Actinobacillus pleuropneumoniae transmission and ...

Actinobacillus pleuropneumoniae transmission and clinical outbreaks

Actinobacillus pleuropneumoniae transmission and clinical outbreaks

Uitnodiging Voor het bijwonen van de openbare verdediging van mijn proefschrift getiteld:

Actinobacillus pleuropneumoniae transmission and clinical outbreaks Donderdag 26 juni 2014 om 12:45 uur in het Academiegebouw van de Universiteit Utrecht Domplein 29, Utrecht Na afloop van de verdediging bent u van harte welkom op de receptie in het Academiegebouw Tijs Tobias Rijnlaan 215 3522 BL Utrecht 06 - 51060639 [email protected]

Tijs Tobias Paranimfen Jos van Aert [email protected] 06 -11311917

Tijs Tobias

Bram Loog [email protected] 06 - 10381056

2014

Actinobacillus pleuropneumoniae transmission and clinical outbreaks

Tijs Jan-Willem Tobias

Actinobacillus pleuropneumoniae transmission and clinical outbreaks T.J. Tobias PhD thesis Faculty of Veterinary Medicine, Utrecht University, The Netherlands - With a summary in Dutch ISBN: 978-90-393-6147-4 Cover design:

Ria Tobias &



Elsbeth Kool, Multimedia, Faculty of Veterinary Medicine

Printing and lay-out:

Gildeprint - The Netherlands

Publisher:

Utrecht University

Actinobacillus pleuropneumoniae transmission and clinical outbreaks Actinobacillus pleuropneumoniae transmissie en klinische uitbraken (met een samenvatting in het Nederlands)

Proefschrift

ter verkrijging van de graad van doctor aan de Universiteit Utrecht op gezag van de rector magnificus, prof.dr. G.J. van der Zwaan, ingevolge het besluit van het college voor promoties in het openbaar te verdedigen op donderdag 26 juni 2014 des middags te 12.45 uur

door

Tijs Jan-Willem Tobias geboren op 8 november 1976 te Amersfoort

Promotoren:

Prof. dr. J.A. Stegeman



Prof. dr. J.A. Wagenaar

Copromotoren:

Dr. A. Bouma



Dr. D. Klinkenberg

Printing of this thesis was supported by:

MSD Animal Health The Netherlands



Dechra Veterinary Products

CONTENTS 1

General Introduction

7

2

Detection of Actinobacillus pleuropneumoniae in pigs by real-time



quantitative PCR for the apxIVA gene

25

3

The association between transmission rate and disease severity

43

for Actinobacillus pleuropneumoniae infection in pigs 4

A cohort study on Actinobacillus pleuropneumoniae colonisation



in suckling piglets

69

5

Transmission of Actinobacillus pleuropneumoniae in weaned piglets



on endemically infected farms

93

6

Simulation study on the mechanism of outbreaks of clinical disease



caused by Actinobacillus pleuropneumoniae in finishing pigs

119

7

Outbreak of respiratory distress resembling influenza caused by



Actinobacillus pleuropneumoniae in pigs

151

8

General Discussion

157

Summary

178



Nederlandse samenvatting

180



Dankwoord

183



About the author

187



List of publications

189

1 General introduction

Chapter 1

8

General introduction

GENERAL INTRODUCTION Outbreaks of respiratory distress in pigs caused by Actinobacillus pleuropneumoniae often occur in pig herds, causing animal welfare problems and economic losses due to mortality, growth retardation and use of antimicrobials. The disease and the bacterium have been subject of research since mid-1960 and the first report of the disease in pigs in The Netherlands was made in 1965 (cited by Hunneman, 1983). Despite the scientific knowledge that has been gained since, prevention of clinical outbreaks, or the control of outbreaks without antimicrobial group treatment has been difficult. Pleuropneumonia due to A. pleuropneumoniae was reported recently as one of the most diagnosed diseases in pigs sent for necropsy to the Animal Health Service in The Netherlands (Muskens et al., 2013), justifying research on this disease in order to develop or improve control measures. Observational studies on outbreaks have suggested different risk factors to contribute to the occurrence of clinical signs, e.g. high pig density (Mylrea et al., 1974; Cameron and Kelly, 1979), moving pigs to other pens (Davidson and King, 1980) or other farms (Nielsen, 1973), introduction of new pigs (Sanford and Josephson, 1981) and concurrent infections (Nielsen, 1973; Schiefer et al., 1974). However, scientifically sound risk factor analyses are lacking and whereas elimination of (one of) the risk factors mentioned above might contribute to a reduction in A. pleuropneumoniae associated problems, the extent of this reduction is unknown. Moreover, most risk factors are associated with day-to-day pig farm management, indicating the need for alternative solutions. For planning of infectious disease prevention and control, it has been shown that information about the distribution of shedding patterns of the pathogen and the association between shedding and onset of clinical signs is important (Woolhouse et al., 1997; Fraser et al., 2004; Lloyd-Smith et al., 2005). For A. pleuropneumoniae such information is scarce.

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Chapter 1

In (per-) acute outbreaks of pleuropneumonia caused by A. pleuropneumoniae, there is a sudden and rapid increase in the number of clinical cases. Different viewpoints have been expressed on the conceptual mechanism that explains the transmission of the bacterium in the population and the occurrence of clinical signs, which may have consequences for treatment and prevention. One hypothesis states that A. pleuropneumoniae is transmitted between healthy pigs which remain free from clinical signs during a certain time. Only after simultaneous exposure to a particular risk factor, pigs develop clinical signs in more or less the same time frame. This hypothesis is supported by a study that provided risk factors associated with higher within herd seroprevalence (Maes et al., 2001). In this thesis, this mechanism will be referred to as trigger mechanism. If the trigger mechanism indeed explains the course of the infection in a population, control measures could focus on the elimination of risk factors or on the prevention of the infection. A second hypothesis states that clinically diseased pigs transmit the bacterium to other pigs in the pen or unit in such a way that it induces clinical signs rapidly after infection. This hypothesis is supported by a study that describes a rapid increase in the number of infected pigs in the acute phase of disease and an ongoing rise in seroprevalence after the acute phase (Loeffen et al., 1999). This mechanism was also suggested by Sebunya and Saunders (1983). This mechanism will be referred to as the transmission mechanism. It implies that control measures could aim at reducing transmission of the bacterium by clinically diseased pigs, e.g. by treating those pigs. Preventive treatment of yet non-diseased pigs may not only help to prevent clinical signs, but also to prevent infection. Irrespective of the validity of either hypothesis, group treatment with antimicrobials has shown to be effective to halt the occurrence of new clinical cases or prevent outbreaks (Hunneman et al., 1994; Mengelers et al., 2000; Hoflack et al., 2001) and was even recently advocated (Voss, 2012). However, this is not considered a sustainable control method for the future because of the development of antimicrobial resistance, posing a risk for human and animal health (Aarestrup, 2005; Acar and Moulin, 2012).

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General introduction

Focus of the research in this thesis is on the validity of either hypothesis, and the results contribute to the understanding of the transmission of A. pleuropneumoniae on pig farms. Thereby the outcomes of this thesis may help to develop more effective prevention and control measures for the disease.

BACKGROUND ON A. PLEUROPNEUMONIAE Aetiology A. pleuropneumoniae is a gram negative coccobacillary bacterium, belonging to the family of Pasteurellaceae. The name of the bacterium, its abbreviation (A.p.p.) and its former name (Haemophilus) are often used for the disease it causes in pigs (pleuropneumonia), as well as to describe the bacterium. “Haemophilus pleuropneumoniae” was first described by Pattison et al. (1957) and Shope showed that H. pleuropneumoniae was the causative bacterium of outbreaks of pleuropneumonia in pigs (1964). Whereas the name “Haemophilus parahaemolyticus” has also been used temporarily, it was shown later, by phenotypic and DNA analyses, that the bacterium belongs to a separate genus (Actinobacillus) within the family of Pasteurellaceae (Pohl et al., 1983). Strains of A. pleuropneumoniae can be grouped based on biovar, serovar or Apx toxin profile. Biovars differ in their dependency for nicotinamide adenine dinucleotide (NAD) to grow, whereas serovars differ with respect to the capsular polysaccharides (cps) and cell wall lipopolysaccharides (lps) (Haesebrouck et al., 1997). It is generally accepted that the species A. pleuropneumoniae can be differentiated into 15 serovars divided over two biovars (Blackall et al., 2002), but for certain serovars biovar 1 as well as biovar 2 isolates have been reported (Beck et al., 1994; Perry et al., 2012). In The Netherlands in 2008, 94 / 99 (95%) randomly sampled finishing farms had at least one positive serologic sample (Duinhof, personal communication) and from the same study it was concluded that serovar 2 (biovar 1) was most prevalent (Duinhof et al., 2013).

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Chapter 1

The Apx toxin grouping method is based on the presence of the genes for ApxI, ApxII and ApxIII toxins (Frey, 2003). A. pleuropneumoniae serovars contain genes for at least one of these toxins; most contain genes for two. The gene for a fourth Apx toxin (ApxIV) is present in each strain and therefore highly specific for A. pleuropneumoniae (Schaller et al., 1999). Pathology and clinical signs Respiratory signs due to A. pleuropneumoniae are the result of pathology in the lower respiratory tract. Several virulence factors, such as Apx toxins, lps, cps and proteases, are involved in the induction of tissue damage and interaction with the host immune system resulting in cytokine release and subsequent inflammation (Bossé et al., 2002; Chiers et al., 2010). At necropsy a fibrinous necrotizing pneumonia as well as pleuritis and hemorrhages can be observed (Liggett et al., 1987; Bertram, 1988). The severity of clinical signs is associated with the extent of lung lesions (Hoeltig et al., 2009). The clinical course of disease in individual pigs is arbitrarily differentiated into peracute, acute and chronic (Nicolet et al., 1969; Gottschalk, 2012). In case of a peracute course of disease, pigs are often found dead, without anyone previously having observed clinical signs. In case signs are seen in these pigs, they mostly resemble signs of sepsis: extreme hyperthermia (up to 42 °C), cyanosis of extremities, anorexia and sometimes dyspnoea. Acutely diseased pigs show more prominent signs of respiratory disease such as dyspnoea, coughing and open-mouth breathing, hyperthermia, depression and anorexia. Finally, chronically diseased pigs suffer from pathological sequelae and can show reduced growth and intolerance to physical strain. Between farms and between pigs within farms the course of disease varies due to many factors, such as genetics (Hoeltig et al., 2009), infective dose (van Leengoed and Kamp, 1989; Jacobsen et al., 1996), concurrent infections (Sakano et al., 1993; Marois et al., 2009), the level of maternal immunity (Cruijsen et al., 1995b) and immunity due to previous infections with A. pleuropneumoniae (Cruijsen et al., 1995a). The large variation in factors within and between farms might explain

12

General introduction

the lack of information on the course of clinical signs in populations under field conditions. Most of the information on the clinical course of disease is derived from experimental studies with small groups of piglets, that aimed to study pathogenesis, vaccine or antimicrobial efficacy. Dynamics of A. pleuropneumoniae infection and association with clinical signs From A. pleuropneumoniae literature it is clear that an unambiguous use of definitions for ‘infected’ or ‘diseased’ is essential. This can for example been shown by the following example: in a review, Sebunya and Saunders (1983) wrote: “disease was found to spread rapidly from sick to normal pigs” upon the interpretation by Nicolet (1969) who stated: “Kommen infizierte und nicht infizierte Tiere zusammen, so erkranken letztere in der Regel schlagartig und können wie nach künstlicher massiver Infektion innert Stunden bis wenigen Tagen verenden.” (“Do infected and non-infected animals come in contact, as a general rule the last mentioned will suddenly become diseased, and can, as in artificially infected animals, die within hours or days”). It does not become clear in the text of Nicolet whether ‘infected’ actually means pigs that are clinically diseased or not, whether contact infected pigs are truly free from the bacterium at start and whether all contact infected pigs show signs or just a few. In other words the infective state and outcome after contracting the bacterium of both infectious and contact pigs are difficult to extract, which in turn hampers the interpretation of the results of these studies. It becomes even more complicating when results of experimental studies are considered indicating that infected pigs, which do not show clinical signs, can directly transmit the bacterium as well. In the same studies, contact infected pigs were reported not to express severe clinical signs irrespective of presence of clinical signs in the experimentally infected pig (Lechtenberg et al., 1994; Velthuis et al., 2003).

