chlamydial infections of livestock and poultry
- Host Animals
- Hosts/Species Affected
- Systems Affected
- Distribution Table
- List of Symptoms/Signs
- Disease Course
- Impact: Economic
- Zoonoses and Food Safety
- Disease Treatment
- Prevention and Control
- Distribution Maps
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PicturesTop of page
IdentityTop of page
Preferred Scientific Name
- chlamydial infections of livestock and poultry
International Common Names
- English: abortion caused by chlamydial infection; air sac inflammation caused by chlamydial infection; Chlamydia muridarum infections; Chlamydia suis infections; Chlamydia trachomatis infections; chlamydial infection in cattle; chlamydial infection in goats; chlamydial infection in pigs; chlamydial infection in poultry; chlamydial infection in sheep; Chlamydophila abortus infections; Chlamydophila caviae infections; Chlamydophila felis infections; Chlamydophila pneumoniae infections; Chlamydophila psittaci infections; conjunctivitis caused by chlamydial infection; enteritis caused by chlamydial infection; ornithosis; pneumonia caused by chlamydial infection; polyarthritis caused by chlamydial infection; psittacosis; sporadic bovine encephalomyelitis
Pathogen/sTop of page Chlamydia muridarum
OverviewTop of page
Bacteria of the order Chlamydiales are obligate intracellular parasites of eukaryotic cells. They have a distinctive developmental cycle with two distinct morphological forms: the elementary body (EB) and the reticulate body (RB). EBs are typically 0.2-0.6 µm in diameter with spherical shape, while RBs are larger, up to 1.5 µm in diameter and more amoeboid in shape. Chlamydial infections cause a wide variety of clinical diseases in animals, which do not show any obvious signs. Chlamydial infections become clinically manifest as a disease syndrome called ornithosis or psittacosis in birds and mammals that includes pneumonia, air sac inflammation, and enteritis. Other distinct chlamydial disease mainfestations include abortion and conjunctivitis, encephalomyelitis and polyarthritis in mammals. When activated by stress factors, chlamydial infection may take a severe, systemic and sometimes fatal course of disease. Classification of chlamydiae had traditionally been based on host and/or disease association. In a new proposal for classification, similarities of the 16S and 23S ribosomal RNA genes are the basis of a new taxonomy for the order Chlamydiales (Everett et al., 1999). The new classification separates the family Chlamydiaceae, which contains all classical pathogenic chlamydiae, into two genera, Chlamydia and Chlamydophila, with a total of nine species. It also adds three new families, the Parachlamydiaceae, Waddliaceae, and Simkaniaceae. The genus Chlamydophila consists of 6 species. C.psittaci in birds has been recognized since the 1870s as causative agent of a disease termed psittacosis (Meyer, 1965). Abortion in sheep caused by C. abortus was first described in Scotland in 1936 (Stamp et al., 1952; Storz, 1971). C. felis was isolated from cat pneumonia in 1944 and is associated with pneumonia and conjunctivitis in cats (Baker, 1944; Cello, 1967; Shewen et al., 1978), and C. caviae was originally isolated from conjunctival scrapings of guinea pigs (Murray, 1964). C. pecorum, a relatively new species, is associated with polyarthritis, enteritis, pneumonia, and urogenital infections in cattle, sheep, goats, koala, and swine (Kaltenboeck et al., 1991; Fukushi et al., 1992; 1993; Girjes et al., 1993; Anderson et al., 1996). C. pneumoniae is mainly a human pathogen, but also infects koalas, horses, and frogs (Kaltenboeck et al., 1993; Reed et al., 2000).
Chlamydia, the second genus of the family Chlamydiaceae, consists of 3 species: C. trachomatis is the classic human pathogen causing ocular and urogenital disease. Chlamydia muridarum had long been known as the mouse pneumonitis biovar of C. trachomatis (Nigg, 1942). C. suis has been identified only recently as a common pathogen in pigs associated with fertility disorders and perinatal mortality (Kaltenboeck et al., 1993; Schiller et al., 1997; Woollen et al., 1991).
Different strains of the three other families of the order Chlamydiales have been sporadically associated with different diseases in a variety of mammalian and cold-blooded hosts.
Host AnimalsTop of page
|Animal name||Context||Life stage||System|
|Bos indicus (zebu)||Domesticated host, Wild host||Cattle & Buffaloes: All Stages|
|Bos taurus (cattle)||Domesticated host, Wild host||Cattle & Buffaloes: All Stages|
|Bubalus bubalis (Asian water buffalo)||Domesticated host, Wild host||Cattle & Buffaloes: All Stages|
|Capra hircus (goats)||Domesticated host, Wild host||Sheep & Goats: All Stages|
|Equus caballus (horses)||Domesticated host, Wild host||Other: All Stages|
|Gallus gallus domesticus (chickens)||Domesticated host, Wild host||Poultry: All Stages|
|Ovis aries (sheep)||Domesticated host, Wild host||Sheep & Goats: All Stages|
|Sus scrofa (pigs)||Domesticated host, Wild host||Pigs: All Stages|
Hosts/Species AffectedTop of page
Chlamydia have one of the widest host range of any type of microorganism, affecting animal species from Amoeba and Hydra through arthropods, insects, molluscs, marsupials, birds, reptiles, amphibians and mammals, including humans. Among mammals, Chlamydophila spp. have been isolated from ruminants, pigs, horses, koalas, dogs, rabbits, ferrets and opossums (Storz, 1988). Chlamydia spp. have been found in rodents (C. muridarum; Eddie et al., 1969) and pigs (C. suis; Kaltenboeck et al., 1997b). There are a number of reports of chlamydial infections in buffaloes (Gupta et al., 1976; Dhingra et al., 1980; Rowe et al., 1978).
