Chlamydia psittaci infections: a review with emphasis on avian chlamydiosis

veterinary microbiology ELSEVIER

Veterinary

Microbiology

45 ( 1995) 93-l 19

Review

Chlamydia psittaci infections: a review with emphasis on avian chlamydiosis D. Vanrompay

*, R. Ducatelle, F. Haesebrouck

Department of Avian Medicine and Pathology and Laboratory for Veterinary Bacteriology and Mycology, Faculty of Veterinary Medicine, R.U.G., Salisburylaan 133 B-9820 Merelbeke, Belgium Received 2 May 1994; accepted

11 January

1995

Abstract In the first part of this article the general characteristics of Chlamydia psittaci namely the history, taxonomy, morphology, reproductive cycle, metabolism and genetics are reviewed. For the taxonomy in particular, a considerable amount of new information has become available in recent years, following the application of monoclonal antibodies and restriction enzymes. Using these techniques isolates of Chlamydia psittuci from birds have been subdivided in different serovars, a number of isolates have been classified in a new species (Chlamydia pecorum) and isolates from animals have been classified as Chlamydia trachomatis. In the second part of the article, the current knowledge on avian chlamydiosis is summarized. Emphasis is put on clinical signs, lesions, pathogenesis, epizootiology, immunity, diagnosis, prevention and treatment. Also the public health considerations are reviewed. It is concluded that the diagnosis of avian chlamydiosis is laborious and that there is still a need for more accurate, simple and rapid diagnostic tools, both for antigen and antibody detection in various species of birds. Keywords: Chlamydia psittaci; Avian; Public health

1. General characteristics

of Chlamydiu psittaci

1.1. History In 1893, in Paris,

transmission

of an infectious

agent from parrots

to humans,

causing

flu-like symptoms was reported. The disease was named psittacosis after the Latin word for parrot,psittucus (Morange, 1895). Nocard isolated a bacterium belonging to the Salmonella group

which

* Corresponding

was generally

regarded

author. Tel 0032-09-2647443

as the causative

agent

Fax 0032-09-2647494.

0378-l 135/95/$09.50 0 1995 Elsevier Science B.V. All rights reserved SSD10378-1135(95)00033-X

of psittacosis.

Salmonella

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psittacosis described by Nocard is now known to have been Salmonella typhimurium, a chance contaminant (Schachter and Dawson, 1978). Investigation of chlamydiae actually started in 1907 by Halberstaedter and von Prowazek, who found characteristic intracytoplasmic micro-organisms within a vacuole in cells of conjunctival scrapings from human patients with trachoma, a chronic infectious disease of the conjunctiva and the cornea (Halberstaedter and von Prowazek, 1907). They thought these “mantled organisms” to be protozoa and conferred the name Chlamydozoaceae after the Grecian word “chlamys” which means mantle. Thus the term chlamydia is actually grounded on a basic misconception. At that time the relationship to psittacosis was not understood. During the winter of 1929-1930, in Europe and the United States, a pandemic of psittacosis in humans occurred and stimulated the introduction of cultural methods for studying chlamydiae. The causative agent of psittacosis was isolated from humans and affected birds and the outbreak was traced to green Amazon parrots imported from South America (Coles, 1930). Also in 1930, a micro-organism resembling the one causing psittacosis, namely the causative agent of lymphogranuloma venereum (LGV) in humans was isolated (HellerStrom and Wassen, 1930). Both micro-organisms were classified as viruses belonging to the Psittacosis-lymphogranuloma venereum group. Until then, avian psittacosis was believed to occur only in psittacine birds. However, in 1932, Meyer and Eddie ( 1932) reported on a case of human psittacosis due to transmission from diseased chickens. In 1938, Haagen and Mauer ( 1938) identified the psittacosis agent in a fulmar petrel. In 1940, domestic pigeons were found to be infected with the psittacosis micro-organism (Pinkerton and Swank, 1940). Wolins ( 1948) reported on a number of cases in domestic ducks, In non-psittacine birds, the disease was called ornithosis. The importance of poultry, particularly turkeys and ducks as sources of human infection became evident in the 1950’s, when important outbreaks in humans due to contact with infected poultry occurred (Meyer and Eddie, 1953; McCulloh, 1955; Andrews et al., 1981; Dickinson et al., 1957: Graber and Pomeroy, 1958). In 1957, the causative organism of trachoma, the trachoma inclusion conjunctivitis (TRIC) agent, was finally isolated. The relation to the “viruses” of the psittacosis-lymphogranuloma group was obvious and the trachoma agent was added to the psittacosislymphogranuloma venereum group (T’ang et al., 1957). In 1960, Meyer ( 1960) suggested the name Bedsonia for all the members of the psittacosis-lymphogranuloma venereum-trachoma group. The name Bedsonia referred to Sir Samuel Bedson, who first described the developmental cycle of the psittacosis-lymphogranuloma venereum-trachoma agents (Bedson and Gostling, 1954). In 1965, Gordon and Quan ( 1965) described a tissue culture method for isolation of agents causing trachoma. This technical achievement gave the investigators more capability to focus on microbiology, epizootiology, serology, pathogenesis, therapy and control. Until then chlamydiae were classified as viruses, but in 1966 it became clear that they really were bacteria (Moulder, 1966). Page ( 1966) then proposed that all organisms of the psittacosislymphogranuloma venereum-trachomagroup be gathered together in one genus, Chlamydia. This proposal was supported by the Taxonomy Committee of the American Society for Microbiology.

