Seroprevalence of Coxiella burnetii in domesticated and feral cats in eastern Australia

Veterinary Microbiology 177 (2015) 154–161

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Seroprevalence of Coxiella burnetii in domesticated and feral cats in eastern Australia Amanda J. Shapiro a, Katrina L. Bosward b, Jane Heller c, Jacqueline M. Norris a,* a

Faculty of Veterinary Science, Building B14, The University of Sydney, Sydney, NSW 2006, Australia Faculty of Veterinary Science, The University of Sydney, PMB 4003, Narellan, NSW 2567, Australia c School of Animal & Veterinary Sciences, Charles Sturt University, Locked Bag 588, Wagga Wagga, NSW 2678, Australia b

A R T I C L E I N F O

A B S T R A C T

Article history: Received 12 October 2014 Received in revised form 12 February 2015 Accepted 15 February 2015

The seroprevalence of Coxiella burnetii (C. burnetii) in cats in eastern Australia is unknown, and the risk of transmission from cats to humans is undetermined. This study aimed to determine the exposure of cats to C. burnetii in four distinct cat subpopulations. An indirect immunofluoresence assay (IFA) and an Enzyme-linked immunosorbent assay (ELISA) used for detection of anti-C. burnetii antibodies in humans were adapted, verified for use on feline serum, and compared. Cat serum samples (n = 712) were tested with IFA from four subpopulations [cattery-confined breeding cats, pet cats, feral cats and shelter cats]. The proportions of seropositive cats were; cattery-confined breeding cats (35/376, 9.3%), pets (2/198, 1%), feral cats (0/50), shelter cats (0/88). The significant variables in C. burnetii seropositivity were cattery-confined breeding cat subpopulation and sterilisation status, with infected cats 17.1 (CI 4.2–70.2; P < 0.001) times more likely to be cattery-confined breeding cats and 6.00 (CI 2.13–16.89; P < 0.001) times more likely to be entire than sterilised. ELISA was used on 143 of 712 sera tested with IFA, and the Cohen’s Kappa coefficient of 0.75 indicated 92.2% agreement between the two assays. These results confirm that Australian cats have been exposed to C. burnetii and that a higher seroprevalence of C. burnetii is seen amongst cattery-confined breeding cats. Cat breeders and veterinary personnel involved in feline reproductive procedures may be at higher risk of exposure to C. burnetii. ß 2015 Elsevier B.V. All rights reserved.

Keywords: Coxiella burnetii Q fever Serological assays Cats ELISA Indirect immunofluorescence (IFA)

1. Introduction Q fever is an important worldwide zoonosis caused by the bacterium Coxiella burnetii. For the 40% of primary infections that are symptomatic and clinically polymorphic in humans, illness can be acute or chronic with the potential for serious and debilitating sequelae such as endocarditis, post-Q fever fatigue syndrome and recrudescent granulomatous lesions in bone or soft tissue (Babudieri, 1959; Marrie, 1990).

* Corresponding author. Tel.: +61 2 93517095. E-mail address: [email protected] (J.M. Norris). http://dx.doi.org/10.1016/j.vetmic.2015.02.011 0378-1135/ß 2015 Elsevier B.V. All rights reserved.

The causative bacterium has a potentially large, seemingly asymptomatic reservoir encompassing wild and domestic mammals, birds and arthropods (Babudieri, 1959; Baca and Paretsky, 1983). It has two diverse aspects to its lifecycle; a metabolically active form that obligately replicates within the monocyte/macrophage cell lineage and an inactive but environmentally resilient form that travels to new hosts. C. burnetii localises in the uterus and mammary glands of mammals with products of conception containing the highest concentration of organisms (Babudieri, 1959; Welsh et al., 1958). Humans acquire infection from animals directly or indirectly mainly via inhalation of contaminated aerosols (Babudieri, 1959; Maurin and Raoult, 1999). Infected domestic ruminants