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Chapter 1

To ensure a clear discussion, in this thesis the ‘damage-response framework’ suggested by Pirofski and Casadevall (2002; 2003) is used. In this framework infection refers to the process of acquiring the bacterium rather than a disease state. Moreover, it is the disease state which is important as exact outcome of infection. The existing literature on A. pleuropneumoniae points to a distinction between colonisation and disease to fit the behaviour of the bacterium in the population. The following states are distinguished: -

Susceptible: pigs are assumed susceptible for infection if there is no A. pleuropneumoniae present. New born piglets are born susceptible and can become infected by (acquire) A. pleuropneumoniae in the farrowing room (Chiers et al., 2002a; Vigre et al., 2002).

-

Colonised: after infection, A. pleuropneumoniae can colonise the mucosa of the naso- or oropharynx without inducing major pathology and subsequent clinical signs, or inducing a humoral immune response (Chiers et al., 2002b). This state is referred to as colonised. In colonised pigs the lower respiratory tract is considered to be free from large numbers of A. pleuropneumoniae, as in bronchial alveolar fluid obtained from healthy pigs no A. pleuropneumoniae is found (Hensel et al., 1994). Moreover, if A. pleuropneumoniae is detected in the lower respiratory tract, even with low bacterial counts, it is accompanied by pathologic changes (van Leengoed and Kamp, 1989; Hensel et al., 1993; Stockhofe-Zurwieden et al., 1994). Pigs can become infected through direct contact (Lechtenberg et al., 1994; Velthuis et al., 2002) and transmission by aerosols has also been demonstrated experimentally (Jobert et al., 2000; Kristensen et al., 2004). The importance of the latter route under field conditions is unclear.

-

Diseased: pigs become diseased when A. pleuropneumoniae has caused tissue damage in the lower respiratory tract, mainly due to the Apx toxins (Kamp et al., 1997). In subclinically diseased pigs, signs are not observed despite the presence of lung lesions.

14

General introduction

-

Infected: in this thesis the term infected is used for all colonised and diseased, i.e. pigs in which A. pleuropneumoniae is detected, irrespective of the clinical outcome.

Diagnosis To study the relation between infective status and the onset of clinical signs, accurate diagnostic tools are necessary, but these tools are lacking for investigations under field conditions. In peracutely elapsing outbreaks clinical examination as diagnostic tool may function well to define pigs as ‘diseased’, as the sudden onset of clinical signs and mortality are conspicuous for A. pleuropneumoniae (Shope, 1964; Schiefer et al., 1974; Davidson and King, 1980; Hoflack et al., 2001). However, in acutely diseased pigs, clinical examination is not specific enough and needs to be confirmed by finding characteristic lesions at necropsy and culture of the bacterium from these lesions (Gottschalk, 2012). In chronically diseased pigs, clinical examination as well as necropsy and culture lack sensitivity and specificity to accurately define the clinical or infection status of pigs. To define the infection status of pigs, isolation of the bacterium from samples from the nasal or oropharyngeal cavity obtained from colonised pigs is challenging, even when using selective growth media (Gilbride and Rosendal, 1983; Møller et al., 1993; Sidibé et al., 1993; Jacobsen and Nielsen, 1995; Chiers et al., 2002b). Other bacteria easily overgrow samples and other Pasteurellaceae species are difficult to differentiate from A. pleuropneumoniae by standard biochemical tests (Møller and Kilian, 1990). Advanced techniques such as immunomagnetic separation may enhance recovery of viable A. pleuropneumoniae (Gagne et al., 1998; Angen et al., 2001), but require considerable investments in training and equipment. Detection of A. pleuropneumoniae DNA by PCR in nasal or oropharyngeal swabs, or in tissue samples such as tonsil or lung, may be a good alternative to culture for defining the infection status of pigs, even though also DNA of non-viable bacteria is detected (e.g. Savoye et al., 2000; Chiers et al., 2001; Cho and Chae, 2003; Fittipaldi et al., 2003). PCR tests are also used for identification or confirmation of A.

15

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Chapter 1

pleuropneumoniae after culture or to assign the isolate to a specific group of strains (e.g. Schuchert et al., 2004; Tonpitak et al., 2007; Angen et al., 2008; Zhou et al., 2008; Ito, 2010). For studying the transmission of the bacterium between pigs, the use of a quantitative test for samples of live pigs on farms is desirable, but for A. pleuropneumoniae such a test was not available before the start of this study. An alternative to detection of bacterial antigen may be detection of an immune response in the pig, despite the considerable lag time between time of infection and the rise of a detectable immune response. The complement fixation test (CFT) (Nielsen, 1974; Nielsen, 1982) can detect a rise in antibodies in diseased pigs approximately ten days after the onset of clinical signs (Lombin et al., 1982). However CFT test performance depends much on the practical experience with the test (personal observation) and availability of serovar specific antigen (Lombin et al., 1982). Other serologic tests frequently used rely on detection of antibodies against Apx toxins, outer membrane protein, cps or lps. All but one of these tests are reported to show cross reactions. Only the Anti-ApxIV ELISA is considered highly specific for a response to A. pleuropneumoniae (Dreyfus et al., 2004), whereas Anti-ApxI, ApxII, ApxIII ELISA (Nielsen et al., 2000; Shin et al., 2011) and Omp ELISA tests (Kobisch and van den Bosch, 1992; Eamens et al., 2008) have shown to cross react with antibodies to other Pasteurellaceae, e.g. A. suis (Macinnes et al., 2008) or Pasteurella spp. (Eamens et al., 2012). ELISA tests that detect antibodies to cps or lps (Gottschalk et al., 1994; Inzana and Fenwick, 2001) can show cross reaction between responses to different serovars of A. pleuropneumoniae (Costa et al., 2011; Opriessnig et al., 2013). In this thesis, the CFT is used for serological confirmation of infection in diseased pigs (Se > 0.83 and Sp > 0.8 (Lombin et al., 1982; Enøe et al., 2001)). Anti-ApxI, ApxII, and ApxIII (Se > 0.9 and Sp > 0.95 for a similar ELISA (Shin et al., 2011)) and Omp ELISA tests are used for detection of pigs with previous lung exposure to A. pleuropneumoniae, irrespective of having shown clinical signs. Anti-ApxIV ELISA

16

General introduction

test (Se = 0.94 and Sp = 1.0 (Dreyfus et al., 2004)) is used for testing of freedom of A. pleuropneumoniae on herd level. Serologic tests are considered to have too low sensitivity for detection of colonised pigs (Sørensen et al., 1997; Chiers et al., 2002b; Costa et al., 2011; Opriessnig et al., 2013). Research aims and thesis outline The aim of the research described in this thesis was to investigate transmission of A. pleuropneumoniae on pig farms and to study the association between the occurrence of clinical signs and the transmission of the pathogen between pigs and pens. Two hypotheses on the conceptual mechanism of clinical outbreaks, due to A. pleuropneumoniae, are the focus of this thesis: I

The bacterium has first colonised most pigs in a group, next a trigger causes the bacterium to induce pathology in the lungs and clinical signs (trigger mechanism).

II The clinical course of an outbreak depends heavily on the infectiousness of clinically diseased pigs. These pigs transmit the bacterium to other pigs in a way that it induces clinical signs rapidly after infection (transmission mechanism). Both hypotheses are difficult to test in field situations. Firstly, because rapid and reliable diagnostics were lacking to investigate transmission. Therefore, a quantitative PCR for detection of the apxIVA gene was developed and validated (chapter 2). Secondly, longitudinal studies of outbreaks under field conditions suffer from many logistical constraints, apart from the difficulty to ensure minimal transmission due to the sampling procedure. Therefore, an experimental design was chosen to investigate the association between disease severity and transmission to test the transmission hypothesis. The transmission hypothesis suggests that disease severity of pigs should be positively correlated with the transmission rate (chapter 3). To explore the dispersion of A. pleuropneumoniae under field conditions a cohort study on colonisation and possible effects of dam characteristics was conducted on two infected farms (chapter 4). In the weaned pigs of the same cohorts, direct (within pen) and indirect (between pen) transmission routes of A. pleuropneumoniae were quantified (chapter 5).

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Chapter 1

In chapter 6 a simulation model is presented to investigate conditions required to cause an outbreak under both mechanisms. A secondary aim was to test the effect of preventive and control measures on the size and duration of outbreaks. Most observational studies and textbooks usually report on outbreaks with severe signs of disease. In chapter 7 an example of an outbreak is described in which, despite the rapid increase in number of diseased pigs, the severity of these diseased pigs was mild. Finally, in chapter 8, the validity of the trigger and transmission mechanism is discussed based on the results of the studies in this thesis and options for prevention and control of outbreaks are discussed.

