Open Access

Chlamydiaceae infections in pig

Veterinary Research201142:29

DOI: 10.1186/1297-9716-42-29

Received: 30 June 2010

Accepted: 17 January 2011

Published: 7 February 2011

Abstract

Chlamydiaceae are Gram-negative obligate intracellular bacteria. They are responsible for a broad range of diseases in animals and humans. In pigs, Chlamydia suis, Chlamydia abortus, Chlamydia pecorum and Chlamydia psittaci have been isolated. Chlamydiaceae infections in pigs are associated with different pathologies such as conjunctivitis, pneumonia, pericarditis, polyarthritis, polyserositis, pseudo-membranous or necrotizing enteritis, periparturient dysgalactiae syndrome, vaginal discharge, return to oestrus, abortion, mummification, delivery of weak piglets, increased perinatal and neonatal mortality and inferior semen quality, orchitis, epididymitis and urethritis in boars. However, Chlamydiaceae are still considered as non-important pathogens because reports of porcine chlamydiosis are rare. Furthermore, Chlamydiaceae infections are often unnoticed because tests for Chlamydiaceae are not routinely performed in all veterinary diagnostic laboratories and Chlamydiaceae are often found in association with other pathogens, which are sometimes more easily to detect. However, recent studies have demonstrated that Chlamydiaceae infections in breeding sows, boars and piglets occur more often than thought and are economically important. This paper presents an overview on: the taxonomy of Chlamydiaceae occurring in pigs, diagnostic considerations, epidemiology and pathology of infections with Chlamydiaceae in pigs, public health significance and finally on prevention and treatment of Chlamydiaceae infections in pigs.

Table of contents

  1. 1.

    Introduction

     
  2. 2.
    Taxonomy of chlamydial species occurring in pig
    1. 2.1

      Chlamydia abortus (ruminant C. psittaci serovar1)

       
    2. 2.2

      Chlamydia pecorum

       
    3. 2.3

      Chlamydia psittaci

       
    4. 2.4

      Chlamydia suis

       
     
  3. 3.

    Diagnostic considerations

     
  4. 4.
    Epidemiology of infections with Chlamydiaceae in pigs
    1. 4.1

      Serology

       
    2. 4.2

      Molecular diagnosis

       
     
  5. 5.
    Pathogenicity
    1. 5.1.

      Experimental infections

       
    2. 5.2.

      Studies on natural infections

       
     
  6. 6.

    Public health significance

     
  7. 7.

    Prevention and treatment

     
  8. 8.

    Authors’ contribution

     
  9. 9.

    Competing interests

     
  10. 10.

    Acknowledgements

     
  11. 11.

    References

     

1. Introduction

Chlamydiaceae are Gram-negative obligate intracellular bacteria. They are responsible for a broad range of diseases in animals and humans. Chlamydiaceae primary replicate in mucosal epithelial cells of the conjunctivae, the respiratory, urogenital and gastrointestinal tract. They can survive and replicate in monocytes and macrophages and are characterized by distinct extracellular and intracellular forms. The infective elementary body (EB) is metabolically inactive. It reorganizes to a reticulate body (RB) after entering the host cell. These RBs are metabolically active and divide by binary fission within phagocytotic vesicles. Condensation to EBs and the subsequent lysis of the host cell completes the chlamydial replications cycle. Several studies, however, described the occurrence of alternative developmental stages consisting of abnormal sized, mostly enlarged RB-like structures called aberrant bodies (ABs) and their association with persistence of Chlamydiaceae[1].

Chlamydiaceae infections in pigs have been known to occur since 1955 when Willingan and Beamer [2] first isolated chlamydia from cases of arthritis and pericarditis in U.S. pigs. Massive outbreaks of chlamydiosis associated with bronchopneumonia or abortion in pigs kept under intensive animal production systems were reported in Eastern European countries and Russia between 1960 and 1970 [35]. In 1969, the first pig chlamydial strains were isolated in Western Europe from numerous Austrian pigs with polyarthritis, polyserositis, pneumonia, conjunctivitis or enteritis, from sows that aborted and from pigs with inapparent intestinal tract infection [6, 7]. In the 1980s, chlamydial strains were isolated from healthy and sick German pigs [8, 9]. In the 1990s, chlamydia was consistently isolated from pig herds in Nebraska and Iowa or detected in conjunctival specimens from pigs affected with conjunctivitis or keratoconjunctivitis in all phases of production [10]. Many of the nursing and nursery pigs with conjunctivitis from these and other herds had diarrhoea, and at necropsy most of the diarrheic pigs also showed pneumonia. Although known pathogens were believed to be the cause of the diarrhoea and the pneumonia, chlamydia was isolated from or detected in the intestines and lungs of affected pigs. During the 1990s, chlamydia was also often isolated and/or detected in German and Swiss pig herds and infections were associated with return to oestrus, abortion, enteric disease and asymptomatic intestinal infections [1115].

Thus, as reported in the literature, chlamydial disease in pigs includes conjunctivitis, pneumonia and pseudomembranous or necrotizing enteritis, as demonstrated by experimental reproduction of infection in gnotobiotic pigs using clinical isolates [1618]. In addition, C. suis is associated with pericarditis, polyarthritis and polyserositis in piglets [2] and numerous reproductive problems such as vaginal discharge, return to oestrus, abortion, mummification, delivery of weak piglets, increased perinatal and neonatal mortality [19] as well as orchitis, epididymitis and urethritis in boars [20]. For most of these disorders, the exact role of Chlamydiaceae still has to be determined. Eggeman et al. [21] noticed a correlation between the occurrence of the periparturient dysgalactiae syndrome (PDS) in sows and the number of animals being seropositive for Chlamydiaceae. However, evidence that C. suis is causing PDS is lacking.

Chlamydiaceae were and still are considered as non-important pathogens of pigs because tests for Chlamydiaceae are not normally performed at most veterinary diagnostic laboratories and Chlamydiaceae are often found in association with other pathogens.

Chlamydial infections in breeding sows, boars and piglets occur more often than originally thought. Chlamydia abortus, Chlamydia pecorum, Chlamydia psittaci and Chlamydia suis can infect pigs.

2. Taxonomy of chlamydial species occurring in pig

In 1999, Everett et al., [22] proposed a reassignment from the single genus Chlamydia into two genera, Chlamydia and Chlamydophila, based on clustering analyses of the 16S rRNA and 23S rRNA genes (Table 1). However, comparative genome analysis of strains mentioned in Table 2 is consistent with the conclusion that host-divergent strains of chlamydia are biologically and ecologically closely related. The previous taxonomic separation of the genus based on ribosomal sequences is neither consistent with the natural history of the organism revealed by genome comparisons, nor widely used by the chlamydia research community eight years after its introduction. Consequently, it was proposed to reunite the Chlamydiaceae into a single genus, Chlamydia[23] (Table 1). Accordingly, the chlamydia nomenclature is used here.
Table 1

Taxonomy of chlamydial organisms.

 

Chlamydial taxonomy before 1999

Chlamydial taxonomy since 1999 (Everett et al., 1999[22])

Chlamydial taxonomy used in the Twenty-first century (Stephens et al., 2009[23])

Order

Chlamydiales

Chlamydiales

Chlamydiales

Family

Chlamydiaceae

Chlamydiaceae, Simkaniaceae, Parachlamydiaceae, Waddliaceae

Chlamydiaceae, Simkaniaceae, Parachlamydiaceae, Waddliaceae

Genus

Chlamydia

 

Chlamydia

 

Chlamydophila

 

Chlamydia

 

Species

C. trachomatis

Trachoma biovar

C. trachomatis

Trachoma biovar

  

C. trachomatis

Trachoma biovar

  

LGV biovar

 

LGV biovar

   

LGV biovar

  

Murine biovar

C. muridarum

   

C. muridarum

 
  

Porcine biovar

C. suis

   

C. suis

 
 

C. pneumonia

Human biovar

  

Cp. pneumonia

TWAR biovar

C. pneumonia

TWAR biovar

  

Koala biovar

   

Koala biovar

 

Koala biovar

  

Equine biovar

   

Equine biovar

 

Equine biovar

 

C. psittaci

Avian subtype

  

Cp. psittaci

 

C. psittaci

 
  

Abortion subtype

  

Cp. abortus

 

C. abortus

 
  

Feline subtype

  

Cp. felis

 

C. felis

 
  

Guinea-pig subtype

  

Cp. caviae

 

C. caviae

 
 

C. pecorum

   

Cp. pecorum

 

C. pecorum

 
Table 2

Completed Chlamydia genomes.

