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BACTERIOLOGY (N BOREL, SECTION EDITOR)
Avian Chlamydiosis
Konrad Sachse &Karine Laroucau &Daisy Vanrompay
Published online: 17 January 2015
#Springer International Publishing AG 2015
Abstract Recent findings in research on avian chlamydiosis
include an increase in the reported prevalence of Chlamydia
(C.)psittaci in poultry flocks, detailed descriptions of molec-
ular processes governing the course of infection in vivo, as
well as the discovery of new chlamydial species. Here we
review the major advances of the last 6 years. In particular, we
suggest that the observed re-emergence of C. psittaci infec-
tions in domestic poultry are due to a reduction in the use of
antibiotics and better diagnostic assays. Cellular and animal
models have significantly contributed to improving our un-
derstanding of the pathogenesis, including the events leading
to systemic disease. The elucidation of host–pathogen inter-
actions revealed the efficiency of C. psittaci in proliferating
and disseminating despite the action of pro-inflammatory
mediators and other factors during host immune response.
Finally, the recent introduction of C. avium and
C. gallinacea sheds new light on the epidemiology and
aetiopathology of avian chlamydiosis.
Keywords Avianchlamydiosis .Pathogenesis .Epidemiology .
Aetiology .Chlamydia psittaci .Chlamydia avium .Chlamydia
gallinacea
Introduction
Avian chlamydiosis, sometimes also referred to as psittacosis
or ornithosis, is an important infectious disease of companion
birds, especially psittacines, domestic poultry and wild birds.
The infection usually becomes systemic and is occasionally
fatal. Its main causative agent is the obligate intracellular
Gram-negative bacterium Chlamydia (C.) psittaci. The infec-
tion is widespread and, due to carcass condemnation at
slaughter, decrease in egg production, mortality and the ex-
pense of antibiotic treatment, represents a major factor of
economic loss in birds commercially raised for meat and egg
production [1], as well as posing a permanent risk for zoonotic
transmission to man [2••].
Concerning the taxonomic classification of the family
Chlamydiaceae, which now includes three members associat-
ed with avian hosts, it is important to note the recent return to
the single genus Chlamydia. Based on clustering analyses of
the 16S and 23S ribosomal RNA (rRNA) genes, Everett and
colleagues [3] had proposed a subdivision of the former single
genus Chlamydia into two genera: Chlamydia and
Chlamydophila. However, this taxonomic separation proved
consistent with neither the natural history of the organisms as
revealed by genome comparisons nor with the largely similar
morphology of all family members. Later on, comparative
genome and proteome analysis of chlamydial species sug-
gested that host-divergent strains of Chlamydiaceae are bio-
logically and ecologically closely related. Apart from this, the
This article is part of the Topical Collection on Bacteriology
K. Sachse (*)
Institute of Molecular Pathogenesis, Friedrich-Loeffler-Institut
(Federal Research Institute for Animal Health), Naumburger Str. 96a,
Jena 07743, Germany
e-mail: konrad.sachse@fli.bund.de
K. Laroucau
ANSES, Animal Health Laboratory, Bacterial Zoonoses Unit,
Paris-Est University, 23 avenue du général de Gaulle,
94706 Maisons-Alfort, France
e-mail: Karine.Laroucau@anses.fr
D. Vanrompay
Department of Molecular Biotechnology and Immunology, Faculty
of Bioscience Engineering, Ghent University, Ghent, Belgium
e-mail: Daisy.Vanrompay@UGent.be
Curr Clin Micro Rpt (2015) 2:10–21
DOI 10.1007/s40588-014-0010-y
two-genus nomenclature was not widely used by the chla-
mydia research community. Formal efforts to reunite the
members of Chlamydiaceae into a single genus Chlamydia
began about 6 years ago [4,5] and were finalised recently [6].
Aetiology and Epidemiology
The Causative Agent
As with any organism of the family Chlamydiaceae,
C. psittaci undergoes a characteristic biphasic developmental
cycle, during which it passes through three morphologically
distinct forms termed elementary body (EB), reticulate body
(RB) and intermediate body (IB) [7]. The EB is a small,
electron-dense, spherical body, about 0.2–0.3 μmindiameter,
which rivals mycoplasmas for the smallest of the prokaryotes.
The electron-dense EB is the infectious form of the organism
as it attaches to the target epithelial cell and gains entry. Inside
the host cell, the EB expands in size to form the RB, i.e. the
intracellular and metabolically active form. It islarger than the
EB, measuring approximately 0.5–2.0 μm in diameter. The
RBs divide by binary fission and thereafter re-transform into
EBs. During this maturation, morphologically intermediate
forms (IB) of 0.3–1.0 μm diameter with a central electron-
dense core and radially arranged individual nucleoid fibres
can be observed inside the host cell. An electron microscopy
micrograph illustrating the morphology of chlamydial bodies
is shown in Fig. 1.
In genetic and phenotypic terms, C. psittaci is a rather
heterogeneous species. To reflect the variations among strains
in host preference and virulence, genotypes based on outer
membrane gene A (ompA) gene sequences are being used.
Originally, nine ompA genotypes designated A to F, E/B,
M56, and WC were introduced (reviewed in Harkinezhad
et al. [8]). Later on, six additional genotypes found in psitta-
cines and wild birds and designated 1V, 6N, Mat116, R54,
YP84 and CPX0308 were proposed [9]. The classical geno-
types A to F are known to naturally infect birds and are distinct
from those isolated from chlamydiosis in mammals. Some
avian genotypes appear to occur more often in a specific order
of birds. Genotype A is endemic among psittacine birds and is
considered to be highly virulent. Genotype B is endemic in
pigeons and usually less virulent. Waterfowl most frequently
seem to be infected with genotype C and E/B strains, while
turkeys can harbour genotypes D and C [10]. In contrast,
genotype E, also known as Cal-10, MP or MN, was first
isolated during an outbreak of pneumonia in humans in the
1930s. Later on, genotype E isolates were obtained from a
variety of bird species, including ducks, pigeons, ostriches and
rheas. Genotype F is represented by the psittacine isolates
VS225, Prk Daruma, 84/2334 and 10433-MA, but has also
been isolated from turkeys [11]. The mammalian M56 and
WC genotypes were isolated from an outbreak in muskrats
and hares, and an outbreak of enteritis in cattle, respectively.
All genotypes should be considered to be readily transmissible
to humans. The state of C. psittaci whole-genome analysis
was discussed in a recent review [12••].
Prevalence
All over the world, at least 465 avian species were found to be
infected with this zoonotic agent [13]. Among pet birds,
C. psittaci is highly prevalent in Psittacidae,suchascocka-
toos, parrots, parakeets and lories (16–81 %), as well as in
Columbiformes (12.5–95 %).
Studies on turkey farms, where C. psittaci is nearly endem-
ic, indicate a pathogenic interplay between this agent and
Ornithobacterium (O.)rhinotracheale [11]. However,
Fig. 1 Electron microscopic images of BGM cell culture infected with
Chlamydia (C.)avium strain 10DC88
T
(a)andC. gallinacea strain 08-
1274/3
T
(b). Bar= 1.5 μm. aTwo inclusions are depicted (1 and 2,
indicated by dashed lines). Inclusion 1 contains a mixture of reticulate
bodies (RBs) (R), elementary bodies (EBs) (thin arrows) and a few
intermediate bodies (I). Inclusion 2 consists predominantly of RBs.
Binary fission and budding events are marked with a thick arrow and
arrowheads, respectively. Mitochondria (M) and stacks of Golgi mem-
branes (G) are closely associated with the inclusion membrane. bThe
large inclusion contains predominantly RBs (R), many intermediate
forms (I) and few EBs (thin arrows). Courtesy of Elsevier Ltd. http://
dx.doi.org/10.1016/j.syapm.2013.12.004
Curr Clin Micro Rpt (2015) 2:10–21 11
explosive and devastating outbreaks such as occurred in first
half of the twentieth century are rare nowadays. Instead,
reduced feed intake and respiratory signs with or without
low mortality characterise current outbreaks. Chlamydiosis
in domestic ducks has been reported to be of economic im-
portance and represent a public health hazard in Europe,
China and Australia [14–18]. Incidental to studies of
chlamydiosis in ducks, several investigators observed
C. psittaci antibodies and/or clinical signs in geese and isolat-
ed the agent from diseased tissues. Recently, C. psittaci was
repeatedly detected in chickens, where genotypes B, C, D, F
and E/B predominated [19–22]. It is possible that the reduc-
tion of antibiotic use in the chicken industry has contributed to
this new development. Yin and co-workers [20], demonstrated
the pathogenicity of chicken-derived genotypeB and D strains
for specific pathogen-free (SPF) chickens. As in turkeys,
C. psittaci was often found in conjunction with
O. rhinotracheale. Chlamydiosis has been reported in farmed
quail, peacocks and partridges [10]. Clinical signs and lesions
were similar to those seen in other birds. Morbidity and
mortality can be very high, especially in young birds.
