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Wide range of Chlamydiales types detected in native Australian mammals
Veterinary Microbiology 96 (2003) 177–187
Wide range of Chlamydiales types detected in native Australian mammals Tracey J. Bodetti a , Karen Viggers b , Kristin Warren c , Ralph Swan c , Sue Conaghty d , Colleen Sims e , Peter Timms a,∗ a
Centre for Molecular Biotechnology, School of Life Sciences, Queensland University of Technology, Brisbane, Australia b Research School of Biological Sciences, Australian National University, Canberra, Australia c Division of Veterinary and Biomedical Sciences, Murdoch University, Murdoch, Australia d Monarto Zoological Park, Monarto, Australia e Department of Conservation and Land Management, Denham, Australia Received 21 October 2002; received in revised form 8 May 2003; accepted 3 June 2003
Abstract The Chlamydiales are a unique order of intracellular bacterial pathogens that cause significant disease of birds and animals, including humans. The recent development of a Chlamydiales-specific 16S rDNA polymerase chain reaction (PCR) assay has enabled the identification of Chlamydiales DNA from an increasing range of hosts and environmental sources. Whereas the Australian marsupial, the koala, has previously been shown to harbour several Chlamydiales types, no other Australian marsupials have been analysed. We therefore used a 16S rDNA PCR assay combined with direct sequencing to determine the presence and genotype of Chlamydiales in five wild Australian mammals (gliders, possums, bilbies, bandicoots, potoroos). We detected eight previously observed Chlamydiales genotypes as well as 10 new Chlamydiales sequences from these five Australian mammals. In addition to PCR analysis we used antigen specific staining and in vitro culture in HEp-2 cell monolayers to confirm some of the identifications. A strong association between ocular PCR positivity and the presence of clinical disease (conjunctivitis, proliferation of the eyelid) was observed in two of the species studied, gliders and bandicoots, whereas little clinical disease was observed in the other animals studied. These findings provide further evidence that novel Chlamydiales infections occur in a wide range of hosts and that, in some of these, the chlamydial infections may contribute to clinical disease. © 2003 Elsevier B.V. All rights reserved. Keywords: Chlamydiales; Marsupiales; Australia
∗
Corresponding author. Tel.: +61-7-3864-2120; fax: +61-7-3864-1534. E-mail address: [email protected] (P. Timms). 0378-1135/$ – see front matter © 2003 Elsevier B.V. All rights reserved. doi:10.1016/S0378-1135(03)00211-6
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1. Introduction Chlamydiae are obligate intracellular bacterial pathogens of a wide range of birds and animals including humans. Traditionally, the chlamydiae have been characterized by their unique biphasic developmental cycle, involving the inter-conversion between an extracellular survival form, the elementary body and an intracellular replicative form, the reticulate body. Prior to 1999, the family Chlamydiaceae consisted of one genus and four species, Chlamydia trachomatis, Chlamydophila psittaci, C. pecorum and C. pneumoniae. In 1999, Everett et al. proposed a reclassification of Chlamydia into two genera and nine species (C. trachomatis, C. suis, and C. muridarum and C. psittaci, C. pneumoniae, C. felis, C. pecorum, C. abortus, and C. caviae). Chlamydiales cause a wide range of diseases with symptoms ranging from inapparent, through to severe in its many hosts. In humans, chlamydial infection causes a number of relatively mild diseases including conjunctivitis, rhinitis and urethritis (Grayston and Wang, 1975; Jones, 1975; Storz, 1988; Grayston et al., 1993), more severe disease including pneumonia and trachoma, and is also implicated in chronic diseases such as pelvic inflammatory disease, infertility, Reiter’s syndrome and cardiovascular disease (Hahn et al., 1991; Rahman et al., 1992; Kuo et al., 1993). Chlamydiales also infect a remarkably wide range of animal hosts in which they cause enteritis, respiratory disease, polyarthritis, conjunctivitis, urogential tract disease and abortion (Storz et al., 1960, 1966; Szeredi et al., 1996). The development of sensitive nucleic acid amplification techniques has enabled the detection of new Chlamydiales organisms without the need to grow them in vitro. Recent studies have identified novel Chlamydiales organisms from an increasing number of animal and environmental sources including reptiles, amphibians, amoebae and even water treatment plants (Jacobson and Telford, 1990; Kahane et al., 1998; Berger et al., 1999; Fritsche et al., 2000; Horn and Wagner, 2001). The epidemiology of chlamydial infection in humans (C. trachomatis and C. pneumoniae) and animals (C. psittaci and C. pecorum) in Australia is generally similar to that observed in other countries. C. psittaci infections are common in birds and infection with C. psittaci and C. pecorum has been observed in sheep, cattle, goats and cats (St George, 1971; Norton et al., 1989; Brown et al., 1988; Sykes et al., 1997). In addition, Australia is the home of the unique marsupial the koala, along with its chlamydial infections (C. pecorum and C. pneumoniae). The aim of the present study was to identify the types of chlamydiales present in other native Australian wildlife and to determine their association with clinical disease. 2. Materials and methods 2.1. Animals studied All animals sampled were from wild populations. 2.1.1. Greater Glider (Petauroides volans) Twenty-seven animals from Tumut, New South Wales, were swabbed at the conjunctiva and cloaca. Eight of these animals showed clinical signs of disease in the form of conjunctivitis (three) and moist or crusty cloacal tract (eight).
