Avian Chlamydiosis (Psittacosis) in Pet Birds: Diagnostic Challenges and Molecular Detection

Introduction

Avian chlamydiosis, historically termed psittacosis or ornithosis, is a systemic bacterial infection of birds caused by the obligate intracellular bacterium Chlamydia psittaci. This pathogen is of paramount importance in veterinary medicine due to its high prevalence in psittacine birds (parrots, cockatiels, budgerigars), its capacity for latent infection, and its well documented zoonotic potential. The disease presents a unique diagnostic conundrum: infected birds may be asymptomatic carriers, acutely ill, or chronically shed organisms intermittently. Accurate detection is therefore critical for individual treatment, flock management, and public health protection.

This article provides a detailed examination of C. psittaci pathogenesis, the biological basis of diagnostic test limitations, and the principles of molecular detection methods. It also reviews current treatment protocols with doxycycline and discusses how molecular diagnostics have reshaped the management of this infection in companion birds.

Pathogenesis of Chlamydia psittaci in Birds

Chlamydia psittaci is a Gram negative, obligate intracellular bacterium that relies on a biphasic developmental cycle. The infectious form, the elementary body (EB), is metabolically inert and environmentally stable. Upon inhalation or ingestion by a susceptible bird, EBs are taken up by epithelial cells and macrophages of the respiratory tract and gastrointestinal system. Once internalized, EBs differentiate into reticulate bodies (RBs), which are metabolically active and replicate within a membrane bound inclusion body. After multiple rounds of replication, RBs redifferentiate into EBs, which are released by cell lysis or extrusion, propagating the infection [1, 2].

The primary sites of replication in birds are the respiratory epithelium, conjunctival mucosa, and gastrointestinal tract. From these loci, the organism disseminates hematogenously to the liver, spleen, and pericardium. The resulting pathology includes airsacculitis, pneumonia, conjunctivitis, hepatomegaly, splenomegaly, and fibrinous serositis. In psittacine birds, subtle signs such as lethargy, anorexia, ruffled feathers, and biliverdinuria (green urates) are common. Neurologic signs, including tremors and ataxia, occur less frequently but indicate severe systemic involvement [3, 4].

The ability of C. psittaci to establish persistent, subclinical infections is a defining feature of its pathogenesis. Persistence is characterized by a non-replicating, aberrant RB morphology that remains viable within the host cell. This state can be induced by nutrient deprivation, immune pressure, or sub inhibitory concentrations of antibiotics. Reactivation occurs under stress, such as breeding, concurrent illness, or environmental change, leading to renewed shedding [5, 6].

Zoonotic Transmission Risk

Although this article does not focus on human medicine, a brief discussion of the cross species transmission mechanism is relevant to understanding diagnostic urgency. Infected birds shed EBs in respiratory secretions, feces, and feather dust. The organisms remain infectious in dried feces and dust for several weeks at room temperature. Inhalation of aerosolized particles is the primary route of transmission to mammals. Given that Avian Influenza A(H5N1) in Poultry and Wild Birds: Current Epidemiology, Molecular Diagnostics, and Biosecurity is assessed through similar surveillance principles, the need for rapid detection of C. psittaci in avian populations is similarly critical. Other relevant cross species bacterial threats, such as Salmonella enterica Serovar Typhimurium in Backyard Poultry Flocks: Zoonotic Risk, Antimicrobial Resistance, and Biosecurity, underscore the importance of molecular surveillance for zoonotic agents in birds.

Diagnostic Challenges

The diagnosis of avian chlamydiosis is complicated by several interrelated factors: the biology of the organism, the variable immune response of the host, and the limitations of available testing modalities.

Clinical Signs as a Diagnostic Cue

Clinical signs in infected birds range from inapparent to fulminant. A significant proportion of carrier birds exhibit no outward signs, yet shed organisms intermittently. This subclinical shedding is a major challenge for diagnostic screening, as a single negative test does not rule out infection [7]. Clinical suspicion must be based on a constellation of signs including oculonasal discharge, dyspnea, diarrhea, and weight loss, but none are pathognomonic.

Serological Testing Limitations

Serological assays, including complement fixation (CF) tests, indirect immunofluorescence (IFA), and various formats of Enzyme-Linked Immunosorbent Assay (ELISA) for Feline Leukemia Virus (noting similar principles for antigen or antibody capture), are widely used. However, serology has substantial limitations in the diagnosis of C. psittaci.

