Bernhard Kaltenboeck, Dr. med. vet., PhD
Bacteria of the order Chlamydiales are obligate intracellular parasites of eukaryotic cells, infecting single-celled organisms such as amoebae to virtually any cell type of multicellular organisms. Virtually any chlamydial organism can infect any host cell, resulting in ubiquitous infections with these bacteria. Different efficiencies in infectivity and replication determine consistent, but not absolute, associations between chlamydial strain, host, and disease manifestation. These bacteria produce only minimally toxic or non-toxic products and regulate their growth based on the availability of host cytoplasmic nutrients. This results in minimal cytotoxicity of chlamydiae, and almost all diseases are manifestations of aberrant immune responses that require weeks to eliminate the bacteria, if at all. Specific strains of chlamydiae infect livestock, and the focus of the following seminar will be on respiratory and mammary gland infections with these bacteria in livestock, and approaches for control of the mostly subclinical disease caused by these ubiquitous bacteria.
Basic Biology of Chlamydiae
The Chlamydiales, thought to be viruses for a long time, are obligate intracellular bacterial pathogens of higher cells that replicate within host cytoplasmic vacuoles termed inclusions. The chlamydial elementary body (EB) is near the limit of light microscopic visibility with approximately 0.3 microns in diameter, round or occasionally pear-shaped, and contains electron-dense structures comprised of tightly coiled genomic DNA covered by histone-like proteins. It is the infectious stage of the chlamydial developmental cycle, and functions as a tough "spore-like" body whose purpose is to permit chlamydial survival in the non-supportive environment outside the host cell. The ultra-structure of EB has been extensively studied1, and spikes in the membrane are thought to be the tubular apparatus of a chlamydial type III secretion system.
The chlamydial RB is the chlamydial developmental stage during intracellular replication, and it is non-infectious. Typically, the RB has a diameter of approximately 1 µm. The RB is metabolically active, and the cytoplasm is rich in ribosomes, which are required for protein synthesis. As the RB begins to differentiate into an elementary body, sites of re-condensation of nucleic acid appear in its cytoplasm. In the maturing inclusion, chlamydial particles appear to be packed tightly in the inclusion membrane. Development of chlamydiae is highly dependent on nutrient supply and metabolic status of host cells. Nutrient deficiencies such as low glucose, iron, or amino acid levels induced, e.g., by interferon-γ, lead to delayed development and too few, aberrant chlamydial organisms within the inclusion. These aberrant inclusions are thought to represent persistent chlamydial forms that are important for maintaining chlamydial infections in a host population.
Classification of chlamydiae had traditionally been based on host and/or disease association, without a high degree of consistency. Subsequent investigations of phylogenetic relationships between the major outer membrane protein (ompA) genes of chlamydiae laid the groundwork for classification based on genetic relatedness.2 In a proposal for classification, similarities of the 16S and 23S ribosomal RNA genes are the basis of a new taxonomy for the order Chlamydiales (Figure 1).3 The new classification separates the family Chlamydiaceae, which contains all classic 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.
Phylogeny of the order Chlamydiales based on full-length 23S rRNA genes.3 Branch lengths are measured in nucleotide substitutions, and numbers show branching percentages in bootstrap replicates.3
The genus Chlamydophila (C.) consists of 6 species. C. psittaci in birds had been recognized since the 1870s as causative agent of a disease termed psittacosis.4 Abortion in sheep caused by C. abortus was first described in Scotland in 1936.5,6 C. felis was isolated from cat pneumonia in 1944 and is associated with pneumonia and conjunctivitis in cats7,8,9, and C. caviae was originally isolated from conjunctival scrapings of guinea pigs.10 C. pecorum, a relatively new species, is associated with polyarthritis, enteritis, pneumonia, and urogenital infections in cattle, sheep, goats, koala, and swine.11,12,13 C. pneumoniae is mainly a human pathogen, but infects also koalas, horses, and frogs.2,14 The genus Chlamydia contains the classical species C. trachomatis, the agent of the scarring human ocular disease trachoma in tropical countries. Other serovars of C. trachomatis cause human urogenital infections and are the most prevalent sexually transmitted human pathogens.15 C. muridarum was isolated from a mouse colony with pneumonia16, and C. suis is the most prevalent chlamydial agent in swine, identified in cases of conjunctivitis, mastitis, enteritis, and urogenital infection.17
The current separation of Chlamydiaceae into two genera has not met wide approval, particularly from the medical community. For this reason, the next edition of Bergey's Manual of Determinative Bacteriology will retain the 9 species, but will abolish the genus Chlamydophila and will contain only a single genus Chlamydia. This approach will presumably for the foreseeable future remain the classification of the medically and veterinary important chlamydial species.