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2.1.2. Mountain Brushtail Possum (Trichosurus caninus) Twelve animals from Cambarville, Vic., were swabbed at the conjunctiva and cloaca. Clinical signs were not observed in these animals at the time of sampling. 2.1.3. Western Barred Bandicoot (Perameles bouganville) Two populations of bandicoots were examined in this study: 16 animals from Dryandra, Western Australia (WA) and 11 animals from Bernier Island, WA, were swabbed at conjunctiva, cloaca and nasopharynx. Thirteen of these animals showed clinical signs of disease in the form of conjunctivitis and proliferation of eyelid (13) and audible lung sounds (1). Five animals were noted to have cataracts or corneal scarring suggestive of previous ocular disease. 2.1.4. Greater Bilby (Macrotis lagotis) Twenty-one animals from Dryandra, WA, were sampled at the conjunctiva, cloaca and nasopharynx. The animals sampled did not show clinical signs of disease at the time of capture. 2.1.5. Gilberts’ Potoroo (Potorous gilbertii) Nine animals from Albany, WA, were sampled at the cloaca. Clinical disease was not observed in these animals at the time of sampling. 2.2. Sampling method A total of 95 animals from five species were swabbed at several anatomical sites (cloaca, conjunctiva and nasopharynx) for Chlamydiales testing. Following collection, swabs were placed in 1 ml of Sucrose Phosphate Glutamine Chlamydia transport media (SPG; Mass and Dalhoff, 1995) prior to sample concentration and chlamydiales detection (Bodetti et al., 2002). Samples were stored at −80 ◦ C prior to testing. Samples were vortexed to resuspend material from the swab into the SPG media, and 100 l aliquots removed for concentration. Sample aliquots were concentrated by centrifugation for 30 min at 14 000 rpm and the supernatant removed. Pellets were resuspended in 10 l of SPG for nucleic acid detection. 2.3. Nucleic acid detection 2.3.1. Chlamydiales 16S rRNA PCR A 293 bp fragment of the chlamydiales 16S rRNA gene (positions 33–326) was amplified (Everett et al., 1999). We have previously demonstrated that this polymerase chain reaction (PCR) assay sensitively detects 40 chlamydial bodies per reaction (Bodetti et al., 2002). Fifty microlitre reactions included 1 X Qiagen HotStarTaq PCR Buffer (Qiagen, Clifton Hill, Vic.), 200 M dNTPs (Roche, Melbourne, Australia) 1 M primers (16SIGF: 5 CGG CGT GGA TGA GGC AT 3 and 16SIGR: 5 TCA GTC CCA GTG TTG G 3 ), 1 U Qiagen HotStarTaq polymerase (Qiagen) and 4 l of concentrated swab material. PCR cycling conditions consisted of an initial denaturation at 95 ◦ C for 15 min, followed by 45 cycles of denaturation at 94 ◦ C for 30 s, primer annealing at 51 ◦ C for 30 s and extension at 72 ◦ C for
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45 s with a final extension for 5 min at 72 ◦ C. Following amplification, the products were separated by agarose gel electrophoresis and visualized by ethidium bromide staining. 2.4. PCR controls To ensure that the PCR products were not the result of contamination, one negative control (ddH2 O as template) was included for every three test specimens. Two positive controls of cell-cultured Chlamydiales were included with each PCR batch to confirm that amplification had occurred. 2.5. Isolate genotyping 16S rDNA PCR products intended for sequencing were extracted using Qiagen QIAquick gel extraction kit (Qiagen) as per the manufacturers’ protocol. Sequencing of the purified products to determine genotypes was performed using the dideoxynucleotide terminator method (Bodetti et al., 2002; Sanger et al., 1977). 2.6. Antigen detection Ten microlitres of concentrated swab material was spotted onto glass microscope slides and fixed in methanol. Slides were stained using a Chlamydiaceae family-specific antilipopolysaccharide (anti-LPS) monoclonal antibody (Cellabs, Sydney, Australia) according to the manufacturer’s instructions. The presence or absence of characteristic apple-greenfluorescing chlamydial particles was determined by examining duplicate wells by confocal microscopy at a magnification of 960× (TCS 4D; Leica, Leitz, Germany). 2.7. Cell culture isolation Confluent HEp-2 cell monolayers grown on glass coverslips in shell vials were used to isolate viable chlamydiales. Infection of the monolayers involved inoculation with 100 l of concentrated swab material in 1 ml of Dulbecco’s minimal essential medium (DMEM; InVitrogen, Melbourne, Australia) an initial 1 h centrifugation at 800×g. DMEM containing cycloheximide (1 mg/ml; Sigma, Melbourne, Australia) replaced the inoculum after 2 h incubation at 37 ◦ C 5% CO2 . Cultures were incubated for 9 days with repeat centrifugations on days 4–7 post-infection and fresh media on days 4 and 7. Up to three blind passages were performed prior to staining with the previously mentioned anti-LPS antibody to examine growth.