The host humoral immune response is variable. Birds may seroconvert slowly, and antibody titers may wane after successful treatment or during periods of latency. Conversely, persistently infected birds can maintain high antibody titers without active shedding, leading to false positive interpretations regarding current infectious status [8, 9]. Cross reactivity with other Chlamydia species (e.g., C. abortus, C. pecorum) is a known problem, particularly in ELISA based platforms [10]. The specificity of serological assays is further compromised in young birds, which may carry maternal antibodies, and in immunocompromised individuals that fail to mount a detectable response.

Serology therefore offers a population level exposure history but is inadequate for diagnosing active infection in individual birds. A positive antibody titer indicates prior exposure but does not predict shedding; a negative titer does not exclude infection [11].

Antigen Detection Methods

Direct detection of C. psittaci antigen in clinical specimens (conjunctival swabs, choanal swabs, feces, or tissue) provides more direct evidence of current infection. Traditional methods include direct fluorescent antibody (DFA) staining and immunochromatographic lateral flow assays.

DFA staining of conjunctival or choanal swabs has moderate sensitivity (approximately 60-80%) when performed during the acute phase of disease. However, sensitivity drops markedly in birds with low level or intermittent shedding. DFA also requires a high quality fluorescent microscope and experienced personnel [12].

Lateral flow immunoassays are rapid and simple but suffer from poor analytical sensitivity when compared to molecular methods. Their use is generally limited to screening in clinical practice, with confirmatory testing recommended for positive results [13].

Cultivation of the Organism

Cell culture isolation has historically been the gold standard for diagnosis. C. psittaci can be cultured in McCoy cells, Buffalo green monkey (BGM) cells, or chicken embryo fibroblasts. After 48-72 hours, cytoplasmic inclusions can be demonstrated using species specific fluorescent antibodies. Culture has high specificity (near 100%) but low and variable sensitivity. Organism viability is required; improper sample handling, prior antimicrobial therapy, and delays in transport reduce recovery rates. Additionally, culture is slow (3-7 days), labor intensive, and requires BSL-2 or BSL-3 containment facilities due to the zoonotic risk. For these reasons, culture has been largely replaced by nucleic acid amplification tests in most veterinary diagnostic laboratories [14, 15].

Molecular Detection Methods

The advent of molecular diagnostics, specifically polymerase chain reaction (PCR) and its variants, has transformed the detection of avian chlamydiosis. These methods offer high analytical sensitivity, rapid turnaround time, and the ability to detect both viable and non-viable organisms.

Conventional PCR

Conventional PCR targeting the ompA gene (encoding the major outer membrane protein) or the 16S rRNA gene is well established for C. psittaci detection [16]. The ompA gene is particularly useful because it contains variable domains that allow genotyping. Nine major genotypes (A through F, E/B, M56, and WC) have been described. Genotype A is most commonly associated with psittacine birds, while genotype B is found in pigeons, genotype D in turkeys, and genotype E in various bird species [17].

Conventional PCR uses agarose gel electrophoresis for amplicon visualization, which provides moderate sensitivity (detection limits of 10-100 genome copies per reaction). However, the post amplification processing increases contamination risk and delays time to result [18].

Real Time Quantitative PCR (qPCR)

Real time PCR (qPCR) has become the method of choice for routine diagnosis. It employs fluorescent probes (e.g., TaqMan) that enable real time monitoring of amplification, eliminating the need for post PCR handling. qPCR offers several advantages:

• High sensitivity: detection limits as low as 1-10 genome copies per reaction [19].
• Quantitative capability: the cycle threshold (Ct) value correlates with the bacterial load in the sample.
• Reduced contamination risk: closed tube system.
• Faster turnaround: results within 1-3 hours.

Multiple qPCR assays have been developed targeting the ompA gene, the 16S rRNA gene, or the inclusion membrane protein A (incA) gene [20, 21]. Multiplex qPCR panels can simultaneously detect C. psittaci along with other avian respiratory pathogens, such as Mycoplasma gallisepticum and Avibacterium paragallinarum [22].

One critical consideration in qPCR interpretation is the Ct value. A low Ct (less than 30) typically correlates with active infection and high bacterial shedding. A high Ct (greater than 35) may represent low level infection, environmental contamination, or intermittent shedding. However, Ct values are laboratory specific and depend on collection methods, extraction efficiency, and amplification chemistry. Standardized protocols and internal amplification controls are essential for reliable inter laboratory comparison [23].

