Toxoplasma and Chlamydia trachomatis are obligate intracellular pathogens that have evolved analogous strategies to replicate within mammalian cells. Both pathogens are known to extensively remodel the cytoskeleton, and to recruit endocytic and exocytic organelles to their respective vacuoles. However, how important these activities are for infectivity by either pathogen remains elusive. Here, we have developed a novel co-infection system to gain insights into the developmental cycles of Toxoplasma and C. trachomatis by infecting human cells with both pathogens, and examining their respective ability to replicate and scavenge nutrients. We hypothesize that the common strategies used by Toxoplasma and Chlamydia to achieve development results in direct competition of the two pathogens for the same pool of nutrients. We show that a single human cell can harbour Chlamydia and Toxoplasma. In co-infected cells, Toxoplasma is able to divert the content of host organelles, such as cholesterol. Consequently, the infectious cycle of Toxoplasma progresses unimpeded. In contrast, Chlamydia's ability to scavenge selected nutrients is diminished, and the bacterium shifts to a stress-induced persistent growth. Parasite killing engenders an ordered return to normal chlamydial development. We demonstrate that C. trachomatisenters a stress-induced persistence phenotype as a direct result from being barred from its normal nutrient supplies as addition of excess nutrients, e.g. amino acids, leads to substantial recovery of Chlamydia growth and infectivity. Co-infection of C. trachomatis with slow growing strains of Toxoplasma or a mutant impaired in nutrient acquisition does not restrict chlamydial development. Conversely, Toxoplasma growth is halted in cells infected with the highly virulent Chlamydia psittaci. This study illustrates the key role that cellular remodelling plays in the exploitation of host intracellular resources by Toxoplasma and Chlamydia. It further highlights the delicate balance between success and failure of infection by intracellular pathogens in a co-infection system at the cellular level.
Obligate intracellular pathogens represent a subset of microbes that have lost the capacity to live independently of their host. A hallmark of these pathogens is a significantly reduced genome size highlighted by deletions of DNA segments that encode entire metabolic pathways. Essential metabolic and biosynthetic intermediates that would normally be provided by these pathways must then be retrieved from the host by the pathogen (reviewed in Fraser-Liggett, 2005). Indeed, although separated from the nutrient-rich cytosol by a membrane, these pathogens have nevertheless evolved efficient strategies to co-opt multiple host cellular pathways and hijack host organelles in order to acquire essential nutrients. Among obligate intracellular bacteria, Chlamydia trachomatis infects a wide range of cell types but replicates primarily within mucosal epithelial cells. A chlamydial infection is initiated by the internalization of the extracellular form of the bacterium, the ‘elementary body’ (EB; summarized in Rockey and Matsumoto, 2000). This process, previously termed parasite-specified phagocytosis (Byrne and Moulder, 1978), involves interactions between various chlamydial ligands and receptors at the host cell surface (reviewed in Dautry-Varsat et al., 2005). The membrane of the nascent Chlamydia-containing vacuole (‘the inclusion’) is derived from the host plasma membrane, whose components are promptly lost and replaced by chlamydial proteins (Scidmore et al., 2003), resulting in the disengagement of the inclusion from host vesicular trafficking pathways (reviewed in Fields and Hackstadt, 2002). In the inclusion, the EB differentiates into the replicating form of the bacterium, the ‘reticulate body’ (RB). After multiple binary fissions, the RBs differentiate to progeny EBs that are then released from the cell upon lysis or exocytosis, with each new EB capable of eliciting infection of a new cell in the same host or a new host.
In comparison with C. trachomatis, the obligate intracellular protozoan parasite Toxoplasma gondii is adapted for invasion and multiplication in virtually all nucleated mammalian cells (reviewed in Sibley, 2003). Unlike Chlamydia that exploits the host endocytic machinery for internalization into a vacuolar compartment, Toxoplasma actively invades cells and creates its own membrane-bound compartment that is immediately impervious to host microbicidal mechanisms. Many parasite proteins are incorporated into the membrane of the Toxoplasma-containing vacuole (‘the parasitophorous vacuole’, PV), making this compartment a unique ‘organelle’ devoid of any salient host features. Within the PV, Toxoplasma replicates by endodyogeny, a process by which two daughter cells are formed within the parent cell; upon host cell lysis, the newly formed parasites are released to the surrounding environment and able to invade new cells.