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1.2. Taxonomy Chlamydiae are obligate intracellular Gram-negative bacteria. There are two main reasons why the chlamydiae are not longer classified as viruses; they possess both DNA and RNA and they have a unique developmental cycle completely different from the viral replication mechanism (Moulder, 1966). For some time chlamydiae were considered among the Rickettsia but have been differentiated from these organisms because they lack a system for electron transport, have no cytochromes and cannot synthesise ATP and GTP. Chlamydiae are classified as Gram-negative bacteria because they possess a number of properties that are highly characteristic to bacteria; a) division by binary fission, b) cell walls comparable in structure to those of Gram-negative bacteria, c) DNA-containing nucleoids with no membranes between the nucleoid and the cytoplasm, d) the presence of ribosomes with antibiotic susceptibilities characteristic for prokaryotic ribosomes (Moulder, 1966). However, Chlamydiae differ from other Gram-negative bacteria because of their unique morphology and developmental cycle and because of their common group antigens. Because of these unique characteristics they are placed in a separate order, the Chlamydiales, which contains the family Chlamydiaceae and one genus Chlamydia, with three species, C. trachomatis, C. psittaci and C. pneumoniae. (Storz and Page, 1971; Campbell et al., 1987; Grayston et al., 1989). Recently a fourth species, C. pecorum, has been proposed (Fukushi and Hirai, 1992). C. trachomatis, infects primarily oculogenital epithelium. The species can be divided into 3 biovars. The mouse pneumonitis biovar is antigenically distinct from the more closely related trachoma and LGV biovars. The trachoma biovar contains 12 serovars (A, B, Ba and C through K) and the LGV biovar contains 3 serovars (Ll, L2 and L3), (Wang et al., 1985). C. pneumoniae is a common causative agent of respiratory infections world-wide. The first C. pneumoniae isolate, TW-183 was isolated in 1965 from the eye of a child in Taiwan. The first pharyngeal isolate, AR-39 was obtained from a university student with pharyngitis in Seattle, in 1983. Both strains were then called TWAR (TW-183 from Taiwan and AR39 from acute respiratory). Only one serovar had been identified by Grayston et al. in 1989, but more recently evidence was provided of several new serovars of C. pneumoniae (Black et al., 1991). C. psittaci strains have been isolated from a wide range of avian and mammalian hosts, including man. C. psittaci infections have also been reported in the bay scallop (Leibovitz, 1989), fish (Wolke et al., 1970), frogs (Xenopus Lueuis) ( Newcomer et al., 1982), the puff adder (Jacobson et al., 1989) and in Moorish tortoises (Vanrompay et al., 1994a). Strains from mammals have been divided into 9 immunotypes (Perez-Martinez and Storz, 1985) using polyclonal sera. Avian strains are classified into 6 serovars (A to F) using serovar-specific monoclonal antibodies ( Andersen, 1991) . C. pecorum, proposed as the fourth chlamydial species, causes pneumonia, polyarthritis, encephalomyelitis and diarrhoea in cattle and sheep (Fukushi and Hirai, 1992). Species identification can be carried out using iodine staining of inclusions, susceptibility of growth to sulfa drugs, ultrastructural morphology of the elementary bodies, serology and deoxyribonucleic acid (DNA) analysis (Chi et al., 1987; Kaltenboeck et al., 1993).

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Recently, DNA sequences coding for 8 1% of the ompA gene from 24 chlamydial strains, representing all chlamydial species, were determined (Kaltenboeck et al., 1993). The ompA sequences were segregated into 23 different ompA alleles and phylogenetic relationships among them were inferred. A single phylogram was produced which was rooted at the branch between two major clusters. One cluster included all C. trachomatis ompA alleles (trachoma group). The second cluster was composed of three major groups of ompA alleles: psittacosis group (alleles MN, 6BC, A22/M, B577, LW508, PEPN and GPIC), pneumonia group ( C. pneumoniae AR388 with the allele KOALA), and polyarthritis group (ruminant and porcine chlamydial alleles LW6 13,66P130, L7 1 and 17 10s with propensity for polyarthritis) . Consistent with the OMPA phylogeny, the porcine isolate S45, previously considered to be C. psittuci, was identified as C. trachomatis. The koala isolates, previously considered to be C. psittaci, were identified as C. pneumoniae. A similar finding was made for the equine isolate N16 (Yates et al., 1992). The phylogenetic tree constructed by Kaltenboeck et al., ( 1993) has revealed that 6 of the 9 immunotypes described by PCrezMartinez and Storz ( 1985) as C. psittaci actually belong to the species C. pecorum. 1.3. Morphology C. psittuci has an extracellular infectious form, the elementary body (EB) and an intracellular replicating form, the reticulate body (RB) , The EB is a small, electron dense, spherical body, approximately 0.2 to 0.3 pm in diameter. The RB, the intracellular metabolically active form is also spherical but larger than the EB and measuring approximately 0.6 to 0.8 pm in diameter. As in gram-negative bacteria, chlamydiae are surrounded by a trilaminar cytoplasmic membrane and a trilaminar outer membrane. The outer membrane of the EB has been investigated intensively. The outer membrane is composed of phospholipids, lipids, lipopolysaccharides and proteins but, unlike other gram-negative bacteria, muramic acid has not been detected in chlamydia (Barbour et al., 1982). In gram-negative bacteria an important fraction of the cell wall is insoluble in ionic detergents such as sarkosyl. This fraction is composed of peptidoglycan with covalently bound lipoprotein. A fraction of the chlamydial cell wall is also insoluble in sarkosyl even though no peptidoglycan has been detected. This fraction is called “chlamydia outer membrane complex” (COMC). The COMC is composed of the major outer membrane protein (MOMP) and some minor proteins. The MOMP is a cysteine-rich protein, which represents approximately 60% of the weight of the outer membrane. The MOMP has a molecular weight of 40 kDa (C. psittuci 6BC strain). Although no direct evidence has been presented, the MOMP probably is cross-linked to extraordinarily cysteine-rich outer membrane proteins of 60 kDa (C. psittuci meningopneumonitis strain) or 59 and 62 kDa (C. psittaci 6BC strain) and 12 kDa (C. psittaci 6BC strain) or 15 kDa (C. psittuci meningopneumonitis strain). The MOMP appears to be important in maintaining the structural rigidity of the elementary body, acting as an adhesin and having pore-forming capabilities. The MOMP is also important in determining genus, species and serovar specificity (Caldwell et al., 1981). Matsumoto et al. ( 1976) and Stokes ( 1978) described the presence of hemispheric projections on the surface of EB’s and spikelike projections on the surface of intermediate

D. Vanrompay et al. /Veterinary Microbiology 4.5 (1995) 93-l 19 0

97

elementary body (EB) reticulate body (RB) host cell

\ chment and cell entry

4

release of new infectious

EB to RB

/

l2l.l DNA, RNA and protein synthesis has started.

multiplication by binairy fission /

2eh

Fig. 1. The developmental

inclusion is formed

cycle of chlamydiae.

forms of C. psittuci. Nichols et al. ( 1985)) observed surface projections on C. truchomutis. These projections may be involved in the attachment mechanism or in transport of nutrients, but their function is still unknown. The outer membrane of chlamydiae contains an antigenically important lipopolysaccharide (LPS) with a molecular weight of 10 kd. The chlamydial LPS is chemically and serologically related to the LPS of enterobacterial rough mutants of the Re chemotype. It contains several antigenic determinants cross-reacting with LPS of enterobacterial Re mutants and Acbzetobacter culcoaceticus or with the lipid A component. The chlamydial LPS contains in its saccharide portion, a trisaccharide of 3-deoxy-D-manno-2-octulosonic acid (Kdo) of the sequence oKdo (2-8) -nKdo- (2-4) -cyKdo (2). This structure represents a genus-specific, periodate-sensitive epitope (Nurminen et al., 1983; Caldwell and Hitchcock, 1984; Nurminen et al., 1984; Nurminen et al., 1985; Brade et al., 1987; Lukacova et al., 1994). 1.4. Developmental

cycle

During the reproductive cycle of chlamydiae, as shown in Fig. 1, at least three distinct morphological forms are present; the reticulate body (0.5-2.0 pm), the intermediate body (0.3-l .O pm) and the elementary body (0.2-0.3 pm). The reproductive cycle starts with