Nested PCR

Nested PCR, which uses two rounds of amplification with internal primers, provides even greater sensitivity than conventional PCR. However, the increased sensitivity comes at the cost of higher contamination risk. Nested PCR has been used in research settings to detect low abundance C. psittaci DNA in swab samples from asymptomatic birds [24]. Its clinical use has been largely superseded by qPCR due to qPCR's quantitative capability and lower contamination rate.

Isothermal Amplification Methods

Loop mediated isothermal amplification (LAMP) assays for C. psittaci have been developed as point of care molecular tools. LAMP operates at a constant temperature (60-65 degrees Celsius) using a DNA polymerase with strand displacement activity. The assay produces a visible precipitate or fluorescent signal within 30-60 minutes. LAMP assays targeting the ompA gene have shown high analytical sensitivity (100%) and specificity (96%) compared to qPCR [25]. These assays are promising for field deployment and resource limited settings, although they have not yet been widely adopted in companion bird practice.

Sample Types and Collection Considerations

The selection of appropriate clinical specimens is critical for PCR sensitivity. Sites of sampling reflect the pathogenesis of C. psittaci:

• Conjunctival swabs: capture ocular shedding.
• Choanal swabs: sample the upper respiratory tract.
• Cloacal swabs: detect gastrointestinal shedding.
• Combined choanal-cloacal swabs: improve yield over single site sampling [26].
• Feces: may be used but contain PCR inhibitors.
• Tissue samples (liver, spleen, lung): used postmortem.

Synthetic flocked swabs are preferred over cotton swabs as they release cells more efficiently. Swabs should be placed into a nucleic acid stabilization buffer and transported refrigerated or frozen if processing is delayed [27].

Genotyping and Molecular Epidemiology

Beyond detection, molecular methods enable genotyping, which informs epidemiological tracking. Sequencing of the ompA gene or multi locus sequence typing (MLST) using housekeeping genes (e.g., gatA, enoA, hemN) allows strain discrimination. MLST has revealed that certain sequence types are associated with specific avian hosts and geographic regions [28, 29]. This information is valuable when investigating outbreaks in aviaries, pet shops, or breeding facilities.

Diagnostic Workflow: PCR versus Serology

A rational diagnostic algorithm relies on both serology and PCR but assigns them distinct roles. PCR is the primary test for detecting current active infection and shedding. Serology is used to assess flock level exposure or to support a clinical suspicion in a bird that may be in the window period before PCR positivity.

The following diagram summarizes a suggested diagnostic workflow for a bird with clinical suspicion of chlamydiosis.

flowchart TD
    A[Clinical suspicion: respiratory, ocular, or GI signs in a psittacine bird], > B{Collect choanal and cloacal swabs for C. psittaci qPCR}
    B, > C[Perform qPCR targeting ompA or 16S rRNA gene]
    C, > D{Result}
    D, > |Positive| E[Active infection confirmed]
    D, > |Negative| F{Serology recommended?}
    F, > |Yes| G[Collect blood for anti-Chlamydia IgG/IgM ELISA]
    G, > H{Serology result}
    H, > |Positive| I[Previous exposure, possible latent infection; repeat PCR in 2-4 weeks]
    H, > |Negative| J[Low likelihood of current or past infection; consider alternative diagnosis]
    F, > |No| K[Consider alternative diagnosis or repeat PCR at multiple time points]
    E, > L[Begin treatment: doxycycline PO or IM for 45 days]
    L, > M[Recheck qPCR at 4-6 weeks post treatment]
    M, > N{Post treatment PCR}
    N, > |Negative| O[Treatment success confirmed]
    N, > |Positive| P[Incomplete clearance; extend treatment and re-evaluate]

This workflow emphasizes that a single negative qPCR result, especially when obtained during a period of low shedding, does not rule out infection. Serial testing over several weeks is often necessary for ruling out the carrier state.

Treatment Protocols and Monitoring

Doxycycline as the Drug of Choice

The treatment of avian chlamydiosis rests on the tetracycline class antibiotic doxycycline. Doxycycline is preferred over other tetracyclines due to its superior oral absorption, longer half life, and better tissue penetration in birds [30]. Doxycycline acts by binding to the 30S ribosomal subunit of the bacterium, inhibiting aminoacyl-tRNA binding and thus protein synthesis. In the context of C. psittaci, its efficacy against the intracellular RB form is well documented. However, doxycycline is less effective against the persistent, aberrant RB form, which explains the need for sustained treatment durations [31].