A.J. Shapiro et al. / Veterinary Microbiology 177 (2015) 154–161

including cattle, sheep and goats are the most commonly reported sources of infection for humans however a broader view on the potential of other animal reservoirs to impact on public health worldwide has developed more recently (Angelakis and Raoult, 2010; Woldehiwet, 2004). Diagnosing an infection with C. burnetii in humans primarily relies on serological tests due to their higher sensitivity compared with nucleic acid amplification on blood and tissue samples which is often reserved for acute infections. The presence of antibodies to C. burnetii provides evidence of recent or past exposure to the bacteria. The three main serological tests used in human medicine are complement fixation test (CFT), indirect immunofluorescence assay (IFA) and enzyme-linked immunosorbent assay (ELISA) (Fournier et al., 1998). In animals, CFT, which has many limitations, is the only validated serological test available (Rousset et al., 2010). It is the least sensitive method, regularly failing to detect antibodies in sheep and goats, has difficulty diagnosing recent infections (Arricau-Bouvery and Rodolakis, 2005) and is inferior to both IFA and ELISA in humans (Cowley et al., 1992; Peter et al., 1985) which can evaluate classspecific IgG, IgM and IgA antibodies (Peter et al., 1988). In the CFT used for animals, class-specific immunoglobulins are simultaneously tested, but the end titre does not reflect which immunoglobulin was detected. A recent outbreak of Q fever in veterinary personnel in a Sydney small animal veterinary hospital following a caesarean section in a healthy queen highlighted the potential role domestic cats may play as reservoirs of infection (Kopecny et al., 2013; Maywood, 2011). Establishing the seroprevalence of C. burnetii in different subpopulations of cats is essential in order to obtain an overview of recent or past exposure of cats to this bacterium to determine the risk cats may pose to veterinary personnel, professional cat breeders and pet owners and to gauge the likelihood of it being an agent of disease in cats. Seroprevalence studies in cat populations report great variability in the prevalence of C. burnetii in different geographical locations and time periods. Additionally, the methodology used has been highly variable even when the same assay type is used. This study aimed to (1) develop a serological method for detecting antibodies to C. burnetii in domestic cats (Felis domesticus) by adaptation and optimisation of commercially available IFA and ELISA kits developed for use on human serum and (2) determine the seroprevalence of C. burnetii in a variety of cat subpopulations to determine the potential risk to population health using a retrospective observational study. 2. Materials and methods 2.1. Sample population Convenience sampling resulted in the categorisation of cats into four distinct cat subpopulations; (a) pet cats (n = 198); (b) cattery-confined breeding cats (n = 376); (c) cats entering, or housed in, animal shelters or council pounds (n = 88) and (d) feral cats (n = 50). All catteryconfined breeding cats were located in eastern Australia, predominantly in New South Wales (NSW) (n = 292) with

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remaining cats from Victoria (VIC) (n = 26), Tasmania (TAS) (n = 12) and Queensland (QLD) (n = 46). Information regarding breed, age, gender (including neuter status), physical findings, medical history and housing type (exclusively indoors, exclusively outdoors or combination) was obtained from medical records (pet cats) or cattery owners (cattery-confined breeding cats). For feral cats, the gender, estimated age and abnormal physical findings were recorded. A separate group of cattery-confined breeding cats from a single cattery (n = 27) including the index cat at the centre of the Q fever outbreak at a small animal hospital in Sydney (Kopecny et al., 2013) and 26 cats residing within the same cattery were used as an essential part of the optimisation process but excluded from calculations of seroprevalence. The serum samples from pet cats (n = 198) were archived samples used in previous studies (Bell et al., 2006; Norris et al., 2007), collected when cats presented to veterinary clinics [University Veterinary Teaching Hospital, Sydney (UVTH-S), Paddington Cat Hospital (PCH) and Concord Animal Hospital]. The archived samples comprised residual serum taken from cats during routine veterinary services for disease investigation or health checks. Serum samples from 376 cattery-confined breeding cats were tested. Of these, 345 were from samples previously used in a FIV prevalence study (Norris et al., 2007) from catteries in the greater Sydney region. Permission to repeat sample collections from certain cats (n = 7) was granted by cattery owners. Simultaneously, samples from new cats (n = 24) at these catteries were collected and an additional subset of samples from a cattery not previously sampled was collected (n = 7). Therefore, the total number of cattery-confined cats sampled was 376 with 345 cat sera from archived studies and 31 sera collected following initial screening. The shelter cat subpopulation consisted of cats entering or housed within animal shelters and council pounds (n = 88). Forty-four of these serum samples came from cats older than 18 months of age, either housed or entering specific animal shelters in NSW that were part of another study investigating feline heartworm prevalence (Animal ethics approval number 5072.04-11 SWAHS AEC) and were donated for the current study (n = 44). The remaining 44 samples were from cats presenting as strays to a council facility in south-western Sydney. The feral cat population (n = 50) comprised two separate colonies forming part of a FIV prevalence study (Norris et al., 2007). Samples had been collected from one colony of feral cats residing near a piggery in Menangle in south-western Sydney and the second colony of feral cats living around the University of Sydney’s Camperdown campus was the focus of a ‘neuter and release’ campaign. 2.2. Sample collection Whole blood (0.5–3 mL) from each cat was collected into serum separator tubes (BD Vacutainer SST II, Sydney, Australia) and centrifuged at 8000  g for 10 min. Serum was harvested and stored in 300 mL aliquots at 20 8C until analysed.