Serological examinations show that chlamydial infections also exist in other mammals, including monkeys, wild boar, hedgehogs (Storz, 1988), deer (Taylor et al., 1996; Giovannini et al., 1988), reindeer (Neuvonen, 1976), fur seals (Eddie et al., 1966), and birds (Meyer, 1965). Chlamydial infections were reported in four species of wild ruminants (fallow deer, mouflon, red deer, and Spanish ibex) from a nature reserve in Spain. These animals might act as reservoirs of chlamydial infection (Cubero-Pablo et al., 2000). Serological surveys also provide evidence of chlamydial infection in antelope (Mansell et al., 1995), captive Arabian oryx (Greth et al., 1992), and snowshoe hares and muskrats in Saskatchewan, Canada (Spalatin et al., 1966).
Chlamydial infection and disease was also confirmed in reptiles such as chameleon, lizard, sea turtles, and crocodiles (Homer et al., 1994; Huchzermeyer et al., 1994). A pneumonia and anaemia disease syndrome in giant barred frogs of Australia is probably caused by the koala biovar of C. pneumoniae. Chlamydial infections have also been confirmed in another amphibian species, the African clawed frog (Newcomer et al., 1982; Howerth, 1984; Reed et al., 2000). Chlamydial infection were reported to cause chronic gill disease in Connecticut striped bass and white perch (Wolke et al., 1970), and high mortality in cultured pacu, a tropical fish species in Brazil (Szakolczai et al., 1999).
Chlamydial infections also occur in invertebrates such as in ticks and fleas (Weyer, 1970; Eddie et al., 1966), and lice (McKercher et al., 1980). Chlamydia-like organisms, now classified as Parachlamydiaceae, have been isolated from amoebae (Amman et al., 1997), were detected in the ovaries of the spider, Segestria senoculata (Traciuc, 1985), and caused fatal disease in the spider Pisaura mirabilis (Morel, 1978).
Harshbarger et al. (1977) found Chlamydia agents in the digestive systems of hard clams and oysters from the Chesapeake Bay area of the USA. Similar organisms were detected in the digestive cells of approximately 5% of mussels from the Basque Coast of Spain (Cajaraville et al., 1991). Over one year, Svardh (1999) examined the prevalence of disease-associated organisms in blue mussel populations in Denmark. Chlamydiae were the only bacteria detected.
Systems AffectedTop of page digestive diseases of large ruminants
digestive diseases of pigs
digestive diseases of poultry
digestive diseases of small ruminants
mammary gland diseases of large ruminants
mammary gland diseases of pigs
mammary gland diseases of small ruminants
multisystemic diseases of large ruminants
multisystemic diseases of pigs
multisystemic diseases of poultry
multisystemic diseases of small ruminants
nervous system diseases of large ruminants
nervous system diseases of pigs
nervous system diseases of poultry
nervous system diseases of small ruminants
reproductive diseases of large ruminants
reproductive diseases of pigs
reproductive diseases of poultry
reproductive diseases of small ruminants
respiratory diseases of large ruminants
respiratory diseases of pigs
respiratory diseases of poultry
respiratory diseases of small ruminants
urinary tract and renal diseases of large ruminants
urinary tract and renal diseases of pigs
urinary tract and renal diseases of poultry
urinary tract and renal diseases of small ruminants
DistributionTop of page
Chlamydial diseases of animals have been described in all continents, and 22% of countries and regions around the world have reported incidences of animal chlamydial infections. This number is probably a substantial underestimate of the true incidence. All data suggest that chlamydial infections of animals are actually ubiquitous. The typically chronic nature of chlamydial diseases, the difficulty in detecting this intracellular pathogen and the inconsistent use of high-sensitivity detection methods have led to underdiagnosis of chlamydial infections in animals. In humans, 73% of all countries and regions worldwide report chlamydial infections, predominantly with C. trachomatis, associated with sexually transmitted diseases and blindness (Resnikoff et al., 2004; Mak et al., 2005).