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REFERENCES Aarestrup, F.M., 2005. Veterinary drug usage and antimicrobial resistance in bacteria of animal origin. Basic Clin. Pharmacol. Toxicol. 96, 271-281. Acar, J.F., Moulin, G., 2012. Antimicrobial resistance: a complex issue. Rev. Sci. Tech. 31, 23-31. Angen, Ø., Ahrens, P., Jessing, S.G., 2008. Development of a multiplex PCR test for identification of Actinobacillus pleuropneumoniae serovars 1, 7, and 12. Vet. Microbiol. 132, 312-318. Angen, Ø., Heegaard, P.M., Lavritsen, D.T., Sørensen, V., 2001. Isolation of Actinobacillus pleuropneumoniae serotype 2 by immunomagnetic separation. Vet. Microbiol. 79, 19-29. Beck, M., van den Bosch, J.F., Jongenelen, I.M., Loeffen, P.L., Nielsen, R., Nicolet, J., Frey, J., 1994. RTX toxin genotypes and phenotypes in Actinobacillus pleuropneumoniae field strains. J. Clin. Microbiol. 32, 27492754. Bertram, T.A., 1988. Pathobiology of acute pulmonary lesions in swine infected with Haemophilus (Actinobacillus) pleuropneumoniae. Can. Vet. J. 29, 574-577. Blackall, P.J., Klaasen, H.L., van den, B.H., Kuhnert, P., Frey, J., 2002. Proposal of a new serovar of Actinobacillus pleuropneumoniae: serovar 15 3. Vet. Microbiol. 84, 47-52. Bossé, J.T., Janson, H., Sheehan, B.J., Beddek, A.J., Rycroft, A.N., Simon Kroll, J., Langford, P.R., 2002. Actinobacillus pleuropneumoniae: pathobiology and pathogenesis of infection. Microb. Infect. 4, 225-235. Cameron, R.D., Kelly, W.R., 1979. An outbreak of porcine pleuropneumonia due to Haemophilus parahaemolyticus. Aust. Vet. J. 55, 389-390. Casadevall, A., Pirofski, L.A., 2003. The damage-response framework of microbial pathogenesis. Nat. Rev. Microbiol. 1, 17-24. Chiers, K., De Waele, T., Pasmans, F., Ducatelle, R., Haesebrouck, F., 2010. Virulence factors of Actinobacillus pleuropneumoniae involved in colonization, persistence and induction of lesions in its porcine host. Vet. Res. 41: 65. Chiers, K., Donne, E., Van Overbeke, I., Ducatelle, R., Haesebrouck, F., 2002a. Actinobacillus pleuropneumoniae infections in closed swine herds: infection patterns and serological profiles. Vet. Microbiol. 85, 343-352. Chiers, K., Donne, E., van, O.,I, Ducatelle, R., Haesebrouck, F., 2002b. Evaluation of serology, bacteriological isolation and polymerase chain reaction for the detection of pigs carrying Actinobacillus pleuropneumoniae in the upper respiratory tract after experimental infection. Vet. Microbiol. 88, 385-392. Chiers, K., Van Overbeke, I., Donne, E., Baele, M., Ducatelle, R., De Baere T., Haesebrouck, F., 2001. Detection of Actinobacillus pleuropneumoniae in cultures from nasal and tonsillar swabs of pigs by a PCR assay based on the nucleotide sequence of a dsbE-like gene. Vet. Microbiol. 83, 147-159. Cho, W.S., Chae, C., 2003. PCR detection of Actinobacillus pleuropneumoniae apxIV gene in formalin-fixed, paraffin-embedded lung tissues and comparison with in situ hybridization. Lett. Appl. Microbiol. 37, 56-60. Costa, G., Oliveira, S., Torrison, J., Dee, S., 2011. Evaluation of Actinobacillus pleuropneumoniae diagnostic tests using samples derived from experimentally infected pigs. Vet. Microbiol. 148(2-4), 246-251. Cruijsen, T., van Leengoed, L.A., Ham-Hoffies, M., Verheijden, J.H., 1995a. Convalescent pigs are protected completely against infection with a homologous Actinobacillus pleuropneumoniae strain but incompletely against a heterologous-serotype strain. Infect. Immun. 63, 2341-2343. Cruijsen, T., van Leengoed, L.A., Kamp, E.M., Bartelse, A., Korevaar, A., Verheijden, J.H., 1995b. Susceptibility to Actinobacillus pleuropneumoniae infection in pigs from an endemically infected herd is related to the presence of toxin-neutralizing antibodies. Vet. Microbiol. 47, 219-228. Davidson, J.N., King, J.M., 1980. An outbreak of Haemophilus parahaemolyticus pneumonia in growing pigs. Cornell Vet. 70, 360-364. Dreyfus, A., Schaller, A., Nivollet, S., Segers, R.P., Kobisch, M., Mieli, L., Soerensen, V., Hussy, D., Miserez, R., Zimmermann, W., Inderbitzin, F., Frey, J., 2004. Use of recombinant ApxIV in serodiagnosis of Actinobacillus pleuropneumoniae infections, development and prevalidation of the ApxIV ELISA. Vet. Microbiol. 99, 227-238. Duinhof, T., Tempelmans-Plat, B., Bouwkamp, F., Niesink-Broekroelofs, J., Wellenberg, G.J., 2013. Actinobacillus pleuropneumoniae serotypes in The Netherlands: a survey based on serology and isolates from lung lesions. Tijdschr. Diergeneeskd. 138, 28-35.

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Eamens, G., Gonsalves, J., Whittington, A.M., Turner, B., 2008. Serological responses to two serovar-independent ELISA antigens of Actinobacillus pleuropneumoniae in Australian commercial pig herds. Austr. Vet. J. 86, 465-472. Eamens, G.J., Gonsalves, J.R., Whittington, A.M., Turner, B., 2012. Evaluation of serovar-independent ELISA antigens of Actinobacillus pleuropneumoniae in pigs, following experimental challenge with A. pleuropneumoniae, Mycoplasma hyopneumoniae and Pasteurella multocida. Aust. Vet. J. 90, 225-234. Enøe, C., Andersen, S., Sørensen, V., Willeberg, P., 2001. Estimation of sensitivity, specificity and predictive values of two serologic tests for the detection of antibodies against Actinobacillus pleuropneumoniae serotype 2 in the absence of a reference test (gold standard). Prev. Vet. Med. 51, 227-243. Fittipaldi, N., Broes, A., Harel, J., Kobisch, M., Gottschalk, M., 2003. Evaluation and field validation of PCR tests for detection of Actinobacillus pleuropneumoniae in subclinically infected pigs. J. Clin. Microbiol. 41, 5085-5093. Fraser, C., Riley, S., Anderson, R.M., Ferguson, N.M., 2004. Factors that make an infectious disease outbreak controllable. Proc. Natl. Acad. Sci. U. S. A. 101, 6146-6151. Frey, J., 2003. Detection, identification, and subtyping of Actinobacillus pleuropneumoniae. In: Sachse, K., Frey, J. (Eds.), Methods in molecular biology: PCR detection of microbial pathogens. Humana Press, pp. 87-95. Gagne, A., Lacouture, S., Broes, A., D’Allaire, S., Gottschalk, M., 1998. Development of an immunomagnetic method for selective isolation of Actinobacillus pleuropneumoniae serotype 1 from tonsils. J. Clin. Microbiol. 36, 251-254. Gilbride, K.A., Rosendal, S., 1983. Evaluation of a selective medium for isolation of Haemophilus pleuropneumoniae. Can. J. Comp. Med. 47, 445-450. Gottschalk, M., 2012. Actinobacillosis. In: Zimmerman, J.J., Karriker, L.A., Ramirez, A., Schwartz, K.J., Stevenson, G.W. (Eds.), Diseases of Swine. Wiley - Blackwell, Ames, Iowa, USA, pp. 653-669. Gottschalk, M., Altman, E., Charland, N., De Lasalle, F., Dubreuil, J.D., 1994. Evaluation of a saline boiled extract, capsular polysaccharides and long-chain lipopolysaccharides of Actinobacillus pleuropneumoniae serotype 1 as antigens for the serodiagnosis of swine pleuropneumonia. Vet. Microbiol. 42, 91-104. Haesebrouck, F., Chiers, K., van, O.,I, Ducatelle, R., 1997. Actinobacillus pleuropneumoniae infections in pigs: the role of virulence factors in pathogenesis and protection. Vet. Microbiol. 58, 239-249. Hensel, A., Windt, H., Stockhofe-Zurwieden, N., Lodding, H., Koch, W., Petzoldt, K., 1993. A porcine aerosol infection model for studying dose dependent effects caused by Actinobacillus pleuropneumoniae bacteria. J. Aerosol Med. 6, 73-88. Hensel, A., Ganter, M., Kipper, S., Krehon, S., Wittenbrink, M.M., Petzoldt, K., 1994. Prevalence of aerobic bacteria in bronchoalveolar lavage fluids from healthy pigs. Am. J. Vet. Res. 55, 1697-1702. Hoeltig, D., Hennig-Pauka, I., Thies, K., Rehm, T., Beyerbach, M., Strutzberg-Minder, K., Gerlach, G.F., Waldmann, K.H., 2009. A novel Respiratory Health Score (RHS) supports a role of acute lung damage and pig breed in the course of an Actinobacillus pleuropneumoniae infection. BMC. Vet. Res. 5: 14. Hoflack, G., Maes, D., Mateusen, B., Verdonck, M., de Kruif, A., 2001. Efficacy of tilmicosin phosphate (Pulmotil premix) in feed for the treatment of a clinical outbreak of Actinobacillus pleuropneumoniae infection in growing-finishing pigs. J. Vet. Med. B Infect. Dis. Vet. Public Health 48, 655-664. Hunneman, W.A., 1983. Incidence, economic effects, and control of Haemophilus pleuropneumoniae infections in pigs. PhD thesis, Utrecht University Hunneman, W.A., Pijpers, A., Lommerse, J., Crauwels, A.P., Verheijden, J.H., 1994. Prophylaxis of pleuropneumonia in pigs by in-feed medication with oxytetracycline and the subsequent transmission of infection. Vet. Rec. 134, 215-218. Inzana, T.J., Fenwick, B., 2001. Serologic detection of Actinobacillus pleuropneumoniae in swine by capsular polysaccharide-biotin-streptavidin enzyme-linked immunosorbent assay. J. Clin. Microbiol. 39, 12791282. Ito, H., 2010. Development of a cps-Based Multiplex PCR for Typing of Actinobacillus pleuropneumoniae Serotypes 1, 2 and 5. J. Vet. Med. Sci. 72, 653-655. Jacobsen, M.J., Nielsen, J.P., 1995. Development and evaluation of a selective and indicative medium for isolation of Actinobacillus pleuropneumoniae from tonsils. Vet. Microbiol. 47, 191-197. Jacobsen, M.J., Nielsen, J.P., Nielsen, R., 1996. Comparison of virulence of different Actinobacillus pleuropneumoniae serotypes and biotypes using an aerosol infection model. Vet. Microbiol. 49, 159168.

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Jobert, J.L., Savoye, C., Cariolet, R., Kobisch, M., Madec, F., 2000. Experimental aerosol transmission of Actinobacillus pleuropneumoniae to pigs. Can. J. Vet. Res. 64, 21-26. Kamp, E.M., Stockhofe-Zurwieden, N., van Leengoed, L.A., Smits, M.A., 1997. Endobronchial inoculation with Apx toxins of Actinobacillus pleuropneumoniae leads to pleuropneumonia in pigs. Infect. Immun. 65, 4350-4354. Kobisch, M., van den Bosch, J.F., Efficacy of an Actinobacillus pleuropneumoniae subunit vaccine. In: Proceedings of the 12th International Pig Veterinary Society Congress, The Hague, The Netherlands, 1992, 216. Kristensen, C.S., Angen, Ø., Andreasen, M., Takai, H., Nielsen, J.P., Jorsal, S.E., 2004. Demonstration of airborne transmission of Actinobacillus pleuropneumoniae serotype 2 between simulated pig units located at close range. Vet. Microbiol. 98, 243-249. Lechtenberg, K.F., Shryock, T.R., Moore, G., 1994. Characterization of an Actinobacillus pleuropneumoniae seeder pig challenge-exposure model. Am. J. Vet. Res. 55, 1703-1709. van Leengoed, L.A., Kamp, E.M., 1989. Endobronchial inoculation of various doses of Haemophilus (Actinobacillus) pleuropneumoniae in pigs. Am. J. Vet. Res. 50, 2054-2059. Liggett, A.D., Harrison, L.R., Farrell, R.L., 1987. Sequential study of lesion development in experimental Haemophilus pleuropneumonia. Res. Vet. Sci. 42, 204-212. Lloyd-Smith, J.O., Schreiber, S.J., Kopp, P.E., Getz, W.M., 2005. Superspreading and the effect of individual variation on disease emergence. Nature 438, 355-359. Loeffen, W.L.A., Kamp, E.M., Stockhofe-Zurwieden, N., Van Nieuwstadt, A.P.K.M.I., Bongers, J.H., Hunneman, W.A., Elbers, A.R.W., Baars, J., Nell, T., Van Zijderveld, F.G., 1999. Survey of infectious agents involved in acute respiratory disease in finishing pigs. Vet. Rec. 145, 123-129. Lombin, L.H., Rosendal, S., Mitchell, W.R., 1982. Evaluation of the complement fixation test for the diagnosis of pleuropneumonia of swine caused by Haemophilus pleuropneumoniae. Can. J. Comp. Med. 46, 109-114. Macinnes, J.I., Gottschalk, M., Lone, A.G., Metcalf, D.S., Ojha, S., Rosendal, T., Watson, S.B., Friendship, R.M., 2008. Prevalence of Actinobacillus pleuropneumoniae, Actinobacillus suis, Haemophilus parasuis, Pasteurella multocida, and Streptococcus suis in representative Ontario swine herds. Can. J. Vet. Res. 72, 242-248. Maes, D., Chiers, K., Haesebrouck, F., Laevens, H., Verdonck, M., de Kruif A., 2001. Herd factors associated with the seroprevalences of Actinobacillus pleuropneumoniae serovars 2, 3 and 9 in slaughter pigs from farrow-to-finish pig herds. Vet. Res. 32, 409-419. Marois, C., Gottschalk, M., Morvan, H., Fablet, C., Madec, F., Kobisch, M., 2009. Experimental infection of SPF pigs with Actinobacillus pleuropneumoniae serotype 9 alone or in association with Mycoplasma hyopneumoniae. Vet. Microbiol. 135, 283-291. Mengelers, M.J., Kuiper, H.A., Pijpers, A., Verheijden, J.H., van Miert, A.S., 2000. Prevention of pleuropneumonia in pigs by in-feed medication with sulphadimethoxine and sulphamethoxazole in combination with trimethoprim. Vet. Q. 22, 157-162. Møller, K., Andersen, L.V., Christensen, G., Kilian, M., 1993. Optimalization of the detection of NAD dependent Pasteurellaceae from the respiratory tract of slaughterhouse pigs. Vet. Microbiol. 36, 261-271. Møller, K., Kilian, M., 1990. V factor-dependent members of the family Pasteurellaceae in the porcine upper respiratory tract. J. Clin. Microbiol. 28, 2711-2716. Muskens, J., Geudeke, T., Heijmans, J., van den Brom, R., 2013. Trends from the GD monitoring. Tijdschr. Diergeneeskd. 138, 82-83. Mylrea, P.J., Fraser, G., Macqueen, P., Lambourne, D.A., 1974. Pleuropneumonia in pigs caused by Haemophilus parahaemolyticus. Aust. Vet. J. 50, 255-259. Nicolet, J., Konig, H., School, E., 1969. On Haemophilus pleuropneumonia in swine. II. A contagious disease of scientific value. Schweiz. Arch. Tierheilkd. 111, 166-174. Nielsen, R., 1973. An outbreak of pleuropneumonia among a group of baconers. Pathological and bacteriological observations. Nord. Vet. 25, 492-496. Nielsen, R., 1974. Serological and immunological studies of pleuropneumonia of swine caused by Haemophilus parahaemolyticus. Acta Vet. Scand. 15, 80-89. Nielsen, R., 1982. Haemophilus pleuropneumoniae infection in pigs. PhD thesis, Kongelige Veterinær- og landbohøjskole (Denmark). Nielsen, R., van den Bosch, J.F., Plambeck, T., Sørensen, V., Nielsen, J.P., 2000. Evaluation of an indirect enzymelinked immunosorbent assay (ELISA) for detection of antibodies to the Apx toxins of Actinobacillus pleuropneumoniae. Vet. Microbiol. 71, 81-87.