Despite the different mechanisms used by C. trachomatis and T. gondii to achieve cell entry and to avoid recognition and degradation by host lysosomes, these phylogenetically distant microbes have developed similar strategies to subvert host cell functions for their own benefit. For example, both pathogens target the host microtubular cytoskeleton to facilitate entry into mammalian cells (Clausen et al., 1997; Sweeney et al., 2010). Once internalized, both travel on microtubules to the host perinuclear area towards the microtubule-organizing centre (MTOC), where their vacuole is encased in a scaffold of host cytoskeletal structures (Schramm and Wyrick, 1995; Grieshaber et al., 2003; Coppens et al., 2006; Walker et al., 2008; Johnson et al., 2009). Both pathogens have a marked requirement for host cell lipids, such as sphingolipids and cholesterol (Hackstadt et al., 1995; 1996; de Melo and de Souza, 1996; Coppens et al., 2000; Carabeo et al., 2003; Capmany and Damiani, 2010; Derré et al., 2011; Elwell and Engel, 2012). From their respective compartments, both chlamydiae and Toxoplasma parasites are equally proficient at selectively attracting host endocytic and exocytic organelles and retrieving their lipid content (Hackstadt et al., 1996; Beatty, 2006; 2008; Coppens et al., 2006; Romano et al., 2008).
Instances of cells dually infected with different prokaryotic and/or eukaryotic pathogens have been previously reported (Meirelles and De Souza, 1983; Black et al., 1990; Rabinovitch and Veras, 1996; de Chastellier et al., 1999; Sinai et al., 2000; Vanover et al., 2008; Borel et al., 2010). Various cell types can temporally shelter two different pathogens that may even reside within the same vacuolar compartment as in the case of Mycobacterium avium- and Coxiella burnetti-infected macrophages (de Chastellier et al., 1999). Although these represent artificially constructed in vitro model systems, dually infected cells may provide unique opportunities not only to evaluate the compatibility of two different pathogens during co-infection but also to gain fundamental knowledge on each of the pathogens. The goal of this study is to evaluate the important contribution of host organelles – and their nutrient-rich content – to the intracellular development of C. trachomatis and Toxoplasma, and therefore the necessity for these pathogens to re-route host organelles to their vacuoles. Our hypothesis is that the shared abilities of Chlamydia and Toxoplasma to usurp host organelles would make these pathogens competing co-occupants in the same cell. To verify this assumption, we have examined whether the presence of Chlamydia and Toxoplasma within infected cultured fibroblasts alters the intracellular fate of either pathogen-containing vacuole. The pathogenic strategies that are essential to either Chlamydia or Toxoplasma infectivity may also be revealed by comparing the behaviour of either pathogen in dually versus singly infected cells. Specifically, the following questions will be addressed: Can Chlamydia and Toxoplasma simultaneously invade and remain within the same cell? If present in the same cell, are these pathogens confined to and replicating normally within their usual vacuole? Do these pathogens occupy segregated compartments or do their vacuoles interact with each other during co-infection? If nutrient depletion is induced by one or both pathogens, can the growth of either pathogen be enhanced by intracellular nutrient supplementation? Alternatively, does the stress of competition for the same nutrient pool severely alter either pathogen's scavenging activities?
Our studies have revealed that a single mammalian cell can harbour both C. trachomatis and Toxoplasma, and that the two pathogens reside in distinct vacuolar compartments throughout co-infection. In co-infected cells, the infectious cycle of Toxoplasma progresses normally, and the parasite proficiently exploits host organelles, independently of the presence of chlamydial inclusions. In contrast, normal chlamydial development is arrested in Toxoplasma-infected cells. Selective elimination of the parasite restores normal development, suggesting that the disruption is directly caused by co-infecting Toxoplasma. Chlamydial developmental arrest presumably results from global nutrient deficiency due to competition with Toxoplasma as replenishment of the medium with selected nutrients restores productive chlamydial growth and development to infectious progeny.
Fibroblasts can be co-infected with Chlamydia and Toxoplasma in vitro
We have established an in vitro cell culture model whereby fibroblasts are exposed simultaneously to C. trachomatis (serovar E) and T. gondii (RH strain) and infected with both pathogens. This system is suitable for investigating potential interactions between these two pathogens, and possible alterations to the developmental biology of either pathogen as a result of their co-occurrence in the same cell (see details in Experimental procedures). Human fibroblasts were chosen for these studies because both Chlamydia and Toxoplasma are able to efficiently infect these cells, and infectious progeny is produced within 3 days of the onset of infection in vitro. Culture conditions were first evaluated using fibroblasts incubated in the presence of either Chlamydia or Toxoplasma. Essential features of the growth of either pathogen in these cells were comparable to those reported in other cells routinely used to cultivate these two pathogens (not shown).
