Lymphogranuloma venereum (LGV) is caused by the L1, L2 and L3 serovars of C. trachomatis. In contrast to other serovars, LGV is an invasive biovar, spreading to the lymphatic and subepithelial tissues. The infection reaches the lymphatic channels after entry through skin lesions. It starts with a lymphangitis before migrating to regional lymph nodes causing lymphadenitis and sometimes to suppurative necrosis. Since 2003, LGV proctitis cases have emerged in Europe, North America and Australia in a population of men who have sex with men. A majority of the patients were HIV-infected. It is important to note here that the ulcerative nature of LGV could facilitate both acquisition and transmission of HIV and other STDs « #$%&'()*+,$-./!.&!$/01!23435.
Diagnosis and treatments
Diagnoses are adapted to the different diseases caused by Chlamydia trachomatis. Concerning trachoma, an initial physical examination of the eye with a magnifying glass by a doctor or qualified paramedic allows the grading of the disease from one to five « 678/.9:%;1! <$=;:(1! >:(.;1! ?.;&1! @! 6$8/:%1! 4ABC5. Confirmation that Chlamydia infection is at the origin of the disease can be obtained by nucleic acid amplification tests (NAATs) – although C. trachomatis detection is sometimes impossible (Tang & Bavoil, 2012). Treatments range from surgery (to reorient the eyelid) to antibiotic treatment. Tetracycline or azithromycin administrated locally for a minimum of six weeks is recommended (Hu et al., 2010). Efforts to improve the hygienic conditions, for instance by limiting the proximity between cattle/fly companions and households, are currently the most efficient strategy to decrease the incidence of conjunctival Chlamydia infection.
Detection of Chlamydia in urogenital tracts is easily performed via NAATs on urine samples or patient-collected vaginal swabs. The typical treatment for serovars D to K is a single dose of azithromycin or seven days of doxycycline. Observational studies showed that azithromycin efficiency is about 92% (Batteiger et al., 2010).
Real-time multiplex polymerase chain reaction (PCR) allows the discrimination between LGV and non-LGV (C.-Y. Chen et al., 2006). Treatment for LGV is doxycycline twice-daily for three to four weeks. Differences in length and strength of the treatment in genital Chlamydia infection show how the precise identification of the serotypes is important.
Up to now, the best strategy to limit the spreading of Chlamydia genital infections remains prevention. Use of sexual protection and detection of potential STDs are necessary to prevent and cure the mainly asymptomatic Chlamydia infections.
Developing a vaccine remains the most appealing strategy to fight against Chlamydia infections. Difficulties due to the biology of the bacterium make the search for a protective vaccine particularly challenging. Several strategies focus on finding the best chlamydial antigen to generate a strong immune response. Outer-membrane proteins and well-studied secreted proteins are the main candidates. It is generally accepted that the best strategy is to induce both local neutralizing antibodies to prevent infection by the extracellular microbe and a cell-mediated immune response to target the intracellular infection (Hafner, Wilson, & Timms, 2014).
A bit of history: once upon a time…
Chlamydia infection is one of the oldest documented human infectious diseases. It may have originated in Central Asia based on the observations that the rate of trachoma was higher in people with Central Asian ancestry (Taboriski, 1952). In 1990, an Australian archeologist named Stephen Webb published what could be the first evidence of trachoma on an Australian aborigine dated to around 8000 years before Christ (B.C.) (Webb, 1990). Some Chinese reports from 2600 B.C. described therapies against trachoma. Other descriptions of the disease itself and related treatments were found in different civilizations: The Egyptians in their Ebers papyrus (1500 B.C.) as well as in Ancient Greece and Rome (1200 B.C. to 400 A.D.). It is in this period that the term “trachoma” (from a Greek word meaning “roughness”) was first used by the Greek doctor Dioscoride. Spreading of trachoma over the years is largely linked to wars. Crusades in the medieval age and the Napoleonic campaign in Egypt contributed to the spread of trachoma throughout Europe (Schlosser, 2013).