3. Results 3.1. Nucleic acid detection The 16S rDNA PCR assay detected chlamydial DNA in a total of 67% (62/95) of the animals. The individual animal species positivity was quite high for all five hosts, ranging
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Table 1 Detection of Chlamydiales in a range of Australian mammals using a 16S rDNA PCR assay Host
Number of animals tested
16S rRNA PCR-positive
Clinical disease Disease sitea
16S rRNA PCR-positiveb
Greater Glider (P. volans)
27
20/27 (74%)
Mountain Brushtail Possum (T. caninus) Western Barred Bandicoot (P. bouganville) Greater Bilby (M. lagotis) Gilberts’ Potoroo (P. gilbertii)
12
6/12 (50%)
Ocular (3/27) Ocular (3/3) None
Urogenital (8/27) Urogenital (4/8) N/A
26
19/26 (73%)
21 9
11/21 (52%) 4/9 (44%)
Ocular (13/26) Respiratory (1/26) None None
Ocular (13/13) Respiratory (0/1) N/A N/A
Total
95
62/95 (64%)
Ocular (16/95) Urogenital (8/95) Respiratory (1/95)
Ocular (16/16) Urogenital (4/8) Respiratory (0/1)
a b
Signs of clinical disease present (number with disease/total number of animals). Number of PCR-positive results in animals with clinical disease.
from 44% (potoroos) to 74% (bandicoots) (Table 1). Clinical signs of disease in the form of conjunctivitis or urogenital tract disease symptoms were present in 23% (22/95) of the total animals examined, with conjunctivitis the most common sign observed (73%, 16/22). A high proportion (32%; 20/62) of the PCR-positive animals had signs of clinical disease, compared to only 6% (2/33) of the PCR negative animals. Of the 95 animals tested, the conjunctiva was the site most commonly found to be PCR-positive and at this site, PCR positivity correlated strongly with clinical disease. Forty-five percentage (42/95) of animals were positive by PCR at the ocular site, including 100% of those with clinical ocular disease. This contrasted quite strongly with the urogenital site, where a significant portion of the animals were PCR-positive (36%) but only half of the eight animals with clinical signs of urogenital disease (crusty cloaca) were PCR-positive. 3.2. Speciation by 16S rDNA genotyping: Eighteen 16S rDNA positive PCR products were selected for direct sequencing. Sequencing identified similarities to several chlamydiales isolates/species currently listed in GenBank, including C. pecorum (4/18), endosymbiont of Acanthamoebae (2/18), Parachlamydia sp. Hall’s coccus (1/18) and Waddlia chondrophila (1/18). The most common identification however was of novel uncultured Chlamydiales (10/18) (Table 2; Fig. 1). 3.3. Antigen detection Twenty-nine 16S rDNA PCR-positive swab samples from 22 animals were selected for antigen detection by direct immunofluorescence. Thirteen (45%) of these swab samples showed characteristic fluorescence of spherical particles of approximately 0.3–2.0 M (data
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Fig. 1. Alignment of 183 bp of the 16S rRNA gene sequences determined in this study (underlined) with previously determined sequence types; N. hartmanellae (GenBank AF177275); C. pecorum bird isolate (GenBank E17343); uncultured Chlamydiales isolates CRG3, CRG23, CRG45 (GenBank AF097186, AY013399, AY013421). Possum and WBB Parachlamydia: sequences isolated from a Mountain Brushtail Possum and Western Barred Bandicoot, respectively, with similarity to sequence from an isolate of Parachlamydia acanthamoebae; Possum, Glider, WBB C. pecorum: from a bird isolate of C. pecorum; Possum, Glider, Bilby and WBB Uncultured: sequences isolated from a Mountain Brushtail Possum, Greater Glider, Greater Bilby and two Western Barred Bandicoots, respectively, with similarity to sequence from uncultured Chlamydiales.
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Fig. 1. (Continued ).
not shown). Structures consistent with chlamydiales inclusions of approximately 5–9 M diameter were also observed (Fig. 2a). 3.4. Cell culture isolation Positive cell culture isolates were obtained from six of the 16S rDNA PCR-positive and LPS antigen-positive swab samples. Chlamydia-specific antigen staining of the cultures Table 2 Genotypes of chlamydiales PCR-positives in Australian mammals Host
Number of PCR-positives genotyped
Swab site
Species similarity by 16S rRNA sequencing
Evidence of clinical disease
Greater Glider (P. volans)
2
Mountain Brushtail Possum (T. caninus)
4
Greater Bilby (M. lagotis)
3
Western Barred Bandicoot (P. bouganville)
8
Gilberts’ Potoroo (P. gilbertii)
1
Ocular Urogenital Ocular Ocular Urogenital Urogenital Ocular Ocular Urogenital Throat Throat Throat Throat Ocular Ocular Ocular Urogenital Urogenital
Uncultured Chlamydiales C. pecorum C. pecorum Uncultured Chlamydiales Uncultured Chlamydiales Endosymbiont of Acanthamoebae Uncultured Chlamydiales Parachlamydia sp. Halls’ coccus Uncultured Chlamydiales C. pecorum Uncultured Chlamydiales Uncultured Chlamydiales Uncultured Chlamydiales Endosymbiont of Acanthamoebae Uncultured Chlamydiales Uncultured Chlamydiales C. pecorum W. chondrophila
No Ocular No No No No No No No No No No Ocular Ocular Ocular Ocular Ocular No
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Fig. 2. Fluorescent antibody staining (FITC-conjugated Chlamydia-specific lipopolysaccharide monoclonal antibody) of urogenital swab from a Mountain Brushtail Possum (a) and HEp-2 cell culture from a urogenital swab from a Western Barred Bandicoot (b). Arrows indicate positive staining chlamydial inclusions. Magnification 960×.