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the attachment of the elementary bodies to the host cell membrane, followed by intemalisation in the host cell. In the host cell chlamydiae are found within vacuoles derived from invagination of the host cell membrane during intemalisation. The infectious elementary body differentiates into the non-infectious reticulate body whereafter binary fission starts. After multiplication, chlamydiae are reorganised, through the intermediate form, into new elementary bodies which are released from the host cell (Bedson and Bland, 1932; Litwin et al., 1960). Chlamydiae need to enter cells to propagate, yet, how chlamydiae attach to and enter eukaryotic cells remains enigmatic. C. psittuci seems to enter the cell by a process resembling receptor-mediated endocytosis, although some studies suggest that chlamydiae enter cells by a microfilament-dependent zipper mechanism. It is likely that chlamydiae have more than one mechanism of cell entry (Ward and Murray, 1984; Hodinka and Wyrick, 1986; Hodinka et al., 1988). A solubilized C. psittuci membrane protein with a molecular weight in the range of 17 to 19 kilodaltons has been shown to bind to susceptible host cell membranes (Hackstadt, 1986). Evidence was presented that this protein is indeed a “sticky” protein (Barry et al., 1992; Perara et al., 1992). Recently, Stephens ( 1992) described a mechanism of interaction between chlamydiae and eukaryotic cells in which a eukaryotic-like glycosaminoglycan (GAG) on the surface of chlamydia mediates attachment. Chlamydiae are believed to synthesise a molecular mimic of heparan sulphate and use this for attachment to eukaryotic cells by binding heparan sulphate receptors. The chlamydial particle enters the cell within a phagosome and stays inside this phagosome through its entire life cycle. Phagosome-lysosome fusion does not take place (Friis, 1972). Zeichner ( 1983) identified a 70 kilodalton protein on the surface of vesicles containing infectious Ebs, that might play a role in preventing fusion of the phagosome and lysosome. The origin and the function of the 70 kDa protein remains unknown. The mechanism of release of chlamydiae is not clear. Chlamydial particles may be released after cell lysis. In vivo, infected intact cells can be sloughed off (Doughri et al., 1972; Soloff et al., 1985). Intact inclusions are occasionally found free in human conjunctival smears. Todd and Caldwell ( 1985) have presented morphological evidence of inclusion release without apparent cell lysis. 1.5. Metabolism

As the elementary body (EB) is intemalised, alteration of the EB cell wall occurs. These alterations primarily involve reduction of disulfide bond cross-linking among the major outer membrane proteins (Newhall and Jones, 1983; Hatch et al., 1984). The reticulate bodies (RBs) synthesise their own DNA, RNA and proteins, but they cannot complete the pentose cycle and do not utilise pyruvate by way of the tricarboxylic acid cycle (McClarty and Fan, 1993; McClarty and Qin, 1993). They can, however, catabolise pyruvic, aspartic, and glutamic acids, generating COP and 2- and 4-carbon residues (Moulder, 1969). The RB cannot generate high-energy phosphate bonds via the metabolism of various sugars and amino acids (Weiss, 1965). Chlamydiae depend on eukaryotic cells for their energy and therefore have been called ‘energy parasites’ (Moulder, 1969). Mitochondria are often found in juxta-position to the C. psittuci inclusion (Matsumoto et al., 1991) .

D. Vanrompay et al. /Veterinary

Microbiology 45 (1995) 93-l 19

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Hatch et al. ( 1982) describe the mechanism of Rbs parasitizing mitochondrial adenosine triphosphate ( ATP) via a chlamydial ATP-adenosine diphosphate ( ADP) translocase. The ATP is broken down to ADP by specific RB ATPase, and the resultant proton motive force helps drive the transport of nutrients. Inside the host cell, chlamydial Rbs perform independent nucleic-acid synthesis, protein synthesis and amino acid and pyruvate metabolism via the Krebs cycle, they have their own lipid metabolism and they metabolise exogenously supplied substrates optimally at neutral pH (Hatch et al., 1982; Moulder, 1985).

1.6. Genetics

Using pulse-field-gel electrophoresis, the genome size of Chlamydia psittaci and Chiamydia trachomatis was estimated to be 2,650 and 1,450, respectively (Frutos et al., 1989; Birkelund and Stephens, 1992). The presence of ribosomal RNA (rRNA) in the EB of C. psittaci was the first indication that the chlamydiae may differ from viruses (Tamura and Higashi, 1963). The first analysis of the RNA in the C. psittaci EB (meningopneumonitis strain) was carried out by Tamura and Iwanaga ( 1965). Three RNA species with sedimentation coefficients of 21S, 16s and 4S were found. The 4s RNA constituted most of the agent’s RNA. The base composition of the total EB-RNA was reported to be; A (25.5%), U (22.8%), G (32.3%) and C ( 19.2%). The sequence of the 16s rRNA gene was compared to that of other eubacteria. The closest relationship was seen with the planctomyces, a group of budding eubacteria. A common feature shared by chlamydiae and planctomyces, but not by other eubacteria, is the lack of a peptidoglycan layer in the outer cel1 wall (Weisburg et al., 1986). All 15 serovars of C. trachomatis contain a small plasmid of 4.4 X lo6 daltons (7.5 kb) (Lovett et al., 1980; Hyypil et al., 1984; Joseph et al., 1986; Palmer and Falkow, 1986) while in all avian C. psittaci strains, a 7.9 kb plasmid has been found. The smaller size of the 7.5 plasmid and the single restriction site for EcoRZ distinguishes this plasmid from the avian plasmid (McClenaghan et al., 1984). The avian plasmid is identical irrespective of avian host. A 5.9 kb fragment of the avian plasmid was cloned, mapped and used to screen a range of chlamydial strains. Hybridizing DNA was absent from ovine abortion and an arthritis isolate and also from the Cal 10 strain, but related sequences were detected in C. psittaci strains of feline pneumonitis, guinea-pig inclusion conjunctivitis, ovine conjunctivitis and C. trachomatis serovar L2 (McClenaghan et al., 1988). The feline, equine and guinea pig inclusion conjunctivitis strains each contain a distinct plasmid with different restriction endonuclease profiles from that of the avian strains ( McClenaghan et al., 1988; Herring et al., 1986; Lusher et al., 1992). The functions of the plasmid genes are not known, although some of the gene products are expressed during infection in cell culture. Evidence was presented that C. psittaci may have bacteriophages (Richmond et al., 1982). A polyhedral virus has been seen in RBs of a C. psittaci strain from a duck. This phage, Chpl, replicates in C. psittaci reticulate bodies. Chpl was found to be a 22 nm icosahedral virus with three structural polypeptides and a single stranded circular DNA of about 4800 bases. The complete DNA sequence of Chpl has been described (Storey et al., 1989).