For a long time trachoma was an infection with no causal agent identified. It was the pioneering work of Halberstaeter and Von Prowasek in 1907, that first showed through Giemsa staining evidence of microcolonies (inclusions) observed in conjunctival epithelial cells. Visualization of transmission to baboons confirmed that it was a pathogenic agent (Freney, Renaud, Hansen, & Bollet, 1994). Around that time, it was also shown that newborns could develop trachoma because of a genital infection present in parents (Borriello et al., 2005). In 1933, the epidemiologist Bedson studied the outbreak of atypical pneumonia in Hamburg. He showed that it originated from parrots and that the infection could spread to other animals. He next qualified it as the psittacosis “virus” and during the following two decades the infectious agent was thought to be a virus (Bedson, 1936). Descriptions of cases and different serovars accumulated during the 20th century. In vitro cultures of Chlamydia were initially performed in eggs (Tang, Chang, & Huang, 1957) but, rapidly, growth of the “trachoma agent” in cell culture was introduced (Gordon, Quan, & Trimmer, 1960). Observations of the “trachoma virus” by electron microscopy suggested a bacteria-like behavior (Litwin, 1962) and lead, with many other observations, to the confirmation of Chlamydia being a bacterium in 1966 (Moulder, 1966).
Over the years, the understanding of Chlamydia biology has increased slowly. The application of novel techniques from the first sequencing of its genome (Stephens et al., 1998) to the recent ability to genetically manipulate it (Y. Wang et al., 2011) have opened many novel avenues of research.
Chlamydia, a bacterium apart from others
After the difficulty to recognize Chlamydia as a bacterium came that of its classification among bacteria. Evolutionary studies seem to date the first Chlamydiales around 700 million years ago (mya). Concerning C. trachomatis, the species appeared around 6 mya at the same time as the “human/chimpanzee” divergence (Nunes & Gomes, 2014; Steiper & Young, 2006). This obligate intracellular pathogen has a genome of about one million base pairs supplemented or not by a seven to eight thousand base pairs plasmid. Chlamydia has a rather small genome (with around 1000 genes) compared to other human bacterial pathogens such as E. coli, which has around 4000 genes (Subtil, Collingro, & Horn, 2014).
In terms of phylogeny, C. trachomatis belongs to the Chlamydiaceae family and has been recently reunited into the single genus “Chlamydia” (Stephens, Myers, Eppinger, & Bavoil, 2009). Nowadays, the classification of Chlamydia is based on whole genome sequences. Inside the C. trachomatis species, the phylogeny divergence between serovars mirrors their tissue tropism (Figure 2) (Nunes & Gomes, 2014).
Figure 2: Phylogeny of the Chlamydia genus (Nunes & Gomes, 2014). A) Phylogenic link between Chlamydia species and their respective main host and capacity to infect human. B) Phylogeny of C. trachomatis grouped by tissue tropism.
The Chlamydia special: a biphasic developmental cycle
Figure 3: Developmental cycle of C. trachomatis. Adapted from Abdelrhaman & Belland. (Abdelrahman & Belland, 2005). EBs attach via reversible and irreversible interactions and induce their entry secreting effectors like TARP. EBs are subject to a primary differentiation into RBs, which start to divide and multiply. Along the cycle, the bacteria interact with the host via effector secretion. Bacteria differentiate a second time back to the condense EB state. Under stress condition generated by different inducers, Chlamydia grow abnormaly and turn into a persistent state generating aberrant RBs. At the end of the cycle, bacteria exit in two different ways: host-cell lysis or extrusion. TTSS: type three secretion system, IF: intermediate form.
Characteristics of EBs and RBs
EBs are the infectious forms of Chlamydia. They are roughly spherical, with a diameter of about 0.3 « m, and electron dense (Matsumoto, 1981). For a long period, it was believed that EBs were metabolically inert and described as “spore-like” forms of Chlamydia. Two recent papers have demonstrated that EBs show some reduced metabolic activity (Omsland, Sixt, Horn, & Hackstadt, 2014; Sixt et al., 2013).
