, as well as combining or repeating

antigen and antibody tests (Berdoulay et al., 2004; Nelson, 2008; Snyder et al., 2000). A

necropsy may provide a definitive answer; however D. immitis have been known to migrate to

aberrant locations (Nogami and Sato, 1997; Oh et al., 2008) and necropsy may not detect

immature infections.

Heartworm Treatment

Although canine heartworm infection is very common, it is also 100% preventable.

Preventives must be administered monthly starting approximately one month after the beginning

of the transmission season and ending approximately one month after the end of the season

according to the American Heartworm Society (AHS) (Nelson et al., 2005a, b). Prevention is

most important during the transmission period but to insure client compliance, and to account for

transmission variability in microclimates, treatment year-round is recommended.

Currently there are no FDA-approved drugs available for adulticide treatment in cats and

because infected cats usually lack circulating microfilaria, they do not need microfilaricide

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treatment. Adulticide therapy in cats has been associated with a mortality rate of 20-30%. AHS

recommends prednisone and supportive therapy to ameliorate clinical signs of disease in cats.

Surgical removal of the worms can be attempted in cats with uncontrolled signs. Aspirin is no

longer recommended as a treatment for heartworm disease in cats. FDA-approved heartworm

preventives for cats include ivermectin, milbemycin oxime, and selamectin. The preventives

should be administered 30 days after transmission season begins and 30 days after it ends on a

monthly basis (Nelson et al., 2005a).

There are more treatment options for dogs than for cats. Dogs should begin preventives at

8 weeks of age. Adult dogs starting preventives for the first time should be tested for D. immitis.

Ivermectin, milbemycin oxime, moxidectin, and selamectin are macrocyclic lactones most

effective against mff, L3s, and L4s. Daily diethylcarbamazine (DEC) can also be used as a

preventive but is rarely used due to the availability of monthly preventives. Testing for

microfilariae is required before treating with DEC to avoid adverse reaction in infected dogs. Re-

testing is recommended for all the preventives 7 months after the end of the transmission season

to assure prevention of infection. The only FDA-licensed adulticide treatment for dogs is

melarsomine dihydrochloride. Using low-dose ivermectin as an adulticide can take several years

to cure infection and is not generally recommended. Microfilaricide treatment should be

performed after adulticide therapy. Re-testing for microfilariae and D. immitis antigens are

recommended after adulticide therapy (Nelson et al., 2005b).

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CHAPTER 2 WOLBACHIA

Biology

The genus Wolbachia belongs to the order, Rickettsiales. Wolbachia is phylogenetically

related to Ehrlichia spp. and Anaplasma spp. The only species identified in the genus is

Wolbachia pipientis. These gram-negative bacteria are found in 100% of the D. immitis

population; all individuals are infected. The bacteria are present in all developmental stages and

are located in the hypodermal cells of the lateral chords (Kozek, 2005; Kramer et al., 2003).

Wolbachia are transmitted maternally (female to offspring). After the first week when L3s

entered the host, bacteria growth increases until L4 development. The amount of bacterial cells

remains constant in male worms, but as the females begin to mature, the bacteria increase in

number and infect the ovaries, oocytes, and embryonic stages inside the uteri. They have not

been reported from the male reproductive system.

Researchers have discovered Wolbachia surface proteins (WSP) circulating in heartworm-

infected dogs, localized in the lungs and kidneys where microfilariae are common (Kozek, 2005;

Kramer et al., 2003). Macrophages containing Wolbachia have been identified in the lung, liver,

and kidneys of dogs with natural D. immitis infection (Kramer et al., 2005b). Dirofilaria immitis-

infected animals also have circulating WSP antibodies (Bazzocchi et al., 2000a; Kozek, 2005;

Kramer et al., 2008; Kramer et al., 2005b; Morchon et al., 2004).

Endosymbiotic Relationship with D. immitis

In recent years, it has been recognized that a large array of helminths and arthropods are

colonized by Wolbachia (Bandi et al., 1998; McCall, 2005). Human and animal filarial

nematodes, including the agents of human river blindness (Onchocerca volvulus), lymphatic

filariasis (Brugia malayi), and feline and canine heartworm disease (Dirofilaria immitis), harbor

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Wolbachia in a mutually dependent manner (Bazzocchi et al., 2000b Kozek, 2005; Kramer et al.,

2003; McCall, 2005; Taylor, 2003). Wolbachia aid in the parasite’s survival and reproduction

and can also influence the sex of the filarial parasites (Bandi et al., 1998; Bandi et al.,

1999).Filariae rendered free of Wolbachia by treatment with tetracycline antibiotics show

inhibition of maturation, survival, and reproduction (Bandi et al., 1999; Casiraghi et al., 2002;

Genchi et al., 1998; Makepeace et al., 2006; Pfarr and Hoerauf, 2006).

