Mauersegler (Apus apus)
Foto: Nailia Schwarz -

Berliner und Münchener Tierärztliche Wochenschrift

Monitoring of the infectious agent Chlamydia psittaci in common swifts (Apus apus) in the area of Hannover, Lower Saxony, Germany


Untersuchungen zum Vorkommen von Chlamydia psittaci bei Mauerseglern (Apus apus) in der Region Hannover, Niedersachsen, Deutschland

Berliner und Münchener Tierärztliche Wochenschrift 134, 1-5

DOI: 10.2376/1439-0299-2020-32

Publiziert: 05/2021


The occurrence of the zoonotic pathogen Chlamydia (C.) psittaci in the common swift (Apus apus) with a high prevalence was reported in the literature. These long-distance migrant, which only consume aerial plankton, can reach high population densities in places with suitable breeding sites. Dedicated competent private individuals take part in the hand-rearing of juvenile common swifts in wildlife rescue centres, which unavoidably results in close contact with the avian patients. For this reason, we monitored the infectious agent C. psittaci in common swifts in the area of Hannover, Lower Saxony, Germany, from 2009 to 2018. Pooled organ samples (liver, spleen, kidney, lungs, trachea) of fatally injured swifts (n = 243) were examined using real-time PCR. In conclusion, specific sequences of C. psittaci (examined in the years 2009–2018) as well as Chlamydia spp. (examined in the years 2016–2018) could not be detected in any common swift within the investigation period and the studied area. These results show that the common swift is unlikely to be a reservoir for the zoonotic agent C. psittaci in the Hannover area.

wild birds


Das Vorkommen des Zoonoseerregers Chlamydia (C.) psittaci bei Mauerseglern (Apus apus) mit hoher Prävalenz ist in der Literatur beschrieben. Der Langstreckenzieher Mauersegler, der sich ausschließlich von Luftplankton ernährt, kann in Gebieten mit geeigneten Brutplätzen hohe Bestandsdichten in Deutschland erreichen. In Wildvogelpflegestationen übernehmen auch engagierte, fachkundige Privatpersonen die Aufzucht juveniler Mauersegler mit einem unvermeidlichen engen Kontakt zum Pflegling.

Aus diesem Grund sollte das Vorkommen von C. psittaci bei Mauerseglern untersucht werden. Hierfür wurden in den Jahren 2009 bis 2018 gepoolte Organproben (Leber, Milz, Niere, Lunge, Trachea) tödlich verunfallter Mauersegler (n = 243) aus dem Raum Hannover, Niedersachsen, mithilfe von real-time PCR untersucht. 
Im gesamten Untersuchungszeitraum konnten keine spezifischen Sequenzen von C. psittaci (untersucht in den Jahren 2009 bis 2018) sowie von Chlamydia spp. (untersucht in den Jahren 2016 bis 2018) bei Mauerseglern im Raum Hannover, Niedersachsen, nachgewiesen werden. Somit ist es unwahrscheinlich, dass der Mauersegler ein Reservoir für den Zoonoseerreger C. psittaci im Untersuchungsgebiet Hannover ist.



The common swift (Apus apus), a long-distance migrant, is the most frequently encountered species of the order Apodiformes in Germany. This swift has been known to travel across Europe to winter in sub-Saharan Africa from autumn to spring, and to spend over 10 months of the year flying (Åkesson et al. 2012, Hedenström et al. 2016, Holmgren 2004, Muijres et al. 2012, Rattenborg et al. 2016, Weitnauer and Scherner 1980, Wellbrock et al. 2017).

From April to September, swifts make up large populations in urban areas in Germany (Gedeon et al. 2014, Tigges 2006). In the region of Hannover, the population was estimated to consist of 700 to 900 breeding pairs in 2006 (Wendt 2006). The German population of common swifts is estimated at 215,000 to 395,000 breeding pairs (Gedeon et al. 2014). In extreme weather situations, such as hot spells or prolonged periods of cold, hundreds of swifts, especially juveniles, are submitted to wildlife rescue centres for rehabilitation (Haupt 2009). Dedicated competent private individuals take part in the hand-rearing of juvenile swifts, and close contact between the patient swifts and humans is unfortunately unavoidable (Haupt 2009, Matthes 2006). 