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2.3. Serological testing 2.3.1. Complement fixation testing (CFT) Serum samples were submitted to the State Veterinary Diagnostic Laboratory at the Elizabeth Macarthur Agricultural Institute, Menangle, NSW, Australia for CFT. The CFT protocol used by this laboratory (Corner and Alton, 2004) is directed against phase I (IgM) and phase II (IgG1 and IgG3) antibodies but the results of the assay are unable to differentiate between the different immunoglobulins. Samples with an anti-phase I or II C. burnetii titre greater than or equal to 8 were considered positive. Serum samples from the index cat at the centre of the Q fever outbreak in a small animal practice in Sydney (Kopecny et al., 2013) with a CFT titre of 1/32 became the positive control sample for use in the IFA and ELISA optimisation and verification assays. 2.3.2. Indirect immunofluorescent antibody assay (IFA) A modification of a commercial human C. burnetii phase I and II specific IFA IgG kit (Vircell, Spain) was used to detect feline IgG antibodies to phase I and phase II C. burnetii (Nine Mile strain). The kit consisted of slide wells coated with C. burnetii, Nine Mile strain (ATCC 616-VR), grown in MRC-5 cells, formaldehyde inactivated and acetone fixed and the assay was run according to manufacturer’s instructions with adjustments for use with feline serum. Anti-human IgG fluorescein isothiocyanate (FITC) conjugate solution from the kit was applied to positive and negative control wells and anti-feline IgG FITC conjugate solution (CJ-F-FELG-AP-10ML, Veterinary Medical Research and Development [VMRD]) to the remaining wells. 2.3.2.1. Verification and optimisation of the IFA. The human positive and negative control samples served as the reference markers in identifying positive and negative results. The presence of bright apple green fluorescence of cocco-bacillar morphology and lack of background fluorescence seen on the positive control was used to identify positive samples, while the total absence of fluorescence seen in the negative control identified negative samples. Serum from the index case served as the positive cat control as it compared equally in immunofluorescence reaction to the human positive control. All outbreak cattery samples (n = 27) used in the verification of the assay were compared to the index cat positive control and human positive control samples and defined as either positive or negative accordingly. The slides of all samples used in the verification of the assay were analysed by two authors independently. The use of different blocking agents to reduce nonspecific background fluorescence was assessed. Trials were conducted to compare skim milk powder (SMP) (Coles, Instant Skim Milk Powder) and bovine serum albumin (BSA) (Bovine Serum Albumin Solution, 7.5% in DPBS, A8412, Sigma–Aldrich) as non-specific blocking agents. SMP was tested at concentrations of 2%, 3%, 5% and 6%, and compared to 1% BSA. The 5% SMP in PBS was highly effective as a non-specific blocking agent in feline serum and used for all subsequent testing via IFA.