Wolbachia are released in large numbers during parasite molts, during production of

microfilariae, and at death of the parasite (Bandi et al., 1999; Taylor, 2003; Taylor et al., 2005).

Wolbachia-associated-molecules (WAMs) have recently been shown to be associated with

inflammation in the parasitized mammalian hosts (Bazzocchi et al., 2000a; Bazzocchi et al.,

2003; Kozek, 2005; Kramer et al., 2005a; Kramer et al., 2005b; McCall, 2005; Morchon et al.,

2004; Taylor, 2003). These inflammatory responses appear to be most profound when the

parasite dies naturally or as a result of anti-filarial drug treatment. In contrast, the death of

parasites rendered free of Wolbachia stimulates less intense inflammation in the host. This

observation has led to treatment strategies for river blindness and lymphatic filariasis that include

pretreatment with tetracycline antibiotics to reduce the burden of Wolbachia, followed by

ivermectin to eliminate the parasites (Makepeace et al., 2006; Taylor, 2003).

Wolbachia surface protein (WSP) stimulates canine neutrophil chemotaxis and IL-8

production (Bazzocchi et al., 2003). Stimulation of canine vascular endothelial cells with WSP

resulted in production of cyclooxygenase-2, 5-lipooxygenase, leukotriene B4, intracellular

adhesion molecules, E-cadherin, and vascular endothelial growth factor (Simon et al., 2008). In

cats, experimental D. immitis infection resulted in production of antibodies against both D.

immitis and Wolbachia within two months of exposure to infective larvae (Morchon et al., 2004).

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Treatment with ivermectin 30 days later to abort D. immitis infection was associated with

disappearance of D. immitis antibodies, but persistence of Wolbachia antibodies. Dogs

experimentally infected with D. immitis and treated with doxycycline in combination with

ivermectin and melarsomine had less pulmonary pathology than dogs treated with melarsomine

alone (Kramer et al., 2008). Taken together, these studies suggest a role for Wolbachia in the

pathogenesis of heartworm disease.

If recently reported experiences with antibiotic treatment of filarial diseases prove to be

predictive of responses in naturally infected animals, then treatment with doxycycline prior to

adulticide therapy in dogs may sterilize D. immitis of Wolbachia, leading to fewer inflammatory

side-effects when the adult worms die. Doxycycline therapy may be especially helpful for cats,

which suffer substantial and life-threatening effects of heartworm disease, but for which

adulticide therapy is associated with high mortality rates and is generally contraindicated.

Elimination of inflammatory Wolbachia organisms may allow cats to coexist more comfortably

with their parasites, even if an actual cure is not feasible. However, it is not currently known

what role Wolbachia plays in the pathogenesis of natural heartworm disease.

The purpose of this study was to determine the contribution of Wolbachia to the

pathogenesis of natural heartworm disease in cats and dogs. Since all D. immitis parasites harbor

Wolbachia, it is difficult to differentiate the effects of the parasite from those of its endosymbiont

bacteria. We hypothesized that animals having sufficient Wolbachia burden to be detected in

pulmonary tissue by immunohistochemistry and/or by PCR would have more severe pulmonary

disease than those with bacteria below the limits of detection. We further hypothesized that

animals that were immunoreactive to pro-inflammatory WSP would have more severe

pulmonary lesions than those that were seronegative for WSP antibodies.

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CHAPTER 3 EXPERIMENTAL DESIGN

Materials and Methods

Animals

Over a two year period (2005-2007), dog (n=90) and cat (n=90) cadavers were collected

from Alachua County Animal Services (ACAS) in Gainesville, Florida following euthanasia for

reasons unrelated to the study. Cadavers were transported to the University of Florida for

necropsy.

For this study, samples were collected from cats and dogs representing three different D.

immitis infection statuses. The heartworm-free (HW-free) groups were composed of 30 cats and

30 dogs that were free of any evidence of current or previous D. immitis infection (seronegative

for D. immitis antigen and antibodies and negative for parasites at necropsy). The heartworm-

exposed (HW-exposed) groups were composed of 30 cats and 30 dogs with evidence of either

larval-stage infection or past infection (seropositive for D. immitis antibodies), but no evidence

of current adult parasite infection (seronegative for D. immitis antigen and negative for parasites

at necropsy). The heartworm-infected (HW-infected) groups were composed of 30 cats and 30

dogs with adult parasites in the heart or pulmonary arteries at necropsy (regardless of serological

status). The severity of pulmonary lesions in each group was correlated with the detection of

Wolbachia proteins or nucleic acid sequences and with the presence of Wolbachia antibodies.