The specific role of common swifts as a source of zoonotic diseases remains unknown. Gerbermann et al. (1994) detected Chlamydia (C.) psittaci in 7 out of 28 tested common swifts in Munich, Germany. In other studies, C. psittaci could not be detected in common swifts (Krawiec et al. 2015, Legler et al. 2011). This Gram-negative and obligate intracellular bacterium is the causative agent of chlamydiosis in birds and an important zoonotic agent (Rohde et al. 2010). 

C. psittaci can be detected in pet parrots, poultry and racing pigeons, as well as in different wild bird species (Andersen and Franson 2007, Beckmann et al. 2014, Kaleta and Taday 2003, Teske et al. 2013, Zweifel et al. 2009). Chlamydiosis in birds can take markedly different courses depending on the virulence of the C. psittaci strain and the immune status of the avian host. In wild birds, subclinical or inapparent courses are often described. However, acute to chronic respiratory and gastrointestinal diseases with spleno- and hepatomegaly can also occur (Vanrompay et al. 1995). Transmission of C. psittaci from birds to humans takes place through contaminated excretions and secretions direct and indirect in the form of aerosols (Balsamo et al. 2017). C. psittaci can cause in humans a disease known as ornithosis, psittacosis or parrot fever. The symptoms of ornithosis range from inapparent to systemic influenza-like illness with severe pneumonia and nonrespiratory health problems. Rarely, ornithosis can result in death (Balsamo et al. 2017, Petrovay and Balla 2008, Rohde et al. 2010). The literature shows that in most cases in humans, contact with pet birds, wild birds or poultry is decisive for the outbreak of ornithosis (Branley et al. 2008, Dickx et al. 2010, Hafez 2011, Heddema et al. 2006, Kalmar et al. 2014, Lagae et al. 2014, Petrovay and Balla 2008, Tiong et al. 2007, Van Droogenbroeck et al. 2009). 

In this study, we monitored the occurrence of C. psittaci in injured adult and juvenile common swifts in the area of Hannover, Lower Saxony, Germany, over 9 years to provide more information about the infection status of swifts. 

Materials and Methods


From 2009 to 2018, a total of 243 common swifts from the Hannover area were sampled in the period from April to September for the detection of Chlamydiaceae (Table 1). The total number of examined swifts consisted of 123 adults (>2 years) and 120 juvenile or fledged swifts (<5 month; Table 1). The age of the swifts was ascertained from the typical colour and shape of the feathers and the moulting pattern, with a completed feather growth in adult birds (Weitnauer and Scherner 1980). Immature birds in the second year can be distinguished from adult birds by the shape and colour of the flight feathers and from fledglings in their first year by the colour of the flight feathers (bleached feathers in the second year). However, birds in their second year were not seen in this study.
All the swifts were handed in by concerned people at the Clinic for Small Mammals, Reptiles and Birds, University of Veterinary Medicine Hannover, Foundation, after they had been grounded and injured. The birds had to be euthanised after incurable injury, severe fractures in most cases, without the possibility of successful rehabilitation (Haupt 2009). 


Each dead common swift was stored at –20°C and then thawed at room temperature for a full necropsy. Samples of lungs, trachea, kidney, liver and spleen, each approximately 20 mg, were taken from each bird using sterile equipment. The organs of each bird were pooled into a microtube with 0.05 mL 0.9% sodium chloride injection solution and frozen at –70°C until further examination. 

DNA extraction

Total DNA was obtained from the pooled organ samples using the illustra™ Tissue & Cells genomicPrep Mini Spin Kit (GE Healthcare Life Sciences, Freiburg, Germany) in the years 2009–2015 and the Kylt® RNA/DNA Purification Kit (AniCon Labor GmbH, Hoeltinghausen, Germany) in the years 2016–2018, following the manufacturer´s instructions. The DNA samples were stored at –20°C until further examination. 