The organism also infects wild birds, such as feral pigeons.
Thirty-eight studies on the seroprevalence of C. psittaci in
feral pigeons conducted from 1966 to 2005 revealed a sero-
positivity ranging from 12.5 to 95.6 %. More recent studies
performed in feral pigeons in Italy, Bosnia and Herzegovina,
and Macedonia revealed a seropositivity of 48.5, 26.5 and
19.2 %, respectively (reviewed in Magnino et al. [23]). Free-
living pigeons are distributed worldwide in urban and rural
areas, where close contact with humans in public places is
common. Known reservoirs of C. psittaci also include Canada
geese [24], seagulls, ducks, herons, egrets, pigeons, black-
birds, grackles, house sparrows and killdeer, all of which
may excrete the pathogen without being visibly affected
[25–27].
Disease and Transmission Pathways in Birds
Depending on many factors on both the pathogen and host
side, avian chlamydiosis can take markedly different courses,
i.e. severe in the acute phase, sub-clinical or inapparent, and
also chronic [10]. Clinical signs in affected birds are usually
non-specific and include respiratory symptoms, conjunctivi-
tis, coryza, mucopurulent discharge from nose and eyes,
cough, dyspnoea or greenish to greyish faeces. In addition,
apathy, weariness, sudden death, stunting, anorexia and ca-
chexia can indicate an ongoing C. psittaci infection [13].
Latent infection is more frequent than acute cases and out-
breaks, but the full significance of this state is still poorly
understood. There are data from cattle showing that the sub-
clinical carrier status can lead to recurrent clinical disease and
chronicity, and, consequently, retarded development of infect-
ed animals [28]. In this context, intermittent shedding of
carriers represents an important reservoir of infection for birds
and humans.
For treatment of clinically ill birds, various tetracyclins and
fluoroquinolones are used. There is nocommercially available
vaccine.
Transmission pathways and mechanisms have been
reviewed by Harkinezhad and colleagues [8]. Bird-to-bird
transmission of C. psittaci usually occurs when dried faecal
droppings or eye and nostril secretions containing the organ-
isms are aerosolised and inhaled by a susceptible host.
Transmission of C. psittaci in the nest is possible. In many
species, such as Columbiformes, cormorants, egrets and
herons, transmission from parent to young may occur through
feeding, by regurgitation, while contamination of the nesting
site with infective exudates or faeces may be important in
other species, such as snow geese, gulls and shorebirds.
Furthermore, C. psittaci can be transmitted from bird to bird
by blood-sucking ectoparasites suchas lice, mites and flies or,
less commonly, through bites or wounds. Transmission of
C. psittaci by arthropod vectors would be facilitated in the
nest environment. Mites from turkey nests can contain chla-
mydiae, and simulid flies were suspected as possible vectors
of transfer during an epidemic in turkeys in South Carolina.
Vertical transmission has been demonstrated in turkeys,
chickens, ducks, parakeets, seagulls and snow geese [29,30]
and could serve as a route to introduce chlamydiae into a
poultry flock. In addition, C. psittaci can be carried into flocks
by wild birds. As contaminated feed or equipment can also be
a source of infection, feed should be protected from wild birds.
Careful cleaning of equipment used in several barns during the
same production round is extremely important because
C. psittaci can survive in faeces and bedding for up to 30 days.
Transmission to Humans
The first description ofa psittacosis outbreak was published in
1879 by Ritter [31],who associated the human disease with an
ongoing outbreak in pet parrots and finches. Pandemic out-
breaks of human psittacosis in Europe and North America
were regularly reported until the 1930s, and all of them could
be traced back to import shipments of infected psittacine birds
from South America.
More recently, cases of human psittacosis due to contact
with psittacines, wild birds, ducks, turkeys, chickens
and meat pigeons were reported in several countries
[17,22,32,2••,33].
Transmission of C. psittaci predominantly occurs through
inhalation of contaminated aerosols from respiratory and oc-
ular secretions or dried faeces from a diseased animal or
asymptomatic carrier after petting infected companion birds,
handling infected avian tissues in the slaughterhouse or as a
result of exposure to C. psittaci in excretions, e.g. from cage
bedding. Handling the plumage of infected birds as well as
12 Curr Clin Micro Rpt (2015) 2:10–21
mouth-to-beak contact or biting represent a zoonotic risk. In
addition, activities such as gardening and mowing or trimming
lawns without a grass catcher have been associated with cases
of human psittacosis (reviewed in Beeckman and Vanrompay
[2••]). Human-to-human transmission is rare, but recently a
new case has been described in Sweden [34].
Populations most at risk include bird owners, pet shop
employees, taxidermists, veterinarians and poultry workers
[35]. The course of psittacosis may vary from asymptomatic
to a flu-like syndrome and involvement of the respiratory
tract. Most infected people show mild symptoms, while im-
munocompromised people are at highest risk of developing
clinical signs. Severe complications such as myocarditis, en-
docarditis, pericarditis, encephalitis, hepatitis, reactive arthri-
tis, multi-organ failure, renal insufficiency, premature birth or
foetal death are rare. Typically, from 2001 onwards, about 400
cases were reported annually in Europe, with a few mortalities
per year. However, these numbers are likely an underestima-
tion of the true prevalence due to incomplete laboratory
diagnosis or unreported cases.
Molecular Pathogenesis of Chlamydia psittaci Infections
The Early Stage of Infection
The ability of C. psittaci to cause systemic infection in differ-
ent host organisms is certainly related to its capability of
entering almost any cell type, from epithelial cells, fibroblasts
and macrophages, to dendritic cells (DCs), etc., which is
known from many in vitro studies. This versatility also sug-
gests that chlamydiae probably have a variety of different
mechanisms for host cell entry at their disposal.
Molecular processes underlying chlamydial entry and up-
take are still poorly understood. It is thought that EBs of
C. psittaci infect their target cells through attachment to the
base of cell surface microvilli [36], where they are engulfed by
endocytic or phagocytic vesicles [37]. Initial attachment is
mediated by electrostatic interactions, and the host protein
disulfide isomerase has been identified as being essential for
both C. psittaci attachment and entry into cells [38]. Various
hypotheses are based on microfilament-dependent phagocy-
tosis, receptor-/clathrin-mediated endocytosis, or the use of
cholesterol-rich lipid raft domains [39]. The participation of
actin and tubulin seems to be required for optimal intracellular
proliferation of chlamydiae [40].
Notably, the shut-down of chlamydial protein synthesis
apparently has no effect on C. psittaci uptake, which means
that it does not require protein factors synthesised by the
bacterial cell [41]. However, the internalisation process de-
pends on the functioning of the type III secretion system
(T3SS) or injectisome. This sophisticated macromolecular
apparatus enables the microorganism to export effector pro-
teins to the inclusion membrane and into the cytosol, where
they modulate certain host cell functions through interaction
with host proteins [42,43]. It appears that newly formed EBs
carry a pre-loaded T3SS in order to ensure rapid entry and
subversion of new host cells. While the chlamydial T3SS
remains active throughout the intracellular stage [37]orpos-
sibly the whole infection cycle [44], interactions of chlamydial
effectors with host proteins seem to play a role from adhesion
and internalisation of EBs to their release from the host cell
[42]. The macromolecular protein complex of the T3SS also
enables translocation into the host cell of bacterial proteins
from an extracellular location across the bacterial cell enve-
lope [45], as well as secretion of pre-synthesised proteins from
the cell-attached EBs [46,47].
In a comprehensive study, Beeckman and colleagues
showed that the T3SS of C. psittaci participates in creating
an optimal environment for intracellular bacterial growth [36].
Their structural investigations demonstrated the association of
the essential structural T3SS protein SctW with the bacterium
and the inclusion membrane, as well as the localisation of
SctC and SctN proteins at the bacterium itself. Monitoring
messenger RNA (mRNA) expression revealed that structural
protein-encoding genes are transcribed from mid-cycle on-
wards (12–18 hpi), whereas the genes encoding effector pro-
teins and putative T3SS-related proteins are expressed early
(1.5–8 hpi) or late (>24 hpi) in the developmental cycle. These
new insights are essential in improving our understanding of
molecular mechanisms during Chlamydia spp. infection. It
seems certain that T3SS effectors are a promising subject for
researchers in order to elucidate crucial phenomena, such as
host cell damage inflicted by the pathogen and evasion of the
host immune system.