Species

Strains

Completed in

Group

Reference

C. trachomatis

D/UW-3/Cx

1998

Berkeley/Stanford

[79]

C. trachomatis

A/Har-13

2005

RML NIAID

[80]

C. trachomatis

LGV L2/434

2008

Sanger Institute

[81]

C. trachomatis

LGV L2/UCH-1

2008

Sanger Institute

[81]

C. pneumoniae

CWL029

1999

Berkeley/Stanford

[82]

C. pneumoniae

J138

2000

Yamaguchi University

[83]

C. pneumoniae

AR39

2000

TIGR

[84]

C. pneumoniae

TW-183

2007

ALTANA

[85]

C. pneumoniae

Koala

2008

University of Maryland

[86]

C. muridarum

Nigg

2000

TIGR

[84]

C. caviae

GPIC

2003

TIGR

[87]

C. abortus

S26/3

2005

Sanger Institute

[88]

C. felis

Fe/C-56

2006

Yamaguchi University

[89]

C. psittaci

6BC

2008

University of Maryland

[86]

C. pecorum

 

2008

University of Maryland

[86]

Protochlamydia

UWE25

2004

University of Vienna

[90]

Simkania

 

2008

University of Maryland

[49]

Adapted from Stephens et al. [23].

2.1. Chlamydia abortus (ruminant C. psittaci serovar 1)

Chlamydia abortus was formerly classified as ruminant C. psittaci serotype 1. This species has a distinct serotype and the ribosomal and outer membrane protein A (ompA) sequences are nearly 100% conserved. An extra chromosomal plasmid has not been identified in any of the C. abortus strains. The ruminant strain B577 (ATCC VR 656) is regarded as type reference strain.

Chlamydia abortus is one of the main infectious causes of abortion in sheep, cattle and goats in many countries around the world [24]. In addition, the pathogen has also been associated with abortion in horses, rabbits, guinea pigs, mice and pigs. Furthermore, pregnant women working with animals infected with C. abortus are at risk, as C. abortus is also a zoonotic agent able to cause abortion in humans [25]. Chlamydia abortus strains in pigs are primarily associated with abortion and weak neonates. So far, transmission of C. abortus from pigs to humans has not been reported.

2.2. Chlamydia pecorum

Chlamydia pecorum strains are serologically and pathogenically diverse and have been isolated only from mammals. Two strains, E58 and Koala II, have an extra chromosomal plasmid, pCp. The type strain for C. pecorum is E58 (ATCC VR 628).

The presence of C. pecorum has been established in ruminants [26], pigs and koalas (marsupials) [22]. Chlamydia pecorum has been associated with a wide range of diseases. In koalas, C. pecorum causes conjunctivitis, reproductive disease, infertility and urinary tract disease. Additionally, C. pecorum has been associated with urogenital tract infections, inapparent intestinal infections, abortion, conjunctivitis, mastitis, encephalomyelitis, enteritis, pneumonia, polyarthritis, pleuritis, pericarditis in sheep, goats, cattle, horses and pigs [2628].

2.3. Chlamydia psittaci

Chlamydia psittaci primarily infects birds, but has caused sporadic zoonotic infections in humans. The bacterium is spread between birds mainly by inhalation of contaminated aerosols of ocular or nasal secretions and contaminated dust from feathers and faecal material. The transmission can also be vertical through the egg. Chlamydia psittaci has nine known genotypes (A-F, E/B, M56 and WC) which are all considered to be transmissible to humans. Several strains have an extra chromosomal plasmid. The type strain for C. psittaci is 6BC (ATCC VR 125).

In pigs, C. psittaci ompA genotype A has been isolated from the genital tract of a Swiss breeding sow [29]. Additionally, C. psittaci has been isolated from the lungs of a Belgian sow [30]. In the latter study, the genotype could not be defined as the nucleotide sequence of the cloned C. psittaci ompA gene (Genbank Accession No. AY327465) was found to be 99.3, 99.1 and 98.9% identical to that of C. psittaci CP3, 6BC and MN Zhang, respectively. A significant relationship was found between C. psittaci infections in pigs and keeping poultry on the farm [21, 30].

2.4. Chlamydia suis

Before 1999, Chlamydia suis strains were referred to as C. trachomatis because of ompA DNA sequence homology [28]. Currently, the only known natural host of C. suis is the pig. Several strains have an extra chromosomal plasmid, pCs [25]. The type strain of this species, S45 (ATCC VR 1474) was isolated in Europe in the late 1960s from faeces of an asymptomatic pig in Austria [6]. This strain is tetracycline sensitive (TcS), as other chlamydial species. Chlamydia suis strains expressing a stable tetracycline resistant phenotype associated with the presence of a resistance gene, tet(C), in the chromosome, have been isolated in farms in Iowa and Nebraska [31], in Italy [32] and in Belgium [33]. Chlamydia suis in pigs has been associated with conjunctivitis, rhinitis, pneumonia, enteritis, PDS, reproductive disorders such as return to oestrus (early embryonic dead in more than 50% of the sows) and inferior semen quality (decrease of sperm cell motility and dead of more than 50% of the sperm cells) and apparently asymptomatic infections [10, 1518, 21, 3436]. A high degree of genetic diversity was observed in C. suis when compared to other chlamydial species [22, 37]. Incongruent reports on the pathology of Chlamydiaceae in pigs and variations in virulence support the theory of genetic diversity [15, 21, 34, 38, 39].

3. Diagnostic considerations

Diagnostic laboratories do not routinely test for Chlamydiaceae in pigs. Cell cultures are the most convenient method for the isolation of Chlamydiaceae. However, with pig strains, this method has been only partially successful because it is difficult to grow the bacteria on the established transformed cell lines (HeLa and McCoy) for chlamydial culture and techniques had to be modified. Guseva et al. [40] used pig genital epithelial cells which were cultured ex vivo for dissecting the hormonal modulation of several aspects of C. trachomatis pathogenesis and infection. Chlamydia suis (S45) was used in this in vitro model. Consequently, these pig primary cells could be suitable in theory for chlamydial diagnosis in pigs. Moreover, they can be frozen at -80°C in DMEM-F-12 medium with 10% dimethyl sulfoxide (DMSO) for several weeks and they can reform monolayers in 3 to 5 days after thawing. However, continuous cell lines are still more convenient and less expensive. Schiller et al. [41] cultivated porcine Chlamydiaceae under various conditions. The combination of centrifugation assisted cell culture infection and cycloheximide treatment of cell coverslip cultures provided the highest inclusion numbers with all chlamydial strains. Interestingly, the use of Iscove's modified Dulbecco's medium instead of Eagle's minimal essential medium significantly increased C. suis inclusion counts in Vero cells. However, C. suis and C. pecorum inclusion numbers were markedly increased in CaCo cells when compared to Vero cells.

The enzyme-linked immunosorbent assay (ELISA) has been very popular for chlamydial antigen detection because it is easy to use. Kits designed to detect C. trachomatis in humans have been used extensively in veterinary medicine because most of them, (especially the earlier developed kits) detect the chlamydial family-specific LPS antigen and therefore will detect all chlamydial strains. However, the most important drawback of these tests are the cost and the lack of sensitivity and specificity [21, 42].