showed inclusions of approximately 6–13 M in diameter and particles of approximately 0.4–1.7 M (Fig. 2b). While isolation was successful for these six samples, the level of infection observed was very low, on average 5–15% of HEp-2 cells and despite several attempts, only one isolate was successfully subcultured (to five passages).
4. Discussion Recent studies have identified Chlamydiales organisms in a number of new hosts (Jacobson and Telford, 1990; Kahane et al., 1998; Berger et al., 1999; Crespo et al., 1999; Fritsche et al., 2000; Horn and Wagner, 2001). In our present study we analysed ocular, respiratory and urogenital swabs from five different Australian native marsupials (95 animals) by 16S rDNA PCR and have identified five new chlamydial hosts and nine new chlamydiales types. Direct sequencing of the Chlamydiales signature sequence enabled identification of the infecting strain. Alignment of the sequences obtained showed similarities of 85–99% to several existing chlamydiales species, including four strains of C. pecorum (94–98% similarity to bird/cattle/sheep strains) and a number of completely new, uncultured Chlamydiales strains (Table 2). In two of the species sampled, bandicoots and gliders, a strong association was observed between ocular infection and disease. Chlamydiales was detected most commonly at the ocular site for all the hosts, although clinical signs of ocular disease, specifically conjunctivitis, were observed only in the bandicoots and gliders (30% of 53 animals). All of these 16 animals with clinical disease tested positive for the presence of chlamydial DNA in the ocular swab. By comparison only half of the animals with urogenital tract disease were
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Chlamydiales PCR-positive, although it is difficult to draw conclusions from only eight animals. Development of sensitive, broad-spectrum nucleic acid detection methods has enabled much easier detection of chlamydial DNA sequences from a wide range of sources (Corsaro et al., 2001; Michel et al., 2001; Fritsche et al., 2000). While previous studies have identified novel Chlamydiales DNA sequences, protein and antigen detection and isolation approaches have not previously been attempted. We confirmed a significant proportion of our 16S rDNA PCR-positive results by immunofluorescent antibody staining of chlamydial antigens, and also by in vitro isolation. Antigen staining of the swab samples identified characteristic Chlamydiales particles in 45% (13/29) of the PCR-positive samples. While in vitro isolation was difficult and the maintenance of the isolates proved equally difficult, we were nevertheless able to obtain some primary cell culture isolations for six nucleic acid and antigen-positive specimens. The organisms observed in both the direct swab and cell-cultured materials were found both intracellularly and also free from host cells. Inclusion-like structures were identified in most of the cell culture isolates and also in 2 of the 29 direct swab samples. The lack of inclusion-like structures in most of the direct swab samples is likely due to the fact that the swabs were frozen dry, in comparison with the two inclusion-positive specimens which were placed in SPG following collection, and then frozen (thawing may then have disrupted any inclusions). The lack of LPS staining in 55% of the PCR-positive swab samples may be due either to (a) the fact that no antigen or whole Chlamydiales were present in these samples or (b) the absence of characteristic chlamydial LPS, as has been reported for Parachlamydiaceae (Everett et al., 1999). Over the past 5 years the use of 16S rDNA PCR and direct sequencing has led to the discovery of many new genotypes of uncultured Chlamydiales in an increasing range of hosts. This has included the identification of novel Chlamydiales in humans (Kahane et al., 1998; Ossewaarde and Meijer, 1999), as well as lineages previously thought to be human-restricted (e.g. C. pneumoniae) in non-human hosts (Berger et al., 1999; Reed et al., 2000; Bodetti et al., 2002). Our work has identified further novel Chlamydiales organisms in additional hosts. What remains uncertain at this time is whether: (a) these infections only occur at low levels in these hosts, (b) these new Chlamydiales can cause disease in animal and human hosts, and (c) whether these strains are host restricted or are capable of cross infecting humans and animals, if given suitable conditions. These powerful and yet difficult to verify molecular techniques are challenging previous ideas of microorganism diversity and host restriction for these important human and animal pathogens.
Acknowledgements We thank Tony Friend and Neil Thomas, Department of Conservation and Land Management, Denham, Australia and June Butcher and Stephanie Hill, Kanyana Wildlife Rehabilitation Centre Inc. for invaluable assistance with the collection of specimens. We thank Felicity Horne and Christina Theodoropoulos for assistance with confocal microscopy. This work was partly supported by the Australian Koala Foundation and the Ruth Oulton Whyte Bequest fund.