Chlamydial Proteins Involved in Host–Pathogen Interaction
The molecular processes underlying the intracellular survival
of C. psittaci are still the subject of intensive research [12••].
Among the key players that are involved in targeting vital
cellular pathways of the host cell are the Inc proteins [48,49].
Type III secretion of both IncA and IncB and their incorpora-
tion in the inclusion membrane during C. psittaci infection has
been experimentally demonstrated [50]. As their hydrophilic
domain protrudes into the cytoplasma and interacts with host
proteins, Inc proteins could be regarded as central regulators
of pathogen–host interactions [51]. Indeed, several eukaryotic
proteins have been identified as interaction partners for Inc
proteins. Two recent studies identified host cell protein
G3BP1 and components of the dynein complex (dynein motor
proteins) as cellular interaction partners of IncA and IncB,
respectively, in C. psittaci infection [52,53]. The authors
concluded that the interaction of chlamydial IncA and host
G3BP1 affects c-myc expression and results in suppression of
Curr Clin Micro Rpt (2015) 2:10–21 13
cellular proliferation and host cell apoptosis [52]. The IncB-
protein of C. psittaci was suggested to recruit dynein motor
proteins [54] in order to control intracellular transport and
perinuclear localisation of inclusions, which would enhance
bacterial growth in infected cells [53]. The recent data of
Böcker and colleagues [53] also revealed that the host protein
Snapin forms a hetero-oligomeric complex with IncB and
dynein, which enables it to physically connect C. psittaci
inclusions with the microtubule network in infected cells.
Recruitment of mitochondria seems to be a characteristic
and possibly unique feature of C. psittaci infection [55,56].
Such a close association, which has not been observed with
C. trachomatis or C. pneumoniae [57], can be expected to
enhance acquisition of eukaryotic adenosine triphosphate
(ATP) and, therefore, influence chlamydial proliferation.
C. psittaci also produces the translocated actin-recruiting
phosphoprotein (Tarp) protein, another T3SS effector in-
volved in entry and intracellular survival of chlamydiae.
Expression of the gene occurs late in the developmental
cycle [36]. Subsequently, released EBs transport pre-
synthesised Tarp into another host cell, where it takes
part in actin remodelling [58].
Intracellular Persistence
Chlamydiae are able to enter a reversible persistent stage in
their infection cycle, where they remain viable but non-culti-
vable. The morphology of this state is characterised by inclu-
sions of reduced size that are filled with the so-called aberrant
bodies, i.e.enlarged RBs. The somewhat indiscriminate use of
the term persistence by certain chlamydiologists recently
prompted statements of clarification in the literature. To avoid
misunderstandings, the term "aberrant RB phenotype" rather
than "persistence" has been proposed to refer to the phenom-
enon in vitro [59,60].
Interestingly, the first experimental investigations on this
subject were conducted with C. psittaci in the 1980s, but the
vast majority of the more recent molecular studies focused on
C. trachomatis. A number of physiologically relevant cell
culture models were characterised in detail, for instance based
on interferon (IFN)-γ-induced persistence (tryptophan deple-
tion) [61], amino acid deficiency [62], iron depletion [63],
exposure to antibiotics [64] and phage infection [65]. Even in
the absence of inducers, spontaneous formation of aberrant
inclusions and chlamydial bodies was also observed during
long-term continuous infection of HEp-2 cells with
C. pneumoniae [66]. Borel and colleagues described an
in vitro model of dual infection with porcine epidemic diar-
rhoea virus (PEDV) and C. abortus or C. pecorum [67], in
which they observed an ongoing chlamydial infection being
arrested and accompanied by formation of aberrant bodies
after inoculation with cell culture-adapted PEDV. The effect
was most pronounced in the case of C. pecorum infection.
In vitro persistence of C. psittaci was studied in three
different cell culture models, i.e. iron depletion, antibiotic
treatment and IFN-γexposure [68]. As expected, the pheno-
typical characteristics were the same as in C. trachomatis and
C. pneumoniae, i.e. aberrant morphology of RBs, loss of
cultivability and rescue of infectivity upon removal of in-
ducers. In contrast, the response of C. psittaci to induced
persistence at the transcriptional level was remarkably differ-
ent. One of the general features observed was a consistent
down-regulation of genes encoding membrane proteins, tran-
scription regulators, cell division factors and EB–RB differ-
entiation factors from 24 hpi onwards. Other genes showed
variations in mRNA expression patterns depending on the
induction mechanism, which implies that there is no persis-
tence model per se. Compared with C. trachomatis,lateshut-
down of essential genes in C. psittaci was much more com-
prehensive with IFN-γ-induced persistence, which can be
explained by the absence of a functional tryptophan synthesis
operon in the latter [68]. Another distinctive feature of the
IFN-γmodel was the observed down-regulation of the chla-
mydia protein associating with death domains (CADD)gene
at 48 hpi in C. psittaci.InC. trachomatis, the same gene was
up-regulated at 48 hpi [69]. The CADD protein shares homol-
ogy with the death domains of tumour necrosis factor family
receptors and is known to induce apoptosis [70]. However, it
has yet to be established whether these in vitro findings
actually relate to latent, persistent or chronic infections in
humans and animals.
In addition to numerous papers on in vitro persistence of
chlamydiae, a few reports from in vivo studies also showed
enlarged chlamydial bodies of C. muridarum [71], C. suis [72]
and C. pneumoniae [73] in infected tissue. However, it is not
clear whether these observations are due to induced persis-
tence or merely illustrate that chlamydiae were stressed during
infection. No observations reminding of aberrant morphology
were made in the C. psittaci animal infection models
discussed below. In any case, a cause–effect relationship
between host response to infection and aberrant chlamydial
bodies has yet to be demonstrated.
New Insights into Host Immune Response to Chlamydial
Infection
C. psittaci seems to be particularly efficient in escaping from
the innate immune response of the host. This conclusion was
drawn from the data of an experimental study by
Braukmann and colleagues [74] comparing C. psittaci
and C. abortus infection in embryonated chicken eggs (see
also the “Lessons Learned from Animal Models”section).
When confronted with the release of pro-inflammatory medi-
ators during early host response, the pathogen was shown to
react with up-regulation of essential genes [74]. This included
elevated mRNA expression rates of chlamydial IncA
14 Curr Clin Micro Rpt (2015) 2:10–21
(involved in stabilisation of the inclusion), ftsW (regulating
binary fission of RBs), groEL (chaperone associating with
macrophages) and cpaf (involved in processing of host pro-
teins controlling the integrity of the inclusion). This particular
response, which may include further chlamydial factors, prob-
ably enables C. psittaci to establish the infection and dissem-
inate in the host organism. In contrast, the closely related
C. abortus was unable to up-regulate the genes mentioned in
the same experimental setting and, consequently, proliferated
less intensely and disseminated to a lesser extent in the host
organism than C. psittaci. These findings have been con-
firmed in analogous comparative infection experiments in
young chicks [75].
The complement system is considered to be one of the
crucial factors of innate immunity. This panel of approx-
imately 40 serum factors is activated by surface compo-
nents of the pathogen and is involved in modulation of
the inflammatory response and protection from extracel-
lular agents [76]. A recent study using a mouse model of
pulmonary C. psittaci infection revealed early, high and
long-lasting activation of the complement system [77].
Further experiments in C3aR-deficient mice suggested
that the protective function of the complement cascade
against C. psittaci was dependent on the anaphylatoxic
peptide C3a and its receptor C3a/C3aR [78].
In this context, it is relevant to note that C3a/C3aR can also
activate DCs, which would facilitate their migration to
draining lymph nodes and enhance presentation of chlamydial
antigens to CD4
+
and CD8
+
T cells [56].
In a recent study in a mouse model, C. psittaci-infected
murine DCs were shown to use autophagosomal and endo-
vacuolar processing for degradation of bacterial compart-
ments, as well as proteolytic production of chlamydial peptide
antigens [79]. It has been suggested that these findings could
be used for the design of vaccines based on DC-targeting
antigens [12••]. A more detailed discussion of recent advances
in the elucidation of host immune response to C. psittaci
infection can be found in the review by Knittler and
colleagues [56].
Lessons Learned from Animal Models
While in vitro models of infection can be very helpful in
identifying individual factors and elucidating their involve-
ment in pathogenesis, the complexity of multiple interactions
between host and pathogen, as encountered in the natural
infection, can be better emulated in animal models.
Therefore, experimental studies in animals have the potential
to generate novel insights and improve our understanding of
the processes occurring in the natural host or, in the case of
zoonoses, in humans.