To examine whether individual fibroblasts can be co-infected with C. trachomatis and T. gondii, we incubated the cells simultaneously with the two pathogens for 24 h (Fig. 1A). Immunofluorescence microscopy was used to detect the infecting microorganisms using antibodies against Toxoplasma GRA7 (a marker of the PV membrane) and Chlamydia EF-Tu (a cytoplasmic marker of EBs and RBs). Results reveal that the two pathogens could invade and remain within the same fibroblast. Cells infected with C. trachomatis and T. gondii were observed with high frequency. Interestingly, several inclusions and PVs could often be detected in the same cell, usually gathered in the host perinuclear region as reported for mono-infections (Grieshaber et al., 2003; Coppens et al., 2006). Although they were inoculated simultaneously, Chlamydia and Toxoplasma never shared the same vacuolar compartment after invasion, consistent with their distinct modes of cell entry and with their dissimilar vacuolar membrane composition precluding recognition and fusion. The process of active invasion mediated by Toxoplasma was substantiated by the detection of a typical PV membrane surrounding the parasite containing GRA7, a protein secreted into the PV 20 min post infection (p.i.). Throughout infection, the two pathogen-containing vacuoles never interact directly with one another, likely as a consequence of not having a ‘matching’ recognition signal.
During co-infection, inclusions display altered morphology whereas the size of the PV is normal
During a 24 h period of intracellular growth, Toxoplasma normally undergoes three cycles of endodyogeny (doubling time ∼ 8 h) and a PV may contain up to eight daughter cells (7 × 2 μm in size). In the same time span, RBs (1 μm in diameter) within a chlamydial inclusion replicate by binary fission (doubling time ∼ 2–3 h; Wilson et al., 2004) while attached to the limiting membrane of the inclusion to a number of 100–200. We next analysed in detail the course of Toxoplasma and Chlamydia development during their co-habitation in the same cell.
In human foreskin fibroblasts (HFF) doubly infected for 24 h, large PVs containing dividing parasites were clearly observed as in mono-infections (Fig. 1A). Some smaller PVs were visible, likely owing to asynchronous invasion events that are known to be frequent in mono-infected cells. In contrast to single infections, the size and the content of chlamydial inclusions were abnormal in mixed infections regardless of the number of parasites per cell. Inclusions were smaller and contained relatively few typical RBs juxtaposed to the inclusion membrane. Moreover, several anomalously enlarged bacterial forms up to 4 μm in diameter were observed in place of 1 μm RBs, particularly in inclusions in close proximity to a PV. These aberrant inclusions morphologically resemble altered chlamydial inclusions that appear under certain stressful conditions (e.g. nutrient starvation or exposure to antibiotics) and that contain RBs that have limited or no capability to divide (Beatty et al., 1994; Skilton et al., 2009; Wyrick, 2010).
Altered inclusion development in the presence of Toxoplasma is independent of host cell type
We extended our co-infection studies using epithelial cells as these cells are preferentially infected by chlamydiae in vivo. HeLa cells and HCT-8 cells were co-infected for 24 h with C. trachomatis and RFP-expressing T. gondii (Fig. 1B). Results illustrate that inclusions in epithelial cells harbouring parasites also contained enlarged, morphologically aberrant chlamydiae, suggesting that chlamydial stress occurs regardless of the nature of the cell type. Quantitative image analysis was used to compare inclusion sizes in mono-infected versus co-infected HFF, HeLa cells and HCT-8 cells with Toxoplasma at 24 h p.i. (Fig. S1). Results show that the inclusions were reduced in size by approximately twofold in dually infected cells (2.3-fold for HFF versus 1.75-fold for epithelial cells). In parallel, we examined the capability of C. trachomatis to produce infectious progeny after 24 h co-cultivation with Toxoplasma. The infectious yield was determined for a control mono-infection for comparison. Re-infection analysis indicates that ∼ 30% of the infectious organisms could be rescued from co-infected cultures (Fig. 1C). This suggests that, although the production of infectious chlamydial progeny was largely diminished due to the presence of PVs compared with non-parasitic infected cells, a proportion of the developmentally aberrant bodies progress to infectious EBs once the parasites are removed.
The parasite growth measured by enumeration of organisms per PV was identical in mono-infected and co-infected fibroblasts or epithelial cells at 24 h p.i. (Fig. 1D). Beyond 72 h p.i., host cell lysis mediated by Toxoplasma occurred and progeny was released in the environment (not shown). This demonstrates that the parasites were able to undergo normal endodyogeny independently of the presence of chlamydial inclusions. Taken together, these results indicate that the simultaneous co-infection of fibroblasts or epithelial cells by Chlamydia and Toxoplasma is most detrimental to the bacterial pathogen. Chlamydial development was altered as illustrated by the presence of abnormally enlarged, non-dividing chlamydial forms in dually infected cells.