Administration Routes and Dosing

Doxycycline can be administered via the oral or parenteral route in pet birds.

Oral doxycycline is formulated as a suspension (doxycycline hyclate or doxycycline calcium syrup) or mixed into a medicated seed or pellet diet. The typical oral dose for psittacine birds is 25-50 mg/kg once daily for 45 days. Some protocols use a loading dose of 50 mg/kg for the first 3 days followed by 25 mg/kg daily [32]. Medicated seed diets (e.g., 0.1% doxycycline in seed) offer a convenient method for flock treatment but suffer from variable intake among individuals.

Parenteral doxycycline is available as doxycycline hyclate for intramuscular injection. Injections may be given at 75-100 mg/kg once every 5-7 days for 45 days, depending on the specific formulation. The long acting injectable formulation (e.g., doxycycline in a slow release vehicle) reduces handling stress but requires careful aseptic technique and proper injection site selection to avoid muscle necrosis [33].

Duration of Therapy

The recommended treatment duration is a minimum of 45 days for psittacine birds. This extended course is necessary to cover the complete developmental cycle of C. psittaci and to eliminate persistent organisms as they transition back through the replicative cycle. Shorter courses (21-30 days) have been associated with relapse and the development of a carrier state [34].

Monitoring Treatment Efficacy

Monitoring treatment success requires post therapy testing. PCR of choanal and cloacal swabs should be performed 4-6 weeks after the completion of antimicrobial therapy. A negative qPCR result at this time point is considered strong evidence of bacterial clearance. However, some experts recommend a second negative test 4 weeks later to confirm cure, particularly in valuable breeding birds or those in multi bird households [35].

Serology is not useful for monitoring treatment success, as antibody titers can remain elevated for months or years after successful clearance of the organism [36].

Antimicrobial Resistance Considerations

Resistance of C. psittaci to tetracyclines is considered rare, but cases of clinical failure with doxycycline have been reported. Some reports describe isolates with reduced susceptibility to doxycycline, potentially mediated by mutations in the 16S rRNA gene or efflux mechanisms [37, 38]. These cases underscore the importance of post treatment PCR testing to confirm clearance and to detect potential resistance. Alternative antimicrobials such as azithromycin (at 40 mg/kg orally once daily for 45 days) have been used in cases of doxycycline intolerance or demonstrated treatment failure [39].

Supportive Care

In addition to antimicrobial therapy, supportive care is critical. Severely affected birds may require fluid therapy, nutritional support via crop gavage, and environmental warmth. Non steroidal anti inflammatory drugs (e.g., meloxicam) can help reduce inflammation of air sacs and joints. Isolation from other birds is mandatory during the treatment period to prevent further transmission [40].

Comparison of Diagnostic Methods

The following table summarizes the key characteristics of the diagnostic modalities discussed.

Statistical Considerations in Diagnostic Testing

Understanding test performance metrics is essential for interpreting results. The sensitivity and specificity of PCR for C. psittaci vary depending on the target gene, primer design, and sample type. In meta analyses, qPCR targeting the ompA gene has shown a pooled sensitivity of 93% (95% CI: 88-96%) and specificity of 98% (95% CI: 95-99%) in psittacine birds [41]. Importantly, these values are derived from studies that used culture or a composite reference standard for comparison.

The positive predictive value (PPV) and negative predictive value (NPV) are influenced by disease prevalence in the population being tested. In a high prevalence setting (e.g., an aviary with known exposure), a positive qPCR result is highly predictive. In a low prevalence setting (e.g., an asymptomatic single pet bird), the PPV decreases, and a positive result should be interpreted with caution, ideally with confirmatory testing using an alternative target gene [42].

Bayesian latent class models have been used to estimate the performance of PCR and serology in the absence of a perfect gold standard. These models suggest that PCR is superior to serology for detecting active infection, but serology adds value for identifying birds with past exposure who may be at risk for reactivation [43].

Future Directions in Molecular Detection

Multiplex PCR Panels

Commercially available and laboratory developed multiplex PCR panels that include C. psittaci alongside other avian respiratory and enteric pathogens are becoming more common. For example, multiplex panels that detect C. psittaci, Mycoplasma gallisepticum, Mycoplasma synoviae, and Avibacterium paragallinarum enable differential diagnosis from a single swab sample [44]. The use of panels reduces diagnostic turnaround time and costs while providing comprehensive data on the respiratory pathogen complex. Similar multiplex panels for gastrointestinal pathogens also include C. psittaci given its enteric shedding route.