PCR in the years 2009–2015

The DNA samples were screened for C. psittaci by a species-specific PCR. Previously published primers (CACTATGTGGGAAGGTGCTTCA, CTGCGCGGATGCTAATGG) and probe (FAM-CGCTACTTGGTGTGAC-TAMRA; FAM: 6-carboxyfluorescein; TAMRA: carboxytetramethylrhodamin) were used to detect the ompA gene of C. psittaci (Pantchev et al. 2009, 2010). The real-time PCR was modified by introducing a suitable internal control system by using a HEX-labelled probe (Hoffmann et al. 2006). The Ambion® Path-ID™ qPCR Master Mix (Life Technologies, Carlsbad, California, USA) was used with a final volume of 25 µL reaction mixture, containing 12.5 µL of reaction buffer (2X qPCR Master Mix), and 2 µL of template DNA. A concentration of 0.6 µM of forward primer, 1 µM of reverse primer and 0.3 µM of probe were used. Nuclease-free water was added to this mixture to reach the final volume. Additionally, a positive amplification control was included, containing 0.3 µM of each primer, 0.2 µM of probe and 0.5 µL of internal control DNA (intype IC-DNA, QIAGEN Leipzig, Leipzig, Germany). Real-time PCR was conducted using the Stratagene MX 3005P detection system (Stratagene, La Jolla, California, USA) with the following set-up: initial denaturation for 10 min at 95°C, followed by 45 cycles of denaturation at 95°C for 15 sec, and annealing with elongation at 60°C for 1 min. A sample was considered positive for C. psittaci if the FAM-curve was positive (10 < FAM-cycle threshold (CT)-value <35) independent of the HEX-curve; negative if the HEX-curve was positive (10 < HEX-CT-value ≤40) but the FAM-curve was negative, and inhibited if neither the FAM-curve nor the HEX-curve was positive (FAM-CT-value ≥35, HEX-CT-value >40). All samples were tested in duplicate.

PCR in the years 2016–2018

The samples in the years 2016–2018 were screened for C. spp. by the Kylt® Chlamydiaceae Screening Real-Time PCR Detection Kit (AniCon Labor GmbH, Hoeltinghausen, Germany). This PCR is family-specific and according to the manufacturer, detects C. psittaci, C. avium and C. gallinaceae, among others. A final volume of 20 µL reaction mixture, containing 16 µL of total Master-Mix (10 µL BCD 2X qPCR-Mix, 6 µL Detection-Mix) and 4 µL of sample DNA (diluted DNA with DNase-free water; Fisher BioReagents®) was used. This particular PCR assay was also run on the Stratagene MX 3005P detection system with initial denaturation for 10 min at 95°C, followed by 42 cycles of denaturation at 95°C for 15 sec and annealing with elongation at 60°C for 1 min. The target genes for Chlamydiaceae in the samples and the positive control included in the commercial kit were amplified and then detected via fluorescently labelled probes (dyes FAM: Chlamydiaceae-specific status and HEX: hexachlorofluorescein: internal control). A sample was considered positive for Chlamydiaceae if the FAM-curve was positive (10 < FAM-CT-value ≤42) independent of the HEX-curve; negative if the HEX-curve was positive (10 < HEX-CT-value ≤40) but the FAM-curve was negative; and inhibited if neither the FAM-curve nor the HEX-curve was positive (FAM-CT-value >42, HEX-CT-value >40). All samples were tested in duplicate.

Results and Discussion

During the investigation period of 9 years, specific sequences of C. psittaci (2009–2018), as well as Chlamydia spp. (2016–2018), could not be detected in any common swift (0 out of 243 sampled swifts) in the described study area of Germany. Conjunctivitis, upper respiratory disease, hepatomegaly and splenomegaly, important findings in chlamydiosis (Vanrompay et al. 1995), could not be detected in the pathological macroscopic examinations of the dissected swifts. 
A study of the prevalence of C. psittaci in wild birds in Poland supports our negative findings in common swifts (Krawiec et al. 2015). The negative results of our study on necropsied swifts are also supported by the examination of swift patients in surveys of different wildlife rehabilitation centres in Hesse and Lower Saxony, Germany. Over a period of two years, specific sequences of C. psittaci were not detected in pooled choanal and cloacal swabs of clinical examined 74 swifts (5 adult; 69 juvenile) using PCR (Legler et al. 2011).