Co-infection with Toxoplasma induces ultrastructural changes in inclusions similar to those from chlamydial persistent forms
The production of infectious chlamydial EBs isolated from co-infected cells in subculture demonstrates that these bacteria exist in a viable, yet non-cultivable (persistent) state. The altered morphology of in vitro persistent aberrant forms of C. trachomatis has been well described at the ultrastructural level (Matsumoto and Manire, 1970; Beatty et al., 1994; Byrne et al., 2001). Under exposure to penicillin, aberrant RBs are characterized by an irregular shape, an enlarged non-dividing body, and/or the shedding of excess membrane as outer membrane blebs or intra-periplasmic vesicles.
To assess possible morphological resemblance of RBs in co-infected cells with penicillin-induced aberrant RBs, we examined the ultrastructure of chlamydiae grown in cells containing Toxoplasma for 24 h (Fig. 2). When bacteria were allowed to develop alone in host cells, we observed as expected large inclusions containing many actively dividing RBs that were homogenous in size and attached to the inclusion membrane (Fig. 2A and B). In contrast, chlamydiae residing in PV-containing fibroblasts were impaired in their replication as illustrated by the presence of enlarged bacterial bodies and the lack of morphological evidence of septum formation or cell division (Fig. 2C–E). A wide range of chlamydial inclusion sizes (1–10 μm in diameter) was noted, suggesting an asynchronous and dysregulated development of the inclusions. The aberrant bodies had variable electron-density and empty envelopes (ghosts) were often visible (Fig. 2D), suggestive of the blebbing of excess chlamydial outer membrane. Interestingly, host lipid bodies were detected in the inclusion lumen (Fig. 2E), indicating that some scavenging activities were retained by the bacteria (Cocchiaro et al., 2008). Moreover, Golgi ministacks were visible around the inclusions in PV-containing cells (Fig. 2F), suggesting that the bacteria were able to fragment the Golgi apparatus as in mono-infected cells (Heuer et al., 2009; inset in Fig. 2F). Finally, we observed ER tubules in close apposition to the inclusion membrane (Fig. 2G) as has recently been described during C. trachomatis infection (Derré et al., 2011). No cytopathies were observed for Toxoplasma in co-infected cells.
These results confirm that the normal developmental cycle of Chlamydia is disrupted in PV-containing cells. The chlamydiae exhibit morphological features of the aberrant RB phenotype that results from exposure to stress. This suggests that the presence of Toxoplasma in the same cell is by some means stressing for the chlamydiae.
Toxoplasma causes stress-induced persistence of Chlamydia in co-infected cells
To further confirm a bacterial shift from normal development to a state of stress-induced persistence in Toxoplasma-infected cells, we monitored the level of chlamydial proteins that are normally expressed during late development. Among these, the polymorphic membrane proteins (Pmps) are expressed at the surface of chlamydiae, and are implicated in virulence (Tan et al., 2010) and immune protection (Crane et al., 2006). As a potent inducer of aberrant RBs of C. trachomatis in vitro, penicillin causes the global downregulation of late-expressed genes, including several pmp genes (Skilton et al., 2009; Carrasco et al., 2011).
As a positive control in our studies, we exposed Chlamydia-infected cells to 100 units ml−1 of penicillin, a concentration that has previously been shown to be optimal for blocking chlamydial late differentiation when added at the mid-development stage (24 h p.i.; Lambden et al., 2006), before immunostaining with anti-PmpB antibodies. Our results confirm the negligible production of PmpB in chlamydiae exposed to penicillin (Fig. S2). In mono-culture conditions, late differentiating chlamydiae specifically distributed in the centre of the inclusion produced PmpB, whereas replicating RBs positioned at the inclusion periphery did not (Fig. S2). However, in co-infections, the production of PmpB was minimal: large aberrant bodies were negative for PmpB whereas normal-sized RBs were stained modestly (Fig. 3A, compare the staining pattern in mono- and co-infected cells from the same monolayer). Measurement of fluorescence intensity confirms a substantial reduction by ∼ 90% of the PmpB signal in inclusions cultivated with Toxoplasma (Fig. 3B). This result further strengthens our previous findings that in Toxoplasma-infected cells, the chlamydial inclusion enters a typical stress-induced persistent state.