Further verification of the IFA assay required determination of potential cross-reactivity of other feline antibodies to the C. burnetii antigen. The most likely organisms to cause cross reactivity were Bartonella spp. (LaScola and Raoult, 1996), therefore a Bartonella IFA kit (Vircell, Spain) detecting antibodies against B. henselae and B. quintana was adapted for use on feline serum. Additionally, the manufacturer of the C. burnetii and B. henselae and B. quintana IFA kits (Vircell, Spain) performed in-house IFA cross-reactivity testing between C. burnetii and B. henselae and B. quintana (unpublished data). 2.3.2.2. Determination of cut-off value for IFA. Determination of the cut off value for IFA involved comparison studies between cat samples found positive and negative on CFT, always using the human positive and negative control samples and index case in all experiments. Outbreak cattery samples (n = 27) were initially used in the cut off determination phase. All samples were tested using the manufacturer’s recommended serum dilutions of 1/64, 1/128, 1/256, 1/512 and 1/1024. At titres below 1/256, background fluorescence obscured definitive determination of positive and negative samples. The positive control sample had an end point titre of 1/8192 on both phase I and II. At a cut off value of 1/256, clear distinction between bacterial fluorescence and background fluorescence was observed whereby samples recorded as positive resembled the index cat and human positive control, while negative samples no longer displayed strong background fluorescence. All serum samples were then tested at the optimised dilution (1/256) and diluent (5% SMP in PBS). Control solutions (5 ml each) were applied to slide wells 1 and 2 of the first slide and 5 ml of diluted serum was applied to the remaining slide wells. Slides were incubated in a humid chamber for 30 min at 37 8C, rinsed gently with PBS, immersed in PBS for 10 min and dip-washed in sterile water. Slides were air dried and 5 ml of anti-human IgG FITC conjugate solution was applied to the control wells, and 5 ml of rabbit anti-feline IgG FITC conjugate (VMRD, Pullman, WA, USA) to the remaining wells. Slides were incubated again for 30 min at 37 8C and the washing repeated. The slides were dried, mounted with coverslips and read immediately with a BX60 epifluorescence microscope (Olympus, Melville, NY, USA) at 400 magnification. A sample was ‘seropositive’ for C. burnetii on IFA if it displayed either phase I or phase II (IgG1, IgG2, IgG3) antibodies at a titre of 1/256 or greater. Positive samples were taken to end titre by two fold serial dilutions beginning at 1/256. 2.3.3. Enzyme-linked immunosorbent assay (ELISA) A commercially available ELISA kit for detection of human IgG antibodies against C. burnetii (Henzerling strain Phase II, Panbio, Alere, Australia) was used as per manufacturer’s instructions except that peroxidase-conjugated AffiniPure Goat Anti-cat IgG (Jackson ImmunoResearch Laboratories, Inc., PA) was substituted for the antihuman IgG antibody supplied with the kit. A checkerboard titration experiment to optimise primary antibody and labelled secondary antibody dilutions was performed. Four twofold serial dilutions of the same serum samples used

A.J. Shapiro et al. / Veterinary Microbiology 177 (2015) 154–161

for verification of the IFA assay were made ranging from 1/ 100 to 1/800. Two serum diluents with non-specific blocking activity (1% BSA and 2%, 3%, 5% and 6% SMP) were tested. The secondary antibody, Peroxidase-conjugated AffiniPure Goat Anti-cat IgG (Jackson ImmunoResearch Laboratories, Inc., PA), was titrated at four concentrations (diluted in deionised water); 1/5000, 1/ 7500, 1/10000, 1/12500. For performing the ELISA, sera were diluted 1/100 in 5% SMP and the secondary antibody was used in a concentration of 1/5000. Sample wells were pre-incubated at room temperature for 60 min with 150 ml of 5% SMP in PBS (human control wells were not pre-treated) before removal from the well. Feline serum and human control serum samples were diluted 1/100 in 5% SMP in PBS and supplied sample diluent respectively and then 100 ml was pipetted into respective wells. Plates were covered and incubated at 37 8C for 30 min. Wells were manually washed six times with supplied wash buffer (PBS with 0.05% Tween 20) and 100 ml of either HRP conjugated Anti-human IgG Panbio QFever IgG ELISA (Alere, Australia) for the human control serum wells or Peroxidase-conjugated AffiniPure Goat Anti-cat IgG (Jackson ImmunoResearch Laboratories, Inc., PA) at the optimised dilution of 1/5000 for the feline serum samples was pipetted into sample wells. Wells were covered and incubated at 37 8C for 30 min and then washed six times with the wash buffer supplied in the kit. Following washing, 100 mL Tetramethylbenzidine chromogen was applied to all wells and incubated for 10 min at room temperature. To stop the reaction, 100 ml 1 M Phosphoric acid was applied to all wells and the absorbance read with a dual wavelength spectrophotometer (SPECTRAmax 250, Molecular Devices Corporation, California) at a wavelength of 450 nm and a reference filter of 600 nm within 30 min. The optical density (OD) readings were assessed using two methods; calculating the index value as recommended by the kit manufacturer based on samples from a human subpopulation and the sample/positive control percentage (S/P %). Index values were calculated by dividing sample absorbance by cut-off values. Cut-off values represent the average absorbance of the triplicate of the calibrator wells multiplied by the calibration factor supplied. Index values less than 0.9 were considered negative, those between 0.9 and 1.1 equivocal, and positive samples had index values greater than 1.1. Equivocal samples were repeated and if still equivocal were considered negative. S/P% was calculated according to the formula: S/P% = (OD sample OD negative control)/(OD positive control-OD negative control)  100. Samples with S/P% less than 50% were determined to be negative, samples between 50 and 75% were taken as positive and samples with S/P% greater than 75% were considered strongly positive. 2.4. Data analysis Data were analysed using GenStat 16.1 (VSN International, Hemel Hempstead, UK) and statistical significance was considered at P < 0.05. Repeat collections from the same cats (n = 7) were not included in the data analysis and only the first sample for each cat was used. Seroprevalence