This study was approved by the University of Florida Institutional Animal Care and Use

Committee (IACUC approval number E12).

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Sample Collection

Blood (4 mL) was collected by transthoracic cardiocentesis into EDTA and serum

separator tubes from each animal within 2 hours after death. Aliquots of EDTA whole blood and

serum were stored at -20°C pending analysis.

The heart and lungs were removed and dissected for the detection of D. immitis. Sections

of the right caudal lung lobe were preserved in 10% formalin for histomorphometry and

immunohistochemistry. Additional sections of fresh lung tissue were stored at -70°C for PCR

analysis. Any recovered D. immitis were preserved in 70% ethanol at room temperature for

immunohistochemistry and PCR.

Feces were collected from the large intestine of each animal and stored at 4°C pending

flotation analysis.

Serology

Cat and dog EDTA whole blood samples were tested by ELISA for D. immitis antigen

(Feline SNAP® Heartworm Antigen Test Kit, Canine SNAP® Heartworm Antigen Test Kit;

IDEXX Laboratories Inc., Westbrook, ME, USA; IDEXX). Feline serum samples were also

screened for D. immitis antibodies with a lateral flow immunoassay (HESKA Solo StepTM FH,

Heska Corporation, CO, USA). These serological results were used to aid in the selection of

cadavers for dissection. Dirofilaria immitis antibody titers and Wolbachia surface protein (WSP)

antibody titers were determined for both cats and dogs as described by Morchon et al., (2004).

Cat serum samples were also tested for FIV antibodies and FeLV antigen by ELISA (PetChek

ELISA, IDEXX Laboratories Inc.)

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Parasitology

Dirofilaria immitis collected at necropsy were counted and examined with a dissecting

microscope to determine sex and age (adult, juvenile, or unknown) of recovered worms. Male

adult worms were identified by their cork-like tails.

Several common parasites of cats and dogs migrate through the lungs as part of their life

cycle and can induce damage that is indistinguishable from that induced by HW infection. To

control for this, feces were examined for ova and larvae. Feces (approximately 1 g) were mixed

with sodium nitrate solution (1.200 specific gravity; Fecasol Solution, Evsco Pharmaceuticals,

Buena, NJ, USA) in a standardized flotation device (Fecalyzer, Evsco Pharmaceuticals),

incubated with a cover slip for 15 minutes, and then evaluated qualitatively by light microscopy

at 100X for the presence of any ova or larvae.

Pulmonary Histologic Morphometry

Formalin-fixed and paraffin-embedded 5 µm sections of tissue collected from the right

caudal lung lobe were stained with hematoxylin and eosin. Digital images were acquired at 100-

400X magnification (Q-Capture software, Q-Imaging Corp, Burnaby, BC, Canada). For each

animal, 10 arterioles, 10 bronchioles, five areas of interstitium, and five areas of alveoli were

assessed. Arteriolar and bronchiolar wall areas were quantified in two-dimensional cross-

sections of the structures as a measure of wall thickness using computer software (Image J,

National Institutes of Health, Bethesda, Maryland, USA). Wall area was calculated by digitally

tracing the entire external and luminal surfaces of the structures and then subtracting the luminal

area from the total area as described in Browne et al. (2005). Pulmonary arteriolar occlusive

hypertrophy was defined as an arteriolar wall that exceeded 95% of the total area, resulting in a

reduced luminal area < 5% of the vessel area.

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In addition to morphometric evaluation of arteriolar and bronchiolar thickness, pulmonary

tissues were scored for severity of histologic lesions on a 0 (normal) to 3 (severely abnormal)

scale. Lesions of the arterioles and bronchioles were scored according to the systems described

in Tables 3-1 and 3-2 and are shown in Figures 3-1 and 3-2. Lesions of the interstitium were

scored based on the frequency of inflammatory cells and the presence of edema within alveolar

septa, whereas lesions of the alveoli were scored according to the relative quantity of smooth

muscle nodules within the tips of alveolar septa as described in Table 3-3 and shown in Figures

3-3 and 3-4. When various components of a structure had different severity grades, the score was

based on the predominant lesion. Mean pulmonary lesion scores for each animal were calculated.

For determination of lesion prevalences, mean scores ≤ 1 were considered to be free of lesions

and mean scores > 1 were considered to have lesions.