Immunohistochemical staining of histological sections is often used as more veterinary diagnostic laboratories are using equipment to automate the staining. A Chlamydiaceae family-specific mouse monoclonal antibody directed against the chlamydial lipopolysaccharide (LPS; Clone ACI-P, Progen, Heidelberg, Germany) and a chlamydial genus-specific mouse monoclonal antibody (IgG1) directed against recombinant C. trachomatis HSP60 (clone A57-B9, Milan Analytica AG, La Roche, Switzerland) have been used with detection by the streptavidin-biotin method (Dako ChemMate™ detection kit; Dako Diagnostics, Heverlee, Belgium) [43]. Unfortunately, aberrant bodies are stained less, especially when using anti-HSP60 antibodies.

PCR techniques have been developed for use in veterinary medicine. They are replacing isolation for the detection of Chlamydiaceae in animals. If properly designed, the specificity is excellent and the sensitivity equals or exceeds well-controlled isolation procedures. For bio-safety reasons, the sample can be inactivated prior to testing. Current PCR tests for detection of Chlamydiaceae species occurring in pigs, target the ompA, the 16S-23S rRNA or the incA gene [22, 38, 44, 45]. Targeting the 16S-23S genes increases sensitivity, as multiple copies of those genes are usually present in the organism. However, cross-reactions with other bacteria can interfere. The sensitivity and specificity of PCR also varies on sample preparation. Reagents designed to stabilise the DNA should be considered when a delay in processing the sample is anticipated. DNA samples can be prepared using inexpensive reagents or using commercial available kits. Sensitivity increases by targeting a relatively short DNA segment, using a nested procedure or using real-time PCR techniques. The nested PCR procedure is very sensitive. However, extreme care should be taken when manipulating the reaction in order to decrease the risk of contamination. The real-time PCR requires a labelled probe and special equipment, which increases costs. The sensitivity of this test is approximately the same as the nested PCR and contamination problems and labour are reduced as it consists of only one reaction in a closed system, often provided with uracil-N-glycosylase (UNG) to prevent post PCR carry over.

Some years ago, Sachse et al. [46] designed a microarray hybridization assay for the identification of chlamydial species using the ArrayTube™ platform (Clondiag Chip Technologies, Jena, Germany). The test proved suitable for unambiguous species identification of chlamydial cell cultures and showed potential for direct detection of these bacteria from clinical tissues. Unfortunately, routine testing for Chlamydiaceae in pigs is not possible because the price (24 € per sample) is not feasible.

Sequencing of PCR products can allow comparison between the sequences of reference chlamydial isolates and this information can be used in phylogenetic analysis for classification and epidemiological purposes. However, sequence analysis of the outer membrane protein A (ompA) gene is to our opinion not advisable for pig isolates, because of high sequence homology between the ompA gene of C. psittaci and C. abortus[47]. Multi locus sequence typing (MLST) has also been used for typing chlamydial species which occur in pigs. Multi locus sequence typing has been described for C. abortus, C. pecorum and C. psittaci[48, 49].

4. Epidemiology of infections with Chlamydiaceae in pigs

4.1. Serology

The earliest serological data on the occurrence of chlamydial organisms in European pigs originate from 1966. At that time, Wilson and Plummer [50] discovered antibodies against Chlamydiaceae by capillary agglutination testing in 23% of sera derived from pigs in Great Britain.

At present, chlamydial infections are endemic in the Belgian pig population [30] as 240 (96.5%) of 249 examined fattening pig farms were seropositive by use of a recombinant ELISA detecting Chlamydiaceae family-specific antibodies. In this ELISA the plate was coated with the recombinant Major outer membrane protein (MOMP) of an avian C. psittaci genotype D strain [51]. Recombinant MOMP was prepared in COS-7 cells [52] instead of E. coli[53], in order to mimic eukaryotic glycosylation of MOMP and avoid E. coli fragments (LPS) which will be unavoidably present in purified recombinant proteins expressed in prokaryotes. In Germany, Chlamydiaceae seroprevalence rates in sows and boars are 33 to 72% and 10 to 47%, respectively [21]. For Switzerland, seroprevalence rates of 62% in sows, 6.9% in piglets younger than 4 weeks and 48.1% in piglets older than 4 weeks are reported [54]. These studies used an LPS-based ELISA described by Wittenbrink et al. [14]. In Italy, 63.5 to 80.3% of the finishing pigs were serologically positive when using a purified elementary body-based micro-immunofluorescence test (MIF) [55]. Bagdonas et al. [42] found a seroprevalence rate of 7.7% in Lithuanian pigs when using the complement fixation test (CFT). However, serological results obtained by using chlamydial LPS or whole elementary bodies should be handled with some reservations, as serological cross-reactions with antibodies against other pathogens do occur [5659]. Moreover, the CFT is less sensitive and less specific than ELISA [60].

Qiu [61] and Zhou and Qiu [62], reported a seroprevalence rate in china of 11% and 80% in piglets and sows, respectively. However, most recent data originate from the Guangdong Province in Southern China revealing a seroprevalence rate of 63.38%, 41.10% and 36.25% in breeding boars, breeding sows and fattening piglets, respectively, when using a commercially marketed indirect haemaglutination assay (IHA kit; Lanzhou Veterinary Research Institute, Lanzhou, China) [63]. To our knowledge, there are no recent serological data on Chlamydiaceae in pigs from other countries.

4.2. Molecular diagnosis

Serology is useful for monitoring the Chlamydiaceae status in pigs, at least when using a Chlamydia-specific target antigen. However, all current serological tests fail to identify the causative chlamydial species. Thus, more detailed information on the prevalence of Chlamydiaceae in pigs originates from newly developed species-specific molecular diagnostic research.

Recently, species-specific nucleic acid amplification tests (NAATs) such as real-time PCR and microarray, detecting the ribosomal intergenic spacer and domain I of the 23S rRNA gene, have been designed [46, 64]. Moreover, Dugan et al. [65] designed a PCR for detecting the tetracycline resistance (TcR) gene, tet(C). These methods have been recently applied to examine the occurrence of different chlamydial species in pigs.

Chlamydia psittaci DNA was only sporadically found in pigs [29, 30, 33]. Chlamydia pecorum was also less frequently found and has been identified, using ompA gene sequencing, in 2% of ompA gene-positive boar sperm samples, in 5% of ompA gene-positive foetuses and in 9% of ompA gene-positive gut tissues [45]. Chlamydia abortus was identified in the lungs of a Belgian pig that accidently died after blood sampling [33]. Although, C. abortus is mainly linked to reproductive failure and abortions in pigs, it has previously been identified in lungs of pigs [46].

Involvement of C. suis was reported in the vast majority of chlamydial intestinal infections in Belgian, German and Swiss pigs. Chlamydia abortus was rarely found in these studies [11, 12, 15, 29]. Chlamydia suis was predominantly associated with conjunctivitis in intensively kept German, Swiss and Estonian pigs [38]. Furthermore, a German study reported a high prevalence of mixed infections with C. suis and C. abortus in the lung and gut of pigs [39].

Chlamydial DNA has been discovered by nested PCR in 57.1% of the animals of a German wild boar population in Thuringia [44]. Organisms were predominantly detected in the lung. Sequencing of the amplified ompA segments revealed C. psittaci, C. abortus and C. suis in this wild boar population. These findings revealed a possible wildlife reservoir of these bacteria.

5. Pathogenicity

5.1. Experimental infections

The first experimental infection was performed by Pavlov et al. [66] infecting young piglets with chlamydial agents isolated from Bulgarian pigs. The infection resulted in keratoconjunctivitis, fever, anorexia and depression.