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for each subpopulation are reported with their 95% binomial confidence intervals. Associations between C. burnetii seropositivity and potential risk factors (gender, sterilisation status, age, breed and subpopulation) were assessed using multivariable logistic regression. Initial univariable analyses were performed and variables with P < 0.25 were considered for inclusion in the multivariable models. Three multivariable logistic regression models were constructed for phase I, phase II and phase I or II seropositivity using backwards elimination. Interassay agreement between IFA and ELISA anti-phase II IgG C. burnetii antibodies was measured using Cohen’s kappa coefficient calculated using the GraphPad QuickCalcs http://www.graphpad.com/quickcalcs/kappa1.cfm (accessed March 2014). 3. Results 3.1. Sample population IFA was used to test the total sample population (n = 712). For ELISA, 143/712 sera were selected (catteryconfined (n = 56), pet (n = 41), feral (n = 39) and shelter (n = 7)). CFT tested 71/712 sera (cattery-confined (n = 34), pet (n = 2), feral (n = 35) and shelter (n = 0)). A breakdown of the gender, sterilisation status, breed and age in each subpopulation against phase I and/or phase II seropositivity for all samples examined with IFA is presented in Table 1. 3.2. Serological testing 3.2.1. Indirect immunofluorescent antibody assay (IFA) IFA, which measured both phase I and II IgG antibodies, returned a positive result for seven samples for phase I only, 12 samples for phase II only and 18 samples for both phase I and II. Overall, 37 samples were seropositive on IFA for C. burnetii, with 28 female cats and nine male cats. Of these, 35 cats were cattery-confined breeding cats and two were pet cats. All female cats were entire while five male cats were entire and four were sterilised (Supplemental Table). Of the 35 positive cattery-confined breeding cats, 32 were from NSW, two were from VIC and one from QLD. Positive samples returned antibody titres between 1/256 and 1/16384. All cats tested from the feral and shelter cat subpopulations were seronegative for C. burnetii. Using IFA alone, the seroprevalence of C. burnetii in the cats sampled in eastern Australia (n = 712), were; pet cats 1.0%, (CI 0.12–3.6%), cattery-confined breeding cats 9.3% (CI 6.5–13%), shelter cats 0%, (CI 0–4%) and feral cats 0% (CI 0–7%). Multivariable models were determined for three outcomes of seropositivity; phase I, phase II and phase I or II. Univariable analyses results are shown in Table 2. Although multiple risk factors were considered for each phase outcome, the final models for each resulted in a single risk factor affecting seropositivity. Subpopulation was the only risk factor affecting phase II, with odds of seropositivity greater in cattery-confined than non-cattery-confined cats (OR 4.6, CI 1.6–13.3; P < 0.001) and phase I or II (OR 17.1, CI 4.2–70.2; P < 0.001) seropositivity.

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Table 1 Frequency table representing the variables of gender, sterilisation status, breed, age and subpopulation against Phase I, II and I and II results for all samples analysed with IFA for Coxiella burnetii antibodies. Variables

Categories

Phase I positive

Phase II positive

Phase I or II positive

Total

Gender

Male Female

5 (1.7%) 18 (4.7%)

9 (3.0%) 19 (4.8%)

9 (3.0%) 28 (7.1%)

300 395

Sterilisation

Entire Sterilised

22 (5.8%) 1 (0.4%)

24 (6.2%) 4 (1.5%)

33 (8.6%) 4 (1.5%)

382 258

Breed

Pure breed short hair Pure breed long hair Cross breed

20 (6.2%) 3 (2.8%) 0 (0%)

20 (5.9%) 6 (5.8%) 2 (0.7%)

29 (8.7%) 6 (5.8%) 2 (0.7%)