The chlamydial genome is one of the smallest genomes of all bacteria, and is between 1.1 and 1.25 Mb in size. Genome sequence comparison has become the preferred method to deduce evidence for functional genetic differences between chlamydiae, because genetic manipulation systems are not available for chlamydiae. The C. muridarum genome was published in 2000 as the first genome of an animal chlamydial species that is of great comparative interest to human C. trachomatis.18 The C. muridarum genome is highly similar to the C. trachomatis D genome, but has also significant differences that potentially explain the different host and tissue tropism. The main differences of C. muridarum are the loss of tryptophan synthesis genes, different nucleotide salvaging pathway genes, and in particular three copies of a toxin gene tox that is very similar to the cytotoxic enterobacterial Efa1 gene of enterohemorrhagic E. coli or clostridial large cytotxins (LCT). The cytotoxicity of these genes is related to their interaction with actin that causes the disassembly of the actin cytoskeleton, and analysis of the cytotoxicity of C. muridarum and C. trachomatis has demonstrated cytopathic LCT effects in epithelial cells are thus potentially attributable to the tox genes.19
The next animal chlamydial genome determined was that of C. caviae in 2003,20 followed by C. abortus in 2005.21 Interestingly, the genomes of these species, while again highly syntenic, differ critically with respect to the above putative host, tissue and/or disease determinants ("niche-specific" genes): C. caviae encodes a tox-ortho-log, and tryptophan synthesis and nucleotide salvaging genes, while C. abortus encodes none of these. This might explain why C. abortus can 1) be propagated easily in many cultured cells and to higher yields than other chlamydiae; 2) be found in high numbers in infected host tissue; 3) colonizes macrophages effectively; and 4) rapidly spreads systemically after mucosal inoculation.22,23,24 Genome sequencing of the remaining four animal chlamydial species--C. psittaci, C. felis, C. pecorum, and C. suis--is currently underway, and the distribution of these and other potential "niche-specific" genes will be of great interest to explain host, tissue, and disease tropisms of animal chlamydial species. Initial data indicate that C. pecorum carries multiple copies of the tox gene, consistent with its high cytotoxicity.25
Diagnosis of Chlamydial Infections in Life stock
Chlamydial infections in animal are usually asymptomatic and inapparent, therefore diagnosis based on clinical symptoms 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, which includes direct impressions smears and cytological staining; cell culture isolation of the agent; immunofluorescence tests; enzyme immunoassays; and nucleic acid amplification based tests.
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.26,27
The standard method for detection of antibodies against Chlamydiaceae spp. 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 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.26,28 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 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 titers of all but one serum sample were negative. The single positive serum had a low titer of 1:10.28 The high seroprevalence of chlamydial infections poses a problem of defining truly negative control sera that allow for a reliable serological cut-off in antichlamydial antibody ELISA assays.
Detection of Chlamydial Organisms
For many years, the standard method of confirming the presence of chlamydial infections had been the propagation of the infecting organism in yolk sacs of chicken embryos, and the demonstration of characteristic chlamydial inclusions.6 Cultivation in cell culture is now preferred, and the use of appropriate techniques is important for high-sensitivity culture29. Buffalo Green Monkey Kidney (BGMK) cells support chlamydial replication effectively, particularly when cultivated in Iscove's Modified Dulbecco's Medium. Nevertheless, propagation methods require adequate transport and cold-storage facilities in order to maintain the viability of the organism prior to inoculation. Moreover, growth and isolation of the organisms in cell culture is tedious, it is difficult to consistently maintain high quality laboratory methods, and nucleic acid based detection methods exceed culture of chlamydiae in detection sensitivity. Therefore, isolation of chlamydiae is typically only performed during epidemiological studies that aim to recover new isolates.
Detection of Chlamydial Antigens
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-labeled Chlamydia-specific monoclonal antibodies, or on the capture and detection of chlamydial antigen in an extract of clinical material using enzyme immunoassay-based procedures. 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 Chlamydiaceae family-specific monoclonal antibodies against the chlamydial lipopolysaccharide, and have therefore a wide diagnostic spectrum suited for use in animal diagnostics.
Detection of Chlamydiae by Nucleic Acid Amplification Techniques
The direct antigen 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 amplification-based methods are now of prime importance for the diagnosis of chlamydial infections.11,17 Indeed, the development of chlamydial tests based on nucleic acid amplification technology (NAAT) is the most important advance for the detection of chlamydial infections since cell culture. 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 process30, and are considerably more sensitive than culture or antigen detection methods.31
Most of the NAAT platform technologies have been specifically marketed for detection of C. trachomatis, but general considerations about sensitivity and specificity equally apply to the detection of animal Chlamydiaceae spp. Allowing for the problems of discrepant analysis, the true sensitivity of PCR and LCR is of the order of 90 to 97%.32 An integrated nucleic acid isolation and real-time PCR platform was developed to specifically detect, differentiate, and quantify all Chlamydiaceae spp. by fluorescence resonance energy transfer real-time PCR with high sensitivity.33,34 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 Chlamydiaceae 23S rRNA allowed for the differentiation of chlamydial species and was more robust at low target numbers than amplification of the ompA gene. The widespread use of such PCR techniques is the reason for the better understanding and increased detection of chlamydial infections in animal. These methods continue to be improved in several aspects. Sampling into guanidinium-based buffers for maximum preservation of nucleic acids and optimum nucleic acid extraction have vastly improved detection of low numbers of chlamydial genomes in specimens.33,35 Real-time PCR techniques have not only the benefit of determination of copy number of chlamydial targets, but also allow routine high-throughput PCR assays, reduce the risk of false positives through product carry-over contamination by virtue of the single tube method, and typically also increase detection sensitivity.35,36,37 Finally, the combination of PCR amplification and low-density single-tube microarrays has resulted in a rapid, accurate, and flexible typing format for use in routine detection and multiplex typing of chlamydial amplicons.38
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