Distribution TableTop of page
The distribution in this summary table is based on all the information available. When several references are cited, they may give conflicting information on the status. Further details may be available for individual references in the Distribution Table Details section which can be selected by going to Generate Report.
|Continent/Country/Region||Distribution||Last Reported||Origin||First Reported||Invasive||Reference||Notes|
|Bahrain||Present||Greth et al., 1992|
|China||Present||Cyranoski, 2003; Normile, 2003|
|India||Present||Gupta et al., 1976|
|Japan||Present||Mochizuki et al., 2000; Iwamoto et al., 2001|
|Philippines||Present||Asai et al., 1991|
|Saudi Arabia||Present||Greth et al., 1992; Greth et al., 1993|
|Sri Lanka||Present||Sixl et al., 1988|
|Taiwan||Present||Saikku et al., 1985; Liao et al., 1997|
|United Arab Emirates||Present||Wernery and Wernery, 1990|
|Botswana||Present||Mushi et al., 2001; Sharma et al., 2003|
|Cape Verde||Present||Miorini et al., 1988|
|Chad||Present||Lefevre et al., 1979|
|Congo||Present||Grimes and Arizmendi, 1992|
|Côte d'Ivoire||Present||Formenty and Domenech, 1992|
|Egypt||Present||Schmatz et al., 1978; Kay, 1997|
|Guinea||Present||Wesselingh et al., 1988|
|Kenya||Present||Krauss et al., 1971|
|Namibia||Present||Apel et al., 1989; Kolb et al., 1993|
|Somalia||Present||Schmatz et al., 1978|
|South Africa||Present||Huchzermeyer, 1997; Verwoerd, 2000|
|Sudan||Present||Abbas et al., 1979|
|Togo||Present||Espinasse et al., 1980|
|Tunisia||Present||Tarizzo and Nabli, 1967; Rekiki et al., 2002|
|Canada||Present||Chalmers et al., 1976; Rigby et al., 1981|
|Mexico||Present||Escalante-Ochoa et al., 1996; Escalante-Ochoa et al., 1997|
|USA||Present||DeGraves et al., 2003|
|-Alabama||Present||DeGraves et al., 2003|
|Argentina||Present||Fernandez and Ouvina, 1999|
|Brazil||Present||Raso et al., 2002|
|Austria||Present||Kaltenboeck et al., 1997b|
|Belgium||Present||Vanrompay et al., 1997; Vanrompay et al., 2004|
|Bulgaria||Present||Vodas and Elitsina, 1986|
|Croatia||Present||Madic et al., 1993|
|Czechoslovakia (former)||Present||Bazala and Renda, 1992; Cislakova et al., 2004|
|Denmark||Present||Ronsholt, 1977; Mordhorst et al., 1992|
|Finland||Present||Puolakkainen et al., 1988; Laurila et al., 1997|
|France||Present||Berthelon and Tainturier, 1975; Lombard et al., 1987|
|Germany||Present||Schettler et al., 2003; Hotzel et al., 2004|
|Greece||Present||Bougiouklis et al., 2000; Felmingham, 2004|
|Italy||Present||Giovannini et al., 1988; Cavirani et al., 2001|
|Lithuania||Present||Domeika et al., 1983|
|Netherlands||Present||Akkermans and Kreeft, 1992; Vanrompay et al., 1993|
|Norway||Present||Berdal et al., 1997; Akerstedt and Hofshagen, 2004|
|Poland||Present||Truszczynski, 1989; Kita and Anusz, 1991|
|Russian Federation||Present||Blagoveshchenskaia et al., 1977|
|Slovakia||Present||Kocianová et al., 1992; Cislakova et al., 2004|
|Spain||Present||Astorga et al., 1994; Cubero-Pablo et al., 2000|
|Sweden||Present||Fryden et al., 1989; Olsen et al., 1998|
|Switzerland||Present||Chanton-Greutmann et al., 2002; Borel et al., 2004|
|UK||Present||Sykes et al., 1997; McDonald et al., 1998|
|Ukraine||Present||Iatsyshin and Bortnichuk, 1976; Fedorov et al., 1984|
|Australia||Present||Sykes et al., 1997; Devereaux et al., 2003|
|Fiji||Present||Raju et al., 1988|
|New Zealand||Present||Childs et al., 1980; Johnston and Philips, 2004|
|Papua New Guinea||Present||Wesselingh et al., 1988|
PathologyTop of page
Pathological lesions change greatly with different hosts or different species of the same host, and types of Chlamydia spp. In the mouse intranasal inoculation model that uses C. muridarum, formerly termed the mouse pneumonitis (MoPn) strain of C. trachomatis, the organism can be found in alveoli as early as 30 min after inoculation (Weiss, 1949). Between 30 and 36 h, some of the inclusions break up, releasing elementary bodies (EB) into the alveolar space. After between 36 h and 7 days, EB can be detected in increasing numbers. They decrease from 10-13 days, and disappear after around 21 days. The initial cellular response of the mouse to MoPn infection was characterized by an accumulation of heterophils in the infected alveoli (Gogolak, 1953). The interstitial inflammatory infiltrate on day 3 contained primarily mononuclear cells consisting of lymphocytes, monocytes, and macrophages. Usually 4 days after inoculation, large portions of the lung are no longer normal in appearance. The interstitium and alveolar spaces contain large numbers of macrophages, and the bronchioles showed inflammation with both heterophils and a small number of mononuclear cells.