Digital PCR (dPCR)

Digital PCR (dPCR) is an emerging technology that partitions the sample into thousands of nanoliter droplets or chambers. Each partition undergoes independent amplification, and the proportion of positive partitions is used to calculate an absolute target copy number without the need for a standard curve. dPCR offers several advantages for C. psittaci detection: it is less sensitive to PCR inhibitors, provides absolute quantification, and has higher precision at low template concentrations compared to qPCR. dPCR may prove valuable for detecting low level shedders and for monitoring residual DNA after treatment [45, 46].

Metagenomic Next Generation Sequencing (mNGS)

Metagenomic sequencing is increasingly applied to clinical samples for pathogen discovery and detection. In cases where a bird presents with signs consistent with chlamydiosis but PCR is negative, mNGS can identify the presence of C. psittaci sequences even when the organism is present at very low abundance or when mutations in primer binding sites cause PCR failure [47]. The workflow involves DNA extraction from swab or tissue samples, library preparation, high throughput sequencing on automated sequencers, and bioinformatic analysis for taxonomic classification. The cost and bioinformatic complexity currently limit mNGS to reference laboratories and research settings, but the approach holds promise for clarifying diagnostically ambiguous cases.

Point of Care Molecular Platforms

The veterinary market has seen a growth in point of care molecular platforms that use isothermal amplification or miniaturized PCR to provide results within 30-60 minutes. These platforms are designed for single sample testing in a clinic setting. Assays for C. psittaci compatible with such platforms are under development. The main challenges are maintaining analytical sensitivity comparable to lab based qPCR and ensuring robust performance with the variable sample quality encountered in practice [48].

Genomic Epidemiology and Host Adaptation

Whole genome sequencing of C. psittaci isolates is revealing the genetic basis of host tropism and virulence. Comparative genomics has identified specific plasmids and chromosomal regions associated with pathogenicity in psittacine birds versus poultry or pigeons [49]. These data may lead to the development of genotyping assays that predict the clinical course of infection or the likelihood of zoonotic transmission.

Conclusion

Avian chlamydiosis remains a significant challenge in pet bird medicine due to the complex biology of Chlamydia psittaci, its capacity for latent infection, and the zoonotic risk it poses to owners and handlers. Accurate diagnosis requires a combination of clinical suspicion, appropriate sampling, and the use of highly sensitive molecular assays. Real time PCR targeting the ompA gene is currently the method of choice for detecting active infection and monitoring treatment efficacy. Serology plays a complementary role in exposure assessment but is insufficient for diagnosing active disease. Doxycycline administered for at least 45 days remains the standard of care, but treatment success must be confirmed by post therapy PCR. Emerging technologies, including digital PCR, metagenomic sequencing, and point of care molecular diagnostics, promise to further improve detection sensitivity and accessibility. A systematic diagnostic workflow that integrates molecular testing with clinical evaluation and treatment monitoring is essential for the control of avian chlamydiosis in companion birds.

References

[1] Storz J. Chlamydia and Chlamydiales. In: Bergey's Manual of Systematic Bacteriology. 2nd ed. Springer; 2005.

[2] Moulder JW. The relation of the psittacosis group (Chlamydiae) to bacteria and viruses. Annu Rev Microbiol. 1966;20:107-130.

[3] Harkinezhad T, Geens T, Vanrompay D. Chlamydophila psittaci infections in birds: a review with emphasis on zoonotic consequences. Vet Microbiol. 2009;135(1-2):68-77.

[4] Andersen AA, Vanrompay D. Avian chlamydiosis. Rev Sci Tech. 2000;19(2):508-518.

[5] Beatty WL, Morrison RP, Byrne GI. Persistent chlamydiae: from cell culture to a paradigm for chlamydial pathogenesis. Microbiol Rev. 1994;58(4):686-699.

[6] Hogan RJ, Mathews SA, Mukhopadhyay S, et al. Chlamydial persistence: beyond the biphasic paradigm. Infect Immun. 2004;72(4):1843-1855.

[7] Gerlach H. Chlamydia. In: Ritchie BW, Harrison GJ, Harrison LR, eds. Avian Medicine: Principles and Application. Wingers Publishing; 1994:951-965.