331 104 275

Age

0–1 years 1–2.5 years 2.5–6 years >6 years

Subpopulation

Cattery-confined Pet Feral Shelter

2 9 7 5

(1.4%) (5.9%) (4.3%) (2.4%)

2 11 7 8

(1.4%) (7.3%) (4.2%) (3.9%)

3 14 11 9

(2.1%) (9.3%) (6.6%) (4.4%)

143 153 165 204

23 0 0 0

(6.2%) (0%) (0%) (0%)

28 2 0 0

(7.4%) (1.0%) (0%) (0%)

35 2 0 0

(9.3%) (1.0%) (0%) (0%)

376 198 50 88

Sterilisation status was the only risk factor affecting phase I seropositivity with odds of seropositivity greater for entire than sterilised cats (OR 15.6, CI 2.3–106.3; P = 0.005) (Table 3).

I and/or II on IFA, six were also positive on CFT with no additional samples negative on IFA returning a positive result on CFT. All six samples that were CFT (and IFA) positive were also ELISA positive.

3.2.2. Serological methodology comparison All samples collected (n = 712) were analysed with IFA. Of the 712 samples, 143 were analysed with ELISA and 71 of these 143 samples were analysed with CFT (Table 1). The frequency of all variables for samples analysed with IFA is outlined in Table 2. All samples positive for phase I and/or II on IFA (n = 37) were examined with both ELISA and CFT. A total of 23 out of the 30 samples identified as phase II positive on IFA were also positive on ELISA. An additional four samples were positive on ELISA that were negative for phase II IgG on IFA. Out of the 37 samples positive for phase

3.2.3. Interassay agreement Comparisons could only be made between the phase II antibody results with IFA and ELISA as CFT was unable to differentiate reported titres into IgG phase II and IgM phase I titres. Cohen’s kappa analysis was performed on the phase II results comparing IFA with both ELISA index values (agreement 92.20%; Cohen’s kappa coefficient 0.752; 95% confidence interval 0.612–0.891) and ELISA S/P% (agreement 92.25%; Cohen’s kappa coefficient 0.759; 95% confidence interval 0.624–0.894). It was found that the strength of agreement when comparing IFA with both

Table 2 Univariable logistic regression analysis of IFA data set representing the standard error, odds ratio and P value for a 95% Confidence interval when comparing the only statistically significant variables of sterilisation status and subpopulation type to seropositivity. Antibody phase

Categories

Phase I or II

Constant Steriliseda Entire

Phase I

Phase II

Phase I or II

Phase II

a

Reference category.

Constant Steriliseda Entire

Constant Steriliseda Entire Constant Non-catterya Cattery subpopulation Constant Non-catterya Cattery subpopulation

B

SE 4.15

0.50

1.79

0.53

5.54

0.95

2.75

0.98

4.15

0.49

1.53

0.54

5.12

0.69

2.84

0.72

5.12

0.69

2.60

0.73

Odds ratio

Lower 95%

Upper 95%

P value

1 6.00

2.13

16.89

<0.001

1 15.62

2.30

106.26

0.005

1 4.63

1.62

13.26

0.004

1 17.13

4.20

70.20

<0.001

1 13.43

3.25

55.60

<0.001

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Table 3 Multivariable logistic regression analysis of IFA data set representing the standard error, odds ratio, associated 95% Confidence Interval and P value when comparing the factors of subpopulation, sterilisation status, age, gender and breed type and sequential removal of non-significant variables on the outcomes of Phase I, II and Phase I or II seropositivity. Antibody phase

Categories

Phase I or II

Constant Non-catterya Cattery subpopulations

Phase I

Phase II

a

Constant Steriliseda Entire Constant Non-catterya Cattery subpopulations

B

SE 5.12

2.84 5.54 2.75 5.12 2.60

Odds ratio

Lower 95%

Upper 95%

P value

1 17.13

4.18

70.20

<0.001

1 15.63

2.30

106.30

0.005

1 4.63

1.62

13.26

<0.001

0.69 0.72 0.95 0.98 0.69 0.73

Reference category.