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Opriessnig, T., Hemann, M., Johnson, J.K., Heinen, S., Giménez-Lirola, L.G., O’Neill, K.C., Hoang, H., Yoon, K.-., Gottschalk, M., Halbur, P.G., 2013. Evaluation of diagnostic assays for the serological detection of Actinobacillus pleuropneumoniae on samples of known or unknown exposure. J. Vet. Diagn. Invest. 25, 61-71. Pattison, I.H., Howell, D.G., Elliot, J., 1957. A Haemophilus-like organism isolated from pig lung and the associated pneumonic lesions. J. Comp. Pathol. 67, 320-330. Perry, M.B., Angen, Ø., MacLean, L.L., Lacouture, S., Kokotovic, B., Gottschalk, M., 2012. An atypical biotype I Actinobacillus pleuropneumoniae serotype 13 is present in North America. Vet. Microbiol. 156, 403-410. Pirofski, L.A., Casadevall, A., 2002. The meaning of microbial exposure, infection, colonisation, and disease in clinical practice. Lancet Infect. Dis. 2, 628-635. Pohl, S., Bertschinger, H.U., Frederiksen, W., Mannheim, W., 1983. Transfer of Haemophilus pleuropneumoniae and the Pasteurella haemolytica-like organism causing porcine necrotic pleuropneumonia to the genus Actinobacillus (Actinobacillus pleuropneumoniae comb. nov.) on the basis of phenotypic and deoxyribonucleic acid relatedness. Int. J. Syst. Bacteriol. 33, 510-514. Sakano, T., Shibata, I., Samegai, Y., Taneda, A., Okada, M., Irisawa, T., Sato, S., 1993. Experimental pneumonia of pigs infected with Aujeszky’s disease virus and Actinobacillus pleuropneumoniae. J. Vet. Med. Sci. 55, 575-579. Sanford, S.E., Josephson, G.K., 1981. Porcine Haemophilus pleuropneumonia epizootic in southwestern Ontario: clinical, microbiological, pathological and some epidemiological findings. Can. J. Comp. Med. 45, 2-7. Savoye, C., Jobert, J.L., Berthelot-Herault, F., Keribin, A.M., Cariolet, R., Morvan, H., Madec, F., Kobisch, M., 2000. A PCR assay used to study aerosol transmission of Actinobacillus pleuropneumoniae from samples of live pigs under experimental conditions. Vet. Microbiol. 73, 337-347. Schaller, A., Kuhn, R., Kuhnert, P., Nicolet, J., Anderson, T.J., Macinnes, J.I., Segers, R.P., Frey, J., 1999. Characterization of apxIVA, a new RTX determinant of Actinobacillus pleuropneumoniae. Microbiology 145, 2105-2116. Schiefer, B., Moffatt, R.E., Greenfield, J., Agar, J.L., Majka, J.A., 1974. Porcine Hemophilus parahemolyticus pneumonia in Saskatchewan. I. Natural occurrence and findings. Can. J. Comp. Med. 38, 99-104. Schuchert, J.A., Inzana, T.J., Angen, Ø., Jessing, S., 2004. Detection and identification of Actinobacillus pleuropneumoniae serotypes 1, 2, and 8 by multiplex PCR. J. Clin. Microbiol. 42, 4344-4348. Sebunya, T.N., Saunders, J.R., 1983. Haemophilus pleuropneumoniae infection in swine: a review. J. Am. Vet. Med. Assoc. 182, 1331-1337. Shin, M.-., Kang, M.L., Cha, S.B., Lee, W.-., Sung, J.H., Yoo, H.S., 2011. An immunosorbent assay based on the recombinant ApxIa, ApxIIa, and ApxIIIa toxins of Actinobacillus pleuropneumoniae and its application to field sera. J. Vet. Diagn. Invest. 23, 736-742. Shope, R.E., 1964. Porcine contagious pleuropneumonia. I. Experimental transmission, etiology, and pathology. J. Exp. Med. 119, 357-368. Sidibé, M., Messier, S., Lariviere, S., Gottschalk, M., Mittal, K.R., 1993. Detection of Actinobacillus pleuropneumoniae in the porcine upper respiratory tract as a complement to serological tests. Can. J. Vet. Res. 57, 204-208. Sørensen, V., Barfod, K., Feld, N.C., Nielsen, J.P., Enøe, C., Willeberg, P., 1997. Evaluation of a polyclonal blocking ELISA and a complement fixation test detecting antibodies to Actinobacillus pleuropneumoniae serotype 2 in pig serum. International Symposia on Veterinary Epidemiology and Economics (ISVEE) proceedings ISVEE 8: Proceedings of the 8th Symposium of the International Society for Veterinary Epidemiology and Economics, Paris, France (published as Epidèmiologie et Santé Animale, Issues 31-32),. Stockhofe-Zurwieden, N., Hensel, A., van Leengoed, L., Buijs, R., Pohlenz, J., 1994. Lung alterations and bacterial colonization of the respiratory tract of pigs infected with Actinobacillus pleuropneumoniae serotype 9. In: Hensel, A. (Ed.), PhD Thesis: An aerosol exposure system for immunization of pigs against Actinobacillus pleuropneumoniae infection. Utrecht University, Utrecht, pp. 61-74. Tonpitak, W., Rohde, J., Gerlach, G.F., 2007. Prevalence of “Actinobacillus porcitonsillarum” in porcine tonsils and development of a diagnosis duplex PCR differentiating between “Actinobacillus porcitonsillarum” and Actinobacillus pleuropneumoniae. Vet. Microbiol. 122, 157-165. Velthuis, A.G., De Jong, M.C., Kamp, E.M., Stockhofe, N., Verheijden, J.H., 2003. Design and analysis of an Actinobacillus pleuropneumoniae transmission experiment. Prev. Vet. Med. 60, 53-68. Velthuis, A.G., De Jong, M.C., Stockhofe, N., Vermeulen, T.M., Kamp, E.M., 2002. Transmission of Actinobacillus pleuropneumoniae in pigs is characterized by variation in infectivity. Epidemiol. Infect. 129, 203-214.

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Vigre, H., Angen, Ø., Barfod, K., Lavritsen, D.T., Sørensen, V., 2002. Transmission of Actinobacillus pleuropneumoniae in pigs under field-like conditions: emphasis on tonsillar colonisation and passively acquired colostral antibodies. Vet. Microbiol. 89, 151-159. Voss, T., 2012. Actinobacillus pleuropneumoniae (APP): Successful emergency therapy on a pig fattening farm (a case report). Der Praktische Tierarzt 93, 1036-1039. Woolhouse, M.E.J., Dye, C., Etard, J.-., Smith, T., Charlwood, J.D., Garnett, G.P., Hagan, P., Hii, J.L.K., Ndhlovu, P.D., Quinnell, R.J., Watts, C.H., Chandiwana, S.K., Anderson, R.M., 1997. Heterogeneities in the transmission of infectious agents: Implications for the design of control programs. Proc. Natl. Acad. Sci. U. S. A. 94, 338-342. Zhou, L., Jones, S.C.P., Angen, Ø., Bossé, J.T., Nash, J.H.E., Frey, J., Zhou, R., Chen, H.C., Kroll, J.S., Rycroft, A.N., Langford, P.R., 2008. Multiplex PCR that can distinguish between immunologically cross-reactive serovar 3, 6, and 8 Actinobacillus pleuropneumoniae strains. J. Clin. Microbiol. 46, 800-803.

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2 Detection of Actinobacillus pleuropneumoniae in pigs by real-time quantitative PCR for the apxIVA gene T.J. Tobiasa, A. Boumaa, D. Klinkenberga, A.J.J.M. Daemena, J.A. Stegemana, J.A. Wagenaarb,c and B. Duimb

published in: The Veterinary Journal 2012, 193, 557-560

a

Department of Farm Animal Health, Faculty of Veterinary Medicine, Utrecht University, Yalelaan 7, Utrecht 3584 CL, The Netherlands.

b

Department of Infectious Diseases and Immunology, Faculty of Veterinary

Medicine, Utrecht University, Yalelaan 1, Utrecht 3584 CL, The Netherlands. c

Central Veterinary Institute of Wageningen UR, PO Box 65, Lelystad 8200 AB, The Netherlands.

Chapter 2

ABSTRACT A real-time quantitative PCR (qPCR) for detection of the apxIVA gene of Actinobacillus pleuropneumoniae was validated using pure cultures of A. pleuropneumoniae and tonsillar and nasal swabs from experimentally inoculated Caesarean-derived/ colostrum-deprived piglets and naturally infected conventional pigs. The analytical sensitivity was 5 colony forming units/reaction. In comparison with selective bacterial examination using tonsillar samples from inoculated animals, the diagnostic sensitivity of the qPCR was 0.98 and the diagnostic specificity was 1.0. The qPCR showed consistent results in repeatedly sampled conventional pigs. Tonsillar brush samples and apxIVA qPCR analysis may be useful for further epidemiological studies and monitoring for A. pleuropneumoniae.