Pathological responses after respiratory C. pneumoniae or C. abortus infection in mice are analogous to those seen in humans as interstitial pneumonitis and resemble those seen in the MoPn infection model (Yang et al., 1993). Gross pathology includes a patchy distribution of areas of consolidation during the first 2 weeks following the intranasal inoculation. Around 2-4 days after infection, there was extensive infiltration of polymorphonuclear neutrophils (PMNs) with exude in alveolar spaces and bronchial lumens. More severe infiltration was seen by days 7-11 with mixed mononuclear and PMN infiltrates. Perivascular and peribronchial lymphoid cell accumulations persisted between day 11 through to day 60. Chlamydial inclusions can be observed predominantly in macrophages.
Pathogenically diverse, C. pecorum strains are associated with pneumonia, polyarthritis, conjunctivitis, abortion, encephalomyelitis, enteritis and diarrhea (Kaltenboeck et al., 1993). They may also cause metritis, salpingitis and infertility in cattle. In koalas, infection with C. pecorum causes genito-urinary disease (Jackson et al., 1999).
DiagnosisTop of page
Chlamydial infections in animals are usually asymptomatic and inapparent, therefore diagnosis based on clinical signs and pathological lesions and differential diagnosis are of minor importance. However, if abortion in mammals or conjunctivitis in birds is observed, chlamydial infection should be suspected. Confirmation of chlamydial infection usually requires collecting an appropriate clinical sample from the animal followed by the direct detection of the organism using a suitable laboratory-based diagnostic test; direct impression smears and cytological staining, cell culture isolation of the agent, immunofluorescence tests, enzyme immunoassays, or nucleic acid amplification-based tests such as nucleic acid direct hybridization or amplification assays.
Serological detection is generally only suitable for prevalence surveys, less for the retrospective diagnosis of chlamydial infection. Most chlamydial infections do not elicit sufficiently high changes in antibody levels to allow for unambiguous diagnosis of a recent infection. The exception is the diagnosis of chlamydial abortion in ruminants, in which the high exposure to C. abortus elicits an increase in antibody levels that is high enough to allow for unambiguous diagnosis (Perez-Martinez et al., 1986; Griffiths et al., 1996).
The standard method for detection of antibodies againstChlamydiaceae species in animals is the complement fixation test using crude or partially purified preparations of Chlamydiaceae-specific lipopolysaccharide, but numerous ELISA methods have also been introduced. The compliment fixation test (CFT) depends on the binding of anti-Chlamydiaceae antibodies of the host species to guinea pig complement, and has highly variable sensitivity depending on the host species and antibody isotype (Perez-Martinez et al., 1986; Kaltenboeck et al., 1997b). In a random survey of 40 sera from Alabama cattle herds with abortion problems, ELISAs against peptides of the C. abortus major outer membrane protein or against recombinant chlamydial lipopolysaccharides (LPS) invariably detected very high antibody levels. In fact, immunoglobulin-rich sera from gnotobiotic calves challenged with bovine coronavirus had to be used as negative controls because it was impossible to find any other Chlamydia-negative bovine sera. In comparison, the CFT titres of all but one serum sample were negative. The single positive serum had a low titre of 1:10 (Kaltenboeck et al., 1997b). The high seroprevalence of chlamydial infections poses a problem of defining truly negative control sera that allow for a reliable serological cut-off in ELISA assays of antichlamydial antibodies. In cattle it was necessary to obtain sera from gnotobiotic calves as negative controls.
For many years, the best method of confirming the presence of chlamydial infection had been the propagation of the infecting organism in cell culture and the demonstration of characteristic chlamydial inclusions. However, this method requires adequate transport and cold-storage facilities in order to maintain the viability of the organism before inoculation. Moreover, growth and isolation of the organisms in cell culture is relatively tedious, and it is difficult to maintain high quality laboratory methods consistently.
A key advance in the laboratory diagnosis of chlamydial infections has been the development of tests that are not dependent on the viability of the agent and are less demanding with respect to specimen transport. The first of these tests were chlamydial antigen detection tests, which relied either on the direct detection of chlamydial elementary bodies in clinical material using fluorochrome-labelled Chlamydia-specific monoclonal antibodies, or on the capture and detection of chlamydial antigen in an extract of clinical material using enzyme immunoassay-based procedures. These methods are still appropriate and remain in widespread use. However, they are gradually being superseded by newer methods based on the detection of chlamydial nucleic acid, either by direct hybridization or preferably by nucleic acid amplification. The latter use a variety of amplification reactions, including the polymerase chain reaction (PCR), ligase chain reaction (LCR), and strand displacement amplification or transcription-mediated amplification. Nucleic acid-based methods generally offer superior sensitivity and specificity to the antigen detection tests, but at greater cost and a greater requirement for trained staff. However, depending on the prevalence of infection in the test population, costs may be reduced by combining different specimens.
Since the 1960s immunofluorescence using polyclonal antibodies, and since the 1980s, monoclonal antibodies have been used for the detection of chlamydial antigen, both in cell culture and in clinical material. The Pathfinder® EIA (Sanofi/Kallestad) and the Boots-Celltech IDEIA® both use Chlamydiaceaefamily-specific monoclonal antibodies against the chlamydial lipopolysaccharide, and have therefore a wide diagnostic spectrum suited for use in animal diagnostics.