On the other hand, the reticulate bodies (RBs) are larger than EBs with a diameter of 1 « m. The RBs are intracellular and not adapted to the environment outside the inclusion, in which they multiply by binary fission. The characteristic inner and outer membranes of Gram negative bacteria are seen by electron microscopy (Abdelrahman & Belland, 2005). For a long time, the absence of detectable peptidoglycan, together with the sensitivity of Chlamydia to penicillin, and the presence in its genome of all the genes required for peptidoglycan synthesis constituted the “chlamydial anomaly”. However, the presence of peptidoglycan has recently been demonstrated thanks to new technologies (Liechti et al., 2013; Pilhofer et al., 2013).
The very first step: the bacterial adhesion
The first step of chlamydial invasion is its adhesion to a host cell. This process occurs in two steps. Primarily, the bacterium interacts with the host in a reversible manner via specific heparan sulfate glycosaminoglycans (HS-GAG) produced by the bacteria (Zhang & Stephens, 1992). In this paper, authors have shown an inhibition of Chlamydia adhesion by incubating the bacteria with heparin-sulfate receptors, heparin, or heparin sulfate lyase prior to infection. The effect of heparin sulfate lyase treatment was reversed by addition of exogenous heparin sulfate indicating the need of those molecules for Chlamydia adhesion. Other chlamydial adhesins are described to play a role in the attachment of the bacterium such as the major outer membrane protein (MOMP), the proteins of the outer membrane complex (OMC) OmcA and OmcB, or even the polymorphic outer membrane proteins (Pmps) (Tang & Bavoil, 2012).
Carabeo & Hackstadt in 2001 described a secondary binding step coming rather from the host cell, which is irreversible (Carabeo & Hackstadt, 2001). They observed that washing the cells with heparin after 30 min of incubation in a normal medium (without heparin) is not enough to abolish the bacterial adhesion. Thus, the authors also demonstrated that the hallmark of this secondary adhesion is temperature-dependent. Also, the Stephens’ lab described that the protein disulfide isomerase (PDI) is implicated not only in the attachment of Chlamydia (Conant & Stephens, 2007) but also in the entry of the bacterium (Abromaitis & Stephens, 2009). It remains so far the only host eukaryotic protein identified to be necessary for different Chlamydia species adhesion. Finally, it has been shown recently that sulfation is required for Chlamydia attachment (Rosmarin et al., 2012). Authors highlighted the fact that separate knockout of three host genes implicated in sulfation is sufficient to impair the bacterium binding.
Knock-knock: The Chlamydia entry
Entry of bacterial pathogens can occur in two non-mutually exclusive mechanisms: through the activation of host receptors or via the translocation of bacterial activators of endocytosis (Pizarro-Cerdá & Cossart, 2006). Both processes occur in a Chlamydia entry mechanism that is yet not fully understood.
The host cell plays an active role in the uptake of the bacterium. Entry may engage lipid microdomains in the plasma membrane. Indeed, markers of lipid rafts localizes with the bacterium even after five hours of infection by C. trachomatis (Jutras, Abrami, & Dautry-Varsat, 2003). The involvement of the actin cytoskeleton in the entry process is very well established. Inhibitors of actin polymerization such as cytochalasin D completely inhibit invasion (Boleti, Benmerah, Ojcius, Cerf-Bensussan, & Dautry-Varsat, 1999; Coombes & Mahony, 2002; Ward & Murray, 1984). Early work has shown that small GTPases of the Rho (Carabeo, Grieshaber, Hasenkrug, Dooley, & Hackstadt, 2004; Subtil, 2004) and Arf (Balañá et al., 2005) families were activated during entry.
In addition, an active mechanism induced by Chlamydia is at work. The bacterium uses the type three-secretion system (T3SS) to translocate effectors triggering its active entry. The most described secreted protein at the entry site is CT456, the translocated actin-recruiting phosphoprotein (TARP) (Clifton et al., 2004). This protein has been shown to be responsible for actin nucleation and recruitment and to be part of different phosphorylation events at the entry site of Chlamydia. This will be discussed more in detail in the second chapter.