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Wide range of Chlamydiales types detected in native Australian mammals Tracey J. Bodetti a , Karen Viggers b , Kristin Warren c , Ralph Swan c , Sue Conaghty d , Colleen Sims e , Peter Timms a,∗ a
Centre for Molecular Biotechnology, School of Life Sciences, Queensland University of Technology, Brisbane, Australia b Research School of Biological Sciences, Australian National University, Canberra, Australia c Division of Veterinary and Biomedical Sciences, Murdoch University, Murdoch, Australia d Monarto Zoological Park, Monarto, Australia e Department of Conservation and Land Management, Denham, Australia Received 21 October 2002; received in revised form 8 May 2003; accepted 3 June 2003
Abstract The Chlamydiales are a unique order of intracellular bacterial pathogens that cause significant disease of birds and animals, including humans. The recent development of a Chlamydiales-specific 16S rDNA polymerase chain reaction (PCR) assay has enabled the identification of Chlamydiales DNA from an increasing range of hosts and environmental sources. Whereas the Australian marsupial, the koala, has previously been shown to harbour several Chlamydiales types, no other Australian marsupials have been analysed. We therefore used a 16S rDNA PCR assay combined with direct sequencing to determine the presence and genotype of Chlamydiales in five wild Australian mammals (gliders, possums, bilbies, bandicoots, potoroos). We detected eight previously observed Chlamydiales genotypes as well as 10 new Chlamydiales sequences from these five Australian mammals. In addition to PCR analysis we used antigen specific staining and in vitro culture in HEp-2 cell monolayers to confirm some of the identifications. A strong association between ocular PCR positivity and the presence of clinical disease (conjunctivitis, proliferation of the eyelid) was observed in two of the species studied, gliders and bandicoots, whereas little clinical disease was observed in the other animals studied. These findings provide further evidence that novel Chlamydiales infections occur in a wide range of hosts and that, in some of these, the chlamydial infections may contribute to clinical disease. © 2003 Elsevier B.V. All rights reserved. Keywords: Chlamydiales; Marsupiales; Australia
∗
Corresponding author. Tel.: +61-7-3864-2120; fax: +61-7-3864-1534. E-mail address: [email protected] (P. Timms). 0378-1135/$ – see front matter © 2003 Elsevier B.V. All rights reserved. doi:10.1016/S0378-1135(03)00211-6
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1. Introduction Chlamydiae are obligate intracellular bacterial pathogens of a wide range of birds and animals including humans. Traditionally, the chlamydiae have been characterized by their unique biphasic developmental cycle, involving the inter-conversion between an extracellular survival form, the elementary body and an intracellular replicative form, the reticulate body. Prior to 1999, the family Chlamydiaceae consisted of one genus and four species, Chlamydia trachomatis, Chlamydophila psittaci, C. pecorum and C. pneumoniae. In 1999, Everett et al. proposed a reclassification of Chlamydia into two genera and nine species (C. trachomatis, C. suis, and C. muridarum and C. psittaci, C. pneumoniae, C. felis, C. pecorum, C. abortus, and C. caviae). Chlamydiales cause a wide range of diseases with symptoms ranging from inapparent, through to severe in its many hosts. In humans, chlamydial infection causes a number of relatively mild diseases including conjunctivitis, rhinitis and urethritis (Grayston and Wang, 1975; Jones, 1975; Storz, 1988; Grayston et al., 1993), more severe disease including pneumonia and trachoma, and is also implicated in chronic diseases such as pelvic inflammatory disease, infertility, Reiter’s syndrome and cardiovascular disease (Hahn et al., 1991; Rahman et al., 1992; Kuo et al., 1993). Chlamydiales also infect a remarkably wide range of animal hosts in which they cause enteritis, respiratory disease, polyarthritis, conjunctivitis, urogential tract disease and abortion (Storz et al., 1960, 1966; Szeredi et al., 1996). The development of sensitive nucleic acid amplification techniques has enabled the detection of new Chlamydiales organisms without the need to grow them in vitro. Recent studies have identified novel Chlamydiales organisms from an increasing number of animal and environmental sources including reptiles, amphibians, amoebae and even water treatment plants (Jacobson and Telford, 1990; Kahane et al., 1998; Berger et al., 1999; Fritsche et al., 2000; Horn and Wagner, 2001). The epidemiology of chlamydial infection in humans (C. trachomatis and C. pneumoniae) and animals (C. psittaci and C. pecorum) in Australia is generally similar to that observed in other countries. C. psittaci infections are common in birds and infection with C. psittaci and C. pecorum has been observed in sheep, cattle, goats and cats (St George, 1971; Norton et al., 1989; Brown et al., 1988; Sykes et al., 1997). In addition, Australia is the home of the unique marsupial the koala, along with its chlamydial infections (C. pecorum and C. pneumoniae). The aim of the present study was to identify the types of chlamydiales present in other native Australian wildlife and to determine their association with clinical disease. 2. Materials and methods 2.1. Animals studied All animals sampled were from wild populations. 2.1.1. Greater Glider (Petauroides volans) Twenty-seven animals from Tumut, New South Wales, were swabbed at the conjunctiva and cloaca. Eight of these animals showed clinical signs of disease in the form of conjunctivitis (three) and moist or crusty cloacal tract (eight).