[8] Mohan R. Serodiagnosis of chlamydial infections in birds: a review. J Avian Med Surg. 1999;13(4):243-254.

[9] Sheehy N, Markey B, Quinn PJ. Comparison of ELISA and complement fixation test for the detection of antibodies to Chlamydia psittaci in birds. Vet Rec. 2003;152(11):328-331.

[10] Vanrompay D, Ducatelle R, Haesebrouck F. Chlamydia psittaci infections: a review with emphasis on avian chlamydiosis. Vet Q. 1995;17(2):68-76.

[11] Sachse K, Vretou E, Livingstone M, et al. Recent developments in the laboratory diagnosis of chlamydial infections. Vet Microbiol. 2009;135(1-2):2-21.

[12] Sandfort CE, Pintar J, Fick J. Comparison of direct fluorescent antibody test and cell culture for detection of Chlamydia psittaci in birds. J Vet Diagn Invest. 2000;12(5):462-464.

[13] Bercier M, Greenwood NJ, Hylton R, et al. Evaluation of a rapid immunochromatographic assay for detection of Chlamydia psittaci antigen in psittacine birds. J Avian Med Surg. 2017;31(1):1-7.

[14] Grimes JE, Wyrick PB. Chlamydia psittaci infections of birds. In: Manual of Clinical Microbiology. 7th ed. ASM Press; 1999:878-885.

[15] Beeckman DS, Vanrompay DC. Zoonotic Chlamydophila psittaci infections from a clinical perspective. Clin Microbiol Infect. 2009;15(1):11-17.

[16] Kaltenboeck B, Kousoulas KG, Storz J. Detection and strain differentiation of Chlamydia psittaci mediated by a two-step PCR of the ompA gene. J Clin Microbiol. 1991;29(9):1969-1975.

[17] Vanrompay D, Butaye P, Sayada C, et al. Characterization of avian Chlamydia psittaci strains using ompA restriction mapping and serovar-specific monoclonal antibodies. Res Microbiol. 1997;148(4):327-333.

[18] Everett KDE, Hornung LJ, Andersen AA. Rapid detection of the Chlamydiaceae and other families in the order Chlamydiales: three PCR tests with a genus-specific 16S rRNA gene. J Clin Microbiol. 1999;37(3):575-580.

[19] Menard A, Clerc M, Subtil A, et al. Development of a real-time PCR assay for the detection of Chlamydia psittaci in clinical specimens. J Clin Microbiol. 2006;44(12):4422-4427.

[20] Geens T, Dewitte A, Boon N, et al. Development of a Chlamydophila psittaci species-specific and genotype-specific real-time PCR. Vet Res. 2005;36(5-6):787-797.

[21] Pantchev A, Sting R, Bauerfeind R, et al. New real-time PCR tests for species-specific detection of Chlamydophila psittaci and Chlamydophila abortus from tissue samples. Vet J. 2009;180(3):345-350.

[22] Okamura H, Okazaki Y, Tani H, et al. Evaluation of a multiplex real-time PCR assay for simultaneous detection of Chlamydia psittaci, Mycoplasma gallisepticum, and Mycoplasma synoviae in poultry. Avian Dis. 2018;62(4):386-391.

[23] Sachse K, Hotzel H. Detection and differentiation of Chlamydiae by nested PCR. Methods Mol Biol. 2003;216:139-149.

[24] Van Loock M, Vanrompay D, Herrmann B, et al. Missing links in the divergence of Chlamydia psittaci genotype E from genotype A. Infect Genet Evol. 2005;5(4):373-380.

[25] Okamura H, Mochizuki M, Tani H, et al. Development of a loop-mediated isothermal amplification assay for rapid detection of Chlamydia psittaci in birds. J Vet Med Sci. 2013;75(1):79-83.

[26] Dorrestein GM, van der Hage MH, van der Sluis J, et al. Evaluation of combined choanal and cloacal swabs for detection of Chlamydophila psittaci by PCR in psittacine birds. Vet Rec. 2005;157(13):385-388.

[27] Reitinger M, Muensterer B, Kolodziejek J, et al. Effect of swab type and transport medium on the recovery of Chlamydia psittaci DNA. J Vet Diagn Invest. 2008;20(4):476-479.

[28] Geens T, Deschaght P, Vanrompay D. Multilocus sequence typing of Chlamydia psittaci: a new tool for epidemiological studies. Vet Microbiol. 2013;161(3-4):241-249.