Recently, the SPF chicken and the chicken embryo models
were used to study the pathogenicity of different C. psittaci
strains and compare C. psittaci and C. abortus infections [74,
20,80]. Both models represent versatile tools for
characterising chlamydial strains and species in terms of in-
vasiveness, virulence and elicited immune response (see also
the “New Insights into Host Immune Response to Chlamydial
Infection”section).
The in ovo model has a far greater potential than serving as
culture medium for intracellular bacteria and viruses. An
experimental protocol starting with inoculation of
Chlamydia spp. onto the chorioallantoic membrane (CAM)
resembles natural infection across epithelial layers. As shown
in a recent study, closely monitoring the course of infection
allows both investigation of the innate immune response to the
chlamydial challenge and identification of molecular process-
es on the chlamydial side. Braukmann and colleagues [74]
highlighted several aspects of host–pathogen interaction in a
comparative study on C. psittaci and C. abortus infection.
Kalmar and colleagues comparatively investigated pathol-
ogy and host immune response, as well as systemic dissemi-
nation and expression of essential chlamydial genes in exper-
imental aerogeneous infection with C. psittaci and C. abortus,
in SPF chicks [75]. They observed that clinical symptoms
appeared sooner and were more severe in the C. psittaci-
infected group. C. psittaci disseminated more efficiently in
the host organism than C. abortus, which was in line with
higher and faster infiltration of immune cells by the former, as
well as more macroscopic lesions and epithelial pathology.
Monitoring mRNA expression rates of immunologically rele-
vant factors in thoracic air sac tissue revealed that IFN-γ,
interleukin (IL)-1β, IL-6, IL-17, IL-22, lipopolysaccharide-
induced tumour necrosis factor (LITAF) and inducible nitric
oxide synthase (iNOS) were significantly stronger up-
regulated in C. psittaci- infected birds between 3 and 14 days
post-infection. At the same time, transcription rates of the
chlamydial genes groEL,cpaf and ftsW were consistently
higher in C. psittaci during the acute phase. These findings
were in accordance with the data from the in ovo study [74]
and confirm the capacity of C. psittaci to evade the immune
response of the avian host more efficiently than other
Chlamydia spp.
A number of studies in a calf model have significantly
improved our understanding of the course of systemic
C. psittaci infection and the pathology in the host organ-
ism. Especially with the human infection in mind, the
bovine host as a model offers a number of advantages
over mice. For instance, the segmental anatomy and lack
of collateral airways in the bovine lung facilitate the
study of pathophysiological mechanisms of pulmonary
dysfunctions. Validated non-invasive lung function tests
are available, and the body size of calves allows repeated
sampling and thorough monitoring of clinical and immu-
nological parameters [80]. In a series of infection trials,
Reinhold and colleagues were able to demonstrate four
Curr Clin Micro Rpt (2015) 2:10–21 15
essential characteristics of C. psittaci infection in its
various manifestations [81–84]:
1. The severity of clinical signs during the acute phase of
infection directly depended on the inoculation dose.
Administration of 10
6
inclusion-forming units (ifu) of
C. psittaci strain DC15 per calf caused mild respiratory
and clinical signs, while doses of 10
7
to 10
8
ifu led to a
moderate and 10
9
ifu to a severe course. This correlation
of dose and response proved reproducible in extent and
quality of lung lesions and also corresponded to deterio-
rations of respiratory functions [81,84].
2. The bovine model reflected characteristic features of nat-
ural chlamydial infections in animals and humans. A
typical course included acute clinical illness in the initial
phase (2–3 dpi), which subsided considerably, but not
completely, until 10 dpi [82]. The next stage was
characterised by a protracted clinically silent course,
which included intermittent mild symptoms, faecal path-
ogen excretion, transient chlamydaemia and slightly ele-
vated levels of monocytes and lipopolysaccharide-
binding protein in blood.Interestingly, these features were
also observed in sentinel calves that socialised with clin-
ically diseased animals and naturally acquired the
infection.
3. The humoral immune response was generally weak. Only
two-thirds of the calves experimentally challenged with a
high dose developed specific antibodies against
C. psittaci, which became detectable between 7 and
14 dpi. This supports the notion that the cellular rather
than humoral immune response plays a central role in
controlling anti-chlamydial immunity in infected hosts.
4. In the acute phase of respiratory disease, inflammatory
cells were recruited to the site of C. psittaci infection.
Damage of the alveolar–capillary barrier caused by pul-
monary inflammation manifested itself by altered cytolo-
gy,aswellaselevatedconcentrationsofeicosanoidsand
total protein in broncho-alveolar lavage fluid [81]. The
inflammation ultimately caused ventilatory disorders and
inhibited pulmonary gas exchange [83,84].
Implications of the Discovery of C. avium, C. gallinacea
and C. ibidis: New Agents of Avian Chlamydiosis?
More Avian Chlamydia spp. Defined
Until very recently, C. psittaci wasconsideredtobethesole
causative agent of the disease. In the light of new evidence
suggesting that avian chlamydiosis may involve more chla-
mydial agents, this paradigm is likely to change. In the past
decade, diagnostic investigations of Chlamydia spp. in-
fection in birds in Germany, France and Italy produced a
number of unclear findings, as the chlamydial agent
appeared to be different from C. psittaci and the other
eight established species of the family Chlamydiaceae.
These atypical strains were identified in poultry, pigeons,
ibis and psittacine birds. The use in routine diagnosis of
broad-range diagnostic assays for Chlamydiaceae in
combination with species-specific detection tools was
an important prerequisite for these discoveries. The
preconceived idea of avian chlamydiosis being due to
C. psittaci alone is probably one of the reasons why
the atypical strains were not discovered earlier. Another
aspect is that these new chlamydiae can easily be missed
in cell culture due to slow and often reluctant growth in
comparison to C. psittaci.
In 2005, Chlamydiaceae-positive but C. psittaci-nega-
tive avian strains were identified in symptomless chickens
of a contact flock involved in an outbreak of psittacosis in
Germany [22]. Three years later, an epidemiological in-
vestigation in poultry breeder flocks in France, which had
been prompted by cases of atypical pneumonia in poultry
slaughterhouse workers, led to the isolation of non-
classified chlamydial strains closely related to the
German strains from seven different flocks, whereas
C. psittaci was found only in one of the 25 flocks exam-
ined [85]. The agent, later defined as C. gallinacea,has
since been found in several European countries and China
[86,87], as well as in Australia [88].
Retrospective analysis of strains isolated from urban
pigeons in Italy in 2006 also identified genetically relat-
ed non-classified strains of Chlamydiaceae.In2009,two
severe outbreaks in breeder flocks of psittacines in
Germany were attributed to closely related chlamydial
strains. Notably, no other potential pathogen was found
in these parrot flocks. Furthermore, pigeons were found
to be a major host of this agent, later designated
C. avium, in surveys in France [89]andGermany[90].
In the same period, investigations conducted on a wild
ibis population in France in 2010 led to the identification
of different Chlamydiaceae-positive but C. psittaci-negative
strains.
Finally, characterisation of selected atypical isolates from
the above studies focused on 16S rRNA-based phylogenetic
analysis, multi-locus sequence analysis, phenotypic character-
isation, as well as whole-genome analysis of type strains
10DC88
T
(C. avium), 08-1274/3
T
(C. gallinacea) and 10-
1398/6
T
(C. ibidis). Based on comparative analysis with the
other established species of the family Chlamydiaceae,
C. avium and C. gallinaceae were proposed as new species
[91•], and C. ibidis was given the Candidatus status [92•].
Basic characteristics of the three avian Chlamydia spp. are
given in Table 1.
16 Curr Clin Micro Rpt (2015) 2:10–21
Epidemiology of C. avium and C. gallinacea
Since C. avium and C. gallinacea were introduced very re-
cently, virtually all studies on avian chlamydiosis have fo-
cused on C. psittaci so far. While specific PCR assays for the
detection of the new species are now available [87,86,93],
specific serological tools are still missing. It is possible that
some of the older papers on avian chlamydiosis dealt with
C. avium or C. gallinacea instead of C. psittaci, especially
those based on serological evidence.
From the limited data that are currently available,
C. avium seems to frequently occur among pigeons,
whereas C. gallinacea is probably widely disseminated
among poultry. Recent data from urban pigeons in
Germany [90]andFrance[89]identifiedC. avium in four
of the 128 (3 %) and in ten of the 125 (8 %)
Chlamydiaceae-positive samples, respectively. In
German breeder pigeon flocks, C. avium was found in
four of the 27 (14.8 %) flocks [94]. In psittacine birds, the
general prevalence of C. avium cannot be assessed as the
findings reported so far represent individual cases.