We also investigated whether Toxoplasma itself suffers distress while in co-culture with Chlamydia. Exogenous stress factors, i.e. alkaline pH, IFN-γ, heat shock or deprivation of nutrients, can trigger the differentiation of Toxoplasma into a slow replicating cyst form (reviewed in Ferreira da Silva Mda et al., 2008). This stage differentiation is largely viewed as a stress-related response to hostile environmental conditions. A major feature of parasite encystation is the transformation of the PV membrane into a thick cyst wall protecting Toxoplasma from the surrounding environment. Detection of the presence of a cyst wall can be assessed by Dolichos lectin staining upon incubation of infected cells in alkaline conditions (Fig. 3C, compare panels a and b). As implied by the observed normal replication rate of Toxoplasma in cells containing chlamydial inclusions, we did not detect a single PV labelled for the cyst wall marker, indicating that the presence of Chlamydia in the cell does not conjure a harmful environment for the parasite.
We next examined whether the production of chlamydial PmpB during co-infection could be influenced by the number of parasites per cell or by the distance separating a given inclusion from a PV. A single PV per cell was enough to abolish chlamydial PmpB production (Fig. S3A). The levels of PmpB were slightly higher in inclusions that were distant from the parasites than in those in close proximity to the PV, but even in inclusions far away from PV, PmpB production still remained considerably lower than that in inclusions from mono-infections (Fig. S3B).
Toxoplasma killing in co-infected cells restores normal chlamydial growth
Persistence of chlamydiae in vitro (i.e. long-lasting residence within a cultured cell) is defined as the dual ability of these bacteria for prolonged survival within the host cell without overt growth when exposed to stress (up to 40 days), and for reversion to normal growth upon removal of the stressor (Beatty et al., 1994; Robertson et al., 2009; Skilton et al., 2009; Wyrick, 2010). We thus undertook to verify whether the stress-induced persistent-like state of Chlamydia in co-infected cells could be reversed to normal development upon the killing of Toxoplasma. The anti-toxoplasmosis drug pyrimethamine is known to inhibit the growth of T. gondii by blocking the activity of the parasite dihydrofolate reductase, and therefore the production of folates (Derouin and Chastang, 1989). We confirmed that exposure of Toxoplasma-infected cells to 10 μM pyrimethamine for 24 h led to a progressive and irreversible decrease in parasite metabolic activities, as evidenced by striking morphological changes in T. gondii, including fragmented nuclei and abnormal daughter cell formation (Fig. S4). Microscopic observations showed that C. trachomatis development and replication were unaffected by pyrimethamine treatment for 24 h (Fig. S4), and even prolonged incubations with the drug did not delay bacterial egress (data not shown). Verification of the selective effects of pyrimethamine on Toxoplasma was further assessed by infecting cells with Toxoplasma and adding pyrimethamine at the time of invasion for a period of 24 h before Chlamydia infection for 1 day (Fig. 4A, exp. A). Results show the absence of parasite replication and normal chlamydial growth, as expected. When cells were incubated with Toxoplasma for 24 h, then treated with pyrimethamine for an additional 24 h before adding Chlamydia in the culture medium, we observed normal chlamydial development despite the prior transformations of the cells by the parasites and the recruitment of host organelles by the parasite (Fig. 4A, exp. B). Quantitative analyses of these data confirm normal inclusion sizes in Toxoplasma-infected cells exposed to pyrimethamine prior to bacterial infection (Fig. 4B, panel a). In contrast, Chlamydia did not develop normally in cells preinfected with Toxoplasma 24 and 48 h as expected (Fig. S5). These experiments reveal that Chlamydia adeptly multiplies in cells previously infected with Toxoplasma when the parasites are dead prior to bacterial invasion. The observations that the progression of T. gondii infection for 24 h or even 48 h before pyrimethamine addition did not alter chlamydial growth, suggests that an active parasite metabolism is required for the switch of Chlamydia to a stress-induced persistent state.