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2.1.2. Mountain Brushtail Possum (Trichosurus caninus) Twelve animals from Cambarville, Vic., were swabbed at the conjunctiva and cloaca. Clinical signs were not observed in these animals at the time of sampling. 2.1.3. Western Barred Bandicoot (Perameles bouganville) Two populations of bandicoots were examined in this study: 16 animals from Dryandra, Western Australia (WA) and 11 animals from Bernier Island, WA, were swabbed at conjunctiva, cloaca and nasopharynx. Thirteen of these animals showed clinical signs of disease in the form of conjunctivitis and proliferation of eyelid (13) and audible lung sounds (1). Five animals were noted to have cataracts or corneal scarring suggestive of previous ocular disease. 2.1.4. Greater Bilby (Macrotis lagotis) Twenty-one animals from Dryandra, WA, were sampled at the conjunctiva, cloaca and nasopharynx. The animals sampled did not show clinical signs of disease at the time of capture. 2.1.5. Gilberts’ Potoroo (Potorous gilbertii) Nine animals from Albany, WA, were sampled at the cloaca. Clinical disease was not observed in these animals at the time of sampling. 2.2. Sampling method A total of 95 animals from five species were swabbed at several anatomical sites (cloaca, conjunctiva and nasopharynx) for Chlamydiales testing. Following collection, swabs were placed in 1 ml of Sucrose Phosphate Glutamine Chlamydia transport media (SPG; Mass and Dalhoff, 1995) prior to sample concentration and chlamydiales detection (Bodetti et al., 2002). Samples were stored at −80 ◦ C prior to testing. Samples were vortexed to resuspend material from the swab into the SPG media, and 100 l aliquots removed for concentration. Sample aliquots were concentrated by centrifugation for 30 min at 14 000 rpm and the supernatant removed. Pellets were resuspended in 10 l of SPG for nucleic acid detection. 2.3. Nucleic acid detection 2.3.1. Chlamydiales 16S rRNA PCR A 293 bp fragment of the chlamydiales 16S rRNA gene (positions 33–326) was amplified (Everett et al., 1999). We have previously demonstrated that this polymerase chain reaction (PCR) assay sensitively detects 40 chlamydial bodies per reaction (Bodetti et al., 2002). Fifty microlitre reactions included 1 X Qiagen HotStarTaq PCR Buffer (Qiagen, Clifton Hill, Vic.), 200 M dNTPs (Roche, Melbourne, Australia) 1 M primers (16SIGF: 5 CGG CGT GGA TGA GGC AT 3 and 16SIGR: 5 TCA GTC CCA GTG TTG G 3 ), 1 U Qiagen HotStarTaq polymerase (Qiagen) and 4 l of concentrated swab material. PCR cycling conditions consisted of an initial denaturation at 95 ◦ C for 15 min, followed by 45 cycles of denaturation at 94 ◦ C for 30 s, primer annealing at 51 ◦ C for 30 s and extension at 72 ◦ C for
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45 s with a final extension for 5 min at 72 ◦ C. Following amplification, the products were separated by agarose gel electrophoresis and visualized by ethidium bromide staining. 2.4. PCR controls To ensure that the PCR products were not the result of contamination, one negative control (ddH2 O as template) was included for every three test specimens. Two positive controls of cell-cultured Chlamydiales were included with each PCR batch to confirm that amplification had occurred. 2.5. Isolate genotyping 16S rDNA PCR products intended for sequencing were extracted using Qiagen QIAquick gel extraction kit (Qiagen) as per the manufacturers’ protocol. Sequencing of the purified products to determine genotypes was performed using the dideoxynucleotide terminator method (Bodetti et al., 2002; Sanger et al., 1977). 2.6. Antigen detection Ten microlitres of concentrated swab material was spotted onto glass microscope slides and fixed in methanol. Slides were stained using a Chlamydiaceae family-specific antilipopolysaccharide (anti-LPS) monoclonal antibody (Cellabs, Sydney, Australia) according to the manufacturer’s instructions. The presence or absence of characteristic apple-greenfluorescing chlamydial particles was determined by examining duplicate wells by confocal microscopy at a magnification of 960× (TCS 4D; Leica, Leitz, Germany). 2.7. Cell culture isolation Confluent HEp-2 cell monolayers grown on glass coverslips in shell vials were used to isolate viable chlamydiales. Infection of the monolayers involved inoculation with 100 l of concentrated swab material in 1 ml of Dulbecco’s minimal essential medium (DMEM; InVitrogen, Melbourne, Australia) an initial 1 h centrifugation at 800×g. DMEM containing cycloheximide (1 mg/ml; Sigma, Melbourne, Australia) replaced the inoculum after 2 h incubation at 37 ◦ C 5% CO2 . Cultures were incubated for 9 days with repeat centrifugations on days 4–7 post-infection and fresh media on days 4 and 7. Up to three blind passages were performed prior to staining with the previously mentioned anti-LPS antibody to examine growth.