100

D. Vanrompayet al. /Veterinary Microbiology 45 (1995) 93-119 EXTERIOR I

IV

II

LPS

PERIPLASMIC

SPACE

Fig. 2. Schema of MOMP in the outer membrane layer of the chIamydia1 cell wall (Baehr et al., 1988). Solid line, membrane-embedded peptide chain of MOMP (not drawn to scale with respect to thickness of membrane or size of VDs); pointed squares, residues comprising the VDs; open squares, conserved cysteines. The presence of lipopolysaccharide (LPS) structures (solid blocks) in chlamydiae is indicated above the outer membrane (OM) .

Monoclonal antibodies have been used to type isolates of chlamydiae (Wang et al., 1985; Newhall et al., 1986; Andersen, 1991; Vanrompay et al., 1993). These monoclonal antibodies recognise genus-, species- and serovar-specific regions on surface-exposed epitopes on the MOMP of chlamydiae (Fig. 2) (Stephens et al., 1988; Baehr et al., 1988). The MOMP gene (omp 1) contains five conserved- and four variable sequence regions (VS 1 to VS4). VS 1 to VS4 encode for the four variable protein domains (VDI to VDIV) (Baehr et al., 1988; Yuan et al., 1989) which protrude from the chlamydial membrane. Epitope mapping has shown that genus- and species-specific antigenic determinants are located within the conserved regions. However, species-specific antigenic determinants have also been found in the most conserved parts of VDIV. VDI and II contain the serovarspecific antigenic determinants. (Baehr et al., 1988; Conlan et al., 1988; Stephens et al., 1987; Stephens et al., 1988; Yuan et al., 1989; Lampe et al., 1990). For C. psinuci the complete nucleotide sequence of the ompl gene of two strains causing abortion in ewes (A22/M and S26), the 6BC strain isolated from a parakeet in California, the Guinea pig inclusion conjunctivitis (GPIC) strain 1 and the meningopneumonitis (Mn) strain Cal- 10 have been described (Pickett et al., 1988; Herring et al., 1989; Zhang et al., 1989; Everett et al., 1991). The MOMP gene has been intensively studied since MOMP antigens are important not only in serovar epidemiology, but also as important potential targets for vaccine development (Caldwell et al., 1987).

2. Diseases caused by C. pszltuci

Diseases caused by various strains of C. psittaci are listed in Table 1

D. Vanrompay et al. /Veterinary Microbiology 45 (1995) 93-l 19

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Table 1 Diseases caused by Chlamydia psittaci Disease or infection details

Affected host

References

( 1) Psittacosis (2) Psittacosis,

Humans Wild and domestic fowl Sheep

Hagan and Brunner, 1988 Grimes and Wyrick. 199 1

ornithosis

(3) Placentopathy foetopathy

and

(a) Ovine enzootic abortion

Stamp et al., 1950

(OEA) (b) Epizootic bovine abortion (c) Koala bears (d) Abortion in other domestic animals (4) Vesiculitis (5) Sporadic bovine encephalomyelitis (SBE) (6) Pneumonitis

(a) Seminal vesiculitis syndrome

(a) Porcine pneumonitis (b) Feline pneumonitis (c) Ovine pneumonitis (d) Bovine pneumonitis (e) Caprine pneumonitis ( f) Canine pneumonitis (g) Equine pneumonitis

(7) Hepatitis (8) Conjunctivitis

(9) Polyartbritis

Cattle

Hagan and Bruuner, Storz, 1971

Koala bears Pigs, goats, horses,

Canfield et al., 1991 Bliibel and Schliesser,

rabbits, mice Cattle, pigs

Hagan and Brunner, Storz et al., 1968

1988

Cattle, dogs

Hagan and Brunner,

1988

Pigs Cats, Humans? Sheep Cattle

Wittenbrink et al., 1991 McKercher, 1952 Schachter et al., 1969 Hagan and Brunner, 1988 Perez-Martinez and Storz, 1985 Rahman and Singh, 1990 Blijbel and Schliesser, 1985

Goats Dogs, budgerigars, humans? Horses

1988

1985

(h) Murine pneumonitis (i) Pneumonitis in tortoises (b) crocodiles (a) Feline conjunctivitis and keratoconjunctivitis (b) Guinea pig inclusion conjunctivitis (GPIC) (c) Hamster (d) Koala bears

Mice Moorish tortoises

Moorthy and Spradbrow, 1978 McChesney et al., 1982 Rodolakis and Souriau, 1992 Vanrompay et al., 1994a

Nile crocodiles Cats

Huchzermeyer et al., 1994 JCgou, 1991, Jongh, 1991

Guinea pig

Hagan and Brunner, 1988

Hamster Koala bear

(e) Bovine keratoconjunctivitis ( f) Ovine infectious keratoconjunctivitis (g) Crocodiles (a) Porcine (b) Bovine (c) Ovine (d) Eouine

Cattle

Hagan and Brunner, 1988 Canfield et al., 1991 Hirst et al., 1992 Storz, 1971

Sheep

Meagher et al., 1992

Nile crocodile Pigs Cattle Sheep Horses

Huchzermeyer et al., 1994 Hagan and Brunner, 1988 Hagan and Brunner, 1988 Hagan and Brunner, 1988 McChesney et al., 1974

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D. Vanrompay et al. /Veterinary Microbiology 45 (1995) 93-119