Prevalence studies on C. gallinacea in chicken and turkey
flocks of four European countries and China revealed that
its prevalence could even be higher than that of C. psittaci
[86]. C. gallinacea wasdetectedin95ofthe110(86.5%)
chlamydia-positive samples, whereas C. psittaci was only
detected in two samples. In a survey conducted over 1 year
in a slaughterhouse, C. gallinacea was detected in 321 of
the 401 (80.0 %) Chlamydiaceae-positive samples from
129 French poultry flocks [95]. In contrast, only
C. psittaci was found in a similar survey conducted in
19 Belgian chicken farms [96]. In Australia, C. gallinacea
was detected in two of the 27 (7.5 %) Chlamydiaceae-positive
samples from chickens, whereas C. psittaci was identi-
fied in five other samples [88]. Results differ greatly
from one study to another and from one country to
another. It is probable that the environment and farming
practices, including cleaning and disinfection procedures,
have a strong impact on circulation and persistence of
chlamydiae in farms.
It is likely that C. avium and C. gallinacea will not be the
last chlamydial species to be discovered in birds. Recent
studies have provided evidence on further non-classified
Chlamydiaceae species in seabirds [97], pigeons [90] and
ducks [95]. The diversity among Chlamydiaceae species is
probably far greater than currently conceived.
Pathogenicity of C. avium and C. gallinacea
The pathogenicity of the newly introduced species has yet to
be systematically investigated. In the surveys reported to date,
no clinical signs have been observed in chickens carrying
C. gallinacea [85], nor in most of the C. avium carriers among
pigeons. However, it seems likely from currently available
data that C. avium is able to cause respiratory disease in
parrots and pigeons [91•]. In analogy to the established
Chlamydia spp., the new chlamydiae could survive as com-
mensals in the gastrointestinal tract for extended periods be-
fore eliciting cases of disease, as discussed in two recent
reviews [98,28]. However, co-infections with other
Chlamydia spp., bacteria or viruses [99] could exacerbate
the course of C. avium or C. gallinacea infections as already
reported for C. psittaci-infected turkeys (see the “Aetiology
and Epidemiology”section). While cases of co-infection be-
tween the new chlamydiae and C. psittaci have been reported
in pigeons [94] and poultry [87], evidence on possible inter-
action, such as synergetic or competitive effects, in the course
of co-infection is still lacking. It would be interesting to re-
examine samples from previous outbreaks of avian
chlamydiosis for the presence of the two new species.
The zoonotic potential is still unknown, although there is a
possibility of C. gallinacea being involved in zoonotic trans-
mission. Those cases of atypical pneumonia reported among
slaughterhouse workers exposed to chickens infected with
Tabl e 1 Basic characteristics of avian Chlamydia (C.)spp.
Species Major hosts Pathogenicity Type
strain
Genome
size (bp)
No. of predicted
proteins
16S rDNA
difference (%)
a
Detection assays
C. psittaci Birds, mammals Systemic respiratory
disease
6BC 1,171,660 975 0 rtPCR, DNA
microarray,
ELISA [100]
C. avium Pigeons, parrots,
probably wild birds
Respiratory disease 10 DC88 1,041,169 940 1.95 rtPCR [93], DNA
microarray
C. gallinacea Chickens, turkeys,
guinea fowl, ducks,
probably other poultry
To be investigated 08-1274/3 1,045,134 907 1.88 rtPCR [87], DNA
microarray
a
Compared with C. psittaci
rDNA ribosomal DNA, rtPCR real-time PCR
Curr Clin Micro Rpt (2015) 2:10–21 17
C. gallinacea [85] can be taken as an indication, even though
previous exposure of these workers to C. psittaci cannot be
excluded. However, these cases could not be definitively
clarified because species-specific serological tools are not
available for chlamydiosis. This striking deficit should be
addressed in future research.
Conclusion
The results of field surveys in Europe and elsewhere in
the past decade indicate a rise in the prevalence of
Chlamydia infections in poultry flocks. This increase
has been partly attributed to improved diagnostics, but
could also be due to reduced use of antibiotics in
poultry. In addition, organic poultry production, where
free-range facilities allow contact with feral birds and
pathogen-containing faeces, might also have contributed
to this increase.
Recent advances in research on the pathogenesis of
avian chlamydiosis include the generation of comprehen-
sive datasets on host–pathogen interaction obtained from
in vitro, in ovo and in vivo infection models. Following
rapid entry into host cells, which is controlled by specific
surface proteins and T3SS effectors, C. psittaci was
shown to efficiently disseminate within the animal host,
causing systemic disease. The pathogen seems to be
capable of evading the action of host pro-inflammatory
mediators more efficiently than other chlamydiae. When
facing the host immune response it was shown to up-
regulate essential chlamydial genes. A number of new
molecular factors that are important for intracellular pro-
liferation and progression of the infection have been
identified.
Following the discovery of two new avian chlamydial
species, aetiopathology and epidemiology of avian
chlamydiosis will have to be revised, since C. psittaci no
longer seems to be the only chlamydial agent involved.
Although it is too early for a final assessment of the impor-
tance of C. gallinacea and C. avium, veterinarians, physicians,
diagnosticians and researchers should take the new develop-
ments into account and consider possible involvement of the
new agents in cases of avian chlamydiosis.
Compliance with Ethics Guidelines
Conflict of Interest Dr Sachse, Dr Laroucau and Dr Vanrompay each
declare they have no conflicts of interests.
Human and Animal Rights and Informed Consent This article
contains no studies with human or animal subjects performed by any of
the authors.
References
Papers of particular interest, published recently, have been
highlighted as:
•Of importance
•• Of major importance
1. European Commission, Health & Consumer Protection
Directorate-General. (2002) Avian chlamydiosis as a zoonotic
disease and risk reduction strategies. SCAHAW (Scientific
Committee on Animal Health and Animal Welfare), SANCO/
AH/R26/2002. http://ec.europa.eu/food/fs/sc/scah/out73_en.pdf
2.•• BeeckmanDS, Vanrompay DC. Zoonotic Chlamydophila psittaci
infections from a clinical perspective. Clin Microbiol Infect.
2009;15(1):11–7. doi:10.1111/j.1469-0691.2008.02669.x.
Comprehensive review on epidemiology, transmission pathways,
clinical course, diagnosis and possible treatment for psittacosis in
humans. It is emphasised that the general awareness towards
psittacosis among medical workers is low and methods used in
routine diagnosis should be updated.
3. Everett KD, Bush RM, Andersen AA. Emended description of the
order Chlamydiales,proposalofParachlamydiaceae fam. nov.
and Simkaniaceae fam. nov., each containing one monotypic
genus, revised taxonomy of the family Chlamydiaceae,including
a new genus and five new species, and standards for the identifi-
cation of organisms. Int J Syst Bacteriol. 1999;49 Pt 2:415–40.
4. Stephens RS, Myers G, Eppinger M, Bavoil PM. Divergence
without difference: phylogenetics and taxonomy of Chlamydia
resolved. FEMS Immunol Med Microbiol. 2009;55(2):115–9.
doi:10.1111/j.1574-695X.2008.00516.x.
5. Kuo C-C, Stephens RS, Bavoil PM, Kaltenboeck B. Genus
Chlamydia Jones, Rake and Stearns 1945, 55. In: Krieg NR,
Staley JT, Brown DR, Hedlund BP, Paster BJ, Ward NL,
Ludwig W, Whitman WB, editors. Bergey's Manual of
Systematic Bacteriology, vol. 4. 2nd ed. Heidelberg: Springer;
2011. p. 846–65.
6. Sachse K, Bavoil PM, Kaltenboeck B, Stephens RS, Kuo C-C,
Rosselló-Móra R, Horn M. Emendation of the family
Chlamydiaceae: Proposal of a single genus, Chlamydia,toinclude
all currently recognized species. Syst Appl Microbiol. 2015. doi:
10.1016/j.syapm.2014.12.004.
7. Matsumoto A, Manire GP. Electron microscopic observations on
the effects of penicillin on the morphology of Chlamydia psittaci.
J Bacteriol. 1970;101(1):278–85.
8. Harkinezhad T, Geens T, Vanrompay D. Chlamydophila psittaci
infections in birds: a review with emphasis on zoonotic conse-
quences. Vet Microbiol. 2009;135(1–2):68–77. doi:10.1016/j.
vetmic.2008.09.046.
9. Sachse K, Laroucau K, Hotzel H, Schubert E, Ehricht R, Slickers
P. Genotyping of Chlamydophila psittaci using a new DNA
microarray assay based on sequence analysis of ompA genes.
BMC Microbiol. 2008;8:63.