There is no information in the literature about C. psittaci infection with clinical symptoms in common swifts. The positively tested animals from the Munich area showed no signs of chlamydiosis in the clinical and pathological examination (Gerbermann et al. 1994). Subclinical cases can result in a particularly high risk for contact persons, since the risk of infection is not recognised. Shedding of chlamydiae occurs in diseased and asymptomatic infected birds and can be activated by stressful events such as captivity, handling or illness (Knitter and Sachse 2015).

A case report of chlamydia infection in a captive colony of Amazilia Emerald Hummingbirds (Amazilia amazilia) shows the susceptibility of birds of the family Trochilidae, near relatives of the swifts (Apodidae). In this report, 90% of the colony died from emaciation, hepatomegaly and splenomegaly (Meteryer et al. 1992). 

Possible contact between swifts and infected wild birds could occur during the nesting period, when swifts fight with European starlings or house sparrows for suitable breeding sites, and sometimes use the same nesting sites (Kruszewicz and Pasowska 1992). The experimental transmission of C. psittaci from infected starlings to healthy turkeys was reported (Grimes et al. 1979). Surface water contaminated with faeces of infected birds and contaminated food animals could also be a source of infection (Thierry et al. 2016). Possible explanations as to why common swifts are mainly negative could be a pronounced and peracute or acute pathology of chlamydiosis in these birds, resulting in rapid death without clinical signs, and therefore missing any indication for a clinical admission. This assumption contradicts the results of Gerbermann et al. (1994), who did not detect pathomorphological findings of chlamydiosis in post-mortem examinations of chlamydia-positive swifts. However, an acute or chronic chlamydiosis with or without respiratory signs may be an important selection factor in a highly aerial bird and long-distance migrant in the wild. Chlamydial infections may act as a predisposing factor for secondary infections or may debilitate the organism too much to cope with migration. On the other hand, it is possible that swifts are not susceptible to infection with chlamydiae and, thus, do not develop clinical disease. 

For the detection of Chlamydia spp., various test methods with different advantages and disadvantages are available. Real-time PCR is recommended to confirme clinical cases and to investigate the prevalence of chlamydia infections (Sachse et al. 2009, Schnee et al. 2019). The real-time PCRs used in our study have already been successfully used to detect C. psittaci in different birds (Pantchev et al. 2009, 2010). However, a recent study showed that the species-specific PCR did not detect all ompA genotypes found in bird species, and a family-specific PCR showed better results (Stalder et al. 2020). For further investigation of possible chlamydia infections in swifts, serological tests could also be helpful (Grimes 1989, Schnee et al. 2019, Vanrompay 2013). 

In conclusion, our study results indicate that the common swift is unlikely to be a reservoir for C. psittaci in the Hannover area. However, humans who have contact with wild birds, and thus have an increased risk of contracting ornithosis, should protect themselves through appropriate measures.


C. psittaci: Chlamydia psittaci; FAM: 6-carboxyfluorescein; HEX: hexachlorofluorescein; TAMRA: carboxytetramethylrhodamin


The authors would like to thank the veterinarians and staff of the Clinic for Small Mammals, Reptiles and Birds, University of Veterinary Medicine Hannover, Foundation, for their help during sample collection. We are grateful to Dr. Martin Ryll, who gave access to laboratory facilities. Special thanks go to Annemarie Grund and Sandra Hartmann for their cooperation and laboratory guidance. We wish to show our appreciation to Florian Werner (Sales & Tech Support Kylt) for providing the PCR reagents. We are in debt to Dr. Florian Brandes for supporting the examinations and Ms. Sherwood-Brock for the critical revision of the manuscript. Our thanks also go to Dr. Norbert Kummerfeld for scientific support.

