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Lymphogranuloma venereum
The serovars causing lymphogranuloma venereum (LGV) differ from other serovars in that they are much more invasive. Similar to serovar D-K, their transmission is through sexual contact, but they spread to subepithelial tissues, infect monocytes and disseminate to the regional lymph nodes, causing lymphadenitis (inflammation of lymph nodes) and in some cases necrosis with pus formation (Mabey and Peeling, 2002). LGV had been rare in industrialised countries before LGV proctitis started to emerge in populations of men who have sex with men. It is noteworthy that co-infections with HIV occur in the majority of LGV proctitis cases (Van der Bij et al., 2006). The ulcerative nature of LGV can in general facilitate the acquisition and transmission of STIs.
Diagnosis and treatment
Diagnosis and treatment vary depending on the disease elicited by C. trachomatis. The diagnosis of trachoma is generally made on clinical grounds. Considering the regions of highest prevalence, a careful examination of the eye with a binocular is a quick and affordable method compared to laboratory diagnosis. Different grading systems of the progress of the disease help to standardise field surveys and research studies. The gold standard for laboratory diagnosis has long been cell culture, because of its near perfect specificity (Solomon et al., 2004). It has been replaced by nucleic acid amplification tests, which are highly specific too, and have the advantage of a higher sensitivity than cell culture (Johnson et al., 2000). Treatments of the infected eye vary from surgical reorientation of the eye-lid to antibiotic administration. Typically, tetracycline or azithromycin are locally administered for a duration of 6 weeks. A suitable strategy also implies environmental changes and a focus on hygiene to reduce the spread and reoccurrence of the infection (Hu et al., 2010).
Chlamydial urogenital tract and LGV infections are generally detected via nucleic acid amplification tests on urine samples or vaginal swabs. Recommended treatments for non-LGV Chlamydiae are a single dose of azithromycin, or doxycycline twice a day for 7 days (www.cdc.gov), with cure rates of 97 % and 98 %, respectively.
The typical treatment for LGV is 3 weeks of doxycycline administration, twice a day. The different recommendations between LGV and non-LGV strains require genotyping of the infective agent, which is not widely accessible for standard laboratory methods (McLean et al., 2007).
A vaccine against C. trachomatis has yet to be developed. The vaccine-development process is challenged by the nature of the bacterium as well as by other aspects, e.g. the pathogenesis of a chlamydial infection, which is partially induced by the immune response of the patient (Brunham and Rey-Ladino, 2005). The best strategy to avoid an infection with C. trachomatis to date is the use of sexual protection and careful facial hygiene.
pneumoniae
C. pneumoniae is a causative agent of community-acquired pneumonia, pharyngitis, laryngitis, sinusitis and bronchitis. The severity of the disease can range from asymptomatic, in most cases to severe, in rare cases. There is also evidence suggesting that C. pneumoniae could play a role in atherosclerosis, as the bacterium gains access to the vasculature during local inflammation of the respiratory tract (Watson and Alp, 2008). Isolation of the pathogen from a patient remains difficult due to its inaccessibility, so antibody tests using paired acute-and convalescent-phase sera have been used for diagnosis. If the diagnosis is clear and if the symptoms require therapy, then antibiotics such as doxycycline can be administered (www.CDC.gov).
Biphasic developmental cycle
All Chlamydiaceae are obligate intracellular pathogens sharing a biphasic developmental cycle. The gram negative bacteria exist in two different forms, the infectious elementary body (EB) and the replicative reticulate body (RB). They are easily distinguished based on their shapes, the EB being around 0.3 « m and the RB being around 1 « m. The infectious EBs attach and enter the host cell, where they stay in a parasitophorous vacuole, called inclusion, throughout their whole developmental cycle. Four to eight hours post infection (hpi) the EB converts into the RB, which is metabolically highly active and proliferates through binary fission. At around 20 hpi RBs start to convert into EBs in an asynchronous manner. By around 48 hpi most RBs have differentiated to EBs, now prepared to leave the host cell, either through lysis of the host cell or extrusion of the inclusion (Figure 5). A new cycle of infection starts (AbdelRahman and Belland, 2005).