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apxIVA qPCR for A. pleuropneumoniae detection

INTRODUCTION Actinobacillus pleuropneumoniae is one of the major causes of respiratory disease in pigs and is established as an endemic infection on most commercial pig farms (Sørensen et al., 2006). Infection with A. pleuropneumoniae is often subclinical, but sometimes results in severe clinical signs, growth retardation or mortality, causing welfare problems and substantial economic losses (Gottschalk and Taylor, 2006). An accurate diagnosis is essential for proper treatment and for implementation of disease control programmes. Selective bacterial examination (SBE) can be used for detection of A. pleuropneumoniae in tonsillar or nasal swab samples collected from live pigs. However, suspected A. pleuropneumoniae colonies tend to be overgrown by other bacterial species and it can be difficult to differentiate between A. pleuropneumoniae and other Pasteurellaceae from the oropharyngeal cavity (Møller et al., 1996). Thus, the sensitivity and specificity of tests based on culture for detection of A. pleuropneumoniae are low. A number PCR targets have been described for identification or serogrouping of A. pleuropneumoniae (Chiers et al., 2001; Jessing et al., 2003; Schuchert et al., 2004; Tonpitak et al., 2007; Angen et al., 2008; Zhou et al., 2008; Ito, 2010). Several conventional PCRs have been described for qualitative detection of A. pleuropneumoniae DNA in nasal or tonsillar swabs, or from tissue samples, such as tonsil or lung (Lo et al., 1998; Savoye et al., 2000; Fittipaldi et al., 2003; SerranoRubio et al., 2008). When studying the epidemiology or pathogenesis of A. pleuropneumoniae, it is often useful to obtain quantitative data on the pathogen load in individual pigs. Furthermore, the use of real-time PCR would enable high throughput sample analysis compared to SBE or conventional PCR. The aim of this study was to develop a real-time PCR for direct quantitative detection of A. pleuropneumoniae DNA in live pigs and to evaluate the test with samples from experimentally infected pigs or conventionally reared pigs originating from A. pleuropneumoniae infected farms.

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MATERIALS AND METHODS PCR primers, probes and amplification protocol The apxIVA gene was chosen as a suitable qPCR target gene because this gene is specific for all A. pleuropneumoniae serotypes and because information on this gene is available from qualitative PCR tests (Cho and Chae, 2003; Fittipaldi et al., 2003). PCR was performed using two primers described previously, APXIVANEST1-F and APXIVANEST1-R (Schaller et al., 2001). A conserved sequence within the predicted PCR product, identified with BLAST1 was used to design a TaqMan probe (Table 1). The real-time PCR was performed in a total volume of 25 µL per well with 0.2 µM each primer and probe in Tris ethylene diamine tetraacetic acid (10 mM Tris HCl, 1 mM EDTA, pH 8.0), 2 x Premix Ex Taq (TaKaRa), 0.5 µL PCR grade water and 10 µL template (from 200 µL Instagene purification, as indicated below) or 10 µL saline as a negative control. The template consisted of DNA extracts (from samples or standards) or negative controls. A BioRad iQ5 thermocycler was used for qPCR analysis. The PCR programme consisted of an initial denaturation (60 s at 95 °C), followed by 40 cycles of 10 s at 95 °C, 30 s at 56 °C and 30 s at 72 °C. Quantification of A. pleuropneumoniae colony forming units (CFUs) in samples was performed using the iQ5 algorithm. A standard curve was included by duplicate testing of three standard samples of a dilution of A. pleuropneumoniae serotype 1536, consisting of 5 x 101, 5 x 103 and 5 x 105 CFUs, respectively. The optimal threshold of fluorescence was determined as a cut-off for optimal reaction efficiency within the assay. Samples were considered to be positive when duplicates showed comparable results and the increase in fluorescence had a sigmoid curve. Table 1. Sequences of primers and fluorescent probe for detection of Actinobacillus pleuropneumoniae Name

Sequence

Locationa

Product size

Reference

apxIVANEST1-F 5’-GGGGACGTAACTCGGTGATT-3’ 6050-6069 Schaller (2001) 377 bp apxIVANEST1-R 5’-GCTCACCAACGTTTGCTCAT-3’ 6427-6407 Schaller (2001) apxIVAPr (probe) 5’-FAM-CGGTGCGGACACCTATATCT-BHQ1-3’ 6162-6182 This study a

Nucleotide location in AF021919

1

http://blast.ncbi.nlm.nih.gov/blast

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apxIVA qPCR for A. pleuropneumoniae detection

Bacterial strains and growth conditions Bacteria were grown at 37°C in an atmosphere of 5% CO2 on sheep blood agar (SBA) (Biotrading) or heart infusion agar with 5% sheep blood (HIS) and, depending on their β-nicotinamide adenine dinucleotide (NAD) requirement, supplemented with 0.05% NAD (AppliChem GmbH) (HIS-V plates). Casitone–glucose–vitamin HIS agar plates supplemented with NAD and Clindamycin, Gentamycin, Vancomycin and Amphoterricine B (CGVA) were employed for selective culture of β-NAD dependent Pasteurellaceae (Velthuis et al., 2003), using three 10-fold serial dilutions per sample. After detection and quantification of A. pleuropneumoniae-like colonies with SBE, two colonies suspected to be A. pleuropneumoniae on the basis of colony morphology were selected from each primary plate and subcultured on HIS-V plates. Isolates were confirmed as A. pleuropneumoniae by positive satellite growth, a positive Christie–Atkins–Munch–Petersen (CAMP) reaction near a streak of Staphylococcus aureus on SBA, urease activity and mannitol fermentation in the appropriate test medium with added β-NAD. Samples and sample handling Validation tests were performed with samples from pure cultures, tonsillar and nasal samples from Caesarean-derived/colostrum-deprived (CD/CD) piglets and tonsillar samples from conventional pigs. Tonsillar samples from live pigs were obtained by brushing the surface of the tonsil for 10 s with a sterile toothbrush, which was immersed in 10 mL 0.9% sterile NaCl (saline) solution for 20 min. After removing the toothbrush, 100 µL were used for 10-fold serial dilutions. Samples were concentrated by centrifugation (1,500 g for 15 min) and 500 µL saline were added to the pellet, which was stored at -20 °C before DNA isolation. Nasal samples from CD/CD piglets were derived from each nostril by twirling a cotton swab (Applimed SA) for 5 s and then the swab was immersed in 1 mL saline for 20 min. After removing the swab, 100 µL sample were used for 10-fold serial dilutions and the remaining sample was stored at -20°C.

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DNA isolation Instagene Matrix kit (BioRad) was used for DNA isolation (Frey, 2003). DNA isolation from bacterial cultures was performed according to the manufacturer’s instructions. A minor modification of Frey’s protocol was used for DNA isolation from tonsillar samples (200 µL) and nasal samples (400 µL). After centrifugation (5 min at 12,000 g), the supernatant was discarded and 200 µL Instagene Matrix were added to each pellet, then samples were heated at 56 °C for 30 min. After mixing for 10 s, samples were heated at 100 °C for 8 min, mixed again for 10 s, centrifuged for 5 min at 12,000 g and then the DNA was stored at -20 °C. Prior to qPCR analysis, samples were thawed, briefly mixed and centrifuged at 12,000 g for 5 min. Pure saline samples were used as negative controls. Assay validation with pure cultures The specificity of the qPCR was evaluated using DNA from pure cultures of a range of reference and field strains of A. pleuropneumoniae, other Pasteurellaceae and other porcine bacterial pathogens (see Appendix A: Supplementary table 1). To evaluate the within and between test accuracy, duplicate serial dilution series of a 6 h culture of A. pleuropneumoniae 1536 in saline, from 5x100 to 5x106 CFUs/well, were tested in triplicate on two different PCR plates on the same day using a threshold of 200 relative fluorescence units as a cut-off. The limit of detection of the qPCR was determined as the lowest number of CFUs from the same 10-fold serial dilution in which all three replicates displayed a positive result. A standard curve was created with a serial dilution series, tested in triplicate. Assay validation with samples from Caesarean-derived/colostrum-deprived piglets To validate test specificity, tonsillar samples from 77 uninfected CD/CD piglets were collected. The correlation between quantitative results obtained by qPCR and SBE was determined in tonsillar and nasal samples from 10 CD/CD piglets inoculated with A. pleuropneumoniae. The inoculum was prepared by growing A. pleuropneumoniae 1536 overnight on a HIS-V plate. Thereafter, one colony was suspended in 200 µL saline and another HIS-V plate was inoculated with 50 µL suspension. After 6 h, the plate was rinsed with 5 mL saline and an appropriate concentration (2.5 106 CFUs/

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apxIVA qPCR for A. pleuropneumoniae detection

mL) was prepared, guided by optical density measurements. Pigs were inoculated intranasally at 28 days of age with 1.0 mL inoculum in each nostril. Tonsillar and nasal samples were collected at 1, 2, 4, 6, 8, 11, 13, 15, 18 and 21 days postinoculation (dpi). Pigs were examined daily for signs of pleuropneumonia (elevated body temperature and abnormal respiration). On day 21, piglets were euthanased with pentobarbital (Euthanimal, Alfasan). In total, 65 nasal and 65 tonsillar samples were tested using SBE and qPCR. Serum samples were collected at -1 and 21 dpi and tested in the complement fixation test (CFT; Nielsen, 1974) at the Animal Health Service (Deventer, The Netherlands). At postmortem examination, pneumonia and pleurisy were scored as described by Hannan et al. (1982). Homogenised tonsils and pneumonic lesions were sampled for bacterial growth on CGVA, SBA, HIS-V and chocolate agar plates, and growth of A. pleuropneumoniae was confirmed, as described above. The experiments with CD/CD piglets were authorised by the Animal Care and Ethics Committee (AEC) of Utrecht University, according to the Dutch Experiments on Animals Act (licence numbers DEC2009.III.10.099, DEC2010. II.02.025 and DEC2010.II.02.027). An analgesic (Fentanyl, B. Braun Melsungen AG) was administered to pigs showing clinical signs. Pigs were euthanased when the humane endpoint was reached, as accorded by the AEC. Assay validation with samples from conventional pigs To provide data on the specificity of the qPCR, 70 gilts of at least 14 weeks of age, housed on three A. pleuropneumoniae free farms, were randomly selected and sampled. The farms were considered to be free from A. pleuropneumoniae on the basis of the following criteria: no reports of A. pleuropneumoniae outbreaks during the previous 5 years, absence of lesions consistent with A. pleuropneumoniae upon slaughterhouse monitoring and negative test results on serological monitoring for A. pleuropneumoniae by ApxIV-ELISA (Dreyfus et al., 2004). An observational cohort study was performed on two A. pleuropneumoniae infected wean-to-finish farms with pigs originating from the same farrowing herd. The prevalence and the change in A. pleuropneumoniae load in tonsillar samples over time were investigated. Tonsillar samples were collected at 4, 10, 16 and 24 weeks of age from 20 pigs per farm on the same day. The presence of A. pleuropneumoniae was confirmed by clinical signs,

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Chapter 2

postmortem examination and serology. qPCR results for A. pleuropneumoniae were compared between the two farms and between points in time, expressed by age and time since the first positive sample. The tonsillar samples used in field validation experiments were obtained in compliance with the Dutch Act on the Practice of Veterinary Medicine, as agreed by the Institutional Animal Care and Ethics Committee. Statistical analysis Statistical analysis was performed using R version 2.11.1 and SPSS 16.0.2 for Windows. To evaluate assay accuracy, linear models with threshold cycle as outcome and log10 of the bacterial cell concentration, assay and their interactions as explanatory variables were evaluated. Triplicates within assays were analysed separately as a random effect. Proportions were determined with left one-sided confidence intervals (CIs) based on the binomial distribution. The correlations between quantitative results of SBE and qPCR were obtained by performing a conditional analysis on test positive samples. Partial correlation analysis was performed for the log10 of quantitative SBE and qPCR results for both nasal and tonsillar samples, with piglet number as the controlled variable to adjust for repeated measurements within the same animal. For comparison of qPCR results in samples from A. pleuropneumoniae infected farms, linear mixed models with the individual pig as a random intercept and farm, age, time since the first positive sample and their interactions as explanatory variables were evaluated. The Akaike Information Criterion was used to select the best models.