Wolbachia DNA Detection

DNA was extracted from lung tissue (40 mg) and parasite tissue (up to 52 µg) using a

commercial kit (DNeasy Tissue Kit, Qiagen Inc., Qiagen Inc.). DNA quality was confirmed by

spectrophotometry and amplification of the ubiquitous housekeeping gene GAPDH. A

conventional PCR assay amplifying a region of the 16S-23S intergenic spacer gene (5’CTG

GGG ACT ACG GTC GCA AGA C 3’ forward; 5’CTC CAG TTT ATC ACT GGA AGT T 3’

reverse) common to Ehrlichia spp., Anaplasma spp., Neorickettsia risticii, and Wolbachia was

performed as described by Lappin et al. (2004). For amplicons of the appropriate size, genetic

sequencing was performed (Macromolecular Resources, Colorado State University, Fort Collins,

CO), and the resultant sequences analyzed by comparison to sequences in GenBank. BLAST

software (National Institutes of Health, NCBI website) was used to confirm the species source of

amplification products. Samples were considered positive for Wolbachia only if PCR assay

results were confirmed by genetic sequencing.

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Wolbachia Immunohistochemistry

Lung tissues and D. immitis parasites were assessed for Wolbachia antigen (WSP) by

immunohistochemistry as described by Kramer et al. (2003). Formalin-fixed paraffin-embedded

5 µm lung sections and heartworm sections were affixed to polylysinated slides. The sections

were incubated in 3% H2O2 for 10 minutes at room temperature. The slides were then treated

with bovine serum albumin and fetal calf serum with phosphate buffered saline (PBS) for 15

minutes. Wolbachia antigens were detected by incubation with rabbit anti-WSP polyclonal

antibody diluted at 1:200 in PBS for 20 minutes followed by anti-rabbit IgG-streptavidin

complex (LSAB2 kit, DAKO, Italy). Color was developed with biotin-horseradish peroxidase,

and sections were counterstained with hematoxylin (LSAB2 kit, DAKO, Italy). Slides were

reviewed microscopically at 100X and were scored as positive or negative for specific staining.

Statistical Analyses

Animals were grouped by heartworm-infection status (HW-free, HW-exposed, HW-infected)

and by Wolbachia status (DNA-, protein-, or antibody-positive or negative) for statistical

analyses. Correlation between the magnitude of D. immitis antibody titers and WSP antibody

titers was measured with the Pearson’s r test (SigmaStat 3.5 statistical analysis software, SPSS

Inc., Chicago, IL, USA). Chi-square analysis was used to compare prevalences of parasitism and

pulmonary lesions when n ≥ 5 and the Fisher exact test was used to compare prevalences when

any group n < 5 (Epi InfoTM 3.5.1, Centers for Disease Control, Atlanta, Georgia). The mean

values of the pulmonary lesion scores were compared using the Wilcoxon rank rum test.

Differences were considered significant when p<0.05.

25

Table 3-1. Pulmonary arteriolar lesion scoring system. Lumen Intima Muscularis Adventitia

Contents Endothelial and Subendothelial

Changes Thickness of Smooth Muscle

Thickness and Contents

Grade 0 Blood Flattened endothelium and thin

intima 2-4 cells; no occlusion of

lumen Thin and indistinct

Grade 1 Mild margination

of eosinophils Cuboidal endothelium and edema of

subendothelial zone 5-7 cells; muscle thickness and lumen diameter similar

Mild thickening; edema

Grade 2

Margination of eosinophils with

mild fibrin deposition

Folded endothelial surface with edema and eosinophils in

subendothelial zone

Partial occlusion; wall thickness 50-95% of lumen

diameter

Marked thickening; edema; mild increase

in inflammation

Grade 3 Fibrinous

thrombosis with many eosinophils

Severe edema and eosinophil infiltration of subendothelial zone;

dissolution of internal elastic lamina

Occlusive hypertrophy; wall thickness >95% of lumen

diameter

Marked thickening; edema; marked

increase in inflammation; fibrosis

26

Table 3-2. Bronchiolar lesion scoring system.

Lumen Mucosa Submucosa Muscularis

Contents Epithelial Changes

Edema Inflammation Serous Glands Fibrosis Thickness of

Smooth Muscle

Grade 0 None Cuboidal Minimal Minimal Rare None 1-2 cells

Grade 1 Mild

seromucoid secretion

Low columnar with mild

crowding and folding

Mild Mild Mild increase Mild 3-4 cells

Grade 2

Moderate seromucoid

secretion with

inflammation

Tall columnar

with moderate

crowding and folding

Moderate Moderate Moderate increase (nesting)

Moderate 5-6 cells

Grade 3

Severe seromucoid

secretion with

inflammation

Tall columnar

with severe crowding and

folding

Severe Severe Severe increase

(circumferential) Severe >6 cells

27

Table 3-3. Pulmonary interstitial and alveolar smooth muscle lesion scoring system. Interstitium Alveolar Smooth Muscle Grade 0 Normal Normal

Grade 1 Increased inflammatory cells in capillaries/mild thickening within alveolar septa