Even if the mechanism of entry is not completely solved, it seems clear that Chlamydia enters via a receptor, possibly clathrin for C. trachomatis (Hybiske & Stephens, 2007a). Engel’s lab showed that PDGFR receptor and more importantly the FGF2 receptor are two key receptors needed for the binding and the entry of the bacterium (Elwell, Ceesay, Kim, Kalman, & Engel, 2008; Kim, Jiang, Elwell, & Engel, 2011). With all the knowledge available on those first steps, it is still not known how the bacterium escapes from early endo-lysosomal trafficking. Known makers of the early endosomes like EEA1 or Rab5 are not co-localized with the EBs or the inclusion (Rzomp, Scholtes, Briggs, Whittaker, & Scidmore, 2003; Scidmore, Fischer, & Hackstadt, 2003). Nevertheless, if EEA1 staining has been investigated before one hour of infection, it is not the case for Rab5 for which the localization has not been studied before 18 hours post infection (hpi). A rapid response of the bacterium might be the cause of this escape since a very short and transient staining of PIP3 is observed in the first minutes of entry by Lane and colleagues (Lane, Mutchler, Khodor, Grieshaber, & Carabeo, 2008). The precise mechanism still remains to be investigated.
Finally, it is very likely that the receptors and signaling pathways implicated show differences between species. For instance, the EGF receptor might serve C. pneumoniae but not C. trachomatis entry (Mölleken, Becker, & Hegemann, 2013). TARP itself shows high dissimilarities between species.
Home sweet home: Setting up the inclusion
Immediately following entry, EBs remain in a tight membrane-bound vesicle termed “inclusion”. Protected by this vacuole, the EB differentiates into a RB in a window of time comprised between 4 hours and 8 hours post infection (Shaw et al., 2000).
Relaxation of the EB’s crossed-linked membranes by reduction of disulfide bridges in its outer membrane may result from the activity of the host PDI protein (Abromaitis & Stephens, 2009). One other important transformation takes place at the bacterial nucleoid. EB DNA is highly compacted with two histone H1 homologues: Hc1 and Hc2 (Barry, Brickman, & Hackstadt, 1993). Those two proteins, tightly attached to DNA, are surprisingly not regulated by degradation or transcription. In fact, it is the translation of the genes hctA and hctB, coding respectively Hc1 and Hc2 that is impaired. Using E. coli and the lethal phenotype observed by hctA expression, Grieshaber et al. identified two genes of Chlamydia able to rescue E. coli. The first protein, IspE, seems to be indirectly implicated in the production of small metabolites necessary to disrupt the DNA-Hc1 binding (N. A. Grieshaber, Fischer, Mead, Dooley, & Hackstadt, 2004). The other gene codes for a small regulatory RNA named IhtA (inhibitor of hctA translation). This small, non-coding RNA inhibits HctA translation and does not affect transcription or the mRNA stability (N. A. Grieshaber, Grieshaber, Fischer, & Hackstadt, 2005). In a third paper, the same authors demonstrated that Hc2 regulation is mediated via IspE and not via IhtA (N. A. Grieshaber, Sager, Dooley, Hayes, & Hackstadt, 2006).
To better understand the beginning of the development cycle, different teams have studied the transcriptional profiles of Chlamydia. The paper of Shaw and colleagues described three temporal classes of genes wherein the early translocated genes correspond to the EB-to-RB differentiation (Shaw et al., 2000). While Belland et al. confirmed these results in 2003, they added the detection of new transcripts as early as one-hour post infection (Belland et al., 2003). Two different groups of genes belong to the early class: “immediate early” for 29 genes and “early” for another 200 genes.
Chlamydia immediately expresses so many genes at the beginning of its cycle because it needs to “build up” its niche. This parasitophorous vacuole is considered as “non-fusogenic” with the lysosomal pathway because it does not resemble other known eukaryotic intracellular compartments. Already after one hour of infection, markers of plasma membrane or early endosomes are not seen on the nascent inclusion (Scidmore et al., 2003). This requires bacterial activity since when bacterial protein synthesis is inhibited, the inclusion eventually fuses with lysosomes, although that event is surprisingly delayed. Lane and colleagues investigated the recruitment of several host proteins within minutes after infection. They showed that phosphatidylinositol 3,4,5- triphosphate (PIP3) is rapidly and transiently recruited to the inclusion membrane; the PIP3 binding protein Vav2 is also recruited at the entry site (Lane et al., 2008).