3. Results 3.1. Nucleic acid detection The 16S rDNA PCR assay detected chlamydial DNA in a total of 67% (62/95) of the animals. The individual animal species positivity was quite high for all five hosts, ranging
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Table 1 Detection of Chlamydiales in a range of Australian mammals using a 16S rDNA PCR assay Host
Number of animals tested
16S rRNA PCR-positive
Clinical disease Disease sitea
16S rRNA PCR-positiveb
Greater Glider (P. volans)
27
20/27 (74%)
Mountain Brushtail Possum (T. caninus) Western Barred Bandicoot (P. bouganville) Greater Bilby (M. lagotis) Gilberts’ Potoroo (P. gilbertii)
12
6/12 (50%)
Ocular (3/27) Ocular (3/3) None
Urogenital (8/27) Urogenital (4/8) N/A
26
19/26 (73%)
21 9
11/21 (52%) 4/9 (44%)
Ocular (13/26) Respiratory (1/26) None None
Ocular (13/13) Respiratory (0/1) N/A N/A
Total
95
62/95 (64%)
Ocular (16/95) Urogenital (8/95) Respiratory (1/95)
Ocular (16/16) Urogenital (4/8) Respiratory (0/1)
a b
Signs of clinical disease present (number with disease/total number of animals). Number of PCR-positive results in animals with clinical disease.
from 44% (potoroos) to 74% (bandicoots) (Table 1). Clinical signs of disease in the form of conjunctivitis or urogenital tract disease symptoms were present in 23% (22/95) of the total animals examined, with conjunctivitis the most common sign observed (73%, 16/22). A high proportion (32%; 20/62) of the PCR-positive animals had signs of clinical disease, compared to only 6% (2/33) of the PCR negative animals. Of the 95 animals tested, the conjunctiva was the site most commonly found to be PCR-positive and at this site, PCR positivity correlated strongly with clinical disease. Forty-five percentage (42/95) of animals were positive by PCR at the ocular site, including 100% of those with clinical ocular disease. This contrasted quite strongly with the urogenital site, where a significant portion of the animals were PCR-positive (36%) but only half of the eight animals with clinical signs of urogenital disease (crusty cloaca) were PCR-positive. 3.2. Speciation by 16S rDNA genotyping: Eighteen 16S rDNA positive PCR products were selected for direct sequencing. Sequencing identified similarities to several chlamydiales isolates/species currently listed in GenBank, including C. pecorum (4/18), endosymbiont of Acanthamoebae (2/18), Parachlamydia sp. Hall’s coccus (1/18) and Waddlia chondrophila (1/18). The most common identification however was of novel uncultured Chlamydiales (10/18) (Table 2; Fig. 1). 3.3. Antigen detection Twenty-nine 16S rDNA PCR-positive swab samples from 22 animals were selected for antigen detection by direct immunofluorescence. Thirteen (45%) of these swab samples showed characteristic fluorescence of spherical particles of approximately 0.3–2.0 M (data
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Fig. 1. Alignment of 183 bp of the 16S rRNA gene sequences determined in this study (underlined) with previously determined sequence types; N. hartmanellae (GenBank AF177275); C. pecorum bird isolate (GenBank E17343); uncultured Chlamydiales isolates CRG3, CRG23, CRG45 (GenBank AF097186, AY013399, AY013421). Possum and WBB Parachlamydia: sequences isolated from a Mountain Brushtail Possum and Western Barred Bandicoot, respectively, with similarity to sequence from an isolate of Parachlamydia acanthamoebae; Possum, Glider, WBB C. pecorum: from a bird isolate of C. pecorum; Possum, Glider, Bilby and WBB Uncultured: sequences isolated from a Mountain Brushtail Possum, Greater Glider, Greater Bilby and two Western Barred Bandicoots, respectively, with similarity to sequence from uncultured Chlamydiales.
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Fig. 1. (Continued ).
not shown). Structures consistent with chlamydiales inclusions of approximately 5–9 M diameter were also observed (Fig. 2a). 3.4. Cell culture isolation Positive cell culture isolates were obtained from six of the 16S rDNA PCR-positive and LPS antigen-positive swab samples. Chlamydia-specific antigen staining of the cultures Table 2 Genotypes of chlamydiales PCR-positives in Australian mammals Host
Number of PCR-positives genotyped
Swab site
Species similarity by 16S rRNA sequencing
Evidence of clinical disease
Greater Glider (P. volans)
2
Mountain Brushtail Possum (T. caninus)
4
Greater Bilby (M. lagotis)
3
Western Barred Bandicoot (P. bouganville)
8
Gilberts’ Potoroo (P. gilbertii)
1
Ocular Urogenital Ocular Ocular Urogenital Urogenital Ocular Ocular Urogenital Throat Throat Throat Throat Ocular Ocular Ocular Urogenital Urogenital
Uncultured Chlamydiales C. pecorum C. pecorum Uncultured Chlamydiales Uncultured Chlamydiales Endosymbiont of Acanthamoebae Uncultured Chlamydiales Parachlamydia sp. Halls’ coccus Uncultured Chlamydiales C. pecorum Uncultured Chlamydiales Uncultured Chlamydiales Uncultured Chlamydiales Endosymbiont of Acanthamoebae Uncultured Chlamydiales Uncultured Chlamydiales C. pecorum W. chondrophila
No Ocular No No No No No No No No No No Ocular Ocular Ocular Ocular Ocular No
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Fig. 2. Fluorescent antibody staining (FITC-conjugated Chlamydia-specific lipopolysaccharide monoclonal antibody) of urogenital swab from a Mountain Brushtail Possum (a) and HEp-2 cell culture from a urogenital swab from a Western Barred Bandicoot (b). Arrows indicate positive staining chlamydial inclusions. Magnification 960×.