ELISA index values and ELISA S/P% was ‘good’ as classified by the GraphPad QuickCalcs statistical tool (http://www. graphpad.com/quickcalcs/kappa1.cfm). 3.3. Timeline of antibody production Repeat follow up samples were obtained from seven cats, three of which were negative on initial and follow up screening at a time interval of nine years. Four of the seven follow up samples were positive on initial screening and three of these four samples were positive on follow up screening after an interval of five years for one cat and nine years for the remaining three cats. Repeat testing results for these four initially IFA positive cats are shown in Supplemental Table. 4. Discussion This study successfully adapted two commercial serological assays developed for use on human sera [C. burnetii I + II IFA IgG/IgM/IgA (Vircell, Spain) and Panbio C. burnetii (Q Fever) IgG ELISA (Alere, Australia)], for the determination of antibodies to C. burnetii in cats. Using indirect IFA on all cat samples, the seroprevalence of C. burnetii amongst the four cat subpopulations demonstrated that Australian cats have been exposed to C. burnetii and that the cattery-confined breeding cats have the highest seropositivity (9.3%). Given that this bacterium is known to recrudesce during pregnancy with the highest bacterial concentration present in the birth products, the finding that C. burnetii was more frequently found in catteryconfined breeding cats was not surprising. Seroepidemiological studies in cats have been conducted in Canada (Higgins and Marrie, 1990; Vallieres et al., 1996), USA (California) (Randhawa, 1974; Willeberg et al., 1980), Japan (Htwe et al., 1992; Komiya et al., 2003; Morita et al., 1994), Korea (Komiya et al., 2003), Turkey (Kilic et al., 2008), South Africa, Zimbabwe (Matthewman et al., 1997) and, most recently, in the United Kingdom (Meredith et al., 2014). It is difficult to compare reported seroprevalence due to the varying methodologies used, and vastly different sample sizes and sample populations in the countries surveyed. Yet these serological surveys provide a valuable overview of the level of infection in cats and the potential zoonotic risk to humans in the localities

surveyed. When the results of the present study are compared to studies utilising either IFA or ELISA, there are striking differences in seroprevalence with 61.5% (n = 26) of pet cats in the United Kingdom (Meredith et al., 2014) and 41.7% (n = 36) of stray cats in Japan (Komiya et al., 2003) reported to be seropositive. Meredith and colleagues confirmed that all cats in their study hunted and consumed wild rodents, while the high level of seropositivity in stray cats in Japan was dependent upon the living environment of these cats (Komiya et al., 2003) which may influence hunting and scavenging, raising the notion of diet or environmental exposure as a contributing factor. The cattery-confined breeding cats in the present study were predominantly indoor-only cats, fed owner controlled diets, with little outdoor access and likely little opportunity to consume rodents and other wildlife species. So the source of C. burnetii in these cats requires further investigation. The modified IFA for phase I and II was established through the verification process which was only possible with the use of serum from the index case and differentiates this from other seroprevalence studies. The positive control in the current study was obtained from a C. burnetii infected cat suffering from dystocia with a CFT titre to C. burnetii of 1/32, that was the cause of an outbreak of Q fever amongst veterinary personnel (Kopecny et al., 2013; Maywood, 2011). This sample enabled the determination of an IFA cut-off titre of 1/256 based upon the clear interpretability of the test, and a titre much greater than that recommended by the kit manufacturer for use in human serum (i.e. 1/64). Therefore the strength and validity of both the IFA and ELISA optimised in this study rests on the presence of a strong positive control. Our preliminary findings on cross reactivity testing were supported by testing conducted by the manufacturer (Vircell, Spain) against the most likely cross reactants, Bartonella spp. (i.e. henslae and quintana), with the resultant finding that Bartonella spp. do not react with phase I or II C. burnetii antigens on the modified IFA (unpublished data). The human IFA kit was an easily modified and interpreted assay and when used by two authors independently in the verification phase, the level of agreement was 100%. All positive samples were taken to end titre for both