One chlamydial protein, CT147, has been proposed to participate in the regulation of the interaction between the inclusion and early endosomes. CT147 shows some similarities with the early endosome antigen 1 (EEA1), a marker of early endosomes. Like EEA1, it has a zinc-finger domain for PI3P binding, but lacks the Rab5 interacting domain (Belland et al., 2003). This particular Rab protein is known to regulate the fusion of EE with each other or with other vesicles (Stenmark, 2009). In their study, Belland et al. show that CT147 is detected around the inclusion from 8 hours post-infection and not before, even though its transcription is detected from the first hour of infection.
Within two hours after entry, the bacteria are trafficked toward the microtubule-organizing center (MTOC) (Clausen, Christiansen, Holst, & Birkelund, 1997). This phenomenon is dependent on the host cell dynein and on bacterial protein synthesis (S. S. Grieshaber, Grieshaber, & Hackstadt, 2003). Interestingly, the overexpression of the p50 dynamitin, which in other systems is sufficient to block dynein/cargo interaction, had no effect on Chlamydia transport. One hypothesis is that a bacterial protein present on the inclusion membrane links it to dynein to travel to the MTOC.
The presence of bacterial proteins on the inclusion is well documented. It has been estimated that 7% to 10% of Chlamydia proteome is represented by inclusion proteins (Inc) (Dehoux, Flores, Dauga, Zhong, & Subtil, 2011). This family of proteins shares the particular characteristic of having a large hydrophobic domain of sixty residues (Bannantine, Griffiths, Viratyosin, Brown, & Rockey, 2000). Among all the Inc proteins, some are transcribed between the entry moment and the first two hours of infection (Scidmore-Carlson, Shaw, Dooley, Fischer, & Hackstadt, 1999). For example, incD, incE, incF, and incG, which belong to the same operon, are transcribed during this time. The four proteins encoded by these genes are observed at the membrane of inclusions by immunofluorescence. Moreover, IncG is detectable by 2 hpi. Because Inc proteins are exposed to the host cytoplasm, they likely participate in the interactions between the bacteria and the host.
Multiplication and development: the reticulate bodies party
It is considered that each EB has fully differentiated into RB after 8 hpi (Abdelrahman & Belland, 2005). Three phases of gene transcription have been described: early, mid-cycle and late (Shaw et al., 2000). Chlamydia retains the ability to regulate gene expression transcriptionally and post-translationally. Regulation of transcription in Chlamydia remains poorly understood, however, since only a few transcription factors are known (Tang & Bavoil, 2012). Moreover, the recent description of regulating non-coding RNA (ncRNA) gives an additional level of regulation for chlamydial gene expression (Abdelrahman, Rose, & Belland, 2011).
As already mentioned above, the reticulate bodies are metabolically active and are the replicative form of the bacterium. The replication of the bacteria follows first an exponential phase with a doubling time of around 2 hours before slowing down and reaching a plateau at the end of the cycle (Shaw et al., 2000). The division mechanism per se of Chlamydia remains poorly understood. The bacteria lack certain essential genes of bacterial division such as FtsZ (Ouellette, Karimova, Subtil, & Ladant, 2012; Stephens et al., 1998). Intriguingly they express a protein associated with the rod-shape of bacillus bacteria, MreB, when Chlamydia clearly has a coccoid shape (Ouellette et al., 2012). The work of Ouellette and colleagues brought the first clues to this surprising Chlamydia characteristic. Using specific inhibitors of Pbps and MreB, they showed that the penicillin binding proteins Pbp2, Pbp3, and MreB are required for Chlamydia division. Also, the interaction of MreB with another division protein FtsK led to the proposition that MreB acts as a substitute for FtsZ. Thus, Ouellette and colleagues brought the first piece of evidence for the existence of a very unusual division mechanism in Chlamydia.