Nodules Present

Grade 2 Moderate thickening Increase in nodules and size; mild bands

Grade 3 Severe thickening, edema, increased eosinophils within alveolar septa

Large nodules; bands and branching

28

Figure 3-1. Representative feline pulmonary tissues demonstrating lesion scores in arterioles. Scores were assigned based on descriptions in Table 3-1. Grade 0 represented normal tissue (arrow pointing to red blood cells in open lumen area) and Grade 3 was considered the most severe lesion. There is a slight increase in smooth muscle in the Grade 1 arteriole (arrow). Edema (arrow) and the hypertrophied smooth muscle are present in the Grade 2 arteriole. The Grade 3 arteriole is displaying occlusive hypertrophy with no visible lumen (wall area >95% of total area) (arrow) (H&E stain, bar=100 µm).

Grade 0 Grade 1

Grade 2 Grade 3

29

Figure 3-2. Representative feline pulmonary tissues demonstrating lesion scores in bronchioles.

Scores were assigned based on descriptions in Table 3-2. Grade 0 represented normal tissue and Grade 3 was considered the most severe lesion. The muscularis in a normal bronchiole has one to two smooth muscle layers (arrow pointing to smooth muscle cell). The epithelium is cuboidal and not crowded. In a Grade 1 bronchiole, the epithelium begins to crowd (arrow), whereas Grade 2 and 3 have compressed nuclei. Serous glands become dominant in Grade 2 and are located in the submucosa and in a Grade 3 the glands become circumferential (arrow) (H&E stain, bar=100 µm).

Grade 0 Grade 1

Grade 2 Grade 3

30

Figure 3-3. Representative feline pulmonary tissues demonstrating lesion scores in alveolar

smooth muscle. Scores were assigned based on descriptions in Table 3-3. Grade 0 represented normal tissue and Grade 3 was considered the most severe lesion. Smooth muscle nodule (arrow A) size varies with grades. Banding develops in Grade 2, and nodules and bands become undistinguishable in Grade 3 (arrow B) (H&E stain, bar=100 µm).

Grade 0 Grade 1

Grade 2 Grade 3

A

B

31

Figure 3-4. Representative feline pulmonary tissues demonstrating lesion scores in interstitium.

Scores were assigned based on descriptions in Table 3-3. Grade 0 represented normal tissue and Grade 3 was considered the most severe lesion. The arrow is pointing to the thickening within alveolar septa and inflammatory cells within the capillaries. (H&E stain, bar=100 µm).

Grade 0 Grade 1

Grade 2 Grade 3

32

CHAPTER 4 RESULTS

Statistical Analyses

Serology

This study was not designed to report the sensitivity or specificity of the antibody or

antigen tests because serology results were used to select specimens for necropsy.

There was a positive but weak correlation between the magnitude of D. immitis antibody

titers and WSP titers in cats (r=0.57, p<0.001) and in dogs (r=0.39, p<0.001) (Figures 4-1, 4-2).

Antibodies against WSP were detected in a majority of HW-infected cats and dogs and in

variable proportions of animals in other HW-status groups (Table 4-1). Of the 90 cats in the

study, 10 had FIV antibodies (five from the HW-infected group and five from the HW-exposed

group) and one was infected with FeLV (from the HW-exposed group).

Parasitology

Low D. immitis intensities were common in cats in the HW-infected group

(mean=1.7±0.2 worms, range=1-6). Of the 51 D. immitis collected from the 30 HW-infected

cats, most were female (35/51, 69%). Only five cats had both male and female worms. Higher

worm intensities were common in dogs in the HW-infected group (mean=36.3±5.9 worms,

range=3-133). Of the 1,090 D. immitis collected from the 30 HW-infected dogs, most were

female (606/1090, 56%). All dogs had both female and male worms.

Ancylostoma spp. were common in both cats (52%) and dogs (61%). Other

gastrointestinal parasites identified included whipworms (0% of cats, 27% of dogs) and ascarids

(13% of cats, 9% of dogs). There were no significant associations between the overall presence

of pulmonary lesions in cats or dogs and the presence of Ancylostoma spp. or ascarids, two

parasites with a pulmonary migratory phase in their life-cycle (p>0.09) (Table 4-2). When

33

animals were grouped according to subtype of lesion (arteriolar, bronchiolar, alveolar smooth

muscle, or interstitial), cats with alveolar lesions were less likely to have ascarids than those

without lesions (p=0.02) and dogs with interstitial lesions were more likely to have Ancylostoma

spp. than dogs without lesions (p=0.02).