Nucleic acid amplification-based methods are now of prime importance for the diagnosis of chlamydial infections (Kaltenboeck et al., 1991; Lisby, 1999; Huang et al., 2001). Indeed, the development of chlamydial tests based on nucleic acid amplification technology (NAAT) has been considered the most important advance for the detection of chlamydial infections since cell culture (Stary, 2000). These tests amplify either the target nucleic acid, DNA or RNA, or the probe after it has annealed to target nucleic acid. Such tests are generally more sensitive than liquid or solid phase hybridization tests, which do not embody an amplification process (Chernesky, 1999), and are considerably more sensitive than culture or antigen detection methods (Ostergaard, 1999).
Most of the NAAT platform technologies have been specifically marketed for detection of Chlamydia trachomatis, but general considerations about sensitivity and specificity equally apply to the detection of the other Chlamydiaceaespecies. Allowing for the problems of discrepant analysis, the true sensitivity of PCR and ligase chain reaction (LCR) is of the order of 90 to 97% (Cheng et al., 2001). An integrated nucleic acid isolation and real-time PCR platform was developed to specifically detect, differentiate, and quantify all Chlamydiaceae species by fluorescence resonance energy transfer real-time PCR with high sensitivity (Huang et al., 2001; DeGraves et al., 2003, 2004). In this approach, step-down thermal cycling and an excess of hot-start Taq polymerase vastly improved the robustness and sensitivity of the real-time PCR while essentially maintaining 100% specificity. The amplification of Chlamydiaceae23S rRNA allowed for the differentiation of chlamydial species and was more robust at low target numbers than amplification of the omp1 gene. Target-specific reverse transcription before the PCR can potentially also detect ongoing bacterial transcriptional activity (Birch et al., 2001) and increase sensitivity of the PCR. Ribosomal RNA is stable and is particularly useful for diagnosis as there may be several thousand copies per bacterial cell.
The main advantage of the nucleic acid amplification-based diagnosis for chlamydiae is that such methods combine unsurpassed sensitivity with high specificity. However, the greater sensitivity of these assays means that accidental contamination with amplified product is a problem of major importance for kit design, laboratory workflow, and personnel. Nucleic acid amplification tests tend to be more expensive than other laboratory methods of testing for chlamydial infection. The clinic or laboratory contemplating adopting such tests therefore needs to consider not just sensitivity, specificity and the clinical requirements, but also the suitability of the test for the facilities and human resources available. Fortunately, for high throughput testing, kit manufacturers can provide instrumentation to achieve at least partial automation. Alternatively, with a little ingenuity, it may be possible to adapt other programmable laboratory dispensing/assay equipment.
List of Symptoms/SignsTop of page
|Digestive Signs / Anorexia, loss or decreased appetite, not nursing, off feed||Sign|
|Digestive Signs / Diarrhoea||Sign|
|General Signs / Dehydration||Sign|
|General Signs / Fever, pyrexia, hyperthermia||Sign|
|General Signs / Generalized weakness, paresis, paralysis||Sign|
|General Signs / Underweight, poor condition, thin, emaciated, unthriftiness, ill thrift||Sign|
|General Signs / Weight loss||Sign|
|Nervous Signs / Dullness, depression, lethargy, depressed, lethargic, listless||Sign|
|Ophthalmology Signs / Chemosis, conjunctival, scleral edema, swelling||Sign|
|Ophthalmology Signs / Conjunctival, scleral, injection, abnormal vasculature||Sign|
|Ophthalmology Signs / Conjunctival, scleral, redness||Sign|
|Ophthalmology Signs / Lacrimation, tearing, serous ocular discharge, watery eyes||Sign|
|Ophthalmology Signs / Purulent discharge from eye||Sign|
|Reproductive Signs / Abortion or weak newborns, stillbirth||Sign|
|Reproductive Signs / Female infertility, repeat breeder||Sign|
|Reproductive Signs / Foul smelling discharge, vulvar, vaginal||Sign|
|Reproductive Signs / Mucous discharge, vulvar, vaginal||Sign|
|Reproductive Signs / Mummy, mummified fetus||Sign|
|Reproductive Signs / Purulent discharge, vulvar, vaginal||Sign|
|Respiratory Signs / Dyspnea, difficult, open mouth breathing, grunt, gasping||Sign|
|Respiratory Signs / Increased respiratory rate, polypnea, tachypnea, hyperpnea||Sign|
|Skin / Integumentary Signs / Rough hair coat, dull, standing on end||Sign|
Disease CourseTop of page
Disease course and progression of chlamydial infection is highly dependent on host, entry site and quantity of infection, co-infections and/or secondary infection, nutrition and immune status, and other stress factors such as transportation and overcrowding. Chlamydial infection induces mainly asymptomatic and clinically inapparent infections in animals. However, manifest clinical signs or even high mortality were seen in the presence of stress factors in young or pregnant animals (Woollen et al., 1990; Verwoerd et al., 2000).