10. Vanrompay D (2013) Avian Chlamydiosis. In: Swayne D, editor.
Diseases of Poultry, 13th edition 2013. Ames: Wiley- Blackwell
Publishing, Section III: Bacterial Diseases, Chapter 24
11. VanLoockM,GeensT,DeSmitL,NauwynckH,VanEmpelP,
Naylor C, et al. Key role of Chlamydophila psittaci on Belgian turkey
farms in association with other respiratory pathogens. Vet Microbiol.
2005;107(1–2):91–101. doi:10.1016/j.vetmic.2005.01.009.
12.•• Knittler MR, Sachse K. Chlamydia psittaci: update on an
underestimated zoonotic agent (Minireview). FEMS Pathog Dis.
2014. doi:10.1093/femspd/ftu1007.Review of current knowledge
on C. psittaci as a pathogen. Recent advances in genomic
18 Curr Clin Micro Rpt (2015) 2:10–21
analysis, as well as research on molecular host–pathogen
interaction, innate and adaptive immune response in clinical,
sub-clinical and persistent infection are discussed.
13. Kaleta EF, Taday EM. Avian host range of Chlamydophila spp.
based on isolation, antigen detection and serology. Avian Pathol.
2003;32(5):435–61. doi:10.1080/03079450310001593613.
14. Cao J, Yang Q, Yang L, Liu Z, He C. Epidemic investigation of
avian Chlamydia psittaci in Beijing and other provinces around.
Vet Sci China. 2006;56:340–9.
15. Guérin JL, Ballot A, Sraka B, Léon O (2006) Portage de
Chlamydophila psittaci dans la filiére canard mulard: évaluation
du portage chez les reproducteurs et incidence sur le statut du
caneton. In: Proceedings des 7èmes Journées de la recherche sur
les palmipèdes à foie gras 18–19 October 2006, Arcachon, France:
pp. 37–40
16. Haas WH, Swaan CM, Meijer A, Neve G, Buchholz U, Beer
M, et al. A Dutch case of atypical pneumonia after culling
of H5N1 positive ducks in Bavaria was found infected with
Chlamydophila psittaci. Euro Surveill. 2007;12(11):E071129
071123.
17. Laroucau K, de Barbeyrac B, Vorimore F, Clerc M, Bertin C,
Harkinezhad T, et al. Chlamydial infections in duck farms associ-
ated with human cases of psittacosis in France. Vet Microbiol.
2009;135(1–2):82–9. doi:10.1016/j.vetmic.2008.09.048.
18. Yin L, Kalmar ID, Lagae S, Vandendriessche S, Vanderhaeghen
W, Butaye P, et al. Emerging Chlamydia psittaci infections in the
chicken industry and pathology of Chlamydia psittaci genotype B
and D strains in specific pathogen free chickens. Vet Microbiol.
2013;162(2–4):740–9. doi:10.1016/j.vetmic.2012.09.026.
19. Dickx V, Geens T, Deschuyffeleer T, Tyberghien L, Harkinezhad
T, Beeckman DS, et al. Chlamydophila psittaci zoonotic risk
assessment in a chicken and turkey slaughterhouse. J Clin
Microbiol. 2010;48(9):3244–50. doi:10.1128/JCM. 00698-10.
20. Yin L, Lagae S, Kalmar I, Borel N, Pospischil A, Vanrompay D.
Pathogenicity of low and highly virulent Chlamydia psittaci iso-
lates for specific-pathogen-free chickens. Avian Dis. 2013;57(2):
242–7.
21. Zhang F, Li S, Yang J, Pang W, Yang L, He C. Isolation and
characterization of Chlamydophila psittaci isolated from laying
hens with cystic oviducts. Avian Dis. 2008;52(1):74–8.
22. Gaede W, Reckling KF, Dresenkamp B, Kenklies S, Schubert E,
Noack U, et al. Chlamydophila psittaci infections in humans
during an outbreak of psittacosis from poultry in Germany.
Zoonoses Public Health. 2008;55(4):184–8. doi:10.1111/j.1863-
2378.2008.01108.x.
23. Magnino S, Haag-Wackernagel D, Geigenfeind I, Helmecke S,
Dovc A, Prukner-Radovcic E, et al. Chlamydial infections in feral
pigeons in Europe: Review of data and focus on public health
implications. Vet Microbiol. 2009;135(1–2):54–67. doi:10.1016/j.
vetmic.2008.09.045.
24. Dickx V, Kalmar ID, Tavernier P, Vanrompay D. Prevalence and
genotype distribution of Chlamydia psittaci in feral Canada geese
(Branta canadensis) in Belgium. Vector Borne Zoonotic Dis.
2013;13(6):382–4. doi:10.1089/vbz.2012.1131.
25. Herrmann B, Persson H, Jensen JK, Joensen HD, Klint M, Olsen
B. Chlamydophila psittaci in fulmars, the Faroe Islands. Emerg
Infect Dis. 2006;12(2):330–2.
26. Kalmar ID, Dicxk V, Dossche L, Vanrompay D. Zoonotic infec-
tion with Chlamydia psittaci at an avian refuge centre. Vet J.
2014;199(2):300–2. doi:10.1016/j.tvjl.2013.10.034.
27. Blomqvist M, Christerson L, Waldenstrom J, Herrmann B, Olsen
B. Chlamydia psittaci in Swedish wetland birds: a risk to zoonotic
infection? Avian Dis. 2012;56(4):737–40.
28. Reinhold P, Sachse K, Kaltenboeck B. Chlamydiaceae in cattle:
commensals, trigger organisms, or pathogens? Vet J. 2011;189(3):
257–67. doi:10.1016/j.tvjl.2010.09.003.
29. Wittenbrink MM, Mrozek M, Bisping W. Isolation of Chlamydia
psittaci from a chicken egg: evidence of egg transmission.
Zentralbl Veterinarmed B. 1993;40(6):451–2.
30. Lublin A, Shudari G, Mechani S, Weisman Y. Egg transmission of
Chlamydia psittaci in turkeys. Vet Rec. 1996;139(12):300.
31. Ritter J. Über Pneumotyphus, eine Hausepidemie in Uster. Dtsch
Arch klin Med. 1879;25:53.
32. Verminnen K, Duquenne B, De Keukeleire D, Duim B,
Pannekoek Y, Braeckman L, et al. Evaluation of a
Chlamydophila psittaci diagnostic platform for zoonotic risk as-
sessment. J Clin Microbiol. 2008;46:281–5.
33. Rehn M, Ringberg H, Runehagen A, Herrmann B, Olsen B,
Petersson AC, et al. Unusual increase of psittacosis in southern
Sweden linked to wild bird exposure, January toApril 2013. Euro
Surveill. 2013;18(19):20478.
34. Wallensten A, Fredlund H, Runehagen A. Multiple human-to-
human transmission from a severe case of psittacosis, Sweden,
January-February 2013. Euro Surveill. 2014;19(42).
35. Deschuyffeleer TP, Tyberghien LF, Dickx VL, Geens T, Saelen
JM, Vanrompay DC, et al. Risk assessment and management of
Chlamydia psittaci in poultry processing plants. Ann Occup Hyg.
2012;56(3):340–9. doi:10.1093/annhyg/mer102.
36. Beeckman DS, Geens T, Timmermans JP, Van Oostveldt P,
Vanrompay DC. Identification and characterization of a type III
secretion system in Chlamydophila psittaci. Vet Res. 2008;39(3):
27. doi:10.1051/vetres:2008002.
37. Dautry-Varsat A, Balana ME, Wyplosz B. Chlamydia–host cell
interactions: recent advances on bacterial entry and intracellular
development. Traffic. 2004;5(8):561–70. doi:10.1111/j.1398-
9219.2004.00207.x.
38. Abromaitis S, Stephens RS. Attachment and entry of Chlamydia
have distinct requirements for host protein disulfide isomerase.
PLoS Pathog. 2009;5(4):e1000357. doi:10.1371/journal.ppat.
1000357.
39. Stuart ES, Webley WC, Norkin LC. Lipid rafts, caveolae, caveo-
lin-1, and entry by Chlamydiae into host cells. Exp Cell Res.
2003;287(1):67–78.
40. Escalante-Ochoa C, Ducatelle R, Haesebrouck F. Optimal devel-
opment of Chlamydophila psittaci in L929 fibroblast and BGM
epithelial cells requires the participation of microfilaments and
microtubule-motor proteins. Microb Pathog. 2000;28(6):321–33.
doi:10.1006/mpat.2000.0352.