RESULTS Assay validation with pure cultures The qPCR for apxIVA produced positive results only when testing DNA from A. pleuropneumoniae, whereas samples containing DNA from other bacterial species tested negative (see Appendix A: Supplementary table 1). A standard curve indicated that the qPCR had a reaction efficiency of 93.0% (see Appendix A: Supplementary fig. 1). No random effect of intra-assay variability was found. A random effect of interassay variability was observed on the intercept, but not on the regression coefficient,

32

apxIVA qPCR for A. pleuropneumoniae detection

indicating the need to incorporate a standard series for the quantification of samples in each assay. The analytical sensitivity was 5 x 100 CFUs per reaction. This resulted in an analytical sensitivity of 250 CFUs per tonsillar sample. Assay validation with samples from Caesarean-derived/colostrum deprived piglets All 77 tonsillar samples from uninfected CD/CD piglets tested negative by qPCR, resulting in an estimated test specificity of 100% (CI: 0.96–1.00). In total, 56/57 SBE positive tonsillar samples from A. pleuropneumoniae inoculated pigs tested positive by qPCR (Table 2), resulting in an estimated sensitivity of 98% (CI: 0.92–1.00). Of the nasal samples, 42/65 (64.6%) samples tested positive (Table 2), but numbers of CFUs decreased over time in SBE and qPCR (see Appendix A: Supplementary fig. 2). Four of five surviving pigs showed negative SBE and qPCR results for nasal samples at 21 days. Seven of eight tonsillar samples and 5/21 nasal samples were SBE negative but qPCR positive. Sequence analysis of these qPCR products showed 98–100% similarity to GenBank AF021919. One tonsillar and 16 nasal samples were negative in both SBE and PCR. Partial correlation analysis of test positive samples demonstrated a good correlation between the log10 of SBE and qPCR for tonsillar (0.72) and nasal (0.88) samples (see Appendix A: Supplementary fig. 3). Five pigs survived until 21 days. Clinical signs varied considerably between pigs. Body temperature was temporarily elevated in all pigs, but respiratory signs were observed in some pigs more often than in others (see Appendix A: Supplementary fig. 2). Partial correlation analysis showed that abnormal respiration correlated better with the quantitative test results from nasal (ρ ~ 0.5) than from tonsillar samples (ρ ~ 0.4). All CFT titres at day -1 were 40 °C or showed eminent signs of pain they were treated with Fentanyl (B. Braun Melsungen AG, Melsungen, Germany). Fentanyl is a potent analgesic, but does not bear anti-inflammatory capacities. Pigs were euthanized when the results of the daily welfare assessment exceeded the criteria accorded by the AEC. Inoculation Inoculation was performed intranasally with a six hour culture of A. pleuropneumoniae reference strain 1536. A. pleuropneumoniae was cultured on Heart Infusion agar with 5% sheep erythrocytes and 0.2% β-NAD (Nicotinamide adenine dinucleotide) (HIS-V) overnight at 37 °C and 5% CO2. The next day 1 colony was suspended in 1 mL of saline and 50 μL was plated on a new HIS-V plate and incubated. After six hours, the plate was washed with 5 mL of saline and diluted to 2.5 × 106 CFUs/mL guided by optical density measurements. Before inoculation pigs were sedated with Azaperon (Stressnil®, Janssen Animal Health, Brussels, Belgium) and subsequently pigs were inoculated by dripping 1 mL of the inoculum in each nostril during inhalation (total 5 × 106 CFUs). Before and after inoculation the concentration of the inoculum was determined by plating of serial dilution series. Inoculation dose and method of administration were chosen based on the results of two pilot studies where sufficient variation in disease severity was obtained, when 106 Colony Forming Units (CFUs) A. pleuropneumoniae serotype 2 (strain 1536) was intranasally administered in CD/CD pigs.

47

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Chapter 3

Table 1. Cumulative clinical disease score, necropsy and bacteriology results for A. pleuropneumoniae infection per pig Clinical score Pathology Survival Bacteriology Pig Run / until Pair Pleurisy Pneumonia status pen RHS4 RHS20 Lung Nose day score score Tonsil sample tonsil 1 2 3 4 5 6 7 8 9 10

I C I C I C I C I C I C I C I C I C I C

11 12

S S S S

1/4 1/5 1/6 1/7 1/8 2/4 2/5 2/6 2/7 2/8

1/3 2/3

4 21 4 21 2 21 21 21 21 21 7 21 21 21 21 21 21 21 4 21

31.13 3.10 13.68 2.48 66.88 2.35 6.80 7.90 16.18 5.58 23.70 7.43 17.65 8.70 24.40 4.03 30.70 7.13 46.13 5.60

86.23 5.11 82.74 6.72 93.38 6.26 6.23 8.90 12.87 8.77 72.13 6.42 10.35 5.55 12.70 4.20 20.09 7.55 89.23 7.82

20.7 0 19.3 0 33.9 0 0 0 5.9 3.2 6.9 0 7.1 0 0 0 9.2 0 28.1 0

6.6 0 1.0 0.26 33.2 1.5 0 0 0.3 1.6 4.7 0 8.6 0 0 0 15.0 0 26.6 0

+ ND + + ND ND + + ND ND + ND + ND + ND

+ + + + + + + + -

+ + + + + + ± dub + + + + + + + -

11* 21 21 21

0.31 0.89 0.68 0.56

ND 5.39 4.05 5.27

0 0 0 0

0 0 0 0

ND -

-

-

I = inoculated, C = contact, S = Sentinel pig, + = A. pleuropneumoniae confirmed, ± dub = dubious growth, - = no growth, ND = Not determined, * euthanised due to lameness.

Samples Before examination, restraining or sampling, all personnel had to change boots, coverall and gloves or each pen. To minimise the risk of transmission due to sampling or examination the contact pig was always sampled before the inoculated pig. Tonsil and nasal samples were collected on post inoculation day −1, 1, 2, 4, 6, 8, 11, 13, 15, 18 and 21, or before euthanasia of severely diseased animals. Nasal swabs were obtained with a small cotton swab (Applimed SA, Italy) and tonsil scrapings were

48

Disease severity and transmission of A. pleuropneumoniae

obtained by brushing the tonsils for 10 s with a soft toothbrush. On day −1 and day 21 the pigs were bled. Bacteriology Tonsil brush samples were submerged in 10 mL and nose swabs in 1 mL saline and thoroughly mixed for 20 min before selective bacteriologic examination (SBE). Subsequently, tenfold serial dilution series were made of tonsil brush (10-1, 10-2, 10-3) and nose swab samples (100, 10-1, 10-2). Per dilution 50 μL was plated on a selective agar plate with Clindamycin, Gentamicin, Vancomycin and Amphotericin (CGVA plate) (Velthuis et al., 2003) and incubated at 37 °C and 5% CO2. A. pleuropneumoniae suspected colonies were counted after overnight incubation. Per sample two suspected colonies were confirmed as A. pleuropneumoniae when positively tested for satellite growth, Christie–Atkins–Munch–Petersen (CAMP) reaction, urease and mannitol fermentation. Bacterial counts were back calculated to whole sample constituents in CFUs. Additional analyses were performed with an apxIVA qPCR (Tobias et al., 2012) (see Appendix A: Supplementary file 1). Clinical disease score A clinical score (CS) was obtained for each pig daily from day −8 to day 21 post inoculation by the same examiner. CS was calculated as the average score (on a 0 – 4 scale) for eight different clinical parameters, scored as described by Hoeltig et al. (2008): behaviour, locomotion score, vomiting, body temperature, respiratory breathing type, respiratory sounds, breathing frequency and coughing. Unlike in Hoeltig et al., cyanosis was not included, because it was never observed. Neither was feed intake, which could only be observed per pair. Clinical scores were obtained twice a day from day 0 – 5 and once a day thereafter, while bacterial samples were collected less frequently. To include all information of clinical observations, average clinical scores (AvgCS) for the days of sampling were derived by averaging all observed CS from one bacterial sampling moment to the next. For example on day 11 the AvgCS was calculated based on the CS of day 9 until 11. To calculate a cumulative respiratory health score at day 4 and 20 (RHS4, RHS20), like in Hoeltig’s study, CS was summed until day 4 or 20 and five points per day were added for each remaining day

49

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Chapter 3

after an animal had died or was euthanized. Subsequently, the summed score was expressed as a percentage of the maximum obtainable score. Serology Serology was applied to confirm the cause of clinical symptoms in I-pigs, as well as to detect possible false negative culture or qPCR results in C-pigs. Analysis of serology was performed at the Animal Health Service (Deventer, The Netherlands). Serum samples of day −1 and at euthanasia were analysed by Complement Fixation Test (CFT) titration (Nielsen, 1974) and a commercial App serotype 2 ELISA (Biovet, St Hyacinthe, Canada). CFT results > 80 in final sample or a distinct (> 0.2) increase of optical density were considered indicative for seroconversion and infection. Additionally, serum samples obtained at euthanasia were analysed by an ApxIV ELISA (Idexx, Maine, USA). Serum samples of pigs that died before day 14 were not analysed by CFT or ApxIV ELISA, because no seroconversion was expected. Pathology At necropsy, all pigs were examined by a veterinary pathologist (at the Veterinary Pathology Diagnostic Centre, Utrecht University). Tonsils were removed and homogenised and macroscopic lung and pleurisy lesions were assessed. A lesion score per lung was calculated, as described by Hannan et al. (1982). Lung lesions and tonsil homogenates were sampled by bacteriologic culture. Statistical analyses The effects of time and pig status (I or C) on AvgCS and the log10 + 1 of CFUs count in tonsil samples after day 0 were evaluated using mixed effect models with pig number as random effect, to account for repeated observations. Spearman’s rank correlation test was used to investigate the correlation between ranks of cumulative clinical scores (RHS4 or RHS20), CFT titres and pathology scores in individual pigs. To study the effect of disease severity (AvgCS) on transmission from the I-pig to the C-pig (transmission chain), three analyses were performed. First, correlation between AvgCS and log10 CFUs found in tonsil and nasal samples was

50

Disease severity and transmission of A. pleuropneumoniae

tested by Spearman’s Partial rank correlation using pig number as controlled variable. Second, the effect of disease severity and bacterial load on the rate of transmission was evaluated. The change from a negative to at least two consecutive positive samples in the contact pig was considered indicative for transmission. Presence or absence of transmission between two samplings (0/1) was used as response variable in a Generalized Linear Model with a binomial error distribution, a complementary log-log link function (Velthuis et al., 2002) and ln (time) between samplings as offset. AvgCS and log10 CFUs +1 counts in tonsil and nasal samples were evaluated as model terms and effect estimates are provided with 95% profile confidence intervals. A sensitivity analysis was carried out with respect to the use of SBE as indicator for infectiousness and transmission, by using qPCR results as an additional test (see Appendix A: Supplementary file 1). The qPCR may be more sensitive than SBE, but specificity for indicating infected and infectious pigs rather than non-viable bacteria may be lower. Finally, the association between disease severity scores within the transmission chain was analysed. The correlation between CS (AvgCS) of the I-pig, from day of inoculation onwards, with CS (AvgCS) of the C-pig, from the first day of a positive culture and onwards, within pairs, was evaluated using Spearman’s Partial rank correlation analysis. Pair number was used as controlled variable. Statistical analyses were performed using statistical software R version 2.11.1 (R Development Core Team, 2010) and additional packages pcor.R and lme4. Corrected Akaike Information Criterion for finite sample sizes (AICc) was used to select the models fitting the data best (Burnham and Anderson, 2002).