Building the inclusion and survival
Chlamydiae are endocytosed into a membrane-bound vacuole known as the inclusion, which grows throughout the developmental cycle to harbour the increasing number of progeny (Figure 7). Interestingly, the inclusion membrane does not seem to display markers of the endocytic or lysosomal pathway (Scidmore et al., 2003). Bacterial activity is required to inhibit fusion with the lysosomal pathway, as inhibition of bacterial protein synthesis results in degradation of the bacteria in lysosomes (Scidmore et al., 1996). Early gene expression is first detected within an hour after entry, and chlamydial proteins called Inc proteins are produced, that are inserted into the inclusion membrane. Inc proteins probably play a central role in controlling cellular interactions of the nascent inclusion with the host cytoplasm, but the function of only a handful of them (out of more than 50 Inc proteins) has been investigated. One of them is probably responsible for the interaction with host dynein, resulting in the transport of the inclusion along microtubules to the microtubule organizing centre (MTOC) (Clausen et al., 1997; Grieshaber et al., 2003). While establishing the appropriate intracellular niche, the EB converts to an RB within the first few hpi (Shaw et al., 2000), which will be explained in more detail in chapter 3.2. A different set of genes, the midUnpublished image of M. Ward and C. Inman, Southampton.
cycle genes are expressed, controlling bacterial metabolism, replication and interactions with the host (Belland et al., 2003). RBs repeatedly divide by binary fission, yielding up to 1000 bacteria per infected cell (Shaw et al., 2000). At 18 to 24 hpi some RBs start to undergo conversion into EBs in an asynchronous manner, strongly linked to the switch from the transcription of mid-cycle genes to late genes (see chapter 3.2). At around 48 hpi Chlamydia exit the cell through pathogen-driven lysis of the inclusion and the host plasma membrane or through extrusion of the intact inclusion. However, the latter has only been observed in cell culture and not in vivo (Rank et al., 2011).
Ultrastructure of the chlamydial surface
Different methods for purification and fractionation of EBs were pioneered around 50 years ago, giving rise to the first descriptions of chlamydial ultrastructure (Manire, 1966; Moulder, 1962). It became clear that Chlamydiae infected cells contained two morphologically different forms: the EBs, an electron-dense coccoid structure with a diameter of about 0.3 µm, and the RBs being around 1 µm in size (Miyashita and Matsumoto, 1992). RBs also displayed a high fragility compared to EBs (Manire and Tamura, 1967). This fragility challenged (and still does) purification of RBs, shifting emphasis towards EB surface structures in subsequent studies. Manire and colleagues noticed regularly spaced hexagonal lattices covering the surface of EBs (Manire, 1966), the chlamydial outer membrane complex (COMC). It was demonstrated later on that subunits of this complex were heavily intra- and interconnected through disulfide cross-links (Hackstadt and Caldwell, 1985). The COMC is mainly composed of the major outer membrane protein (MOMP) and the cysteine rich proteins OmcA and OmcB. MOMP is also present on the surface of RBs, but in a reduced state, probably accounting for the higher susceptibility of RBs to osmotic shock. It has also been suggested that this change in the redox status contributes to EB-to-RB conversion (Hackstadt et al., 1985). The family of polymorphic outer membrane proteins (Pmps) is another abundant group of proteins on the chlamydial surface. They were first discovered when genome sequencing revealed a surprising number of autotransporters, proteins that insert their ß-barrel domain in a pore like fashion into the outer membrane in order to secrete the functional passenger domain (details can be found in chapter 4.1) (Grimwood and Stephens, 1999). C. trachomatis encodes for 9 different Pmps with a low degree of conservation. They have been proposed to be a potential source of diversity in adhesion of the bacterium to the host cell and to also play a role in chlamydial immune evasion (Becker and Hegemann, 2014). Some of theses Pmps are transcribed throughout the developmental cycle, some of them are only expressed in EBs. Generally, the composition of the enveloppes of extracellular EB and intracellular RB strongly differ and reflect their distinct requirements for the survival in two environmental surroundings. Whereas EBs need osmotic protection, to mask immunodominant epitopes on their surface and to express adhesion molecules, RBs require pores for easy acquisition of nutrients and for secretion systems to communicate with the host cell.