To demonstrate the reversibility of the persistent state of C. trachomatis during co-infection, cells were infected with Toxoplasma and Chlamydia together for 24 h, and then exposed to 10 μM pyrimethamine for an additional 24 h. The inclusion morphology was inspected using anti-EF-Tu antibodies (Fig. 4A, exp. C). Remarkably, the normal development of the bacteria was restored under pyrimethamine treatment as documented by the recovery of normal size RBs and the disappearance of the large aberrant bodies. The resumption of productive development for C. trachomatis upon treatment with the anti-parasitic drug was probed by scoring PV size and progeny formation. Forty-eight hours post infection, no morphological difference in the inclusion content and size was noticeable between cells infected with Chlamydia alone and those containing Toxoplasma and treated with the drug. This was confirmed by quantitative image analysis (Fig. 4B, panel a). This suggests a relatively fast transition of aberrant bodies to RBs, followed by rapid chlamydial divisions concomitant to the progressive deterioration of Toxoplasma by pyrimethamine. This reversion also occurred in co-infected cells containing multiple PVs (Fig. S6). In this case, we observed that the inclusions encircled the PVs containing dead parasites tightly, in sharp contrast to co-infections with live parasites where the inclusions appeared to remain distant from the PVs. The restoration of productive bacterial growth was measured as inclusion-forming units (IFU). Results confirm that C. trachomatis grown in pyrimethamine-treated cells developed into infectious EBs, equally as efficiently as chlamydiae from mono-cultures (Fig. 4B, panel b).
Finally, to further ascertain that the inclusions underwent late developmental differentiation in pyrimethamine-treated co-infected cells, we stained the bacteria with anti-PmpB antibodies (Fig. 4C). Results reveal that, upon drug treatment for 24 or 48 h post co-infection, chlamydial inclusions had regained the ability to produce the late-expressed PmpB similarly to normally growing inclusions in mono-infections (PmpB fluorescence intensity in A.U.: 0.88 ± 0.15 versus 0.79 ± 0.2 in mono-cultures and co-cultures respectively).
Overall, these results indicate that the presence of co-infecting Toxoplasma in a Chlamydia-infected cell induces a typical state of persistence in the bacterium. Indeed, chlamydiae remain dormant thus viable throughout co-infection, and are then able to resume normal development and regain infectivity upon death of the parasites.
Toxoplasma outcompetes Chlamydia for host cholesterol during co-infection
Having established that chlamydial inclusions enter a persistent state in cells co-infected with Toxoplasma, we focused on characterizing the specific mechanism leading to bacterial stress. Several immune system-mediated factors (e.g. IFN, TNF) induce alterations in chlamydiae development similar to those produced by exposure to antimicrobial agents (e.g. penicillin) and contribute to persistent growth (Beatty et al., 1994). The lack of production of cytokines by fibroblasts or epithelial cells infected with Toxoplasma makes a process linked to immunologically induced persistence unlikely. Alternatively, competition between C. trachomatis and Toxoplasma for the same pool of nutrient and the demonstration that the parasite holds a significant competitive advantage over the bacterium leads to the hypothesis that a mechanism of nutrient deficiency-induced persistence likely occurs for Chlamydia.
We first examined whether arrested chlamydial development correlated with a reduced ability of the chlamydiae to retrieve nutrients from host organelles in co-infected cells. Among such nutrients, host lipids, e.g. sterols are important for both C. trachomatis and T. gondii growth and infectivity. Neither chlamydiae nor Toxoplasma parasites can synthesize sterols. These pathogens acquire cholesterol from the extracellular environment via the low-density lipoprotein (LDL) pathway (Coppens et al., 2000; Carabeo et al., 2003). Cholesterol molecules that are produced in the host cell are also transported to the chlamydial inclusion via a Golgi-dependent pathway. The individual requirements for host cholesterol by Toxoplasma and Chlamydia put the two pathogens in direct competition for the acquisition of this lipid from the host cell, at least as it applies to the LDL source of cholesterol. To verify that C. trachomatis can take up sterols directly from the external milieu, we incubated infected cells with fluorescent NBD-cholesterol incorporated into LDL for 2 h (Fig. 5A). Results show that the envelope of individual chlamydiae was intensely stained with the fluorescent lipid. The fluorescent labelling of the inclusions was abrogated by ∼ 85% with exposure to penicillin, confirming the active participation of the bacteria in the process of cholesterol scavenging (Fig. 5B). Cells were co-infected with Chlamydia and Toxoplasma for 24 h before incubation with [NBD]cholesterol-LDL under the same conditions as for mono-infection. In these cells, the PVs exhibited bright fluorescence on the plasma membrane and perinuclear area similar to that detected on parasites in mono-infected cells (Figs S7 and 5A). In contrast, inclusions in PV-containing cells showed negligible NBD staining, corresponding to less than 5% of those in mono-infected cells (Fig. 5A and B). This suggests that the bacteria are unable to scavenge cholesterol-LDL in the presence of Toxoplasma. When cells were co-infected for 24 h, then treated with pyrimethamine for 24 h before a 2 h pulse with [NBD]cholesterol-LDL, NBD fluorescent signal reappeared on the inclusions (Fig. 5A). After 2 h, the fluorescent cholesterol content associated with inclusions was ∼ 30% of that in inclusions in mono-infections (Fig. 5B), suggesting that the chlamydial ability to retrieve sterols from the host cell was restored upon parasite death.