Detection of Wolbachia

Wolbachia DNA was amplified from adult D. immitis collected from 26 of 30 HW-

infected cats, and from D. immitis collected from all of the HW-infected dogs. Three out of the

four Wolbachia-negative D. immitis samples from the HW-infected cats were GAPDH negative,

indicating inadequate DNA quality. WSP was detected by immunohistochemistry in D. immitis

specimens from all cats and dogs. In general, Wolbachia DNA or WSP was identified most

commonly in the lungs of HW-infected animals, but was also present in variable proportions of

lung samples in the other HW-infection status groups (Table 4-1).

Pulmonary Histologic Morphometry

As expected, pulmonary lesions were more common in HW-infected animals than in HW-free

animals (Table 4-1). Pulmonary arteriolar occlusion was more common in HW-infected cats

(57%) (p=0.003) than in HW-infected dogs (17%). In addition, HW-infected cats with occlusion

had more affected arterioles (43±6%) than did HW-infected dogs with occlusion (12± 2%)

(p=0.007). In cats, occlusive hypertrophy was identified most commonly in the HW-infected

group, but was also present in the HW-free and HW-exposed groups. In contrast, arteriolar

occlusion in dogs was present only in the HW-infected group. Although there were differences in

the proportions of cats affected by arteriolar occlusion, there were no significant differences in

the severity of occlusion among the groups. The percentage of occluded vessels in the HW-

exposed group (44±11%) was similar to that of the HW-infected group, and the HW-free group

(28±7%) (p=0.31). Although pulmonary lesions were most common in HW-infected animals,

34

there was no clear additive effect on the prevalence of lesions when either Wolbachia DNA/WSP

was detected in pulmonary tissue or when circulating Wolbachia antibodies were detected (Table

4-3). Similarly, there were no significant differences in the magnitude of pulmonary lesion

scores within each HW-infection status group regardless of whether Wolbachia DNA/WSP or

antibodies were detected (Figures 4-3 to 4-6).

35

Table 4-1. Prevalence of Wolbachia surface protein (WSP) antibodies, Wolbachia DNA, WSP, and pulmonary lesions in cats and dogs with no evidence of D. immitis infection (HW-free), evidence of either larval-stage infection or past infection (HW-exposed), or with adult D. immitis in the heart or pulmonary arteries (HW-infected). Values with different superscript letters within a row are significantly different (p<0.05).

Cats HW-free HW-exposed HW-infected

WSP antibodies 3 (10%)a 10 (33%)b 18 (60%)c Wolbachia DNA in lung 7 (23%) 12 (40%) 12 (40%) WSP in lung 4 (13%) 5 (16%) 10 (33%)

Arteriolar lesions 10 (33%)a 16 (53%) 19 (63%)b Arteriolar occlusive hypertrophy 8 (27%)a 9 (30%) a 17 (57%)b Bronchiolar lesions 8 (27%) 7 (23%) 10 (33%)

Interstitial lesions 14 (47%)a 27 (90%)b 27 (90%)b

Alveolar smooth muscle lesions 6 (20%)a 8 (27%)a 18 (60%)b Dogs

WSP antibodies 14 (46%)a 23 (76%)b 20 (66%)

Wolbachia DNA in lung 0 (0%)a 1 (3.4%)a 14(46%)b

WSP in lung 7 (23%) 4 (13%)a 12 (40%)b

Arteriolar lesions 3 (10%)a 12 (40%)b 18 (60%)b Arteriolar occlusive hypertrophy 0 (0%)a 0 (0%)a 5 (17%)b

Bronchiolar lesions 7 (23%)a 24 (80%)b 30 (100%)c

Interstitial lesions 12 (40%)a 20 (67%)b 30 (100%)c

Alveolar smooth muscle lesions 5 (17%)a 13 (43%)b 16 (53%)b

36

Table 4-2. The prevalence of Ancylostoma spp. and ascarids in cats and dogs with pulmonary lesions. Values with differ superscript letters within a row are significantly different (p<0.05).

Cats (n=90) Lesions No Lesions Ancylostoma spp. 40/81 (49%) 7/9 (78%)

Ascarids 9/81 (11%) 3/9 (33%) Dogs (n=90)

Ancylostoma spp. 48/75 (64%) 7/15 (47%) Ascarids 7/76 (9.2%) 1/13 (7.7%)

37

Table 4-3. Frequency of pulmonary lesions in cats and dogs with and without Wolbachia. Wolbachia-negative animals were free of detectable Wolbachia DNA, WSP, and WSP antibodies. Wolbachia-positive animals were positive for at least one of these tests for Wolbachia. There were no significant differences in the frequency of lesions in the Wolbachia-negative groups vs. the Wolbachia-positive groups (p>0.05).