EpidemiologyTop of page
Psittacosis is the term for chlamydial infection of psittacine birds or man, and ornithosis is the term for the same infection in birds other than psittacines (Storz, 1971). Infections with chlamydial agents have been described in 130 species of birds (Meyer, 1965; Bonnet et al., 2000), and infections of C. psittaci in birds are important because they cause economic loss to the poultry production and represent a biological hazard to human health. Ornithosis in birds involves mainly the gastrointestinal tract, and the pathogen is shed in faeces or through infectious discharges from the respiratory tract. C. psittaci has been isolated from symptomatic and apparently healthy birds. Clinically asymptomatic and latent infections may predominate for C. psittaci infection. Stress factors such as overcrowding, poor nutrition, viral and other bacterial infections, and transportation can precipitate overt clinical disease and mortality. Infectious chlamydiae in respiratory secretions or faeces may remain viable for several months. Transmission of disease is mainly through aerosols of faecal or feather dust. Vertical transmission through eggs has been found in chickens, ducks, seagulls and psittacine birds (Shewen, 1980). Young birds tend to be more susceptible to infection than older birds, and some species seem to be more susceptible than others. It is also possible that feral birds act as natural reservoirs of the agent and introduce chlamydiae into farmed bird populations such as turkeys and ducks. Wild and racing pigeons, the bird trade, and migrations of wild birds such as seagulls, finches, sparrows and waterfowl may all contribute to the dissemination and transmission of C. psittaci throughout avian populations (Travnicek et al., 2002).
Ruminant chlamydial abortion and infertility
Chlamydophila abortus strains have been isolated worldwide in cases of abortion, predominantly in sheep, goats, and cattle, but occasionally also in horses, pigs, rabbits, guinea pigs and mice (McCauley et al., 1968; Jain et al., 1975; Appleyard et al., 1983; Schiller et al., 1997; Everett et al., 1999). In general, chlamydial abortion is commonest in lowland sheep flocks, mainly where sheep are closely confined. Usually, herds become affected after the introduction of asymptomatic carrier animals with chlamydial infection. Infection most likely occurs in uninfected sheep at lambing time by ingestion of C. abortus,which is excreted by infected and aborting ewes in diseased placentas, uterine discharges, and faeces. In sheep, the infection remains present at a subclinical level until the last 4 weeks of the next pregnancy. The major outbreak of disease in a flock tends to occur in the second lambing season after C. abortus infection was contracted. It has also been reported that sheep can acquire C. abortus infection and abort in one lambing season (Storz, 1971). Most affected sheep abort in the last month of pregnancy, and the majority of aborting ewes are young animals, although sheep of all ages are susceptible to infection (Young et al., 1958). In a flock with first-time C. abortus infection, up to 30% of pregnant ewes may abort. In subsequent years, as the infection becomes established as an enzootic disease, between 5-10% of pregnant ewes abort annually. C. abortus infection resulting in abortion leads to effective immunity in affected ewes.
Chlamydial abortion in cattle and other species is similar to enzootic abortion in sheep, but much more sporadic and less common than the disease in sheep and goats. Transmission of the disease in cattle occurs similarly to that in sheep, mostly by ingestion of infected tissues. However, C. abortus has been shown to cause seminal vesiculitis in bulls and rams, may reduce semen quality, and may be transmitted in semen (Storz et al., 1976).
C. pecorum also causes disease of the reproductive tract of cattle and pigs. This might be analogous to the insidious progress of C. trachomatis genital tract infection in humans (Hitchcock, 1999), in which symptoms may go unnoticed for a considerable period, but may lead to chronic sequelae such as pelvic inflammatory disease and infertility. In the USA, a 53% prevalence of vaginal C. abortus or C. pecorum infection has been detected in virgin heifers by quantitative PCR, suggesting that transmission is predominantly extragenital (DeGraves et al., 2003, 2004; Jee et al., 2004). Although C. abortus is primarily associated with spontaneous abortion in cattle and sheep, there is evidence that C. pecorum causes pregnancy wastage (Jones et al., 1999). C. pecorum infection was reported to cause a severe metritis, which would have resulted in at least temporary infertility (Jones, 1999; Magnino et al., 2000). Infertility problems have occurred in dairy cows following abortions (Reed et al., 1975), and it is possible that sporadic C. pecorum abortions are either undiagnosed or have been misdiagnosed as C. abortus abortions.
Ruminant chlamydial enteritis, pneumonia, polyarthritis, and sporadic bovine encephalomyelitis
C. pecorum infections are both endemic and chronic in the intestinal tract of sheep and cattle populations around the world (Griffiths et al., 1992; 1996; Markey, et. al., 1993; Jones et al., 1997; Clarkson et al., 1997). Intestinal carriage and faecal excretion onto pasture probably plays a major role in the maintenance of C. pecorum infection. These infections occasionally result in acute enteritis. Respiratory tract infection with C. pecorum may lead to severe pneumonia associated with lung consolidation. Animal diseases caused uniquely by C. pecorum are polyarthritis in calves and lambs, and presumably also in piglets (Storz, 1971; Kaltenboeck et al., 1993). Bovine encephalitis was one of the first chlamydial diseases in cattle identified by McNutt (1940). This disease occurs worldwide, and C. pecorum has been identified as the aetiologic agent of the disease termed sporadic bovine encephalomyelitis (SBE) (Storz, 1971).