The literature describes four conjunctival C. suis strains: H5, H7, R22 (USA) and DC6 (Germany); 14 C. suis strains obtained from Italian pigs with conjunctival and/or reproductive disorders (MS1 to MS14); five intestinal strains: R19, R27, 130, 132 (USA) and S45 (Austria); and two respiratory strains R24 and R33 (USA) (Table 3). All C. suis strains, with the exception of S45, originated from pigs with clinical disease. Koch's postulates have been fulfilled in gnotobiotic pigs using H7 (conjunctivitis), R19 and R27 (intestinal lesions) and R33 (lung and nasal lesions) [1618]. Strain S45 was isolated from the faeces of an asymptomatic pig, but experimental enteric infection of gnotobiotic piglets with S45 proved its pathogenic potential as it elicited significant enteric disease [67]. Interestingly, Pospischil et al. [43] reported the occurrence of aberrant C. suis developmental forms, indicative for a persistent infection in experimentally infected gnotobiotic pigs.
Table 3

Chlamydia sui s strains isolated from pigs

  

Isolated from

  

Strain

Location

Year

Tissue

Clinical symptoms of the pig(s)

S45

Austria

1969

Intestines (faeces)

Asymptomatic infection

R19

Nebraska

1992

Intestines (faeces)

Pneumonia, enteritis, conjunctivitis

R22

Nebraska

1992

Conjunctiva

Conjunctivitis

R24

Nebraska

1992

Respiratory tract (nasal mucosa)

Upper respiratory tract disease

R27

Nebraska

1993

Intestines (colon)

Enteritis

R33

Nebraska

1994

Respiratory tract (nasal mucosa)

Pneumonia

H5

Iowa

1994

Conjunctiva

Conjunctivitis

H7

Iowa

1994

Conjunctiva

Conjunctivitis

130

Nebraska

1996

Intestines (jejenum)

Asymptomatic infection

132

Nebraska

1996

Intestines (ileum)

Asymptomatic infection

DC6

Germany

2004

Conjunctiva

Conjunctivitis

MS1

Italy

2004-2007

Conjunctiva

Conjunctivitis

MS2

Italy

2004-2007

Conjunctiva

Conjunctivitis

MS3

Italy

2004-2007

Conjunctiva

Conjunctivitis

MS4

Italy

2004-2007

Conjunctiva

Conjunctivitis and return to oestrus

MS5

Italy

2004-2007

Conjunctiva

Conjunctivitis and return to oestrus

MS6

Italy

2004-2007

Conjunctiva

Conjunctivitis and return to oestrus

MS7

Italy

2004-2007

Conjunctiva

Conjunctivitis and return to oestrus

MS8

Italy

2004-2007

Conjunctiva

Conjunctivitis and return to oestrus

MS9

Italy

2004-2007

Conjunctiva

Conjunctivitis and return to oestrus

MS10

Italy

2004-2007

Conjunctiva

Conjunctivitis and return to oestrus

MS11

Italy

2004-2007

Conjunctiva

Conjunctivitis and return to oestrus

MS12

Italy

2004-2007

Conjunctiva

Conjunctivitis

MS13

Italy

2004-2007

Conjunctiva

Conjunctivitis

MS14

Italy

2004-2007

Conjunctiva

Conjunctivitis

Experimental aerosol challenge with C. suis of conventional raised colostrum-fed pigs confirmed the pathogenic potential of C. suis for the porcine respiratory system [46, 68]. In the latter studies, six week-old pigs were infected by aerosol using 109 chlamydial inclusion-forming units. All infected animals developed an acute infection characterized by a dry cough, serous nasal discharge and severe dyspnoea accompanied by wheezing, shortness of breath and breathlessness. The body temperature of the pigs rose above 40°C for at least five days post infection. Clinical signs lasted for seven days post infection.

The role of C. suis and C. abortus in reproductive problems is still controversial. One of the reasons for this is that there are often ethical objections when willing to proof Koch's postulates, as it requires C. suis or C. abortus experimental infections of Chlamydiaceae-negative sows, at different stages of gestation. Moreover, it is extremely difficult to find Chlamydiaceae-negative sows. To our knowledge, there is only one study that deals with an experimental C. abortus infection in sows. Experimental infections of four sows with the BS ruminant C. psittaci serovar 1 strain at 42 days of gestation resulted in infection of fetal membranes, but failed to induce abortion [69].

Guscetti et al. [70] studied the pathogenicity of a C. psittaci isolate of pigeon origin (T49/90) in three day-old gnotobiotic piglets. The animals were infected intragastrically resulting in a productive enteric infection with mild lesions, weak systemic dissemination, and faecal shedding, indicating the pig as a potential host for avian Chlamydiaceae. Oral administration of C. suis was more virulent for three day-old gnotobiotic piglets than oral inoculation of C. psittaci (pigeon strain T49/90) or C. abortus (S26/E) [70, 71].

5.2. Studies on natural infections

The prevalence, the zoonotic risk and the economic impact of different chlamydial species occurring in pigs was difficult to determine because species-specific diagnostic tests were unavailable. Recently, chlamydial species-specific NAATs have been developed. These NAATs are currently being used to study the prevalence and the economic impact of different chlamydial species occurring in pigs. Chlamydiaceae, and especially, C. suis are widespread in pigs. Considering the high degree of genetic diversity observed in C. suis when compared to other chlamydial species [22, 37], incongruent reports on the pathology of Chlamydiaceae in pigs and variations in virulence support the theory of genetic diversity [15, 21, 34, 38, 39]. Intestinal C. suis infections seem to be common in commercial pigs and most, if not all, are believed to be subclinical [12, 15, 17, 34, 72]. However, Evans postulates have been fulfilled for this pathogen [1618], thus it is probably more harmful as some might believe. Chlamydia suis was involved in return to oestrus (early embryonic dead in more than 50% of the pregnant sows) in sows and inferior semen quality (decrease of sperm cell motility and dead of more than 50% of the sperm cells) in boars, as demonstrated on fattening pig farms in Belgium, Cyprus, Estonia, Germany, Israel and Switzerland [14, 21, 33, 36, 39, 54]. Becker et al. [38] found a high prevalence of C. suis in eyes of German (90%) and Swiss (79%) ocular symptomatic pigs. In general, intensive kept pigs seemed to be pre-disposed to ocular chlamydial infection associated with conjunctivitis. Schautteet et al. [36] observed the same, examining conjunctivitis and reproductive failure on a large Estonian pig production plant.

Chlamydia suis, C. pecorum and C. abortus have been detected in aborted foetuses [11, 13]. Eggemann et al., [21] found a significant correlation between chlamydial PCR positivity and the incidence of abortion and litters with stillborn piglets and piglets with low viability. Seropositive farms had statistically less weaned piglets per sow and litter. Hoelzle et al., [39] used two PCRs targeting the ompA (encoding the MOMP; 40 kDa) or ompB (encoding the cys teine rich outer membrane protein; 60 kDa) gene, respectively. The PCR was not species-specific but was able to detect C. suis, C. pecorum as well as C. abortus in lungs, intestines and endocervical swabs of pigs. PCR amplicons were generated from 49 and 60% of pigs with respiratory or reproductive disorders, respectively. Chlamydial DNA was present in 24.5% of respiratory healthy controls and in none of the endocervical swabs from fertile control sows. They found a high prevalence of mixed C. abortus and C. suis infections in lungs and intestines using RFLP and DNA sequence analysis of ompA.

6. Public health significance

The zoonotic potential of C. abortus and C. psittaci is well documented [73]. However, to our knowledge, transmission of these pathogens from pigs to humans has not been reported. Chlamydia suis, previously referred to as C. trachomatis, might be a potential zoonotic pathogen. So far, no reports on zoonotic transmission were published.