Host-pathogen interactions
As an obligate intracellular pathogen with a highly reduced genome of around 900 genes C. trachomatis depends strongly on the host cell (Stephens et al., 1998). Interactions with the host are indispensible for the establishment and maintenance of the niche, and for the acquisition of nutrients. They are aimed at subverting the host defence systems. Specialised secretion systems enable the delivery of bacteria-derived exoproteins that play a pivotal role in regulating host-pathogen interactions, thereby contributing to the virulence of the pathogens.
Chlamydial secretions systems
A wide range of secretion systems have been discovered in bacteria in the last decades enabling them to translocate proteins across several membranes. Chlamydiae possess three different secretion systems: Type II (T2S), type III (T3S) and type V (T5S) (Figure 9). The T2S system, a ubiquitous secretion system in Gram-negative bacteria, secretes folded proteins from the periplasm into the extracellular space. Initially, the effector is synthesized with an N-terminal signal peptide targeting it for translocation to the periplasm through a pore formed by the Sec complex. After cleavage of the signal peptide the already folded protein resides in the periplasmic space, and will subsequently be secreted through a complex of pore forming proteins, the general secretory pathway (Gsp) (Korotkov et al., 2012). The only known chlamydial T2S effector is CPAF, which can be found in the host cell cytosol (Chen et al., 2010). T2S alone does not explain translocation through the inclusion membrane, since it merely provides a release into the extrabacterial space. Small outer membrane vesicles (OMV) have been suggested to deliver T2S proteins into the host cytosol, a mechanism that had been shown to function in other bacterial systems (Ellis and Kuehn, 2010; Giles et al., 2006). Chlamydia also exploits the T5S, or autotransporter, mechanism. Similar to the T2S system, it requires the help of the Sec pathway to deliver the exoproteins to the periplasm. Once there, a C-terminal ß-barrel domain of the effector protein inserts into the outer membrane. The functional passenger domain is subsequently exposed on the bacterial surface and can be cleaved off and released, even though this is not always the case (Saier, 2006). An example of autotransporters is the family of polymorphic outer membrane proteins (Pmp), which cover the surface of EBs, playing a role in chlamydial attachment and niche adaptation (Tanzer et al., 2001). The widest spread mechanism for protein secretion into a eukaryotic cell in Gram-negative bacteria is the T3S system, which has first been shown to exist in Chlamydia in 1997 (Hsia et al., 1997). Secretion occurs through a complex secretory apparatus referred to as « injectisome », which has a very conserved structure. It includes a basal secretory apparatus spanning the inner membrane, the periplasmic space and the outer membrane. The needle complex bridges the space between the bacterial surface and the target membrane, where the tip complex is inserted. Target membranes for translocation are the host cell plasma membrane or the inclusion membrane (Betts-Hampikian and Fields, 2010). T3S systems are active in the early steps of chlamydial invasion, when they inject Tarp and probably other effectors into the host cell, and stay essential throughout the whole developmental cycle. TepP (« translocated early phosphoprotein ») is a recently identified early T3 effector protein that was proposed to act downstream of Tarp and to amplify signalling cascades (Chen et al., 2014). It has equally been shown that CT694 is a chlamydial early T3 effector, which might act upon invasion (Hower et al., 2009). Chaperones are likely to pilot substrates to the injectisome, prevent premature folding or association of proteins participating in interactions, which are to take place outside the bacteria. Few chlamydial chaperones have been discovered so far, and it is likely that not all T3S effectors require one.
In general, T3S systems are supposed to be regulated and stimulated through contact with the host cell membrane or host molecules (Hueck, 1998). While contact with the host cell membrane might trigger the translocation of early effectors such as Tarp and TepP, it is not known how the secretion of effectors from within the inclusion is regulated. Many effector proteins remain to be identified, for which computational prediction of secretion signals depicts a very useful tool. While T2S and T5S signals are well characterised, the T3S signal has not been fully elucidated yet. It could be shown that the T3S signal lies in the N-terminus or 5′ end of an effector or its mRNA (Anderson and Schneewind, 1997; Lloyd et al., 2001). Several teams have developed tools for computational predictions of T3 effectors, but in Chlamydia they are only of limited use because they fail to find T3S signals in many Inc proteins, which are all T3S substrates (Dehoux et al., 2011). Therefore, the best strategy to predict that a given protein might be a T3S substrate remains to test for the presence of a T3S signal directly in vivo. Due to given difficulties to do so in Chlamydia, heterologous secretion systems were largely used (Fields and Hackstadt, 2000; Pais et al., 2013; Subtil et al., 2005; Subtil et al., 2001).