Feline chlamydial infection
Chlamydophila felis is endemic among domestic cats worldwide, the commonest conditions that it causes are conjunctivitis and rhinitis in young cats (Gaillard et al., 1984). The disease is transmitted through infected aerosols and secretions. Since chlamydial strains from mammals tend to be of relatively low infectivity for humans, conjunctival infection in cats is not usually considered to be a major cause of symptomatic human infection. However, zoonotic infection of humans with C. felis has been described.
Impact: EconomicTop of page
Economically, chlamydial infections impact on both the efficiency of animal production and on public health. Costs depend on the prevalence of the disease, the associated costs of detection, vaccination, treatment, management, and the impact on productivity. There are numerous reports and statistical analyses on the economic impact of chlamydial disease on public health (Rieetmeijer et al., 2002; Postma et al., 2002). However, only scant evidence of the economic impact of chlamydial infection in animals is available, even though this pathogen induces numerous diseases in a wide range of animal hosts. Interest in and understanding of the economic importance of animal chlamydial infections is probably biased towards the acute disease manifestations such as avian herd infections, abortion, and sporadically diagnosed diseases such as sporadic bovine encephalomyelitis (SBE), polyarthritis, pneumonia, or enteritis. This bias is most likely enhanced by the underdiagnosis of chronic infections. It is noteworthy that epidemiological surveys suggest that chronic human infection by C. pneumonia might accelerate the development of inflammatory conditions such as cardiovascular disease (Shi et al., 2004; de Kruif et al., 2005).
Recent studies in cattle suggest that chronic chlamydial infections may be more economically important than anticipated, and because of their ubiquitous prevalence, might be more important than the acute chlamydial diseases, which tend to occur infrequently. Application of highly sensitive real-time PCR and ELISA methods for detecting Chlamydophila spp. DNA and of antibodies against Chlamydophila spp., respectively, in a series of prospective cohort studies revealed a high prevalence of Chlamydophila spp. genital infections in female calves (61%) and adult heifers (53%) (Jee et al., 2004; DeGraves et al., 2003). These infections were acquired by extragenital transmission in the first weeks of life, and infection was frequency increased by crowding of the animals. A challenge study demonstrated that infection with C. abortus resulted in decreased fertility of heifers. The experimental use of a C. abortus vaccine provided evidence of immunoprotection against C. abortus-induced suppression of bovine fertility. The results of these investigations suggest that bovine Chlamydophila infection should be viewed more as pervasive, low-level infection of cattle than as rare, severe disease. Such infections proceed without apparent disease or with only subtle expressions of disease, but potentially have a large impact on bovine herd health and fertility.
Zoonoses and Food SafetyTop of page
Chlamydophila psittaci can be transmitted from birds to humans. In humans, the resulting infection is referred to as psittacosis, also known as parrot fever or ornithosis. Psittacosis typically causes influenza-like symptoms and can lead to severe pneumonia and non-respiratory health problems. From 1988 to 2003, the Centers for Disease Control in the USA received reports of 935 cases of psittacosis, which is probably an under-representation of the actual number of cases. Most human cases are associated with exposure to pet birds. A very serious course of psittacosis in pregnancy was also reported (Idu et al., 1998), in which the patient developed adult respiratory distress syndrome complicated by premature birth and perinatal mortality. The diagnosis ‘psittacosis’ was established on clinical grounds, confirmed serologically, and by the case history of contacting infected birds. Pregnant women should be advised to avoid contact with infected birds as they run an increased risk of a severe disease course (Idu et al., 1998).
Human C. psittaci infections are typically acquired by exposure to pet psittacine birds. However, transmission has been documented from poultry and free-ranging birds, including doves, pigeons, birds of prey and shore birds. Infection with C. psittaci usually occurs when a person inhales organisms that have been aerosolized from dried feces or respiratory tract secretions of infected birds. Other means of exposure include mouth-to-beak contact and handling infected birds’ plumage and tissues. Even brief exposures can lead to symptomatic infection; therefore, certain patients with psittacosis might not recall or report having any contact with birds. Infectious respiratory secretions may remain airborne in droplets for a long time. Organisms contained in dust composed of dried droppings represent a threat where bird faeces is not removed regularly. Clinical signs of human psittacosis include fever, headaches, muscle pain and respiratory symptoms. Since the disease is spread primarily through the excretions of infected animals, humans can protect themselves by avoiding inhaling or ingesting any of these excretions. Use of protective clothing and masks, together with normal hygienic practices during contact with infected animals virtually eliminates the risk of infection.
It is important to clean the animal or bird’s environment regularly to prevent dried faeces containing the organism from becoming dust. Chlamydia may live for months in the appropriate environment but are easily killed by most disinfectants. Depopulation and repopulation of a colony is effective only if a clean source of animals is available and when the prevention of re-infection is possible. Re-infection of a colony may occur through the introduction of infected animals or by the transport of organisms between infected and uninfected colonies such as dirty utensils, soiled clothing, and air movement.