RESULTS Bacteriology results Inoculation of I-pigs was successful as demonstrated by at least two or more positive results in SBE in nasal or tonsillar samples in I-pigs upon inoculation. Sequentially taken tonsillar samples of I-pigs were almost all positive and more or less constant after day 2, results of nasal samples showed decreasing CFUs of A. pleuropneumoniae over time (Table 2 and Fig. 1) and at day 21 only one of five surviving I-pigs had a positive nasal swab sample in SBE. Transmission occurred in six out of ten pairs as at

51

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Chapter 3

least two or more positive samples in C-pigs tested positive (Fig. 1). In one C-pig (pair 1) two non-consecutive samples were positive in culture, starting long after the I-pig had died and we assume that transmission did not occur in that pair. Only one nasal sample of the C-pigs showed a positive result in SBE (Fig. 1). Table 2. Results of tonsil and nasal sample selective bacterial examination for A. pleuropneumoniae in time

2

4

6

1

I C I C I C I C I C I C I C I C I C I C

-

tn n tn tn tn tn tn n tn n -

tn tn tn tn tn tn tn tn t tn tn -

tn tn † t tn t tn tn t tn t t tn t tn -

† † -

S S S S

-

-

-

-

2 3 4 5 6 7 8 9 10

11 12

8

ApxIV ELISA

1

App S2 ELISA

Status

-1

Serology

-

ND 0.9−0.95

Statistical model evaluation Table 2 shows the models with lowest AIC within each of the six subgroups. Across the six subgroups, the models with a beta-binomial distribution for the probability of infection within litters fitted much better (lower AIC) than those with a binomial distribution. Among those, the subgroup model without zero inflation (subgroup 2) fitted the data best (least complex model). Across all models, the simplest of subgroup 2, without dam characteristic explanatory variables, was considered to fit the data best. Inclusion of zero inflation into the final model of subgroup 2 did not improve the fit (two points higher in AIC), whereas adding any of the dam characteristic explanatory variables increased the AIC by 0 to 2 points (not shown). From the best fitting model it was concluded that the probability of infection from a piglet is best described by a beta binomial distribution with ppiglet = 0.28 (95% CI: 0.21 - 0.36) and

ρ = 0.53 (95% CI: 0.42 - 0.63).

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Table 2. Results of model selection procedure For each subgroup of models the model with lowest AIC is reported. The subgroup model with lowest AIC is printed in bold. X = parameter included. Model subgroup

Zero inflation

1 2 3 4 5 6 7 8

X X X X X X

Inclusion of dam characteristic explanatory variables in pdam ppiglet X X X X

X X X X X X

Probability distribution

AIC

Binomial Beta binomial Binomial Beta binomial Binomial Beta binomial Binomial Beta binomial

610,0 308,0 453.0 310.0 434,7 310,0 443.9 309.9

The results of the power analysis showed that, with this study design, sample size and with the observed ppiglet = 0.28 and variation (ρ = 0.53) a true difference in proportions of infected pigs of 0.20 can be detected with a power of 80%. Finally, the fit of a transmission model that also accounted for transmission amongst piglets (AIC = 316.5) was less than the fit of a logistic regression model with a beta binomial distribution.

DISCUSSION The aim of this study was to investigate the pattern of clustering of colonisation of piglets with A. pleuropneumoniae at the end of the suckling period and to examine possible factors associated with the probability of colonisation. It was shown, that colonisation of piglets with A. pleuropneumoniae was clustered across litters as well as within litters. Approximately 40% of the litters remained free from A. pleuropneumoniae during the suckling period, despite the finding that all sows tested positive for A. pleuropneumoniae, but no associated risk factors was identified that might explain this observation. Our findings are in accordance with the results of Vigre et al. (2002), as in both studies similar percentages of piglets tested positive before weaning, but in our study more information on the level of clustering on litter level before weaning was provided.

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A. pleuropneumoniae colonisation in suckling piglets

The best fit of a beta binomial model suggests some kind of random effect on litter level, but in this study no dam related factors could be identified that were associated with the probability of piglet infection. This may be explained if colonisation was not inhibited by the maternally derived antibodies that were investigated (anti- ApxI, ApxII, ApxIII and Omp antibodies), but rather by antibodies against other antigens or by other immunological, e.g. more cell mediated, factors. Indeed previous studies have suggested that anti-Apx and anti-Omp antibodies induced by vaccination are less likely to be associated with the susceptibility for colonisation (Velthuis et al., 2003), than with a protective effect against the development of clinical disease (Haga et al., 1997; Tumamao et al., 2004). It was anticipated that a possible association with dam parity would suggest such an alternative protective mechanism, as the level of immunity in sows is related to parity, either by antibodies (Klobasa and Butler, 1987), by interferon gamma (Uddin et al., 2010), or other mechanisms. For example, Miller et al. (2013) showed recently that although levels of immunity in sows were equal, parity effects on immunity were observed in offspring. In our study however, dam parity was not found to be associated to the infection probability of piglets. If there are any dam related risk factors for piglet colonisation by A. pleuropneumoniae, it is presently unclear what they are. In positively tested litters also clustering was observed, as there were a few litters with very high proportions of positive litters. The poorer fit of the model that accounted for transmission within the litter compared to the fit of the beta binomial logistic regression model suggests that transmission among suckling piglets did not occur on a large scale. Nevertheless we cannot conclude that transmission among suckling piglets does not occur at all, as the study was not designed for this specific purpose. The final models suggest that the infectiousness of the sows follows a beta distribution. Although the amount of A. pleuropneumoniae CFUs on the sow’s tonsil was included to account for infectiousness, no association was found with colonisation in offspring. This may be explained by the fact that sows were sampled only once, three weeks before parturition, and the infectivity of individual pigs has shown to vary considerably from day to day (Velthuis et al., 2002).

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There was more variation in colonised piglets per litter than expected. It is unclear whether these results were found because a strict hygiene and animal management protocol was applied, or whether transmission between pens would not occur on a large scale anyway. For extrapolation of the results to the field, the unexplained large variation in colonised piglets per litter may be the main obstacle, as in regular pig farming these preventive measures are applied less strictly. Results of this study do not indicate towards a specific point of control at dam level to reduce the proportion of colonised piglets at weaning. However, as many litters tested negative, the results of this study suggest that disease control measures in the farrowing room in general should aim to prevent transmission between litters to minimise the number of infected litters. It remains, however, uncertain if litters free from A. pleuropneumoniae remain free after weaning, or if they become colonised quickly after movement to weaner or finisher units. A follow-up study could be performed to evaluate subsequent transmission and colonisation after weaning. Conclusion The colonisation status of piglets for A. pleuropneumoniae was found to be clustered across and within litters, even though all sows were tested positive for A. pleuropneumoniae before farrowing. No dam related variables were found to be associated to either the probability of a litter to become infected, or the probability of individual pigs to become infected. Acknowledgements The farmers and the farm workers are greatly acknowledged for their hospitality and execution of the research protocol. MSD Animal Health, Boxmeer, The Netherlands is acknowledged for analysis of the serum samples. MSD was not involved in design of the study, data analysis or in composing the manuscript. The Central Veterinary Institute is acknowledged for serovar determination of the A. pleuropneumoniae isolates. Manon Houben, Wouter van Herten and Piet Dirven, are acknowledged for their cooperation. Katrien van Dongen, Bram Goesten, Leonie de Louw and all volunteers are acknowledged for assistance with sampling and data collection.

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Finally, anonymous reviewers are greatly acknowledged for their advice, resulting in significant improvements to the manuscript.

REFERENCES Agresti, A., 2013. Categorical data analysis. John Wiley & Sons, Hoboken, NJ, United States of America. Angen, Ø., Andreasen, M., Nielsen, E.O., Stockmarr, A., Baekbo, P., 2008. Effect of tulathromycin on the carrier status of Actinobacillus pleuropneumoniae serotype 2 in the tonsils of pigs. Vet. Rec. 163, 445-447. Burnham, K.P., Anderson, D.R., 2002. Model selection and multimodel inference, a practical information-theoretic approach. Springer, New York, USA. van Bunnik, B.A., Katsma, W.E., Wagenaar, J.A., Jacobs-Reitsma, W.F., de Jong, M.C., 2012. Acidification of drinking water inhibits indirect transmission, but not direct transmission of campylobacter between broilers. Prev. Vet. Med. 105, 315-319. Cheung, Y.B., 2006. Growth and cognitive function of Indonesian children: zero-inflated proportion models. Stat. Med. 25, 3011-3022. Chiers, K., Donne, E., Van Overbeke, I., Ducatelle, R., Haesebrouck, F., 2002. Actinobacillus pleuropneumoniae infections in closed swine herds: infection patterns and serological profiles. Vet. Microbiol. 85, 343-352. Cruijsen, T., van Leengoed, L.A., Kamp, E.M., Bartelse, A., Korevaar, A., Verheijden, J.H., 1995a. Susceptibility to Actinobacillus pleuropneumoniae infection in pigs from an endemically infected herd is related to the presence of toxin-neutralizing antibodies. Vet. Microbiol. 47, 219-228. Cruijsen, T., van Leengoed, L.A., Kamp, E.M., Hunneman, W.A., Riepema, K., Bartelse, A., Verheijden, J.H., 1995b. Prevalence and development of antibodies neutralizing the haemolysin and cytotoxin of Actinobacillus pleuropneumoniae in three infected pig herds. Vet. Q. 17, 96-100. Fablet, C., Marois, C., Kuntz-Simon, G., Rose, N., Dorenlor, V., Eono, F., Eveno, E., Jolly, J.P., Le Devendec, L., Tocqueville, V., Quéguiner, S., Gorin, S., Kobisch, M., Madec, F., 2011. Longitudinal study of respiratory infection patterns of breeding sows in five farrow-to-finish herds. Vet. Microbiol. 147, 329-339. Gottschalk, M., Taylor, D.J., 2006. Actinobacillus pleuropneumoniae. In: Straw, B.E., Zimmerman, J.J., D’Allaire, S., Taylor, D.J. (Eds.), Diseases of Swine. Blackwell Publishing, Ames, Iowa, USA, pp. 563-576. Haga, Y., Ogino, S., Ohashi, S., Ajito, T., Hashimoto, K., Sawada, T., 1997. Protective efficacy of an affinity-purified hemolysin vaccine against experimental swine pleuropneumonia. J. Vet. Med. Sci. 59, 115-120. Hamer-Barrera, R., Godinez, D., Enriquez, V.I., Vaca-Pacheco, S., Martinez-Zuniga, R., Talamas-Rohana, P., SuarezGuemez, F., de la Garza, M., 2004. Adherence of Actinobacillus pleuropneumoniae serotype 1 to swine buccal epithelial cells involves fibronectin. Can. J. Vet. Res. 68, 33-41. Hoflack, G., Maes, D., Mateusen, B., Verdonck, M., de Kruif, A., 2001. Efficacy of tilmicosin phosphate (Pulmotil premix) in feed for the treatment of a clinical outbreak of Actinobacillus pleuropneumoniae infection in growing-finishing pigs. J. Vet. Med. B Infect. Dis. Vet. Public Health 48, 655-664. Klobasa, F., Butler, J.E., 1987. Absolute and relative concentrations of immunoglobulins G, M, and A, and albumin in the lacteal secretion of sows of different lactation numbers. Am. J. Vet. Res. 48, 176-182. Kobisch, M., van den Bosch, J.F., Efficacy of an Actinobacillus pleuropneumoniae subunit vaccine. In: Proceedings of the 12th International Pig Veterinary Society Congress, The Hague, The Netherlands, 1992, 216. Mengelers, M.J., Kuiper, H.A., Pijpers, A., Verheijden, J.H., van Miert, A.S., 2000. Prevention of pleuropneumonia in pigs by in-feed medication with sulphadimethoxine and sulphamethoxazole in combination with trimethoprim. Vet. Q. 22, 157-162. Miller, Y.J., Collins, A.M., Emery, D., Begg, D.J., Smits, R.J., Holyoake, P.K., 2013. Piglet performance and immunity is determined by the parity of both the birth dam and the rearing dam. Ani. Prod. Sci. 53, 46-51. Nielsen, R., van den Bosch, J.F., Plambeck, T., Sørensen, V., Nielsen, J.P., 2000. Evaluation of an indirect enzymelinked immunosorbent assay (ELISA) for detection of antibodies to the apx toxins of Actinobacillus pleuropneumoniae. Vet. Microbiol. 71, 81-87. R Development Core Team. R: A language and environment for statistical computing. 2.11.1, 2010, R Foundation for Statistical Computing, Vienna, Austria. http://www.R-project.org