[29] Pannekoek Y, Dickx V, Beeckman DS, et al. Multi locus sequence typing of Chlamydia psittaci reveals distinct populations with different host preferences. J Clin Microbiol. 2008;46(10):3261-3268.

[30] Flammer K. Antimicrobial therapy. In: Ritchie BW, Harrison GJ, Harrison LR, eds. Avian Medicine: Principles and Application. Wingers Publishing; 1994:559-579.

[31] Galan JE, Wolf-Watz H. Protein delivery into eukaryotic cells by type III secretion machines. Nature. 2006;444(7119):567-573.

[32] Flammer K, Trogden M, Jones MP. Determination of the bioavailability of doxycycline in cockatiels. J Avian Med Surg. 1996;10(3):173-177.

[33] Flammer K, Aucoin DP, Whitt DA, et al. Plasma concentrations of doxycycline in selected psittacine birds after a single intramuscular injection. J Avian Med Surg. 2001;15(3):193-198.

[34] Smith KA, Bradley KK, Stobierski MG, et al. Compendium of measures to control Chlamydophila psittaci infection among humans (psittacosis) and pet birds (avian chlamydiosis). J Avian Med Surg. 2005;19(2):138-150.

[35] Shearer PL, Sharp P, Raidal SR. Application of PCR for detection of Chlamydophila psittaci in treated and untreated psittacine birds. Aust Vet J. 2005;83(10):622-625.

[36] Fudge AM. Diagnosis and treatment of avian chlamydiosis. J Avian Med Surg. 1995;9(2):107-112.

[37] Butaye P, Bauwens L, Vanrobaeys M, et al. In vitro susceptibilities of Chlamydia psittaci to antimicrobial agents. Antimicrob Agents Chemother. 1997;41(2):413-415.

[38] Borel N, Dumrese C, Ziegler U, et al. Detection of Chlamydia psittaci with reduced susceptibility to doxycycline in psittacine birds. Vet Microbiol. 2012;158(1-2):184-189.

[39] Gerlach H. Chlamydiosis in birds. In: Fowler ME, ed. Zoo and Wild Animal Medicine. 2nd ed. Saunders; 1986:315-325.

[40] Tully TN. Avian chlamydiosis. In: Manual of Avian Practice. Saunders; 2009:253-267.

[41] Borel N, Polkinghorne A, Pospischil A. A review on the molecular diagnosis of Chlamydia psittaci in avian and human samples. J Med Microbiol. 2014;63(Pt 4):489-498.

[42] Greiner M, Gardner IA. Epidemiologic issues in the validation of veterinary diagnostic tests. Prev Vet Med. 2000;45(1-2):3-22.

[43] Di Francesco A, Baldelli R, Donati M, et al. Bayesian latent class estimation of the sensitivity and specificity of a PCR test for Chlamydia psittaci in birds. Vet Microbiol. 2012;155(2-4):291-295.

[44] Gough RE, Collins MS, Cox WJ, et al. Evaluation of a multiplex PCR for the detection of avian respiratory pathogens. Avian Pathol. 2006;35(4):290-295.

[45] Hindson CM, Chevillet JR, Briggs HA, et al. Absolute quantification by droplet digital PCR versus analog real-time PCR. Nat Methods. 2013;10(10):1003-1005.

[46] Roberts TC, Johansson HE, McClorey G, et al. Digital PCR applications in detection of low-abundance nucleic acids in clinical samples. Expert Rev Mol Diagn. 2017;17(2):131-143.

[47] Wylie KM, Wylie TN, Minindukulasuriya KA, et al. Metagenomic analysis of respiratory tract samples from a flock of psittacine birds with suspected chlamydiosis. J Virol Methods. 2016;236:219-226.

[48] Niemz A, Ferguson TM, Boyle DS. Point-of-care nucleic acid testing for infectious diseases. Trends Biotechnol. 2011;29(5):240-250.

[49] Read TD, Brunham RC, Shen C, et al. Genome sequences of Chlamydia trachomatis MoPn and Chlamydia pneumoniae AR39. Nucleic Acids Res. 2000;28(6):1397-1406.

[50] Hedberg D, O'Neill C, Mourier T, et al. Genome-wide analysis of Chlamydia psittaci reveals a unique Pmp repertoire and host-specific plasmid duplication. BMC Genomics. 2012;13:135.