Mammals occasionally transmit Chlamydiaceae organisms to humans. Chlamydophila abortus and C. pecorum infect sheep, goats, and cattle, causing chronic infection of the reproductive tract, placental insufficiency and abortion. Those chlamydiae may be transmitted to humans when humans are exposed to the birth fluids and placentas of infected animals. Human infection resulting from contact with infected goats has been reported (Villemonteix et al., 1990). It is likely that chlamydial abortion in goats should be treated as a similar threat to human health as infection in sheep. Genetic analysis of the isolates associated with sporadic abortion in women who had contact with infected sheep identified the strains as C. abortus (Herring et al., 1987). Organisms were recovered from the placental tissues and, in rare fatal cases, from additional organs. The incidence of this animal-acquired infection is not known, but several cases of pregnant women who have had spontaneous abortions following exposure to sheep infected with chlamydiae have been reported. Sheep infected with C. abortus strains represent an important potential risk to pregnant women (McKinlayet al., 1985). Furthermore, inhalation of infected material from sheep might also result in chlamydial respiratory disease in non-pregnant humans.
C. felis, the cat keratoconjunctivitis and pneumonitis agent, typically causes rhinitis and conjunctivitis in cats. Transmission of this species from cats to humans may be underreported.
Disease TreatmentTop of page
For many years, tetracyclines have been the main drug for anti-chlamydial therapy. Tetracyclines are inexpensive and still highly adequate for herd animal treatment. Antibiotic treatment should be maintained for at least 10 days. Antibiotic resistance is typically not a problem in chlamydial infections, but tetracycline resistance has been reported for one strain of Chlamydia suis.
Macrolide antibiotics, particularly erythromycin and tylosin have long been used as alternatives to the tetracyclines for the treatment of chlamydial infection. These drugs are far from ideal, with clinical cure rates of only 80% (Ridgway, 1997). The most relaible data are available for antibiotic treatment of human diseases, and these studies will be reported as guidelines for the treatment of animal chlamydial infections.
Recently azithromycin (Zithromax®) has in many ways revolutionized the treatment of chlamydial infections in humans and companion animals. This drug is an azalide and not a macrolide, having a 15-member ring with a methyl-substituted nitrogen in the aglycone ring. This confers acid stability, high tissue penetration, low serum levels and a very long half-life (Bryskier et al., 1993). Anti-chlamydial levels of the drug are readily achieved inside cells or tissues, particularly useful for treating intracellular chlamydial infections. Furthermore, adequate intracellular levels may be sustained for several days because of the slow efflux of the drug from cells.
Telithromycin®, the first of a new family of antibacterials, the ketolides, has recently been evaluated for chlamydial treatment. Telithromycin® has been advocated for the treatment of community acquired pneumonia due to Chlamydia pneumoniae and other respiratory organisms (Hammerschlag et al., 2001; Miyashita et al., 2001). The in vitro pharmacokinetics and chlamydiacidal activity of telithromycin and another ketolide against two strains of C. pneumoniae have been reported (Gustafsson et al., 2000; Miyashita et al., 2001; Strigl et al., 2000; Miyashita et al., 2001).
Gieffers et al. (2001) compared the susceptibility of C. pneumoniae growing in endothelial or smooth muscle cell culture to various quinolones (ofloxacin, levofloxacin, trovafloxacin, moxifloxacin). They found that moxifloxacin and trovafloxacin were as effective as the macrolides. The activity of the new generation fluoroquinolones against C. pneumoniae has recently been reviewed, and their efficiency and activity have been compared (Jones, 2002).
Prevention and ControlTop of page
The general principles and methods to prevent and control infectious diseases are applicable for Chlamydiaceae species. High quality nutrition and husbandry, strict quarantine before grouping animals of different origin, and reduction of stress factors are important to increase resistance of animals and to reduce the mortality of chlamydial infections. If chlamydial infection is confirmed, timely and adequate treatment of sufficient duration with antibiotics is required.
Vaccines have been used against animal chlamydial diseases, but their efficacy is uncertain, and large-scale clinical trials have not been performed. For use in cats, Fel-O-Vax vaccines are designed for prevention of feline pneumonitis caused by Chlamydophila felis in addition to other major feline diseases (Hoven et al., 1993).
Whole chlamydial antigen and subunit vaccines against ruminant chlamydial infections are in development. Stemke-Hale et al. (2005) reported the results of a general protocol that was used to screen the whole genome of Chlamydophila abortus, type strain B577, in a mouse pneumonia model. In this trial, genetic immunization was used to functionally test the genes of C. abortus as vaccines in a mouse challenge system. Nine gene fragments were isolated that conferred protection, with five protecting as effectively as the live-vaccine positive control. Clinically inapparent infection with C. abortus suppresses fertility in heifers, and vaccination with the recombinant C. abortus subunit vaccine improved 6-week pregnancy rates after single artificial insemination to 83% from a 50% pregnancy in mock-vaccinated heifers (Stemke-Hale et al., 2005).
A cohort study of 140 dairy cows in Germany showed that a therapeutic inactivated whole-organism C. abortus-C. pecorum vaccine significantly reduced somatic cell counts in dairy cow milk for about 140 days from 230,000/ml milk, with a peak reduction after 10 weeks to 83,000 cells/ml (unpublished data). These data suggest that subclinical Chlamydophila spp. infections have a continuous, low-level impact on animal health, and that vaccination against Chlamydophila spp. has the potential to improve animal health and production.
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