The late phase in the developmental cycle starts around 20 hpi, when RBs start to differentiate into EBs in an asynchronous manner (Shaw et al., 2000). To date, it is not known what triggers the RB-to-EB differentiation. One possibility would be a quorum sensing system even though there is no evidence of such a system or its sensors in the annotated Chlamydia genome. Still, many genes in its genome remain hypothetical and the existence of quorum sensing in a community inside the inclusion microenvironment is an attractive hypothesis. Another hypothesis of the RB-to-EB transition is that the release of the RBs from the inclusion membrane to the inclusion lumen triggers the differentiation to the EB form. The paper of Wilson and collaborators supports this theory by following the movement of both forms in live imaging (Wilson, Whittum-Hudson, Timms, & Bavoil, 2009). However, this model would only be feasible in C. trachomatis because other species of Chlamydia do not require attachment to the inclusion membrane (e.g. C. pneumoniae). The physical process of differentiation itself is better understood. The transcription of the genes hctA and hctB coding for the histone-like proteins Hc1 and Hc2 is concomitant with the timing of the differentiation (Belland et al., 2003). Moreover the same study shows the concomittant expression of gyrases, which might act on the DNA topology when it needs to be compacted. RB-to-EB transition also involves reassembly of the chlamydial outer membrane complex (COMC) including OmcA and OmcB. Disulfide cross-links are made in the COMC most likely by isomerases also expressed late in the cycle (Tang & Bavoil, 2012). The disulfide bonding of components of the type III secretion apparatus follows the same pattern, with a reduction in EB-to-RB differentiation and an oxidation in RB-to-EB differentiation (Betts-Hampikian & Fields, 2011).
The persistent form: a party forever?
The possibility of persistent C. trachomatis infection arose from the observation of seemingly chronic genital infections. Persistence is defined as a long-term association between Chlamydia and the host in which the bacteria are viable but cannot propagate. As discussed, the same patient can show several episodes of chlamydial infections in his lifetime, which can result from reinfection but also from persistence of the organism after unresolved infection. Early studies of persistence revealed that, in vitro, Chlamydia became abnormal in the presence of IFN- (Shemer & Sarov, 1985), a state later described by Wyrick as: “morphologically enlarged, aberrant, non-dividing, viable but non cultivable”. This “phenotype” can actually be observed following a variety of treatments: iron, amino acids, or nutrient starvation; exposure to penicillin or IFN-; or even during host cell maturation. Transcriptomics studies showed that the gene expression profiles differ with the inducer, suggesting that, although morphologically identical, the bacteria are not in an equivalent state. The main question in the field remains to understand how these in vitro systems reflect clinical situations (Wyrick, 2010).
The persistent stage might allow the bacteria to survive in a transiently unfavorable environment. This is observed in vitro after adding back nutrients in the culture medium for example or in vivo in the case of lack of treatment observance for patients infected by Chlamydia (Hogan, Mathews, Mukhopadhyay, Summersgill, & Timms, 2004).
Parasitism of the host cell by Chlamydia
Chlamydia has a surprisingly low number of genes (about 900 for C. trachomatis) and comparison of the pathogenic strains with their environmental relatives show that, if both have lost several biosynthetic pathways, the pathogenic strain has further reduced its genome (Omsland et al., 2014). As a consequence, Chlamydia relies heavily on the host cell for many metabolites such as amino acids and nucleotides (Collingro et al., 2011; Horn et al., 2004; Stephens et al., 1998). The main metabolites gained from the host will briefly be exposed here. The second chapter will give some details about the pathways involved.
To begin with, it has been described that Chlamydia takes up several host lipids including sphingolipids (Hackstadt, Rockey, Heinzen, & Scidmore, 1996; Hackstadt, Scidmore, & Rockey, 1995; Moore, Fischer, Mead, & Hackstadt, 2008), cholesterol (Carabeo, Mead, & Hackstadt, 2003), and glycerophospholipds (Wylie, Hatch, & McClarty, 1997) mostly for building its own membrane.
Chlamydia is able to import ATP from its host and to hydrolyze it as an energy source (T. P. Hatch, Al-Hossainy, & Silverman, 1982). Nevertheless, it has been shown that the bacterium has the capacity to produce its own ATP from glucose metabolism and possibly even oxidative phosphorylation (Iliffe-Lee & McClarty, 1999). This theory fits with the work of Schwöppe and colleagues which demonstrated the possible uptake of glucose-6-phosphate from the host (Schwoppe, Winkler, & Neuhaus, 2002). Moreover, genome sequencing of C. trachomatis showed the presence of an intact pentose phosphate pathway as well as all the enzymes necessary for glycogen storage and degradation (Stephens et al., 1998). In addition, ribonucleoside triphosphates are derived from the host mainly to synthesize bacterial RNA (T. P. Hatch, 1975).