Ethical Approval

This study was conducted according to the current German and European animal welfare legislation and did not involve any animal experiments or interventions. Data collection was based on the analysis of dead animal carcasses as part of a veterinary investigation into the prevalence of zoonotic disease in wild animals.

Conflict of Interest

The authors confirm that there is no conflict of interest.


This publication was supported by Deutsche Forschungs­gemeinschaft and University of Veterinary Medicine Hannover, Foundation within the funding programme Open Access Publishing.

Authors Contribution

Conception: ML; sample collection: WT, EG, ML; sample examination WT, RL, LM; data interpretation: WT, RL, LM, ML; drafting the article: WT; critical revision of the article: RL, LM, ML, final approval of the version to be published: WT, EG, RL, LM, ML.

Address for correspondence

Dr. Marko Legler
Clinic for Small Mammals, Reptiles and Birds
University of Veterinary Medicine Hannover, Foundation
Bünteweg 9
30559 Hannover 


Åkesson S, Klaassen RR, Holmgren J, Fox JW, Hedenström A (2012): Migration routes and strategies in a highly aerial migrant, the common swift Apus apus, revealed by light-level Geolocators. PLoS One 7: 1–9.
Andersen AA, Franson JC (2007): Avian chlamydiosis. In: Nancy JT, Hunter DB, Atkinson CT (eds.), Infectious diseases of wild birds. Wiley-Blackwell Publishing, Iowa, 303–316.
Balsamo G, Maxted AM, Midla JW, Murphy JM, Wohrle R, Edling TM, Fish PH, Flammer K, Hyde D, Kutty PK, Kobayashi M, Helm B, Oiulfstad B, Ritchie BW, Stobierski MG, Ehnert K, Tully TN Jr (2017): Compendium of measures to control chlamydia psittaci infection among humans (psittacosis) and pet birds (avian chlamydiosis). J Avian Med Surg 31: 262–282.
Beckmann KM, Borel N, Pocknell AM, Dagleish MP, Sachse K, John SK, Pospischil A, Cunningham AA, Lawson B (2014): Chlamydiosis in British garden birds (2005–2011): Retrospective diagnosis and Chlamydia psittaci genotype determination. Ecohealth 11: 544–563.
Branley J, Roy B, Dwyer D, Sorrell T (2008): Real-time PCR detection and quantitation of Chlamydophila psittaci in human and avian specimens from a veterinary clinic cluster. Eur J Clin Microbiol Infect Dis 27: 269–273.
Dickx V, Geens T, Deschuyffeleer T, Tyberghien L, Harkinezhad T, Beeckman DSA, Braeckman L, Vanrompay D (2010): Chlamydophila psittaci zoonotic risk assessment in a chicken and turkey slaughterhouse. J Clin Microbiol 48: 3244–3250.
Gedeon K, Grüneberg C, Mitschke A, Sudfeldt C, Eickhorst W, Fischer S. Flade M, Frick S, Geiersberger I, Koop B, Kramer M, Krüger T, Roth N, Ryslavy T, Stübing S, Sudmann SR, Steffens R, Vökler F, Witt K (2014): Atlas Deutscher Brutvogelarten, Atlas of German Breeding Birds. Stiftung Vogelmonitoring Deutschland und Dachverband Deutscher Avifaunisten, Münster.
Gerbermann H, Korbel R, Kösters J (1994): Zum Vorkommen von Chlamydia psittaci-Infektionen bei verschiedenen einheimischen Wildvögeln. Proceedings of the 9th DVG-Tagung über Vogelkrankheiten, München, Germany, March 3–4, 1994, 130–142. 
Grimes JE (1989): Serodiagnosis of avian chlamydia infections. J Am Vet Med Assoc 195: 1561–1563.
Grimes JE, Owens KJ, Singer JR (1979): Experimental transmission of Chlamydia psittaci to turkeys from wild birds. Avian Dis 37: 817–824.