Table 1 (continued) Disease or infection details ( 10) Enteritis and diarrhea

( 11) Unknown disease condition

(a) Snowshoe hare (b) Muskrat (c) Bovine (d) Porcine Fish

Affected host

References

Snowshoe hares Muskrat Cattle Pigs Wolke et al., 1970

Blobel und Blobel und Doughri et Wittenbrink

African clawed frogs Puff adder Chameleon Bay scallop

Newcomer et al., 1982 Jacobson et al., 1989 Frye, 1991 Leibovitz, 1989

Schliesser, 1985 Schliesser, 1985 al., 1972 et al., 1991

3. Avian Chlamydiosis 3.1. Clinical signs

It is a commonly accepted view that individual bird species may be infected by C. psittaci strains which differ in virulence. This gives rise to different incubation times, signs and recovery. Typical signs of a clinically apparent infection with virulent C. psittaci include “pneumo-enteritis” with respiratory signs, mucopurulent nasal discharge, diarrhoea, polyuria and dullness (fluffed and inactive). Droppings may also be yellow, suggesting high concentrations of bile pigments (Jenkins, 1989). A complete clinical picture of chlamydiosis, is most frequently seen in Amazon parrots and macaws, in which central nervous system disturbances may be occasionally observed (Harrison, 1989). Cockatiels often have unilateral or bilateral conjunctivitis and keratoconjunctivitis. They often produce intermittent yellow urates. Budgerigars additionally often show sinusitis. In pigeons the disease may only become clinically apparent when concurrent infections are present. Signs are largely confined to the upper respiratory tract and there may be decreased flying performance. Chronically infected pigeons may show lameness, torticollis, opisthotonus, tremor and convulsions (Wages, 1987). The incidence of chlamydiosis in canaries and finches is low. Dullness, upper respiratory tract disease, conjunctivitis and wet droppings can be observed (Jenkins, 1989). Clinical signs in turkeys include ruffling of feathers, depression, anorexia, cachexia and mild diarrhoea with yellow droppings. Respiratory signs include ocular and nasal discharge, coughing and dyspnoea (Grimes and Wyrick, 199 1) . In ducks clinical signs may include head tremors, unsteady gait, conjunctivitis, serous to purulent nasal discharge, depression, recumbence and mortality ( et al., In ducks geese, temporary of equilibrium observed (Grimes Wyrick, 1991). are relatively to chlamydial and there few reports the isolation chlamydia from species. It mostly young are affected and Wyrick, although the of chlamydiosis 36-day old has been (Barr et 1986). Clinical in chickens blindness, weight and a increase in rate.

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3.2. Lesions Autopsy and histological findings are not characteristic enough to differentiate chlamydiosis from other systemic diseases. The severity of lesions found in birds with chlamydiosis is dependent upon a number of factors, including the virulence of the strain, the susceptibility of the host, the route of exposure and any concurrent disease (McDonald and Bayer, 198 1). Enlargement of the spleen is generally observed. The spleen is usually softer in consistency and may contain white necrotic foci or petechial hemorrhages on its surface. The liver is generally enlarged, friable, yellowish or green in colour, and small necrotic foci may be present on the capsular or cut surface. The air sac membranes may become thickened and cloudy in appearance and may be sometimes covered with a thick, yellowish, fibrinopurulent exudate. The lungs usually are congested and diffuse congestion is mostly present. The pericardial sac sometimes undergoes a marked inflammatory reaction which may be purulent, serous or fibrinous in nature. Congestion may be evident in the intestinal tract, especially on the serosal surface (Amstein and Meyer, 1969; Bankowski et al., 1977; McDonald and Bayer, 1981; Fudge, 1984; Wages, 1987; Grimes and Wyrick, 1991). Minimal histopathological lesions can occur in acute, fatal cases of chlamydiosis. The extent of the histological lesions depends on the virulence of the strain, the duration of the disease, host susceptibility and route of infection. Concurrent pathology, due to other infectious agents is common (Fudge, 1984). The spleen may show altered architecture associated with histiocytosis (reticular-endothelialcell hyperplasia) . Lymphoid hyperplasia and plasmocytosis may be present and areas of necrosis can be observed. In the liver, periportal heterophilic and mononuclear infiltrates are common. Very acute infections will often show multifocal, coagulative necrosis. More chronic infections show bile duct hyperplasia and histiocytosis. The increase in Kupffer cell activity correlates clinically with the monocytosis seen in more chronic infections. Hemosiderosis may be present. Chronic hepatic infections will show significant fibrosis and mononuclear infiltrates. Lesions in the air sacs are the result of an infiltration of mononuclear cells (especially macrophages) and heterophils. The air sacs may be thickened by a proliferation of the epithelium and connective tissue. Pulmonary pathology is often minimal; however, a mild pneumonitis may be present. Myocarditis can be diffuse and occasionally shows large areas of necrosis that are grossly visible. The intestines often show a plasmacytic, lymphocytic enteritis, which is associated clinically with white blood cells in the stool. Acute necrosis and mixed inflammatory infiltrates may be present in the kidney. Inflammatory adrenal and gonadal disease may be present as well. Brain lesions are rare. Examination of the bone marrow reveals a significant increase in the granulocytic series (McDonald and Bayer, 198 1; Fudge, 1984). 3.3. Pathogenesis The pathogenesis of avian chlamydiosis was first studied in turkeys (Page, 1959). Turkeys were infected by airborne and oral routes using the New Jersey (NJ) 1 isolate, which was recovered from the muscles of an acutely diseased turkey from an outbreak of ornithosis (Page, 1959). The pathogenesis was studied in turkeys kept in open houses and was performed with the techniques available at that time when chlamydiae could only be isolated in mice and embryonated eggs and when isolated strains could not be serotyped. The strain

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used by Page has recently been classified as an avian serovar D strain ( Andersen, 1991). Since 1959, studies on the pathogenesis of chlamydiosis in birds have been rare and incomplete, therefore only the study by Page ( 1959) will be mentioned here. In turkeys, the primary route of infection is by inhalation of contaminated dust from feathers and/or faecal material but transmission can also occur through ingestion of contaminated faeces (Page, 1959). Blood-sucking ectoparasites, which include arachnids, lice and simulid flies, have been shown to transmit chlamydiae in turkeys but probably act as mechanical vectors rather than biological vectors (Shewen, 1980). After inoculation of turkeys with an infectious aerosol, it was demonstrated that chlamydiae were present within four hours multiplying in the lungs, air sacs and pericardial sac. Within 48 hours, chlamydiae were present in the blood, spleen, liver and kidney. At 72 hours, chlamydiae were also found in the bone marrow, the testes or ovary and in the muscle. Furthermore, large numbers of chlamydiae were found at portals of exit, including the nasal turbinates and the cloaca (Page, 1959). Vertical transmission of chlamydiae was described in chickens, ducks, parakeets, sea gulls and snow geese (Meyer and Eddie, 1933; Illner, 1962; Lehnert, 1962; Wilt et al., 1972; Wittenbrink et al., 1993). 3.4. Epizootiology Chlamydiosis in birds is diagnosed world-wide through the demonstration of the microorganism in animals autopsied, in faeces or cloacal swabs or through the demonstration of anti-chlamydial antibodies. In Florida and South California, C. psittuci was isolated from 20 to 50% of post-mortemed pet birds (Schwartz and Frazer, 1982). In the Netherlands and in Germany, chlamydial antigen was demonstrated, by ELISA on cloaca1 samples, in 14 to 16% of locally bred Psittaciforms and in 4 to 5% of imported psittaciforms. No clinical signs had been observed in these birds by the owners (Janeczek and Gerbermann, 1988; Dorrestein and Wiegman, 1989). In Belgium, during 1991,32 out of 307 birds of various species were positive for chlamydiosis using a direct immunofluorescence test of cloacal, conjunctival and/or organ smears (Vanrompay et al., 1991). Chlamydiosis is most often diagnosed in psittacine birds, especially in parrots, parakeets and cockatoos. Several seroepizootiological studies have been carried out. In the U.K. 50% of the investigated racing pigeons showed antibodies against C. psittuci (Alexander et al., 1989). In Japan, 30.5% of the examined pigeon sera showed anti-chlamydia antibodies (Chiba et al., 1984). In the poultry raising industry, chlamydiosis is especially important in turkeys and ducks. In the USA epidemics of chlamydiosis in turkeys have been reported since 1950. These epidemics were economically devastating to producers because of carcass condemnation at slaughter, decrease in egg production, and/or the expense of antibiotic treatment to reduce mortality and allow marketing of poultry (Grimes and Wyrick, 199 1) . In Europe, there is still controversy as to the significance of chlamydiosis in turkeys. Chlamydiosis in ducks became apparent as a problem firstly in Eastern Europe but more recently outbreaks have also been reported in the U.K. (Chalmers, 1986). Captive birds as well as free living wild birds are important as reservoirs of C. psittuci (Brand, 1989). Diseased birds as well as subclinically infected birds, can shed chlamydiae and are therefore a potential threat to both human and animal health (Roberts and Grimes,

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1978; Wobeser and Brand, 1982; Franson, 1993).