Cats Wolbachia-negative (n=37) Wolbachia-positive (n=53) Arteriolar lesions (n=45, 50%)

HW-free (n=10) 7 (37%) 3 (27%) HW-exposed (n=16) 4 (33%) 12 (66%) HW-infected (n=19) 3 (50%) 16 (66%)

Arteriolar occlusive hypertrophy (n=34, 38%) HW-free (n=8) 5 (26%) 3 (27%) HW-exposed (n=9) 4 (33%) 5 (27%) HW-infected (n=17) 3 (50%) 14 (58%)

Bronchiolar lesions (n=25, 28%) HW-free (n=8) 6 (32%) 2 (18%) HW-exposed (n=7) 3 (25%) 4 (22%) HW-infected (n=10) 2 (33%) 8 (33%)

Alveolar smooth muscle lesions (n=32, 36%) HW-free (n=6) 4 (21%) 2 (18%) HW-exposed (n=8) 5 (42%) 3 (16%) HW-infected (n=18) 4 (67%) 14 (58%)

Interstitial lesions (n=68, 76%) HW-free (n=14) 8 (42%) 6 (54%) HW-exposed (n=27) 11 (92%) 16 (88%) HW-infected (n=27) 6 (100%) 21 (87%)

Dogs Wolbachia-negative (n=18) Wolbachia-positive (n=72) Arteriolar lesions (n=33, 37%)

HW-free (n=3) 1 (8%) 2 (11%) HW-exposed (n=12) 1 (100%) 11 (44%) HW-infected (n=18) 0 (0%) 18 (62%)

Arteriolar occlusive hypertrophy (n=5, 5.6%) HW-free (n=0) 0 (0%) 0 (0%) HW-exposed (n=0) 0 (0%) 0 (0%) HW-infected (n=5) 1 (100%) 4 (13%)

Bronchiolar lesions (n=61, 68%) HW-free (n=7) 3 (25%) 4 (22%) HW-exposed (n=24) 5 (100%) 19 (76%) HW-infected (n=30) 1 (100%) 29 (100%)

Alveolar smooth muscle lesions (n=34, 38%) HW-free (n=5) 4 (33%) 1 (5.5%) HW-exposed (n=13) 3 (60%) 10 (40%) HW-infected (16) 0 (0%) 16 (55%)

Interstitial lesions (n=62, 69%) HW-free (n=12) 5 (42%) 7 (38%) HW-exposed (n=20) 2 (40%) 18 (72%) HW-infected (n=30) 1 (100%) 29 (100%)

38

R2 = 0.181

0.00

0.20

0.40

0.60

0.80

1.00

1.20

1.40

1.60

0 20 40 60 80 100

Wolbachia surface protein absorbance

He

art

wo

rm a

nti

bo

dy

ab

so

rba

nc

e

Figure 4-1. Feline D. immitis antibody titer versus Wolbachia surface protein (WSP) antibody titer in all cats (n=90). There was a positive but weak correlation between the magnitude of the titers (p>0.001).

39

R2 = 0.1591

0.00

0.50

1.00

1.50

2.00

2.50

0.00 0.20 0.40 0.60 0.80 1.00 1.20 1.40 1.60

Wolbachia surface protein absorbance

He

art

wo

rm a

nti

bo

dy

ab

so

rba

nc

e

Figure 4-2. Canine D. immitis antibody titer versus Wolbachia surface protein (WSP) antibody titer in all dogs (n=90). There was a positive but weak correlation between the magnitude of the titers (p>0.001).

40

Figure 4-3. Severity of pulmonary lesions in cats with or without detectable Wolbachia DNA and/or Wolbachia surface protein in pulmonary tissues. Each D. immitis infection status group included 30 cats, and histologic lesions were scored on a 0-3 scale. Boxes represent the middle 50th percentile, bars represent the 25th and 75th percentiles, and dots represent outlier data points. There were no significant differences in median pulmonary lesion scores between groups (p>0.05).

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5

Wolbachia Neg Wolbachia Pos Wolbachia Neg Wolbachia Pos Wolbachia Neg Wolbachia Pos

Arteriolar lesion score

HW-free HW-exposed HW-infected

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5


Bronchiolar lesion score


0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5


Alveolar lesion score


0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5


Interstitial lesion score


41

Figure 4-4. Severity of pulmonary lesions in dogs with or without detectable Wolbachia DNA and/or Wolbachia surface protein in pulmonary tissues. Each D. immitis infection status group included 30 dogs, and histologic lesions were scored on a 0-3 scale. Boxes represent the middle 50th percentile, bars represent the 25th and 75th percentiles, and dots represent outlier data points. There were no significant differences in median pulmonary lesion scores between groups (p>0.05).