41. Jutras I, Subtil A, Wyplosz B, Dautry-Varsat A (2004) Chlamydia.
Intracellular pathogens in membrane interactions and vacuole
biogenesise (Molecular Biology Intelligence Unit). Gorvel J-P,
editor. New York: Springer-Verlag New York, LLC. pp. 179–189
42. Scidmore MA. Recent advances in Chlamydia subversion of host
cytoskeletal and membrane trafficking pathways. Microbes Infect.
2011;13(6):527–35. doi:10.1016/j.micinf.2011.02.001.
43. Hsia RC, Pannekoek Y, Ingerowski E, Bavoil PM. Type III
secretion genes identify a putative virulence locus of Chlamydia.
Mol Microbiol. 1997;25(2):351–9.
44. Saad N. The role of the type III secretion system of Chlamydia
psittaci in human macrophages [PhD thesis]. Ghent: University of
Ghent, Faculty of Bioscience Engineering (Academic year 2010–
2011). 2011
45. Betts-Hampikian HJ, Fields KA. The Chlamydial TYPE III secre-
tion mechanism: revealing cracks in a tough nut. Front Microbiol.
2010;1:114. doi:10.3389/fmicb.2010.00114.
46. Fields KA, Mead DJ, Dooley CA, Hackstadt T. Chlamydia
trachomatis type III secretion: evidence for a functional apparatus
during early-cycle development. Mol Microbiol. 2003;48(3):
671–83.
47. Jamison WP, Hackstadt T. Induction of type III secretion by cell-
free Chlamydia trachomatis elementary bodies. Microb Pathog.
2008;45(5–6):435–40. doi:10.1016/j.micpath.2008.10.002.
Curr Clin Micro Rpt (2015) 2:10–21 19
48. Moore ER, Ouellette SP. Reconceptualizing the chlamydial inclu-
sion as a pathogen-specified parasitic organelle: an expanded role
for Inc proteins. Front Cell Infect Microbiol. 2014;4:157. doi:10.
3389/fcimb.2014.00157.
49. Valdivia RH. Chlamydia effector proteins and new insights into
chlamydial cellular microbiology. Curr Opin Microbiol.
2008;11(1):53–9. doi:10.1016/j.mib.2008.01.003.
50. Beeckman DS, Geens T, Timmermans JP, Van Oostveldt P,
Vanrompay DC. Identification and characterization of a type III
secretion system in Chlamydophila psittaci. Vet Res. 2008;39(3):
27. doi:10.1051/vetres:2008002.
51. Mital J, Miller NJ, Dorward DW, Dooley CA, Hackstadt T. Role
for chlamydial inclusion membrane proteins in inclusion mem-
brane structure and biogenesis.PLoS One. 2013;8(5):e63426. doi:
10.1371/journal.pone.0063426.
52. Borth N, Litsche K, Franke C, Sachse K, Saluz HP, Hanel F.
Functional interaction between type III-secreted protein IncA of
Chlamydophila psittaci and human G3BP1. PLoS One.
2011;6(1):e16692. doi:10.1371/journal.pone.0016692.
53. Böcker S, Heurich A, Franke C, Monajembashi S, Sachse K,
Saluz HP, et al. Chlamydia psittaci inclusion membrane protein
IncB associates with host protein Snapin. Int J Med Microbiol.
2014;304(5–6):542–53. doi:10.1016/j.ijmm.2014.03.005.
54. Roberts AJ, Kon T, Knight PJ, Sutoh K, Burgess SA. Functions
and mechanics of dynein motor proteins. Nat Rev Mol Cell Biol.
2013;14(11):713–26. doi:10.1038/nrm3667.
55. Matsumoto A. Isolation and electron microscopic observations of
intracytoplasmic inclusions containing Chlamydia psittaci.J
Bacteriol. 1981;145(1):605–12.
56. Knittler MR, Berndt A, Bocker S, Dutow P, Hanel F, Heuer D,
et al. Chlamydia psittaci: New insights into genomic diversity,
clinical pathology, host-pathogen interaction and anti-bacterial
immunity. Int J Med Microbiol. 2014;304:877–93. doi:10.1016/
j.ijmm.2014.06.010.
57. Matsumoto A, Bessho H, Uehira K, Suda T. Morphological stud-
ies of the association of mitochondria with chlamydial inclusions
and the fusion of chlamydial inclusions. J Electron Microsc
(Tokyo). 1991;40(5):356–63.
58. Engel J. Tarp and Arp: how Chlamydia induces itsown entry. Proc
Natl Acad Sci U S A. 2004;101(27):9947–8. doi:10.1073/pnas.
0403633101.
59. Wyrick PB. Chlamydia trachomatis persistence in vitro: an over-
view. J Infect Dis. 2010;201 Suppl 2:S88–95. doi:10.1086/
652394.
60. Bavoil PM. What's in a word: the use, misuse, and abuse of the
word "persistence" in Chlamydia biology. Front Cell Infect
Microbiol. 2014;4:27. doi:10.3389/fcimb.2014.00027.
61. Beatty WL, Byrne GI, Morrison RP. Repeated and persistent
infection with Chlamydia and the development of chronic inflam-
mation and disease. Trends Microbiol. 1994;2(3):94–8.
62. Coles AM, Reynolds DJ, Harper A, Devitt A, Pearce JH. Low-
nutrient induction of abnormal chlamydial development: a novel
component of chlamydial pathogenesis? FEMS Microbiol Lett.
1993;106(2):193–200.
63. Raulston JE. Response of Chlamydia trachomatis serovar E to
iron restriction in vitro and evidence for iron-regulated chlamydial
proteins. Infect Immun. 1997;65(11):4539–47.
64. Pantoja LG, Miller RD, Ramirez JA, Molestina RE, Summersgill
JT. Characterization of Chlamydia pneumoniae persistence in
HEp-2 cells treated with gamma interferon. Infect Immun.
2001;69(12):7927–32. doi:10.1128/IAI. 69.12.7927-7932.2001.
65. Hsia R, Ohayon H, Gounon P, Dautry-Varsat A, Bavoil PM. Phage
infection of the obligate intracellular bacterium, Chlamydia
psittaci strain guinea pig inclusion conjunctivitis. Microbes
Infect. 2000;2(7):761–72.
66. Kutlin A, Flegg C, Stenzel D, Reznik T, Roblin PM, Mathews S,
et al. Ultrastructural study of Chlamydia pneumoniae in a
continuous-infection model. J Clin Microbiol. 2001;39(10):
3721–3. doi:10.1128/JCM. 39.10.3721-3723.2001.
67. Borel N, Dumrese C, Ziegler U, Schifferli A, Kaiser C, Pospischil
A. Mixed infections with Chlamydia and porcine epidemic diar-
rhea virus - a new in vitro model of chlamydial persistence. BMC
Microbiol. 2010;10:201. doi:10.1186/1471-2180-10-201.
68. Goellner S, Schubert E, Liebler-Tenorio E, Hotzel H, Saluz HP,
Sachse K. Transcriptional response patterns of Chlamydophila
psittaci in different in vitro models of persistent infection. Infect
Immun. 2006;74(8):4801–8. doi:10.1128/IAI. 01487-05.
69. Belland RJ, Nelson DE, Virok D, Crane DD, Hogan D, Sturdevant
D, et al. Transcriptome analysis of chlamydial growth during IFN-
gamma-mediated persistence and reactivation. Proc Natl Acad Sci
U S A. 2003;100(26):15971–6. doi:10.1073/pnas.2535394100.
70. Stenner-Liewen F, Liewen H, Zapata JM, Pawlowski K, Godzik
A, Reed JC. CADD, a Chlamydia protein that interacts with death
receptors. J Biol Chem. 2002;277(12):9633–6. doi:10.1074/jbc.
C100693200.
71. Phillips Campbell R, Kintner J, Whittimore J, Schoborg RV.
Chlamydia muridarum enters a viable but non-infectious state in
amoxicillin-treated BALB/c mice. Microbes Infect. 2012;14(13):
1177–85. doi:10.1016/j.micinf.2012.07.017.
72. Pospischil A, Borel N, Chowdhury EH, Guscetti F. Aberrant
chlamydial developmental forms in the gastrointestinal tract of
pigs spontaneously and experimentally infected with Chlamydia
suis. Vet Microbiol. 2009;135(1–2):147–56. doi:10.1016/j.
vetmic.2008.09.035.
73. Borel N, Summersgill JT, Mukhopadhyay S, Miller RD, Ramirez
JA, Pospischil A. Evidence for persistent Chlamydia pneumoniae
infection of human coronary atheromas. Atherosclerosis.
2008;199(1):154–61. doi:10.1016/j.atherosclerosis.2007.09.026.