1987; Brand, 1989; Gerbermann

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and Korbel,

3.5. Immunity Little is known about protective immunity against chlamydial infection in birds. Information on immunity is mostly obtained from studies in humans and mice and from studies with C. psittuci in non-human mammals. Immunity to chlamydial infection may be regarded as a “double-edged sword” since both protective and pathological immune responses can be elicited. In the majority of chlamydial infections, only relatively small proportions of cells at affected sites are found to be infected. Because each inclusion releases hundreds of viable chlamydiae and not all cells in the surrounding area are infected, there must be control mechanisms that limit infectivity. However, the mechanisms are not clear. Experimental studies of C. truchomatis infections have provided evidence that mucosal serovar-specific IgA antibodies play an important role in acquired resistance to reinfection (Murray et al., 1973; Watson et al., 1973; Barenfanger and MacDonald, 1974; Nichols et al., 1978). It is possible that neutralising antibodies play a role but the exact mechanism by which neutralising antibodies act is not known. Antibodies against MOMP have been shown to neutralise infectivity in vitro and in vivo (Caldwell and Perry, 1982; Peeling et al., 1984; Lucero and Kuo, 1985; Baehr et al., 1988; Byrne et al., 1993). Antibodies to the MOMP, however, do not interfere with chlamydial attachment, ingestion or inhibition of phagolysosomal fusion (Caldwell and Perry, 1982). It is possible that the MOMP antibodies may become cross-linked across MOMP molecules in the outer membrane so that the reorganisation of EBs to RBs is not permissible. Less is known about the neutralizing effect of the non-MOMP antibodies. Tumor necrosis factor (TNF) (Holtmann et al., 1990; Williams et al., 1990) and gamma interferon have been shown to have an inhibitory effect on chlamydiae. Gamma interferon inhibits chlamydiae infection in human macrophages and in mice (Rothermel et al., 1983; Byrne et al., 1986). The interferon appears to delay the developmental cycle so that the RBs persist for a longer period of time (Shemer and Sarov, 1985). This may result in persistent, inapparent infections and may also play a role in possible immunopathogenesis. Chlamydiae do not appear to survive well in polymorphonuclear leukocytes (PMN) (Yong et al., 1982). Chlamydia are rapidly intemalised by human PMNs. The majority are rendered non infectious within 1 hour but a small portion of chlamydial infectivity would seem to persist. The majority of chlamydial particles are found in PMN phagosomes where lysosomal fusion has occurred (Yong et al., 1986). The mechanism of killing by PMNs is not known, but it is likely that both oxygen-dependent and- independent mechanisms are involved. Chlamydiae can act as stimulators of B lymphocytes. Stimulation can be effected in mice that are LPS non-responders, suggesting something other that the genus-specific LPS antigenie determinant is responsible for stimulation of B lymphocytes (Levitt et al., 1986). Chlamydiae do not appear to grow well in monocytes. However, after monocytes have become activated to macrophages, C. psittuci and C. trachomatis LGV biovar infections are highly productive (Manor and Sarov, 1986). In macrophages phagolysosomal fusion

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is successfully prevented and it appears that, in infections in vivo, the macrophages become activated (Williams et al., 1984; Guagliardi et al., 1986). Leukocytes play an important role in resistance to infection and in clearance of primary infection. T-cell deficient mice do not produce significant levels of chlamydia-specific antibody (Williams et al., 1982). Lymphocyte transformation in vitro and delayed-type hypersensitivity reactions in vivo are also T-cell dependent (Williams et al., 1984; Rank et al., 1985). Recently T-cell antigenic determinants from MOMP have been described (Allen et al., 1990; Su et al., 1990). Natural killer (NK) cell activity appears to be increased in mice infected with the mouse pneumonitis agent (Williams et al., 1987). Antigen-dependent cytotoxic cell activity was not found using LGV-infected L cells (mice fibroblasts) (Pavia and Schachter, 1983). Infected cells may be killed by NK cells or perhaps even by cytokines. In conclusion, it seems that cell-mediated immune responses may be responsible for the clearance of initial infection, although the contribution of gamma interferon may actually result in persistent, inapparent infection. Antibody-mediated immunity appears more likely to be involved in resistance to reinfection. However, the current knowledge on control of chlamydial infections in man and other mammals can not necessarily be extrapolated to chlamydiosis in birds. Birds have their own special immune system. So, before studying the control of chlamydial infections in birds, there is still work to be done studying the immune system of different bird species.

3.6. Diagnosis

There is no easy way to diagnose chlamydiosis in birds. Although the history of the bird, clinical examination, radiology and haematology may contribute towards a tentative diagnosis, a confirmation can only be obtained from isolation and/or identification of the organism (McDonald and Bayer, 198 1; Fudge, 1984). As to methods for the detection of infected birds, the first to be considered should be the isolation of the organism from organs, faecal samples or swabs submitted to the laboratory. Avian C. psittaci strains are preferentially cultured in Vero, McCoy or Buffalo Green Monkey cell cultures ( Meissler and Krauss, 1980; Ahrens and Weingarten, 198 1; Vanrompay et al., 1991). The inclusions that develop following inoculation, centrifugation and incubation can be stained in various ways. The most commonly used stains are a&dine orange, Castaneda, Giemsa, Gimenez, Macchiavello, methylene blue, Stamp and immunofluorescence. The reliable detection of inclusions in cell culture confirms the unequivocal existence of viable chlamydiae. However, because chlamydiae are intermittently shed in the faeces, a single, one-time negative culture is not a reliable indication that the bird is not carrying a chlamydial infection. In addition, therapy with inhibitory antimicrobial drugs, 2 to 3 weeks before testing, can give false negative results (Grimes et al., 1987). Furthermore, contaminated, or improperly prepared transport medium, length of transportation time, and overheating, can give negative culture results even though the bird is infected. The success of chlamydia culture also depends on the type and age of the host cell used and on the quality control and expertise of the laboratory culturing the organisms (McDonald and Bayer, 1981; Fudge, 1984; Grimes, 1989; Vanrompay et al., 1991).