7. Prevention and treatment

Chlamydiaceae are highly susceptible to chemicals that affect their lipid content or the integrity of their cell walls. Cleaning of equipment and stables of infected pigs is important because Chlamydiaceae can survive for up to 30 days in faeces and bed materials. Disinfection with most common detergents and disinfectants will inactivate Chlamydiaceae. The following disinfectants can be used to inactivate the organism: 1:1000 dilution of quaternary ammonium compounds, 70% isopropyl alcohol, 1% Lysol, 1:100 dilution of household bleach or chlorophenols [74, 75]. Common infection sources, infection routes, possible vectors and infection kinetics on the farm have not been examined.

Current infections are being treated by means of antibiotics. Generally, tetracyclines (chlortetracycline, oxytetracycline, doxycycline) are the drugs of choice to control the disease because they are most effective. Quinolones (enrofloxacin) or macrolides (erythromycin) can be administered, in case of an infection with a tetracycline resistant C. suis strain. Enrofloxacin might present a solution in case of tetracycline resistant C. suis strains [33].

Pollman et al. [76], demonstrated the beneficial effect of a probiotic strain of Enterococcus faecium (NCIMB 10415) on reducing carryover infections from naturally Chlamydiaceae infected sows to newborn piglets. The probiotic strain is licensed by the European Union as a feed supplement for animals.

So far, no vaccines are available. Chlamydial vaccines produced for Chlamydiaceae in other animals would likely have no efficacy in pigs, as the strains are both genetically and serologically very different. Therefore, Knitz et al. [77] used a herd-specific, formalin-inactivated C. abortus strain (OCHL03/99) originating from vaginal discharge of sows to study the humoral immune response in immunized breeding sows, as a preliminary study towards vaccine development. Compared to the control group, vaccinated animals showed a marked primary and secondary IgG serum antibody response. Protection was not examined. Recently, Zhang et al. [78] presented data on vaccination of mice using a porcine C. abortus strain. They obtained a protective immune response following co-vaccination using a DNA vaccine together with recombinant MOMP.

Nevertheless, it is obvious that we need more progress in understanding protective and (possible) pathological immune mechanisms in pigs, before a potential vaccine candidate for Chlamydiaceae can be generated.

Declarations

10. Acknowledgements

Ghent University (01G00805) is acknowledged for providing a research grant to K. Schautteet.

Authors’ Affiliations

(1)
Department of Molecular Biotechnology, Faculty of Bioscience Engineering, Ghent University