Hafez HM (2011): Chlamydiose (Psittakose/Ornithose). In: Kaleta EF, Krautwald-Junghanns M-E (Hrsg.), Kompendium der Ziervogelkrankheiten. 4. Aufl. Schlütersche, Hannover, 223–229.
Haupt C (2009): Radiologische Diagnostik am Mauersegler (Apus apus Linnaeus, 1758): Anatomie und Pathologie des Skeletts und ein Beitrag zur tierärztlichen Therapie und Prognose. Gießen, Justus-Liebig-Universität, Faculty of Veterinary Medicine, Diss.
Heddema ER, van Hannen EJ, Duim B, de Jongh BM, Kaan JA, van Kessel R, Lumeij JT, Visser CE, Vandenbroucke-Grauls CM (2006): An outbreak of psittacosis due to Chlamydophila psittaci genotype A in a veterinary teaching hospital. J Med Microbiol 55: 1571–1575.
Hedenström A, Norevik G, Warfvinge K, Andersson A, Bäckman J, Åkesson S (2016): Annual 10-Month Aerial Life Phase in the common swift Apus apus. Curr Biol 26: 3066–3070.
Hoffmann B, Depner K, Schirrmeier H, Beer M (2006): A universal heterologous internal control system for duplex real-time RT-PCR assays used in a detection system for pestiviruses. J Virol Methods 136: 200–209.
Holmgren J (2004): Roosting in tree foliage by common swift Apus apus. Ibis 146: 404–416. 
Kaleta EF, Taday EMA (2003): Avian host range of Chlamydophila spp. based on isolation, antigen detection and serology. Avian Pathol 32: 435–462.
Kalmar ID, Dicxk V, Dossche L, Vanrompay D (2014): Zoonotic infection with Chlamydia psittaci at an avian refuge centre. Vet J: 300–302.
Knittler MR, Sachse K (2015) Chlamydia psittaci: Update on an underestimated zoonotic agent. Pathog Dis 73: 1–15.
Krawiec M, Piasecki T, Wieliczko A (2015): Prevalence of Chlamydia psittaci and other Chlamydia species in wild birds in Poland. Vector-Borne Zoonotic Dis 15: 652–655.
Kruszewicz A, Pasowska K (1992): Occurrence of Chlamydia psittaci in House Sparrows. Proceedings of the 8th DVG-Tagung über Vogelkrankheiten, München, Germany, March 5–6, 1992, 124–129.
Lagae S, Kalmar I, Laroucau K, Vorimore F, Vanrompay D (2014): Emerging Chlamydia psittaci infections in chickens and examination of transmission to humans. J Med Microbiol 63: 399–407.
Legler M, Ryll M, Hartmann S, Haupt C, Brandes F, Kummerfeld N (2011): Kommen Mauersegler als Überträger der Chlamydiose in Frage? Zur Prävalenz von Chlamydophila psittaci bei Mauerseglern (Apus apus) in Niedersachsen und Hessen in den Jahren 2009–2010. Proceedings 2nd DVG-Tagung Vogel- und Reptilienkrankheiten, Hannover, Germany, September 16–18, 2011, 272–275.
Matthes H (2006): Recovery of a hand-reared common swift (Apus apus). APUSlife 3035, ISSN: 1438–2261.
Meteryer CU, Chin RP, Castro AE, Woods LW, Gentzler RP (1992): An epizootic of chlamydiosis with high mortality in a captive population of Euphonias (Euphonia violacea) and hummingbirds (Amazilia amazilia). J Zoo Wild Med 23: 222–229.
Muijres FT, Henningsson P, Stuiver M, Hendenström A (2012): Aerodynamic flight performance in flap-gliding birds and bats. J Theor Biol 306: 120–128. 
Pantchev A, Sting R, Tyczka J, Bauerfeind R, Sachse K. (2009): New real-time PCR tests for species-specific detection of Chlamydophila psittaci and Chlamydophila abortus from tissue samples. Vet J 181: 145–150.