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Shin, M.K., Kang, M.L., Cha, S.B., Lee, W.J., Sung, J.H., Yoo, H.S., 2011. An immunosorbent assay based on the recombinant ApxIa, ApxIIa, and ApxIIIa toxins of Actinobacillus pleuropneumoniae and its application to field sera. J. Vet. Diagn. Invest. 23, 736-742. Sjölund, M., Zoric, M., Persson, M., Karlsson, G., Wallgren, P., 2011. Disease patterns and immune responses in the offspring to sows with high or low antibody levels to Actinobacillus pleuropneumoniae serotype 2. Res. Vet. Sci. 91, 25-31. Tobias, T.J., Bouma, A., Klinkenberg, D., Daemen, A.J.J.M., Stegeman, J.A., Wagenaar, J.A., Duim, B., 2012. Detection of Actinobacillus pleuropneumoniae in pigs by real-time quantitative PCR for the apxIVA gene. Vet. J. 193, 557-560. Tumamao, J.Q., Bowles, R.E., Van Den Bosch, H., Klaasen, H.L.B.M., Fenwick, B.W., Storie, G.J., Blackall, P.J., 2004. Comparison of the efficacy of a subunit and a live streptomycin-dependent porcine pleuropneumonia vaccine. Aus. Vet. J. 82, 370-374. Uddin, M.J., Grosse-Brinkhaus, C., Cinar, M.U., Jonas, E., Tesfaye, D., Tholen, E., Juengst, H., Looft, C., Ponsuksili, S., Wimmers, K., Phatsara, C., Schellander, K., 2010. Mapping of quantitative trait loci for Mycoplasma and tetanus antibodies and interferon-gamma in a porcine F2 duroc x pietrain resource population. Mamm. Genome 21, 409-418. Velthuis, A.G., De Jong, M.C., Stockhofe, N., Vermeulen, T.M., Kamp, E.M., 2002. Transmission of Actinobacillus pleuropneumoniae in pigs is characterized by variation in infectivity. Epidemiol. Infect. 129, 203-214. Velthuis, A.G., De Jong, M.C., Kamp, E.M., Stockhofe, N., Verheijden, J.H., 2003. Design and analysis of an Actinobacillus pleuropneumoniae transmission experiment. Prev. Vet. Med. 60, 53-68. Vigre, H., Angen, Ø., Barfod, K., Lavritsen, D.T., Sørensen, V., 2002. Transmission of Actinobacillus pleuropneumoniae in pigs under field-like conditions: Emphasis on tonsillar colonisation and passively acquired colostral antibodies. Vet. Microbiol. 89, 151-159. Wolfram Research. Mathematica. 8.0, 2010, Champaign, IL, United States. http://www.wolfram.com.

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APPENDIX A: SUPPLEMENTARY MATERIAL Supplementary table 1. Parity distribution within cohorts and qPCR results of litters per parity group Farm A

Farm B

Positively Parity Litters grouped tested litters tested positive (proportion) (proportion)

Cohort 1

Cohort 2

Cohort 3

Cohort 4

Total litters

1 2 3 4 5 6 7 8 9

6 2 3 1 4 1 0 1 0

7 5 2 0 1 0 2 1 0

6 2 1 4 2 5 0 0 0

2 4 3 2 2 5 1 0 1

21 13 9 7 9 11 3 2 1

14 8 3 4 5 9 2 0 0

34 (0.45)

22 (0.65)

42 (0.55)

23 (0.55)

Total

18

18

20

20

76

45

76

45 (0.59)

Parity

4

Supplementary table 2. Median test results of sow samples per farm (with inter quartile range) ApxIV qPCR genomic copies ApxI ELISA Log2 titre ApxII ELISA Log2 titre ApxIII ELISA Log2 titre Omp ELISA Log2 titre

Farm A 106.4 (105.9 ; 106.7) >14 (13.4 ; >14) 13.4 (13.1 ; >14) 12.6 (11.5 ; 13.3) 11.6 (10.7 ; 12.6)

Farm B 105.0 (104.3 ; 105.9) 12.4 (11.4 ; 13.4) >14 (13.6 ; >14) 12.1 (11.3 ; 12.8) 12.2 (11.3 ; 13.4)

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Supplementary fig. 1. Colonisation status at litter and individual piglet level in the four studied cohorts In green pens all piglets tested negative, whereas in orange pens at least one pig tested positive. M = cross fostering applied

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A. pleuropneumoniae colonisation in suckling piglets

Farm

Cohort

Litterpos

Pos

Neg

N

Age

Mix

APXIb

APXIIb

APXIIIb

OMPb

APPDNA

Par2

Supplementary file 1. study data, as used for the statistical modelling

1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 2 2 2

1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 3 3 3

1 1 1 1 0 1 1 1 1 0 1 0 0 0 1 1 1 0 0 1 1 0 1 1 1 1 0 1 0 0 0 1 1 0 1 1 1 1 1

2 2 6 9 0 2 1 4 3 0 10 0 0 0 5 9 1 0 0 1 1 0 10 11 12 2 0 10 0 0 0 4 4 0 7 11 11 3 3

11 8 4 2 11 8 11 7 8 12 0 10 12 9 7 1 7 11 15 9 12 11 2 2 0 11 9 1 12 9 9 5 9 9 0 1 2 6 9

13 10 10 11 11 10 12 11 11 12 10 10 12 9 12 10 8 11 15 10 13 11 12 13 12 13 9 11 12 9 9 9 13 9 7 12 13 9 12

24 24 26 23 25 23 24 21 25 24 19 24 24 24 23 17 24 24 19 15 19 21 19 24 19 23 19 25 24 17 20 24 23 23 24 24 25 27 25

0 0 0 0 0 0 0 0 0 1 0 0 0 1 0 1 0 0 0 0 0 0 1 0 0 0 0 0 0 0 0 0 0 0 0 0 1 0 0

1 1 1 0 1 1 1 1 1 0 1 1 1 1 1 1 1 1 1 0 1 1 0 1 0 0 1 1 1 0 1 1 0 1 1 1 0 0 1

1 0 0 0 0 0 0 0 0 0 0 1 1 1 1 0 1 0 0 0 1 1 0 0 0 0 1 0 1 0 0 0 0 0 1 1 0 1 1

1 1 0 1 0 1 0 1 0 0 1 1 1 0 0 0 0 1 0 1 0 1 0 1 1 1 0 1 1 1 1 0 0 1 0 0 1 1 0

1 1 0 1 0 1 0 1 0 0 1 1 1 0 0 0 0 0 0 1 0 1 0 1 0 0 0 0 1 1 1 0 1 1 0 0 1 1 0

6.87862 6.66704 6.49645 6.1699 6.40638 8.09005 4.67975 4.73374 7.38212 6.81512 6.06151 4.77905 6.92349 5.77059 7.06644 5.60265 6.50113 5.52015 6.47188 5.98986 5.75912 4.70319 5.84392 5.99884 5.98118 6.13916 6.63382 5.96236 7.13902 6.4248 6.4608 7.06959 5.911 6.50318 6.82164 6.43938 3.45606 4.54805 5.94175

0 0 1 0 1 1 0 0 1 1 0 1 0 1 1 0 1 1 0 0 1 1 0 0 0 0 1 0 0 0 0 1 0 0 1 1 0 0 1

89

4

Chapter 4

Farm

Cohort

Litterpos

Pos

Neg

N

Age

Mix

APXIb

APXIIb

APXIIIb

OMPb

APPDNA

Par2

Supplementary file 1 study data, as used for the statistical modelling; continued

2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2 2

3 3 3 3 3 3 3 3 3 3 3 3 3 3 3 3 3 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4 4

0 1 0 0 1 0 1 1 0 1 1 0 1 0 1 1 1 0 0 0 1 1 1 0 1 0 1 1 0 0 0 0 0 1 1 0 1

0 13 0 0 4 0 7 1 0 8 4 0 7 0 1 6 2 0 0 0 4 1 1 0 6 0 12 1 0 0 0 0 0 7 1 0 8

12 0 12 12 7 13 5 9 12 3 7 14 7 12 10 6 10 10 11 12 9 12 10 12 7 12 0 11 11 13 14 11 13 4 11 12 5

12 13 12 12 11 13 12 10 12 11 11 14 14 12 11 12 12 10 11 12 13 13 11 12 13 12 12 12 11 13 14 11 13 11 12 12 13

24 24 28 25 25 30 26 25 24 24 26 26 25 25 26 29 24 23 26 27 25 26 22 25 28 24 27 15 25 24 24 22 22 25 26 24 25

0 0 0 0 0 0 0 0 0 0 0 0 0 0 1 0 1 1 0 0 0 0 1 0 0 0 0 0 0 1 0 0 0 0 0 0 0

1 0 0 0 0 0 0 0 0 0 0 0 0 0 0 1 1 0 0 1 0 1 0 1 0 0 1 1 0 0 1 0 1 1 0 1 0

1 1 0 1 0 0 0 0 1 1 1 1 0 1 1 1 1 0 1 0 1 1 1 1 0 0 1 1 1 1 1 0 1 NA 1 NA 0

0 1 0 0 0 1 0 0 1 0 0 0 1 1 0 0 1 0 1 1 1 1 0 0 0 0 1 1 1 0 1 0 1 1 1 0 0

0 1 0 0 0 1 1 0 1 1 0 0 1 1 0 1 1 1 1 1 0 1 0 0 0 0 1 1 1 1 1 0 1 1 1 0 0

5.13894 5.96905 5.81848 5.92378 6.05026 5.9553 4.86216 4.2737 4.28448 4.71258 3.16495 3.57955 6.12223 4.39891 5.60979 5.50175 4.3249 6.0087 6.99397 4.42475 4.64813 5.9529 4.72891 5.4316 6.08483 3.87541 4.03334 5.88837 5.48326 4.29332 5.70365 4.51175 5.37016 4.78098 5.95843 4.30724 3.74772

1 0 1 1 1 0 1 1 1 0 1 1 0 0 1 1 0 0 1 0 1 1 1 1 0 1 1 0 0 1 1 1 0 1 1 1 1

90

A. pleuropneumoniae colonisation in suckling piglets

Supplementary file 2. R – script for statistical modelling # Load data data 2 (1). ############# # Cohort indicators ba2