References

  1. Hammerschlag MR: The intracellular life of chlamydiae. Semin Pediatr Infect Dis. 2002, 13: 239-248. 10.1053/spid.2002.127201.View ArticlePubMedGoogle Scholar
  2. Willingan DA, Beamer PD: Isolation of a transmissible agent from pericarditis of swine. J Am Vet Med Assoc. 1955, 126: 118-122.Google Scholar
  3. Guenov I: Etudes sur la pericardite fibrineuse des porcelets due au virus de l'ornithose. Bull Off Int Epizoot. 1961, 55: 1465-1473.Google Scholar
  4. Popovici V, Hiastru F, Draghici D, Berbinschi C, Dorobantu R: Bedsonia isolierung von Schweinen mit verschiedenen Krankheiten. Lucr Inst Cercet Vet Bioprep Pasteur. 1972, 8: 19-28. (article in German)Google Scholar
  5. Sorodoc G, Surdan C, Sarateano D, Suteo V: Research on the identification of enzootic swine bronchopneumonia virus. Rev Sci Med. 1961, 6: 113-115. (article in French)PubMedGoogle Scholar
  6. Kölbl O: Untersuchungen über das Vorkommen von Miyagawanellen beim Schwein. Wiener Tierärztl Monatssch. 1969, 56: 355-361. (article in German)Google Scholar
  7. Kölbl O, Burtscher H, Hebenstreit J: Polyarthritis bei Schlachtsweinen. Mikrobiologische, histologische und fleischhygienische Untersuchungen und Aspekte. Wiener Tierärztl Monatssch. 1970, 57: 355-361. (article in German)Google Scholar
  8. Leonard I, Wittenbrink MM, Bisping W: Nachweis von Chlamydia psittaci im Kot von Schweinen. Berl Munch Tierärztl Wschr. 1988, 101: 124-128. (article in German)Google Scholar
  9. Plagemann O: Chlamydien als Abortursache beim Schwein und als Differentialdiagnose zum Smedi-Komplex. Tierärztl Umsch. 1981, 36: 842-846. (article in German)Google Scholar
  10. Rogers DG, Andersen AA, Hogg A, Nielsen DL, Huebert MA: Conjunctivitis and Keratoconjunctivitis Associated with Chlamydiae in Swine. J Am Vet Med Assoc. 1993, 203: 1321-1323.PubMedGoogle Scholar
  11. Schiller I, Koesters R, Weilenmann R, Thoma R, Kaltenboeck B, Heitz P, Pospischil A: Mixed infections with porcine Chlamydia trachomatis/pecorum and infections with ruminant Chlamydia psittaci serovar 1 associated with abortions in swine. Vet Microbiol. 1997, 58: 251-260. 10.1016/S0378-1135(97)00154-5.View ArticlePubMedGoogle Scholar
  12. Szeredi L, Schiller I, Sydler T, Guscetti F, Heinen E, Corboz L, Eggenberger E, Jones GE, Pospischil A: Intestinal Chlamydia in finishing pigs. Vet Pathol. 1996, 33: 369-374. 10.1177/030098589603300401.View ArticlePubMedGoogle Scholar
  13. Thoma R, Guscetti F, Schiller I, Schmeer N, Corboz L, Pospischil A: Chlamydiae in porcine abortion. Vet Pathol. 1997, 34: 467-469. 10.1177/030098589703400512.View ArticlePubMedGoogle Scholar
  14. Wittenbrink MM: Detection of antibodies against Chlamydia in swine by an immunofluorescent test and an enzyme immunoassay. Berl Munch Tierärztl Wochenschr. 1991, 104: 270-275. (article in German)PubMedGoogle Scholar
  15. Zahn I, Szeredi L, Schiller I, Kunz US, Burgi E, Guscetti F, Heinen E, Corboz L, Sydler T, Pospischil A: Immunhistologischer Nachweis von Chlamydia psittaci/pecorum und C. trachomatis im Ferkel-Darm. Zentralbl Veterinärmed. 1995, 42: 266-276. (article in German)Google Scholar
  16. Rogers DG, Andersen AA, Hunsaker BD: Lung and nasal lesions caused by a swine chlamydial isolate in gnotobiotic pigs. J Vet Diagn Invest. 1996, 8: 45-55.View ArticlePubMedGoogle Scholar
  17. Rogers DG, Andersen AA: Intestinal lesions caused by two swine chlamydial isolates in gnotobiotic pigs. J Vet Diagn Invest. 1996, 8: 433-440.View ArticlePubMedGoogle Scholar
  18. Rogers DG, Andersen AA: Conjunctivitis caused by a swine Chlamydia trachomatis-like organism in gnotobiotic pigs. J Vet Diagn Invest. 1999, 11: 341-344.View ArticlePubMedGoogle Scholar
  19. Woollen N, Daniels EK, Yeary T, Leipold HW, Phillips RM: Chlamydial infection and perinatal mortality in a swine herd. J Am Vet Med Assoc. 1990, 197: 600-601.PubMedGoogle Scholar
  20. Sarma DK, Tamuli MK, Rahman T, Boro BR, Deka BC, Rajkonwar CK: Isolation of Chlamydia from a pig with lesions in the urethra and prostate gland. Vet Rec. 1983, 112: 525-10.1136/vr.112.22.525-a.View ArticlePubMedGoogle Scholar
  21. Eggemann G, Wendt M, Hoelzle LE, Jager C, Weiss R, Failing K: Prevalence of chlamydial infections in breeding sows and their correlation to reproductive failure. Dtsch Tierarztl Wochenschr. 2000, 107: 3-10. (article in German)PubMedGoogle Scholar
  22. Everett KDE, Bush RM, Andersen AA: Emended description of the order Chlamydiales, proposal of Parachlamydiaceae fam. nov. and Simkaniaceae fam. nov., each containing one monotypic genus, revised taxonomy of the family Chlamydiaceae, including a new genus and five new species, and standards for the identification of organisms. Int J Syst Bacteriol. 1999, 49: 415-440. 10.1099/00207713-49-2-415.View ArticlePubMedGoogle Scholar
  23. Stephens RS, Myers G, Eppinger M, Bavoil PM: Divergence without difference: phylogenetics and taxonomy of Chlamydia resolved. FEMS Immunol Med Microbiol. 2009, 55: 115-119. 10.1111/j.1574-695X.2008.00516.x.View ArticlePubMedGoogle Scholar
  24. Kerr K, Entrican G, McKeever D, Longbottom D: Immunopathology of Chlamydophila abortus infection in sheep and mice. Res Vet Sci. 2005, 78: 1-7. 10.1016/j.rvsc.2004.08.004.View ArticlePubMedGoogle Scholar
  25. Everett KDE: Chlamydia and Chlamydiales: more than meets the eye. Vet Microbiol. 2000, 75: 109-126. 10.1016/S0378-1135(00)00213-3.View ArticlePubMedGoogle Scholar
  26. Fukushi H, Hirai K: Proposal of Chlamydia pecorum Sp-Nov for Chlamydial strains derived from ruminants. Int J Syst Bacteriol. 1992, 42: 306-308. 10.1099/00207713-42-2-306.View ArticlePubMedGoogle Scholar
  27. Andersen AA: Chlamydial diseases in swine. Proc 25th Ann Meet Am Assoc Swine Pract. 1994, 259-263.Google Scholar
  28. Kaltenboeck B, Storz J: Biological properties and genetic analysis of the ompA locus in Chlamydiae isolated from swine. Am J Vet Res. 1992, 53: 1482-1487.PubMedGoogle Scholar
  29. Busch M, Thoma R, Schiller I, Corboz L, Pospischil A: Occurrence of Chlamydiae in the genital tracts of sows at slaughter and their possible significance for reproductive failure. J Vet Med B Infect Dis Vet Public Health. 2000, 47: 471-480.View ArticlePubMedGoogle Scholar
  30. Vanrompay D, Geens T, Desplanques A, Hoang TQT, De Vos L, Van Loock M, Huyck E, Miry C, Cox E: Immunoblotting, ELISA and culture evidence for Chlamydiaceae in sows on 258 Belgian farms. Vet Microbiol. 2004, 99: 59-66. 10.1016/j.vetmic.2003.08.014.View ArticlePubMedGoogle Scholar
  31. Andersen AA, Rogers DG: Resistance to tetracycline and sulfadiazine in swine C. trachomatis isolates. Proceedings of the Ninth International Symposium on Human Chlamydial Infection. Edited by: Stephens. 1998, 313-316. Chlamydial infectionsGoogle Scholar
  32. Di Francesco A, Donati M, Rossi M, Pignanelli S, Shurdhi A, Baldelli R, Cevenini R: Tetracycline-resistant Chlamydia suis isolates in Italy. Vet Rec. 2008, 163: 251-252. 10.1136/vr.163.8.251.View ArticlePubMedGoogle Scholar
  33. Schautteet K: Epidemiological Research on Chlamydiaceae in pigs and evaluation of a Chlamydia trachomatis DNA vaccine. 2010, Ghent, Belgium: Ghent UniversityGoogle Scholar
  34. Nietfeld JC, Janke BH, Leslie-Steen P, Robison DJ, Zeman DH: Small intestinal Chlamydia infection in piglets. J Vet Diagn Invest. 1993, 5: 114-117.View ArticlePubMedGoogle Scholar
  35. Rogers DG, Andersen AA: Intestinal lesions caused by a strain of Chlamydia suis in weanling pigs infected at 21 days of age. J Vet Diagn Invest. 2000, 12: 233-239.View ArticlePubMedGoogle Scholar