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For direct identification without culturing, the following methods can be used: electronmicroscopy (Grimes et al., 1987), cytological staining (Macchiavello, 1937; Stamp et al., 1950; Gimenez, 1964; Campbell, 1988)) immunofluorescence tests (Woods, 1986; Grimes et al., 1987, Palmer et al., 1988; Timms et al., 1988; Vanrompay et al., 1994b), enzymelinked immunosorbent assays (ELISA’s) (Gerbermann, 1989; Brown and Newman, 1990; Fudge and Connely, 1990; Grimes and Arizmendi, 1990; Kingston, 1992; Thiele et al., 1992; Vanrompay et al., 1994b), peroxidase anti-peroxidase tests (Moore et al., 1991)) immunoperoxidase tests (Tappe et al., 1989), polymerase chain reaction (PCR) (Hewinson et al., 1991) and DNA-spot hybridization (Timms et al., 1988). However, ideally chlamydial culture should still be performed even if these tests proved negative. Isolation in vitro indeed still is considered the reference test for antigen detection. In recent years, a number of diagnostic tests and reagents have become commercially available for direct identification of C. truchomaris in human specimens. Some reagents such as monoclonal antibodies against the group-specific chlamydial lipopolysaccharide antigen have been prepared and, in theory, should be suitable for detecting C. psittuci in avian samples. These tests offer the considerable advantage of speed over culture techniques. However, proper evaluation is always required before using these tests for the routine diagnosis of avian chlamydiosis in the laboratory. Some investigators claim that serology is useful in diagnosing avian chlamydiosis, especially at the flock level. However, as with other infections, in the very early stages, serological tests may be negative. Paired sera should be examined in an individual bird to look for at least four fold rising titers, and if only one serum sample is available, serology should be combined with isolation (Grimes, 1989). Different anti-chlamydia antibody detection methods have been described of which complement fixation (CF) (Fudge, 1989; Grimes, 1989)) latex agglutination (LA) (Moore et al., 1991) and ELISA (Schmeer, 1983; Ruppanner et al., 1984; Gerbermann, 1989; Hafez and Sting, 1993) are commonly used. The usefulness of the CF and LA is bird speciesdependent (Fudge, 1989; Grimes, 1989). The LA only detects IgM, which is generally indicative for a current chlamydial infection. However, many chlamydial infections may be are chronic or long-standing. The CF detects both IgM and IgG, however, a problem in the interpretation of the CF can be that, even after apparent cure, following treatment, birds may retain moderate to relatively high CF titers ( 1 / 128 to l/256) for some weeks to months depending on the original titre (Grimes, 1989). Therefore anti-chlamydia antibody detection tests are more suitable for epidemiological studies of avian chlamydiosis rather than for diagnosis of infection.

4. Prevention and treatment All attempts to induce immunity in birds through vaccination have failed so far. Possible reasons for failure include serovar or strain differences, ill-defined virulence variation and incorrect presentation of the antigens to the host. Recovery of chlamydia-infected birds requires a competent immune system to remove the organisms. Therefore, treatment of acutely infected birds requires not only administration of what is believed to be the thera-

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peutic dose of drug, but also supportive care, e.g., intravenous fluid therapy, heated environment and a diet supplemented with lactulose and vitamins (Harrison, 1989). Tetracyclines are the drugs of choice for treating avian chlamydiosis. In the Unites States, chlortetracycline (CTC) medicated feed is the recommended method of treatment. The goal of treatment is to maintain CTC blood concentrations above 1 pg/ml for the entire treatment period (Flammer, 1989). However, investigators in Germany were able to eliminate chlamydiosis from experimentally infected Amazon parrots by maintaining blood concentrations as low as 0.10 pg/ml (Gylstorff et al., 1984). The relationship between maintenance of a particular blood concentration and treatment efficacy are currently illdefined; however, until better pharmacological standards can be defined, maintenance of 1 pg/ml chlortetracycline remains the basis of comparison. Parakeets, budgerigars, finches, canaries and rice birds can be treated through the use of commercially available hulled millet impregnated with CTC (0.5 mg of drug/g of seed) (Keet Life, Hartz Mountain Products Corp, Harrison, NJ, USA) for a minimum of 30 days. For large, psittacine birds feed containing 1% CTC is recommended. Medicated pelleted feeds (5000 ppm CTC) are commercially available (Psittacin, Oberhausener Kraftfutterwerke, Germany; Psittavit, Claus Limburgerhof, Germany; Zeigler Brothers, P.O. Box 95, Gardners, PA, USA) or bird owners can prepare a cooked mash (Wachendorfer and Luthgen, 1974; Cooper, 1980; McDonald and Bayer, 1981) or nectar diet (Amstein and Meyer, 1969) and add CTC powder (Mix 66, Cyanamid) after cooking. This diet should be administered for at least 45 days. Although they can subsist on millet seed, lovebirds apparently do not develop adequate blood levels from millet impregnated with CTC. They must be fed according to directions for large psittacine birds (McDonald and Bayer, 198 1) . For psittacine birds, antibiotics are never administered through the drinking water because these birds do not drink enough to get sufficient medication. Pigeons can be fed with uncooked hen feed supplemented with CTC (5000 ppm) (Mix 66, Cyanamid). Medicated feed is to be administered for 45 days (Wachendijrfer and Luthgen, 1974; Wages, 1987). Medication through the drinking water is usually not effective in pigeons because of inadequate water consumption. Turkeys should be treated with CTC at a concentration of 400 g/ton of pelleted feed. The CTC medicated feed must be given for at least 2 weeks. CTC medicated feed should be replaced by non-medicated feed for 2 days prior to the birds being slaughtered for meat for human consumption (Grimes and Wyrick, 199 1) . Essentially the treatment for turkeys also applies to other fowl infected with C. psirtuci. Ducks can be treated with 750 g oxytetracycline/ton of feed for 3 weeks (Arzey et al., 1990). To reduce interference with CTC absorption, the calcium content of the diet has to be reduced to 0.7%. Although treatment with CTC medicated feed may be successful, significant problems may be associated with its use. These problems include poor acceptance (Flammer, 1989). Chlortetracycline mash is often poorly accepted because of its bitter taste. Furthermore, treatment with medicated pellets is not effective if the birds have not been converted to a pelleted diet prior to treatment. The stress of diet conversion during treatment may precipitate more losses than necessary (Harrison, 1989). In such cases, direct administration of tetracyclines via the oral, subcutaneous or intramuscular route should be considered. The use of intramuscular CTC injections (twice a day 200 mg/kg bw CTC in Sesamoil) in Amazon parrots has been described (Jacoby and Gylstorff, 1983). Oxytetracycline intramuscular administration at a dose of 50 mg/kg bw twice a day is clinically