These data were further confirmed by visualization of cholesterol distribution at steady state on the pathogens using filipin, a fluorescent polyene antibiotic that binds the 3′ hydroxyl group of membrane sterols (Volpon and Lancelin, 2000). The membranes of Toxopasma and C. trachomatis stained consistently and intensely with filipin throughout development in mono-infected cells (Fig. 6A). Filipin staining was also clearly visible on the inclusion membrane. Fibroblasts were next co-infected for 24 h and stained for filipin to monitor cholesterol content associated with the PV and the inclusion (Fig. 6B). In every view, the PVs showed fluorescence intensity similar to that of PVs in mono-infected cells. In contrast, the inclusions exhibited weaker fluorescence staining in co-infections than in single infections, and the labelling was essentially restricted to the inclusion membrane (Fig. 6B). Only 7% of the inclusions displayed staining on both the inclusion membrane and the envelope of some of the chlamydiae (Fig. 6C). Other inclusions contained either a few bacteria that were filipin-positive (49%) or unstained bacteria (44%).
These results indicate that Chlamydia scavenges host cholesterol poorly in co-infected cells. This diminished capability may be due to alterations in sterol trafficking pathways in co-infected cells, thereby blocking cholesterol access to the inclusions, causing a scarcity of available cholesterol as a consequence of its diversion to the Toxoplasma PV, and/or resulting in a reduced need for nutrients by the bacteria owing to their stressed state. Alternatively, the aberrant RB state may be triggered by the restricted supply of nutrients due to the rivalry with Toxoplasma to acquire the same host molecule.
Reintroduction of selected nutrients in the medium results in substantial recovery of chlamydial infectivity
The growth asymmetry between Toxopasma and C. trachomatis during co-infection could be attributed to the more successful sequestration of limited nutritional resources – as shown at least for ldl-cholesterol – by the parasite. To explore the nutrient shortage hypothesis, the culture medium was supplemented with an excess of selected nutrients with the objective to identify the nutrient/s and cofactor/s that is/are critical for Chlamydia development. Apart from lipids, Chlamydia development requires iron, lipoic acid and specific amino acids from the host cell (reviewed in Ramaswamy and Maurelli, 2010; Wyrick, 2010). In a first set of experiments, we supplemented the ‘infection medium’ [Dulbecco's modified Eagle's medium (DMEM) + 10% fetal bovine serum (FBS)] with a large nutrient pool including lipids (cholesterol, sphingomyelin and triacyglycerols), iron provided by ferric chloride and holo-transferrin, lipoic acid, and essential and non-essential amino acids (‘nutrient-enriched medium’) and monitored the inclusion size and progeny formation 24 h post co-infection (Fig. 7 and Table 1). Data in Fig. 7A illustrate that in the ‘nutrient-enriched medium’, chlamydial inclusions could develop, forming typical small RBs, in sharp contrast to chlamydial inclusions maintained in the ‘infection medium’ without any addition. The inclusion size in fibroblasts containing a single PV was almost identical (∼ 90%) to that of bacteria from mono-cultures (Table 1). However, the inclusion size in co-infected cells, i.e. the total number of RBs per inclusion was influenced by the number of PV within the cell (Fig. 7A and B). In cells containing 5–10 PVs, smaller inclusions were visible compared with inclusions grown in cells containing one to five PVs. However, those smaller inclusions contained very few aberrant bodies, suggesting that their occurrence was likely resulting from space constraints imposed by large growing PVs in the cell. Chlamydial IFU measurements further revealed that the production of infectious progeny in co-cultures in nutrient-rich medium was comparable to that in chlamydial mono-cultures, confirming that most RBs in the inclusions were not developmentally arrested but infectious, and had developed to fully infectious EBs (Fig. 7C).
Table 1. Effect of nutrient supplementation in the medium on C. trachomatis growth in co-infected cells
HFF were co-infected with T. gondii and C. trachomatis for 24 h in the presence of various nutrients at the indicated concentrations before scoring the size of chlamydial inclusions. Upon these incubations, the PV size remained statistically unchanged (not shown).