  36. Schautteet K, Beeckman DS, Delava P, Vanrompay D: Possible pathogenic interplay between Chlamydia suis, Chlamydophila abortus and PCV-2 on a pig production farm. Vet Rec. 2010, 166: 329-333. 10.1136/vr.b4714.View ArticlePubMedGoogle Scholar
  37. Bush RM, Everett KDE: Molecular evolution of the Chlamydiaceae. Int J Syst Evol Microbiol. 2001, 51: 203-220.View ArticlePubMedGoogle Scholar
  38. Becker A, Lutz-Wohlgroth L, Brugnera E, Lu ZH, Zimmermann DR, Grimm F, Grosse BE, Kaps S, Spiess B, Pospischil A, Vaughan L: Intensively kept pigs pre-disposed to chlamydial associated conjunctivitis. J Vet Med A Physiol Pathol Clin Med. 2007, 54: 307-313.View ArticlePubMedGoogle Scholar
  39. Hoelzle LE, Steinhausen G, Wittenbrink MM: PCR-based detection of chlamydial infection in swine and subsequent PCR-coupled genotyping of chlamydial omp1-gene amplicons by DNA-hybridization, RFLP-analysis, and nucleotide sequence analysis. Epidemiol Infect. 2000, 125: 427-439. 10.1017/S0950268899004446.PubMed CentralView ArticlePubMedGoogle Scholar
  40. Guseva NV, Knight ST, Whittimore JD, Wyrick PB: Primary cultures of female swine genital epithelial cells in vitro: a new approach for the study of hormonal modulation of chlamydial infection. Infect Immun. 2003, 71: 4700-4710. 10.1128/IAI.71.8.4700-4710.2003.PubMed CentralView ArticlePubMedGoogle Scholar
  41. Schiller I, Schifferli A, Gysling P, Pospischil A: Growth characteristics of porcine chlamydial strains in different cell culture systems and comparison with ovine and avian chlamydial strains. Vet J. 2003, 168: 74-80. 10.1016/S1090-0233(03)00039-X.View ArticleGoogle Scholar
  42. Bagdonas J, Mauricas M, Gerulis G, Petkevicius S, Jokimas J: Evaluation of different laboratory methods for diagnosis of pig chlamydiosis in Lithuania. Pol J Vet Sci. 2005, 8: 49-56.PubMedGoogle Scholar
  43. Pospischil A, Borel N, Chowdhury EH, Guscetti F: Aberrant chlamydial developmental forms in the gastrointestinal tract of pigs spontaneously and experimentally infected with Chlamydia suis. Vet Microbiol. 2009, 135: 147-156. 10.1016/j.vetmic.2008.09.035.View ArticlePubMedGoogle Scholar
  44. Hotzel H, Berndt A, Melzer F, Sachse K: Occurrence of Chlamydiaceae spp. in a wild boar (Sus scrofa L.) population in Thuringia (Germany). Vet Microbiol. 2004, 103: 121-126. 10.1016/j.vetmic.2004.06.009.View ArticlePubMedGoogle Scholar
  45. Kauffold J, Melzer F, Henning K, Schulze K, Leiding C, Sachse K: Prevalence of chlamydiae in boars and semen used for artificial insemination. Theriogenology. 2006, 65: 1750-1758. 10.1016/j.theriogenology.2005.10.010.View ArticlePubMedGoogle Scholar
  46. Sachse K, Hotzel H, Slickers P, Ellinger T, Ehricht R: DNA microarray-based detection and identification of Chlamydia and Chlamydophila spp. Mol Cell Probes. 2005, 19: 41-50. 10.1016/j.mcp.2004.09.005.View ArticlePubMedGoogle Scholar
  47. Van Loock M, Vanrompay D, Herrmann B, Vander SJ, Volckaert G, Goddeeris BM, Everett KD: Missing links in the divergence of Chlamydophila abortus from Chlamydophila psittaci. Int J Syst Evol Microbiol. 2003, 53: 761-770. 10.1099/ijs.0.02329-0.View ArticlePubMedGoogle Scholar
  48. Mohamad KY, Rodolakis A: Recent advances in the understanding of Chlamydophila pecorum infections, sixteen years after it was named as the fourth species of the Chlamydiaceae family. Vet Res. 2010, 41: 27-10.1051/vetres/2009075.View ArticlePubMedGoogle Scholar
  49. Pannekoek Y, Morelli G, Kusecek B, Morre SA, Ossewaarde JM, Langerak AA, van der EA: Multi locus sequence typing of Chlamydiales: clonal groupings within the obligate intracellular bacteria Chlamydia trachomatis. BMC Microbiol. 2008, 8: 42-10.1186/1471-2180-8-42.PubMed CentralView ArticlePubMedGoogle Scholar
  50. Wilson MR, Plummer P: A survey of pig sera for presence of antibodies to psittacosis-lymphogranuloma-venereum group of organisms. J Comp Pathol. 1966, 76: 427-433. 10.1016/0021-9975(66)90064-8.View ArticlePubMedGoogle Scholar
  51. Verminnen K, Van Loock M, Hafez HM, Ducatelle R, Haesebrouck F, Vanrompay D: Evaluation of a recombinant enzyme-linked immunosorbent assay for detecting Chlamydophila psittaci antibodies in turkey sera. Vet Res. 2006, 37: 623-632. 10.1051/vetres:2006023.View ArticlePubMedGoogle Scholar
  52. Vanrompay D, Cox E, Mast J, Goddeeris B, Volckaert G: High-level expression of Chlamydia psittaci major outer membrane protein in COS cells and in skeletal muscles of turkeys. Infect Immun. 1998, 66: 5494-5500.PubMed CentralPubMedGoogle Scholar
  53. Hoelzle LE, Hoelzle K, Wittenbrink MM: Recombinant major outer membrane protein (MOMP) of Chlamydophila abortus, Chlamydophila pecorum, and Chlamydia suis as antigens to distinguish chlamydial species-specific antibodies in animal sera. Vet Microbiol. 2004, 103: 85-90. 10.1016/j.vetmic.2004.07.016.View ArticlePubMedGoogle Scholar
  54. Camenisch U, Lu ZH, Vaughan L, Corboz L, Zimmermann DR, Wittenbrink MM, Pospischil A, Sydler T: Diagnostic investigation into the role of Chlamydiae in cases of increased rates of return to oestrus in pigs. Vet Rec. 2004, 155: 593-596. 10.1136/vr.155.19.593.View ArticlePubMedGoogle Scholar
  55. Di Francesco A, Baldelli R, Cevenini R, Magnino S, Pignanelli S, Salvatore D, Galuppi R, Donati M: Seroprevalence to Chlamydiae in pigs in Italy. Vet Rec. 2006, 159: 849-850.PubMedGoogle Scholar
  56. Brade H, Brade L, Nano FE: Chemical and serological investigations on the genus-specific lipopolysaccharide epitope of Chlamydia. Proc Natl Acad Sci USA. 1987, 84: 2508-2512. 10.1073/pnas.84.8.2508.PubMed CentralView ArticlePubMedGoogle Scholar
  57. Caldwell HD, Hitchcock PJ: Monoclonal antibody against a genus-specific antigen of Chlamydia species: location of the epitope on chlamydial lipopolysaccharide. Infect Immun. 1984, 44: 306-314.PubMed CentralPubMedGoogle Scholar
  58. Nurminen M, Lounatmaa K, Leinonen M, Wahlstrom E: The effect of mercaptoethanol on the solubilization of the 39.5 kDa major outer membrane protein of elementary bodies of Chlamydia trachomatis and purification of the protein. FEMS Microbiol Lett. 1984, 24: 185-191. 10.1111/j.1574-6968.1984.tb01302.x.View ArticleGoogle Scholar
  59. Yuan Y, Lyng K, Zhang YX, Rockey DD, Morrison RP: Monoclonal-antibodies define genus-specific, species-specific, and cross-reactive epitopes of the chlamydial 60-kilodalton heat-shock protein (Hsp60): specific immunodetection and purification of chlamydial Hsp60. Infect Immun. 1992, 60: 2288-2296.PubMed CentralPubMedGoogle Scholar
  60. Henning K, Sachse K, Kirschen P, Bohmer J, Strutzberg-Minder K, Grossmann E: An enzyme-linked immunosorbent assay (ELISA) for the detection of anti-chlamydial antibodies in pig sera. Berl Munch Tierarztl Wochenschr. 2005, 118: 1-7. (article in German)PubMedGoogle Scholar
  61. Qiu QC: Investigation on Chlamydiosis in pigs. Progr Vet Med. 2003, 21: 88-91.Google Scholar
  62. Zhou JZ, Qiu CQ: Epidemic of animal Chlamydiae in China. Chin Husbandry Vet. 2007, 34: 110-113.Google Scholar
  63. Xu MJ, He Y, Liang R, Zhou DH, Lin RQ, Yin CC, He XH, Liang M, Zhu XQ: Seroprevalence of Chlamydia infection in pigs from intensive farms in Southern China. J Anim Vet Adv. 2010, 9: 1143-1145. 10.3923/javaa.2010.1143.1145.View ArticleGoogle Scholar
  64. Pantchev A, Sting R, Bauerfeind R, Tyczka J, Sachse K: Detection of all Chlamydophila and Chlamydia spp. of veterinary interest using species-specific real-time PCR assays. Comp Immunol Microbiol Infect Dis. 2009, 33: 473-484. 10.1016/j.cimid.2009.08.002.View ArticlePubMedGoogle Scholar