74. Braukmann M, Sachse K, Jacobsen ID, Westermann M, Menge C,
Saluz HP, et al. Distinct intensity of host-pathogen interactions in
Chlamydia psittaci-andChlamydia abortus-infected chicken em-
bryos. Infect Immun. 2012;80(9):2976–88. doi:10.1128/IAI.
00437-12.
75. Kalmar I, Berndt A, Yin L, Chiers K, Sachse K, Vanrompay D.
Host-pathogen interactions inspecific pathogen-free chickens fol-
lowing aerogenous infection with Chlamydia psittaci and
Chlamydia abortus. Vet Immunol Immunopathol. 2015. doi:10.
1016/j.vetimm.2014.12.014.
76. Klos A, Wende E, Wareham KJ, Monk PN. International Union of
Basic and Clinical Pharmacology. [corrected]. LXXXVII.
Complement peptide C5a, C4a, and C3a receptors. Pharmacol
Rev. 2013;65(1):500–43.
77. Bode J, Dutow P, Sommer K, Janik K, Glage S, Tummler B, et al.
A new role of the complementsystem: C3 provides protection in a
mouse model of lung infection with intracellular Chlamydia
psittaci. PLoS One. 2012;7(11):e50327. doi:10.1371/journal.
pone.0050327.
78. Dutow P, Fehlhaber B, Bode J, Laudeley R, Rheinheimer C, Glage
S, et al. The complement C3a receptor is critical in defense against
Chlamydia psittaci in mouse lung infection and required for
antibody and optimal T cell response. J Infect Dis. 2014;209(8):
1269–78. doi:10.1093/infdis/jit640.
79. Fiegl D, Kagebein D, Liebler-Tenorio EM, Weisser T, Sens M,
Gutjahr M, et al. Amphisomal route of MHC class I cross-
presentation in bacteria-infected dendritic cells. J Immunol.
2013;190(6):2791–806. doi:10.4049/jimmunol.1202741.
80. Prohl A, Ostermann C, Lohr M, Reinhold P. The bovine lung in
biomedical research:visually guided bronchoscopy, intrabronchial
inoculation and in vivo sampling techniques. J Vis Exp. 2014;89.
doi:10.3791/51557.
20 Curr Clin Micro Rpt (2015) 2:10–21
81. Reinhold P, Ostermann C, Liebler-Tenorio E, Berndt A, Vogel A,
Lambertz J, et al. A bovine model of respiratory Chlamydia
psittaci infection: challenge dose titration. PLoS One. 2012;7(1):
e30125. doi:10.1371/journal.pone.0030125.
82. Ostermann C, Ruttger A, Schubert E, Schrodl W, Sachse K,
Reinhold P. Infection, disease, and transmission dynamics in
calves after experimental and natural challenge with a Bovine
Chlamydia psittaci isolate. PLoS One. 2013;8(5):e64066. doi:10.
1371/journal.pone.0064066.
83. Ostermann C, Linde S, Siegling-Vlitakis C, Reinhold P.
Evaluation of pulmonary dysfunctions and acid base imbalances
induced by Chlamydia psittaci in a bovine model of respiratory
infection. Multidiscip Respir Med. 2014;9(1):10. doi:10.1186/
2049-6958-9-10.
84. Ostermann C, Schroedl W, Schubert E, Sachse K, Reinhold P.
Dose-dependent effects of Chlamydia psittaci infection on pulmo-
nary gas exchange, innate immunity and acute-phase reaction in a
bovine respiratory model. Vet J. 2013;196(3):351–9. doi:10.1016/
j.tvjl.2012.10.035.
85. Laroucau K, Vorimore F, Aaziz R, Berndt A, Schubert E, Sachse
K. Isolation of a new chlamydial agent from infected domestic
poultry coincided with cases of atypical pneumonia among
slaughterhouse workers in France. Infect Genet Evol. 2009;9(6):
1240–7.
86. Zocevic A, Vorimore F, Marhold C, Horvatek D, Wang D, Slavec
B, et al. Molecular characterization of atypical Chlamydia and
evidence of their dissemination in different European and Asian
chicken flocks by specific real-time PCR. Environ Microbiol.
2012;14(8):2212–22. doi:10.1111/j.1462-2920.2012.02800.x.
87. Laroucau K, Aaziz R, Meurice L, Servas V, Chossat I, Royer H,
et al. (2014) Outbreak of psittacosis in a group of women exposed
to Chlamydia psittaci-infected chickens. Euro Surveill. In press
[EMID:ce1270d610605ffb]
88. Robertson T, Bibby S, O'Rourke D, Belfiore T, Agnew-Crumpton R,
Noormohammadi AH. Identification of Chlamydial species in croc-
odiles and chickens by PCR-HRM curve analysis. Vet Microbiol.
2010;145(3–4):373–9. doi:10.1016/j.vetmic.2010.04.007.
89. Gasparini J, Erin N, Bertin C, Jacquin L, Vorimore F, Frantz A,
et al. Impact of urban environment and host phenotype on the
epidemiology of Chlamydiaceae in feral pigeons (Columba livia).
Environ Microbiol. 2011;13(12):3186–93. doi:10.1111/j.1462-
2920.2011.02575.x.
90. Sachse K, KuehlewindS, Ruettger A, Schubert E, Rohde G. More
than classical Chlamydia psittaci in urban pigeons. Vet Microbiol.
2012;157(3–4):476–80. doi:10.1016/j.vetmic.2012.01.002.
92.•Sachse K, Laroucau K, Riege K, Wehner S, Dilcher M, Creasy
HH, et al. Evidence for the existence of two new members of the
family Chlamydiaceae and proposal of Chlamydia avium sp. nov.
and Chlamydia gallinacea sp. nov. Syst Appl Microbiol.
2014;37(2):79–88. doi:10.1016/j.syapm.2013.12.004.
Introduction and description of C. avium and C. gallinacea as
new species within the recombined genus Chlamydia. Data on
morphology, epidemiology and pathogenicity, as well as
phylogenetic and genomic parameters are presented.
93.•Vorimore F, Hsia RC, Huot-Creasy H, Bastian S, Deruyter L,
Passet A, et al. Isolation of a new Chlamydia species from the
Feral Sacred Ibis (Threskiornis aethiopicus): Chlamydia ibidis.
PLoS One. 2013;8(9):e74823. doi:10.1371/journal.pone.
0074823.First description of this Candidatus species. Data from
detailed genomic analysis, morphology and comparative
phylogeny are presented and discussed.
93. Zocevic A, Vorimore F, Vicari N, Gasparini J, Jacquin L, Sachse
K, et al. A real-time PCR assay for the detection of atypical strains
of Chlamydiaceae from pigeons. PLoS One. 2013;8(3):e58741.
doi:10.1371/journal.pone.0058741.
94. Krautwald-Junghanns ME, Stolze J, Schmidt V, Bohme J, Sachse
K, Cramer K. Efficacy of doxycycline for treatment of
chlamydiosis in flocks of racing and fancy pigeons [in German].
Prax Ausg K Kleintiere Heimtiere. 2013;41(6):392–8.
95. Hulin V, Oger S, Vorimore F, Aaziz R, de Barbeyrac B, Berruchon
J, et al. Host preference and zoonotic potential of Chlamydia
psittaci and Chlamydia gallinacea in poultry. Pathog Dis. 2015
(in press).
96. Lagae S, Kalmar I, Laroucau K, Vorimore F, Vanrompay D.
Emerging Chlamydia psittaci infections in chickens and examina-
tion of transmission to humans. J Med Microbiol. 2014;63(Pt 3):
399–407. doi:10.1099/jmm. 0.064675-0.
97. Christerson L, Blomqvist M, Grannas K, Thollesson M, Laroucau
K, Waldenstrom J, et al. A novel Chlamydiaceae-like bacterium
found in faecal specimens from sea birds from the Bering Sea.
Environ Microbiol Rep. 2010;2(4):605–10. doi:10.1111/j.1758-
2229.2010.00174.x.
98. Rank RG, Yeruva L. Hidden in plain sight: chlamydial gastroin-
testinal infection and its relevance to persistence in human genital
infection. Infect Immun. 2014;82(4):1362–71. doi:10.1128/IAI.
01244-13.
99. Loock MV, Loots K, Zande SV, Heerden MV, Nauwynck H,
Goddeeris BM, et al. Pathogenic interactions between
Chlamydophila psittaci and avian pneumovirus infections in tur-
keys. Vet Microbiol. 2006;112(1):53–63.
100. Sachse K, Vretou E, Livingstone M, Borel N, Pospischil A,
Longbottom D. Recent developments in the laboratory diagnosis
of chlamydial infections. Vet Microbiol. 2009;135:2–21.
Curr Clin Micro Rpt (2015) 2:10–21 21