The end of the show: a double way out ticket
At the end of the cycle, the inclusion contains several hundreds of EBs ready to infect new cells (Wyrick, 2000). In vitro, lysis of the infected cell is most commonly observed at the end of the cycle. However, early electron microscopy experiments imaging infected McCoy cells showed evidence of parts of inclusions released outside of the cell as a cell-free inclusion (la Maza & Peterson, 1982). Moreover, authors pictured some intact cells with a “crater”, interpreted as a scar left by the exit of Chlamydia inclusion without cell destruction. Years later, the group of Stephens studied more specifically this phenomenon that has been called extrusion (Hybiske & Stephens, 2007b). Using a stable GFP expressing HeLa cell line, Hybiske followed by live microscopy the evolution of the chlamydial inclusion seen as a growing black hole inside the cell. This approach allowed the authors to distinguish the two different ways of exit, lysis or extrusion, and to establish that both ways can be employed by Chlamydia (Figure 4). The lysis event starts with the rupture of the inclusion membrane followed by the bursting of the cell within a maximum of 15 min in a protease dependent manner. In contrast, extrusion is slower (3 hours) and requires actin polymerization, N-WASP, myosin II, and Rho-GTPase.
Table of contents :
I.! Chlamydia trachomatis, a human pathogen
1.! Pathology & epidemiology: which diseases and where?
b.! Urogenital infections
c.! Lymphogranuloma venereum
d.! Diagnosis and treatments
2.! A bit of history: once upon a time
3.! Chlamydia, a bacterium apart from others!
4.! The Chlamydia special: a biphasic developmental cycle
a.! Characteristics of EBs and RBs
b.! The very first step: the bacterial adhesion
c.! Knock-knock: The Chlamydia entry
d.! Home sweet home: Setting up the inclusion
e.! Multiplication and development: the reticulate bodies party
f.! The persistent form: a party forever?
g.! Parasitism of the host cell by Chlamydia
h.! The end of the show: a double way out ticket
5.! Chlamydia’s swiss knife: the secretion systems
a.! The T2SS: the general secretory pathway (GSP)
b.! The T5SS: the twin-arginine translocation (tat) pathway
c.! The T3SS: the chlamydial “injectisome”
6.! The challenge Chlamydia: from darkness to light
a.! A long time in the darkness
b.! … and then there was light: welcome to a new Chlamydia world.
II.! Interactions between Chlamydia and host membranes
1.! The first Chlamydia interaction: entering the cell
2.! Chlamydia fattens up: delivery of lipids to the inclusion
3.! A sociable guest: multiple interactions between Chlamydia and host organelles
a.! Chlamydia attracts most if not all the host organelles
b.! The Inc proteins: key components of the Chlamydia-host cell interaction
c.! Manipulation of the Rab proteins, key trafficking regulators
4.! Chlamydia trachomatis infection impairs cytokinesis
5.! The defensive answers of the host
6.! Chlamydia and ESCRT
III.! Biology of the ESCRT system
1.! Overview of a machinery with many complexes
2.! One system, many functions
a.! MVB biogenesis
b.! Abscission during cytokinesis
c.! Virus budding
d.! Exosomes secretion
3.! Focus on Hrs
a.! From Hrs to the ESCRT-0 complex
b.! Dissection of Hrs: domains, interactions, regulations
c.! Hrs, a multi-functions protein
4.! Focus on Tsg101: a central protein for the ESCRT system
5.! ESCRT system, where else?
I.! Chlamydia strains
II.! Cell culture, transfection and chemicals
III.! Cloning procedures
IV.! Production and purification of recombinant protein
V.! Yeast-two-hybrid (Y2H) assays
VI.! Immunofluorescence and western blotting analysis
VII.! Quantification of bacterial entry
VIII.! Flow cytometry on infected cells and EBs
IX.! Immunoprecipitation (IP)