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5



HW-free HW-Exposed HW-infected

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5




0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5




0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5




42

Figure 4-5. Severity of pulmonary lesions in cats with or without detectable circulating Wolbachia surface protein antibodies (WSP Ab). Each D. immitis infection status group included 30 cats, and histologic lesions were scored on a 0-3 scale. Boxes represent the middle 50th percentile, bars represent the 25th and 75th percentiles, and dots represent outlier data points. There were no significant differences in median pulmonary lesion scores between groups (p>0.05).

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5

WSP Ab Neg WSP Ab Pos WSP Ab Neg WSP Ab Pos WSP Ab Neg WSP Ab Pos



0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5



HW-free HW-exposed

HW-infected

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5

WSP Ab-Neg WSP Ab Pos WSP Ab Neg WSP Ab Pos WSP Ab Neg WSP Ab Pos



0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5



HW-fee

HW-exposed HW-infected

43

Figure 4-6. Severity of pulmonary lesions in dogs with or without detectable circulating

Wolbachia surface protein antibodies (WSP Ab). Each D. immitis infection status group included 30 dogs, and histologic lesions were scored on a 0-3 scale. Boxes represent the middle 50th percentile, bars represent the 25th and 75th percentiles, and dots represent outlier data points. There were no significant differences in median pulmonary lesion scores between groups (p>0.05).

0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5




0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5


Brochiolar lesion score


0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5




0.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5




44

CHAPTER 5 DISCUSSION

In this study, we identified the presence of Wolbachia in HW-defined groups of cats and

dogs to determine if there was an association between the bacterium and the severity of

pulmonary lesions characteristic of heartworm disease. Although pulmonary lesions were most

severe in animals with adult D. immitis there was no clear additive effect when Wolbachia was

present in sufficient amounts for detection in the lungs or when immunoreactivity against

Wolbachia was detected.

Since all D. immitis carry Wolbachia, it is difficult to separate the effect of the parasite

from the effect of its endosymbiont. In both cats and dogs, the frequency of detection of

Wolbachia in the lung of HW-exposed animals was intermediate between that of HW-free

animals and HW-infected animals. Similarly, the frequency and severity of pulmonary lesions in

HW-exposed animals also tended to be intermediate between that of HW-free animals and HW-

infected animals. Although these findings could suggest that the presence of Wolbachia in

amounts sufficient for detection in the lungs or the immune responses to Wolbachia antibodies

are associated with increased pulmonary disease, it is not possible to completely rule out an

effect of the parasite as well.

Wolbachia was detected in animals with no evidence of adult D. immitis. Studies have

demonstrated bacteria are released in large amounts when the parasite dies (Kramer et al., 2008;

Saint Andre et al., 2002). Therefore it is possible the HW-exposed group of animals had previous

D immitis infection that deposited WSP or that had initiated an anti-WSP response (Morchon et.

al., 2004).

When interpreting the pathological findings of the pulmonary tissues, it was important to

account for the potential contribution by migration of immature intestinal parasites such

45

Ancylostoma spp. and ascarids. A Baermann’s test was not conducted and it is possible other

parasites, such as lungworms, could also contribute to the pulmonary disease but escaped

detection (Patton and McCracken, 1991).

The lack of clear evidence for a role of Wolbachia in heartworm disease creates a dilemma

for veterinarians treating cats and dogs in D. immitis-endemic areas. Although the indiscriminant

use of antibiotics should be avoided, many clinicians prescribe doxycycline based on the

favorable responses observed in human filarial diseases and on promising results from the first

published studies of doxycycline use in D. immitis-infected dogs.

46

CHAPTER 6 CONCLUSIONS

Our study design was did not demonstrate that the presence of Wolbachia or WSP

antibodies was associated with increased pulmonary disease associated with D. immitis infection.

Future studies could further address this potential association by treating HW-exposed and HW-

infected animals with the anti-Wolbachia antibiotic doxycycline and observing the evolution of

pulmonary disease. Alternatively, studies could be performed to compare disease development in

animals experimentally infected with natural D. immitis versus D. immitis rendered free of

Wolbachia by doxycycline treatment.

47

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BIOGRAPHICAL SKETCH

Patricia Ann Dingman was born in 1982, in Angeles City, Philippines. She attended

elementary school in Okinawa, Japan, and Plattsburgh, New York, and finished high school in

Coudersport, Pennsylvania. Patricia graduated from the State University of New York Oswego

with a B.S. in zoology in 2006. She received her M.S. in veterinary medical sciences in 2009

from the University of Florida and plans to continue her education in veterinary medicine.

When Patricia volunteered for Dr. Levy, she had the opportunity to investigate disease

transmission in cats and dogs following Hurricane Katrina and attend monthly trap-neuter-return

clinics for feral cats (Operation Catnip). These experiences have inspired her to continue

pursuing veterinary-related research in cats.

association of wolbachia with heartworm disease in

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