Chlamydia might engage all these different secretion systems as each of them could be adapted to achieve physiologically distinctive goals. T2S could be responsible for bulk transport of effectors, whereas T5S is obviously a straightforward mechanism to anchor proteins to the surface. T3S effectors can be highly localized and regulated, making it the perfect mechanism for fine-tuning of host-Chlamydia interaction.
Recruitment and uptake of host organelles and lipids
Consistent with the obligate intracellular lifestyle, Chlamydiae have lost the ability to synthesize many vital metabolites. They rely on the host cell for essential nutrients, such as amino acids, nucleotides and lipids (Stephens et al., 1998). The acquisition of nutrients occurs hence through a tight interaction with the host and its organelles.
Table of contents :
TABLE OF CONTENTS
TABLE OF FIGURES
ABBREVIATIONS
ABSTRACT
RESUME (UNE PAGE)
RESUME (VERSION LONGUE)
INTRODUCTION
1. Chlamydiae
1.1 Phylogeny
2. Human pathogens
2.1 C. trachomatis
2.1.1 Trachoma, disease and history
2.1.2 Urogenital tract infections
2.1.3 Lymphogranuloma venereum
2.1.4 Diagnosis and treatment
2.2 C. pneumoniae
3. Biphasic developmental cycle
3.1 Entry to Exit
3.1.1 Attachment and Entry
3.1.2 Building the inclusion and survival
3.2 Characteristics of EBs and RBs
3.2.1 Ultrastructure of the chlamydial surface
3.2.2 Transcriptional regulation
3.3 Persistence
4. Host-pathogen interactions
4.1 Chlamydial secretions systems
4.2 Recruitment and uptake of host organelles and lipids
4.2.1 Recruitment of host lipids
4.2.2 Interaction with host organelles
4.3 Interface of interaction: the inclusion membrane
4.4 The host defence and chlamydial immune evasion
5. Chlamydial metabolism
5.1 Energy parasite hypothesis
5.2 Metabolic pathways in the post-genomic era
5.2.1 Substrate level phosphorylation
5.2.2 Oxidative phosphorylation
5.2.3 Other metabolic pathways
5.3 Glycogen
5.3.1 Chlamydial glycogen metabolism
5.3.2 Eukaryotic glycogen metabolism
MATERIAL AND METHODS
Cells and bacteria
Electron microscopy and periodic acid-thiocarbohydrazide-silver proteinate reaction
(PATAg)
Reinfection assay and flow cytometry
Quantitative Reverse Transcription PCR and Reverse Transcription PCR
Transfection
Construction of recombinant plasmids
Immunofluorescence and PAS staining
Quantification of glycogen with CellProfiler
Western Blot and antibodies
Zymogram
Heterologous secretion assay in Shigella flexneri
Transformation of C. trachomatis L2
Glucose uptake assay
RESULTS
1. Glycogen detection in Chlamydia trachomatis inclusion
2. Glucose is essential for Chlamydia trachomatis infection
3. Intraluminal glycogen is not derived from bacterial lysis
4. Kinetics of glycogen accumulation
5. Mechanisms of glycogen accumulation in inclusion lumen
5.1 Hypotheses
5.2 Vesicular import of host glycogen
5.2.1 Part of luminal glycogen is translocated in bulk from the host cytoplasm
5.2.2 Import of host glycogen and glycogen enzymes is autophagosome independent
5.3 Import of host glucose derivative
5.3.1 UDP-glucose is the host sugar transported into the inclusion lumen
5.3.2 UDP-glucose is a substrate for chlamydial GlgA
5.3.3 Identification of UDP-Glc transporter at inclusion membrane
6. Chlamydial glycogen enzymes are secreted for de novo glycogen synthesis
6.1 Heterologous test of secretion in Shigella flexneri
6.2 GlgX is present in the inclusion lumen
6.3 Overexpression of glycogen enzymes in C. trachomatis
6.4 Ectopically expressed GlgA compensates for the plasmid-less deficiency in glycogen accumulation
7. Chlamydia import Glc6P, but not Glc1P nor Glc
DISCUSSION
BIBLIOGRAPHY