showed inclusions of approximately 6–13 M in diameter and particles of approximately 0.4–1.7 M (Fig. 2b). While isolation was successful for these six samples, the level of infection observed was very low, on average 5–15% of HEp-2 cells and despite several attempts, only one isolate was successfully subcultured (to five passages).
4. Discussion Recent studies have identified Chlamydiales organisms in a number of new hosts (Jacobson and Telford, 1990; Kahane et al., 1998; Berger et al., 1999; Crespo et al., 1999; Fritsche et al., 2000; Horn and Wagner, 2001). In our present study we analysed ocular, respiratory and urogenital swabs from five different Australian native marsupials (95 animals) by 16S rDNA PCR and have identified five new chlamydial hosts and nine new chlamydiales types. Direct sequencing of the Chlamydiales signature sequence enabled identification of the infecting strain. Alignment of the sequences obtained showed similarities of 85–99% to several existing chlamydiales species, including four strains of C. pecorum (94–98% similarity to bird/cattle/sheep strains) and a number of completely new, uncultured Chlamydiales strains (Table 2). In two of the species sampled, bandicoots and gliders, a strong association was observed between ocular infection and disease. Chlamydiales was detected most commonly at the ocular site for all the hosts, although clinical signs of ocular disease, specifically conjunctivitis, were observed only in the bandicoots and gliders (30% of 53 animals). All of these 16 animals with clinical disease tested positive for the presence of chlamydial DNA in the ocular swab. By comparison only half of the animals with urogenital tract disease were
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Chlamydiales PCR-positive, although it is difficult to draw conclusions from only eight animals. Development of sensitive, broad-spectrum nucleic acid detection methods has enabled much easier detection of chlamydial DNA sequences from a wide range of sources (Corsaro et al., 2001; Michel et al., 2001; Fritsche et al., 2000). While previous studies have identified novel Chlamydiales DNA sequences, protein and antigen detection and isolation approaches have not previously been attempted. We confirmed a significant proportion of our 16S rDNA PCR-positive results by immunofluorescent antibody staining of chlamydial antigens, and also by in vitro isolation. Antigen staining of the swab samples identified characteristic Chlamydiales particles in 45% (13/29) of the PCR-positive samples. While in vitro isolation was difficult and the maintenance of the isolates proved equally difficult, we were nevertheless able to obtain some primary cell culture isolations for six nucleic acid and antigen-positive specimens. The organisms observed in both the direct swab and cell-cultured materials were found both intracellularly and also free from host cells. Inclusion-like structures were identified in most of the cell culture isolates and also in 2 of the 29 direct swab samples. The lack of inclusion-like structures in most of the direct swab samples is likely due to the fact that the swabs were frozen dry, in comparison with the two inclusion-positive specimens which were placed in SPG following collection, and then frozen (thawing may then have disrupted any inclusions). The lack of LPS staining in 55% of the PCR-positive swab samples may be due either to (a) the fact that no antigen or whole Chlamydiales were present in these samples or (b) the absence of characteristic chlamydial LPS, as has been reported for Parachlamydiaceae (Everett et al., 1999). Over the past 5 years the use of 16S rDNA PCR and direct sequencing has led to the discovery of many new genotypes of uncultured Chlamydiales in an increasing range of hosts. This has included the identification of novel Chlamydiales in humans (Kahane et al., 1998; Ossewaarde and Meijer, 1999), as well as lineages previously thought to be human-restricted (e.g. C. pneumoniae) in non-human hosts (Berger et al., 1999; Reed et al., 2000; Bodetti et al., 2002). Our work has identified further novel Chlamydiales organisms in additional hosts. What remains uncertain at this time is whether: (a) these infections only occur at low levels in these hosts, (b) these new Chlamydiales can cause disease in animal and human hosts, and (c) whether these strains are host restricted or are capable of cross infecting humans and animals, if given suitable conditions. These powerful and yet difficult to verify molecular techniques are challenging previous ideas of microorganism diversity and host restriction for these important human and animal pathogens.
Acknowledgements We thank Tony Friend and Neil Thomas, Department of Conservation and Land Management, Denham, Australia and June Butcher and Stephanie Hill, Kanyana Wildlife Rehabilitation Centre Inc. for invaluable assistance with the collection of specimens. We thank Felicity Horne and Christina Theodoropoulos for assistance with confocal microscopy. This work was partly supported by the Australian Koala Foundation and the Ruth Oulton Whyte Bequest fund.
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