Pantchev A, Sting R, Bauerfeind R, Tyczka J, Sachse K (2010): Detection of all Chlamydophila and Chlamydia spp. of veterinary interest using species-specific real-time PCR assays. Comp Immunol Microbiol Infect Dis 33: 473–484.
Petrovay F, Balla E (2008): Two fatal cases of psittacosis caused by Chlamydophila psittaci. J Med Microbiol 57: 1296–1298.
Rattenborg NC, Voirin B, Cruz SM, Tisdale R, Dell’Omo G, Lipp H-P, Wikelski M, Vyssotski AL (2016): Evidence that birds sleep in mid-flight. Nat Commun 7: 1–9.
Rohde G, Straube E, Essig A, Reinhold P, Sachse K (2010): Chlamydiale Zoonosen. Dtsch Ärztebl Int 107: 174–184.
Sachse K, Vretou E, Livingstone M, Borel N, Pospischil A, Longbottom D (2009): Recent developments in the laboratory diagnosis of chlamydial infections (review). Vet Microbiol 135: 2–21.
Schnee C, Vanrompay D, Laroucau K (2019) Avian Chlamydiosis. In: World Organization for Animal Health (OIE) (ed.), OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals. OIE, Paris, 783–795.
Stalder S, Marti H, Borel N, Sachse K, Albini S, Vogler BR (2020): Occurrence of Chlamydiaceae in Raptors and Crows in Switzerland. Pathogens 9: 724.
Teske L, Ryll M, Rubbenstroth D, Hänel I, Hartmann M, Kreienbrock I, Rautenschlein S (2013): Epidemiological investigations on the possible risk of distribution of zoonotic bacterial through apparently healthy homing pigeons. Avian Pathol 42: 397–407.
Thierry S, Vorimore F, Rossignol C, Scharf S, Sachse K, Berthon P, Durand B, Virlogeux-Payant I, Borel N, Laroucau K (2016): Oral uptake of Chlamydia psittaci by ducklings results in systemic dissemination. PLoS One 11: Doi:10.1371/journal.pone.0154860.
Tigges U (2006): The breeding cycle in calendar form of the common swift Apus apus across its Eurasian breeding range – A testable hypothesis? Podoces 1: 27–33.
Tiong A, Vu T, Counahan M, Leydon J, Tallis G, Lambert S (2007): Multiple sites of exposure in an outbreak of ornithosis in workers at a poultry abattoir and farm. Epidemiol Infect 135: 1184–1191.
Van Droogenbroeck C, Beeckman DS, Verminnen K, Marien M, Nauwynck H, de Boesinghe LT, Vanrompay D (2009): Simultaneous zoonotic transmission of Chlamydophila psittaci genotypes D, F and E/B to a veterinary scientist. Vet Microbiol 135: 78–81.
Vanrompay D (2013): Avian chlamydiosis. In: Swayne D, Glisson JR, McDougald LR, Nolan LK, Suarez DL, Nair VL, (eds.), Diseases of Poultry. 13th ed. Wiley-Blackwell Publishing, Iowa, 1055–1073.
Vanrompay D, Ducatelle R, Haesebrouck F (1995): Chlamydia psittaci infections: a review with emphasis on avian chlamydiosis. Vet Microbiol 45: 93–119 
Weitnauer E, Scherner ER (1980): Mauersegler. In: Glutz von Blotzheim UN (Hrsg.), Handbuch der Vögel Mitteleuropas (Band 9). Akademische Verlagsgesellschaft, Wiesbaden, 669–712.
Wellbrock AHJ, Bauch C, Rozman J, Witte K (2017): “Same procedure as last year?” – Repeatedly tracked swifts show individual consistency in migration pattern in successive years. J Avian Biol 48: 897–903.
Wendt D (2006): Die Vögel der Stadt Hannover. BWH GmbH – Medien Kommunikation, Hannover.
Zweifel D, Hoop R, Sachse K, Pospischil A, and Borel N (2009): Prevalence of Chlamydophila psittaci in wild birds – potential risk for domestic poultry, pet birds, and public health. Europ J Wildl Res 55: 575–581.

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