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effective. However, because prolonged intramuscular injections of oxytetracycline can cause severe local tissue necrosis, injectable medication is not given for longer than one week (Flammeret al., 1990). If, after one week, the bird is not consuming adequate amounts of feed on its own then an oral tetracycline preparation should be given (McDonald and Bayer, 1981). Because of the risk of tissue necrosis it is better to use doxycycline (Vibramycin-IV, Pfizer) by intramuscular or subcutaneous injections instead of CTC or oxytetracycline. Doxycyclin is a semisynthetic tetracycline derivate. A number of characteristics observed in man, distinguish this drug from tetracycline (Leibovitz et al., 1972; Sande and Mandell, 1982). Doxycyclin is more lipophilic and greater tissue concentrations are achieved than with use of other tetracyclines. Serum t ,,2 is prolonged (22 hours versus 8 hours for tetracycline) and absorption from the gastrointestinal tract is rapid and virtually complete (95% versus 25 to 80% for other tetracyclines). Calcium has less effect on doxycyclin absorption from the intestines and doxycyclin can be taken with meals without appreciable reduction in absorptive efficiency. Orally administered doxycyclin has less adverse effect on autochthonous alimentary tract flora. Intramuscularly administered doxycyclin (Vibramycin-IV, Pfizer; Respidox, Kela) is also used for treating avian chlamydiosis, 75 to 100 mg/kg bw are injected into the pectoral muscles and dosage regimens requiring only 8 to 10 injections in a 45-day period have proved to be efficacious in a variety of psittacine species (Wachendbrfer et al., 1982; Gylstorff, 1984). Oral doxycyclin formulations are also available (Vibravet, Pfizer). In several psittacine species 25 to 50 mg/kg bw, when given once a day will maintain adequate plasma concentrations (Flammer, 1987). Doxycyclin must be given on an empty crop so it can be absorbed properly (Spencer, 1989). In humans, the quinolones have been demonstrated to possess antichlamydial activity. A quinolone approved for use in domestic animals, enrofloxacin, is now being used for treatment of avian chlamydiosis. In the United states an injectable as well as an oral enrofloxacin formulation is available. A water soluble formulation is available for birds in Europe (Baytril 1O%, Bayer AG). The pharmacokinetics of this drug were investigated in healthy African Grey parrots (Flammer, 1989). Further investigation is needed to determine pharmacokinetics in sick birds as well as in other species. Preliminary results indicate that treatment with enrofloxacin-medicated food for 3 weeks could be enough to eliminate chlamydiae from parakeets (Dorrestein, 1989). Groups of experimentally infected budgerigars and other psittacines were effectively treated for 14 days with medicated food containing enrofloxacine (Jung, 1992) at 250 ppm for budgerigars and 500 ppm for other psittacine birds. Complete elimination of chlamydiae from a quarantine group of 196 Senegal parrots was achieved only after substituting normal mixed food with medicated corn containing 1000 ppm enrofloxacin (Lindenstruth, 1992). A minimum blood level of 0.5 mg/L enrofloxacin for at least 14 days was considered necessary to control psittacosis (Jung, 1992; Lindenstruth, 1992). During treatment, frequent cleaning to eliminate infected dust and disinfecting of the aviary with quatemary ammonium products is beneficial in eliminating chlamydiae from the environment and preventing reinfection. However, there is growing evidence to suggest that some avian species may not be cured of chlamydiosis, no matter which medication is used.

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5. Public health considerations People usually contract the infection by inhaling an aerosol of the droppings of infected birds. Human psittacosis may occur sporadically or in outbreaks. Psittacine birds still appear to be the major source of infection but outbreaks have occurred where the sources were non-psittacine birds including commercially reared poultry (Irons et al., 1951; Strauss, 1967; Palmer et al., ,1988; De Schrijver, 1987; Yung and Grayson, 1988; Hedberg et al., 1989). Several groups of humans are particularly at risk of contracting psittacosis, such as., pet owners, veterinary surgeons and poultry workers (Grimes and Panigraphy, 1978; Edwards, 1981; Filstein et al., 1981). Psittacosis in man thus has to be considered as an occupational hazard. Clinical psittacosis in man varies in severity from mild, ‘flu’-like symptoms with shivering, fever, headache, anorexia, sore throat and photophobia (Schaffner et al., 1967; Gregg and Wehrle, 1972; Byrom et al., 1979; Eeckhout et al., 1986) to a more serious atypical pneumonia with dry cough and difficult or painful breathing (Ommeslag, 1987; Oldach et al., 1993). There may be diarrhoea early in the course of disease, with nausea and vomiting. Severe systemic illness involving endocarditis, myocarditis (Levison et al., 1971) and renal complications (Byrom et al., 1979; Jeffrey et al., 1992) have been reported but are less common. Encephalitis, meningitis and myelitis due to psittacosis have been described (Wieck and Heerklotz, 1972; &r-Locke and Mair, 1976; Reis et al., 1985; Shee, 1988; Williams and Sunderland, 1989). To prevent human cases of psittacosis, chlamydial infection has to be eliminated from contact birds, aviaries and pets. Following the 1929-1930 psittacosis pandemic, in many countries control measures are taken for all imported psittacine birds (Satalowich et al., 1993). In order to safeguard public health, birds are supposed to be quarantined for 45 days in the country of origin followed by 30 days of quarantine in the importing country. Some authors consider that the investigation of all imported birds, both psittacine and nonpsittacine, using an chlamydia antigen detection ELISA may constitute an important tool in the prevention of human psittacosis. A positive test result should then lead to obligatory antibiotic treatment of the whole group of birds. However, a considerable proportion of the aviary birds and pets may be are carriers of C. psittuci, so control in these animals is also needed. Ideally, this control should be based on voluntary participation.

References Ahrens, M. and Weingarten, M., 1981. Vergleichende Untersuchungen an Buffalo Green Monkey (BGM)-Zellen und Maitsen zur Isolierung van Chlamydiapsittaci aus Kot- und Organproben von Viigeln. Zbl. Vet. Med. B., 28: 301-309. Alexander, D.J., Bevan, B.J., Lister, S.A. and Bracewell, C.D., 1989. Chlamydia infections in racing pigeons in Great Britain: A retrospective serological survey. Vet. Rec., 125: 239. Allen, J.E., Beatty, P.R. and Stephens, R.S., 1990. Recombinant fusion proteins define T-cell antigenic sites on the major outer membrane protein of Chlamydia trachomatis. In: Bowie W.R. et al., (eds.), Chlamydial infections, Cambridge University Press, Cambridge, pp. 101-104. Andersen, A.A., 1991. The serotyping of Chlamydiopsittaci isolates using serovar-specific monoclonal antibodies with the micro-immunofluorescence test. J. Clin. Microbial., 29: 707-7 11. Andrews, B.E., Major, R. and Palmer, S.R., 1981. Ornithosis in poultry workers. Lance& 1: 632-634.

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