  65. Dugan J, Rockey DD, Jones L, Andersen AA: Tetracycline resistance in Chlamydia suis mediated by genomic islands inserted into the chlamydial inv-like gene. Antimicrob Agents Chemother. 2004, 48: 3989-3995. 10.1128/AAC.48.10.3989-3995.2004.PubMed CentralView ArticlePubMedGoogle Scholar
  66. Pavlov P, Milanov M, Tchilev D: Recherches sur la rickettsiose kerato-conjonctivale du porc en Bulgarie. Ann Inst Pasteur (Paris). 1963, 105: 450-454. (article in French)Google Scholar
  67. Guscetti F, Schiller I, Sydler T, Heinen E, Pospischil A: Experimental enteric infection of gnotobiotic piglets with Chlamydia suis strain S45. Vet Microbiol. 2009, 135: 157-168. 10.1016/j.vetmic.2008.09.038.View ArticlePubMedGoogle Scholar
  68. Reinhold P, Kirschvink N, Theegarten D, Berndt A: An experimentally induced Chlamydia suis infection in pigs results in severe lung function disorders and pulmonary inflammation. Vet Res. 2008, 39: 35-10.1051/vetres:2008012.View ArticlePubMedGoogle Scholar
  69. Vazquez-Cisneros C, Wilsmore AJ, Bollo E: Experimental infections of pregnant sows with ovine Chlamydia psittaci strains. Vet Microbiol. 1994, 42: 383-387. 10.1016/0378-1135(94)90069-8.View ArticlePubMedGoogle Scholar
  70. Guscetti F, Hoop R, Schiller I, Corboz L, Sydler T, Pospischil A: Experimental enteric infection of gnotobiotic piglets with a Chlamydia psittaci strain of avian origin. J Vet Med B Infect Dis Vet Public Health. 2000, 47: 561-572.View ArticlePubMedGoogle Scholar
  71. Guscetti F, Schiller I, Sydler T, Corboz L, Pospischil A: Experimental Chlamydia psittaci serotype 1 enteric infection in gnotobiotic piglets: Histopathological, immunohistochemical and microbiological findings. Vet Microbiol. 1998, 62: 251-263. 10.1016/S0378-1135(98)00221-1.View ArticlePubMedGoogle Scholar
  72. Storz J: Overview of animal diseases induced by chlamydial infections. Edited by: Barron AL. 1988, Microbiology of Chlamydia, CRC press, Florida, 167-192.Google Scholar
  73. Beeckman DS, Vanrompay DC: Zoonotic Chlamydophila psittaci infections from a clinical perspective. Clin Microbiol Infect. 2009, 15: 11-17. 10.1111/j.1469-0691.2008.02669.x.View ArticlePubMedGoogle Scholar
  74. Longbottom D: Chlamydial vaccine development. J Med Microbiol. 2003, 52: 537-540. 10.1099/jmm.0.05093-0.View ArticlePubMedGoogle Scholar
  75. Smith KA, Bradley KK, Stobierski MG, Tengelsen LA: Compendium of measures to control Chlamydophila psittaci (formerly Chlamydia psittaci) infection among humans (psittacosis) and pet birds, 2005. J Am Vet Med Assoc. 2005, 226: 532-539. 10.2460/javma.2005.226.532.View ArticlePubMedGoogle Scholar
  76. Pollmann M, Nordhoff M, Pospischil A, Tedin K, Wieler LH: Effects of a probiotic strain of Enterococcus faecium on the rate of natural chlamydia infection in swine. Infect Immun. 2005, 73: 4346-4353. 10.1128/IAI.73.7.4346-4353.2005.PubMed CentralView ArticlePubMedGoogle Scholar
  77. Knitz JC, Hoelzle LE, Affolter P, Hamburger A, Zimmermann K, Heinritzi K, Wittenbrink MM: Humoral immune response in sows vaccinated with a bacterin prepared from a herd-derived Chlamydophila abortus strain. Dtsch Tierarztl Wochenschr. 2003, 110: 369-374. (article in German)PubMedGoogle Scholar
  78. Zhang F, Li S, Yang J, Yang L, He C: Induction of a protective immune response against swine Chlamydophila abortus infection in mice following co-vaccination of omp-1 DNA with recombinant MOMP. Zoonoses Public Health. 2009, 56: 71-76. 10.1111/j.1863-2378.2008.01160.x.View ArticlePubMedGoogle Scholar
  79. Stephens RS, Kalman S, Lammel C, Fan J, Marathe R, Aravind L, Mitchell W, Olinger L, Tatusov RL, Zhao QX, Koonin EV, Davis RW: Genome sequence of an obligate intracellular pathogen of humans: Chlamydia trachomatis. Science. 1998, 282: 754-759. 10.1126/science.282.5389.754.View ArticlePubMedGoogle Scholar
  80. Carlson JH, Porcella SF, McClarty G, Caldwell HD: Comparative genomic analysis of Chlamydia trachomatis oculotropic and genitotropic strains. Infect Immun. 2005, 73: 6407-6418. 10.1128/IAI.73.10.6407-6418.2005.PubMed CentralView ArticlePubMedGoogle Scholar
  81. Thomson NR, Holden MT, Carder C, Lennard N, Lockey SJ, Marsh P, Skipp P, O'Connor CD, Goodhead I, Norbertzcak H, Harris B, Ormond D, Rance R, Quail MA, Parkhill J, Stephens RS, Clarke IN: Chlamydia trachomatis: genome sequence analysis of lymphogranuloma venereum isolates. Genome Res. 2008, 18: 161-171. 10.1101/gr.7020108.PubMed CentralView ArticlePubMedGoogle Scholar
  82. Kalman S, Mitchell W, Marathe R, Lammel C, Fan J, Hyman RW, Olinger L, Grimwood J, Davis RW, Stephens RS: Comparative genomes of Chlamydia pneumoniae and Chlamydia trachomatis. Nat Genet. 1999, 21: 385-389. 10.1038/7716.View ArticlePubMedGoogle Scholar
  83. Shirai M, Hirakawa H, Kimoto M, Tabuchi M, Kishi F, Ouchi K, Shiba T, Ishii K, Hattori M, Kuhara S, Nakazawa T: Comparison of whole genome sequences of Chlamydia pneumoniae J138 from Japan and CWL029 from USA. Nucleic Acids Res. 2000, 28: 2311-2314. 10.1093/nar/28.12.2311.PubMed CentralView ArticlePubMedGoogle Scholar
  84. Read TD, Brunham RC, Shen C, Gill SR, Heidelberg JF, White O, Hickey EK, Peterson J, Utterback T, Berry K, Bass S, Linher K, Weidman J, Khouri H, Craven B, Bowman C, Dodson R, Gwinn M, Nelson W, DeBoy R, Kolonay J, McClarty G, Salzberg SL, Eisen J, Fraser CM: Genome sequences of Chlamydia trachomatis MoPn and Chlamydia pneumoniae AR39. Nucleic Acids Res. 2000, 28: 1397-1406. 10.1093/nar/28.6.1397.PubMed CentralView ArticlePubMedGoogle Scholar
  85. NCBI. genome C. pneumonia TW-183. 2010, [http://www.ncbi.nlm.nih.gov/sites/entrez?Db=genome&Cmd=ShowDetailView&TermToSearch=311]
  86. Myers GS, Mathews SA, Eppinger M, Mitchell C, O'Brien KK, White OR, Benahmed F, Brunham RC, Read TD, Ravel J, Bavoil PM, Timms P: Evidence that human Chlamydia pneumoniae was zoonotically acquired. J Bacteriol. 2009, 191: 7225-7233. 10.1128/JB.00746-09.PubMed CentralView ArticlePubMedGoogle Scholar
  87. Read TD, Myers GS, Brunham RC, Nelson WC, Paulsen IT, Heidelberg J, Holtzapple E, Khouri H, Federova NB, Carty HA, Umayam LA, Haft DH, Peterson J, Beanan MJ, White O, Salzberg SL, Hsia RC, McClarty G, Rank RG, Bavoil PM, Fraser CM: Genome sequence of Chlamydophila caviae (Chlamydia psittaci GPIC): examining the role of niche-specific genes in the evolution of the Chlamydiaceae. Nucleic Acids Res. 2003, 31: 2134-2147. 10.1093/nar/gkg321.PubMed CentralView ArticlePubMedGoogle Scholar
  88. Thomson NR, Yeats C, Bell K, Holden MT, Bentley SD, Livingstone M, Cerdeno-Tarraga AM, Harris B, Doggett J, Ormond D, Mungall K, Clarke K, Feltwell T, Hance Z, Sanders M, Quail MA, Price C, Barrell BG, Parkhill J, Longbottom D: The Chlamydophila abortus genome sequence reveals an array of variable proteins that contribute to interspecies variation. Genome Res. 2005, 15: 629-640. 10.1101/gr.3684805.PubMed CentralView ArticlePubMedGoogle Scholar
  89. Azuma Y, Hirakawa H, Yamashita A, Cai Y, Rahman MA, Suzuki H, Mitaku S, Toh H, Goto S, Murakami T, Sugi K, Hayashi H, Fukushi H, Hattori M, Kuhara S, Shirai M: Genome sequence of the cat pathogen, Chlamydophila felis. DNA Res. 2006, 13: 15-23. 10.1093/dnares/dsi027.View ArticlePubMedGoogle Scholar
  90. Horn M, Collingro A, Schmitz-Esser S, Beier CL, Purkhold U, Fartmann B, Brandt P, Nyakatura GJ, Droege M, Frishman D, Rattei T, Mewes HW, Wagner M: Illuminating the evolutionary history of chlamydiae. Science. 2004, 304: 728-730. 10.1126/science.1096330.View ArticlePubMedGoogle Scholar

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