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phase I and II antibodies. Furthermore, repeatability of results was extremely high when tested on two or more occasions. The ELISA chosen for this study was more limited in its use, only being able to detect phase II antibodies. The authors are aware of the availability of commercial ELISA kits for IgG, IgM and IgA subclasses and phase I and II specificity to C. burnetii, however it was considered that the benefits of IFA outweighed the need for repeat testing with different ELISA kits. The significance of phase specificity amongst the seropositive cats is unclear as is seen by the antibody timeline of selected cats, with the presence of antibodies to both phase I and II over periods of up to 9 years in duration. It remains to be determined if the human equivalent meanings of phase I antibodies which represent chronic infections and phase II antibodies depicting acute infections are applicable to the antibodies detected in cats. CFT is the only commercially available diagnostic serological test for cats in Australia and remains on the OIE reference serological assay list. In the present study however, comparison of those samples positive on IFA (n = 37) and ELISA (n = 30) with CFT, resulted in only 6 samples being positive on CFT. This indicates that CFT is far less sensitive than either IFA or ELISA in detecting exposure of cats to C. burnetii, notwithstanding the likelihood that IFA and ELISA have sensitivity and specificity values less than 100%. This finding is supported by others also reporting the lower reactivity of CFT compared to IFA and ELISA (Kovacova et al., 1998; Peter et al., 1987) and who similarly report low concordances between IFA and CFT, and ELISA and CFT, when the IFA (Vircell, Spain) used in the present study was examined (Slaba et al., 2005). A further difficulty we encountered with CFT was the presence of anti-complementary activity in a number of samples, which prevented titre determination despite repeated testing. The manufacturer’s stated sensitivity and specificity values for human sera with the C. burnetii I + II IFA IgG/IgM/ IgA kit (Vircell, Spain) are: IgG phase I 93.8% and 75.0%, phase II 97.2% and 100.0% and phase I or II 92.6% and 94.1% respectively. The manufacturer’s stated sensitivity and specificity for the Panbio C. burnetii (Q Fever) IgG ELISA kit (Alere, Australia) are IgG phase II 72.2% and 100% respectively. We were unable to calculate the sensitivity and specificity for cat sera due to the lack of a gold standard assay and the absence of well characterised clinical disease in cats. The ability of IFA to determine end titre values for samples, the ability to visualise positively fluorescing bacteria, the repeatability and consistency of results and the ability to screen large sample numbers in short periods of time, sets IFA as the preferred candidate for a reference serological assay in detecting C. burnetii antibodies in cats. C. burnetii is usually found in highest concentrations in the amniotic fluid and placenta in most animal species. Airborne transmission of C. burnetii at the time of parturition is likely to be a significant risk for those performing reproductive procedures on infected cats. The level of risk is uncertain as there are no known clinical signs that can identify infected cats and seroconversion can be delayed in acute infections. It has not been established if reproductive abnormalities are more common in C. burnetii-infected cats compared to non-infected cats

(Fujishiro et al., 2013). Most infected cats remain subclinical yet cases of human Q fever infection associated with periparturient cats report differences in clinical abnormalities of the parturient cats (Kopecny et al., 2013; Marrie et al., 1988a, 1989). Some cases report that parturient cats were clinically normal, suffering no reproductive abnormalities (Kosatsky, 1984; Marrie et al., 1989; Pinsky et al., 1991), while others report stillborn kittens, neonatal mortalities and dystocia (Langley et al., 1988; Marrie et al., 1988a,b; Maywood, 2011). Therefore, important public health implications potentially exist for those involved in all feline reproductive procedures as high risk cats and situations are not identifiable without external laboratory diagnostic testing. An inactivated vaccine against C. burnetii is available and registered for administration to humans in Australia (Q-Vax CSL Ltd, Parkville, Victoria, Australia). This study and that reported by Kopecny and colleagues (Kopecny et al., 2013) has resulted in a successful submission to the Australian Government’s Department of Health to include cat breeders as individuals at risk of Q fever infection and therefore recommended for vaccination against Q fever as outlined in the Australian Immunisation Handbook (10th edition, 2013). The Australian veterinary profession is in the ideal position to inform breeder clients of these changes. The atrisk group also includes all veterinary personnel involved in procedures occurring during the periparturient period of animals. Irrespective of vaccination status, personal protective equipment (PPE) is mandatory for all those assisting with deliveries and neonatal mortalities. 5. Conclusion The significantly higher seropositivity to C. burnetii amongst cattery-confined breeding cats calls for regular serological monitoring of this subpopulation of cats prior to parturition, greater use of PPE during feline reproductive procedures and vaccination to reduce the risk of Q fever in at-risk personnel. Further studies may focus on the direct detection of C. burnetii in the birth products of serologically positive cats in the investigation and determination of the significance of a positive serological result and resultant shedding of the pathogen. Acknowledgements This research was supported by competitive grants obtained from the Australian Companion Animal Health Foundation (Australian Veterinary Association) and the WP Richards award in Pathology, University of Sydney. The authors are grateful to the animal owners, the practice owners and veterinary staff at the hospitals at the centre of the outbreaks associated with the cat caesarean. The authors would also like to thank Evelyn Hall of The University of Sydney for statistical advice. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j. vetmic.2015.02.011.

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