We next sought to identify which of the nutrients added in excess to the medium was responsible for stimulating chlamydial development in co-infected cells. Different classes of nutrients were introduced separately in the culture medium at various concentrations to evaluate their impact on the restoration of chlamydial development (Table 1). Addition of ferric chloride had a modest but significant concentration-dependent effect on the restoration of bacterial growth, with a ∼ 1.8-fold increase of the inclusion size, whereas transferrin saturated in iron was ineffective. Like chlamydiae, Toxoplasma scavenges host lipoate (Crawford et al., 2006), making this cofactor coveted by both pathogens. However, supplementation of lipoic acid up to 10 μM in the medium did not reinstate normal chlamydial development. A mixture of cholesterol, sphingomyelin and four different triacylglycerols (as a source of fatty acids) slightly restored bacterial growth (∼ 1.6-fold) at the highest concentration tested in co-infected cells. Interestingly, addition of excess exogenous amino acids resulted in the near complete reestablishment of C. trachomatis infectivity. In contrast, excess non-essential amino acids did not statistically restore normal chlamydial growth. Conversely, omission of essential amino acids from the ‘nutrient-enriched medium’ led to a ∼ 70% decrease in inclusion size when compared with inclusions grown in ‘nutrient-enriched medium’. These results confirm the importance of essential amino acids for the optimal intracellular development of C. trachomatis.
Altogether, these data support our hypothesis that depletion of selective nutrients from the medium, therefore from the host intracellular pools, are responsible for the interruption of the normal programme of infectivity of C. trachomatis and production of infectious EBs in co-infected cells. This strongly suggests that the parasites are more efficient at scavenging host nutrients, which leads to depletion in the intracellular pools for Chlamydia. Addition of exogenous nutrients allows the host cell to replenish its pools, providing extra nutrients for the bacteria, Toxoplasma having by then presumably engorged itself.
In cells infected with slow growing Toxoplasma strains, C. trachomatis development is moderately affected
To further validate our hypothesis of nutrient deficiency-induced persistence, HFF were co-infected for 24 h and 48 h with C. trachomatis and a cystogenic strain of T. gondii (Prugniaud or 76 K strain) that replicates much slower than the virulent RH strain in mammalian cells. For example, at 24 h p.i., about 60% and 40% of the PVs of the Prugniaud strain typically harbour one or two parasites respectively (not shown). At 48 h, all the parasites are still intracellular with a majority of the PVs containing 8–16 parasites, and lysis of the host cells usually occurs at 3–4 days p.i. (moi of 1). We reasoned that in the presence of a slow growing parasite strain like Prugniaud or 76 K, the bacterium would have the opportunity to mobilize host cell resources and nutrients for its own profit more efficiently than against the fast-growing RH parasites. Figure S8 shows that after 24 h of co-infection, the Prugniaud strain divided normally, without apparent defects in daughter cell formation. In contrast, C. trachomatis inclusions were smaller than inclusions from mono-cultures (inclusion size of 154 ± 25 versus 255 ± 32 μm2 in co-infection and mono-infection respectively) and contained some aberrant bodies. Interestingly, after an additional 24 h of co-culture, the inclusions had expanded in size (inclusion size of 404 ± 66 versus 523 ± 40 μm2 in co-infection and mono-infection respectively). Measurements of infectivity after 48 h of co-culture reveal that ∼ 48% of the bacteria were infectious, compared with ∼ 19% when grown in the presence of the RH strain (Fig. 8A). Among the type II strains of T. gondii, the 76 K strain has one of the slowest rates of multiplication in vitro (Spano et al., 2002; our observations). When this strain was co-cultivated with C. trachomatis, it grew normally during the first 24 h of co-infection while the bacteria had an abnormally small size (inclusion size of 178 ± 33 versus 312 ± 41 μm2 in co-infection and mono-infection respectively; Fig. S8). As observed for co-cultures with the Prugniaud strain after 48 h, the inclusions could develop despite the presence of the 76 K strain and have a nearly normal size (inclusion size of 378 ± 47 versus 412 ± 30 μm2 in co-infection and mono-infection respectively). Measurements of infectivity after 48 h of co-culture show that ∼ 70% of the bacteria were infectious (Fig. 8A). This suggests that while co-infection with the Prugniaud or 76 K strain for 24 h was detrimental to the bacteria, chlamydiae were able to recover after 48 h of co-infection. In contrast, while an increased number of bacteria produced infectious progeny after 48 h of co-infection with Prugniaud strain, even more pronounced with the 76 K strain, these parasites, unlike the RH strain, experienced a growth delay in its replicative cycle with PVs containing only six to eight parasites.
To further probe the role of nutrient scavenging in the competition between Toxoplasma and C. trachomatis, we used a Toxoplasma mutant (Δgra7) that is defective in nutrient acquisition (Coppens et al., 2006). GRA7 is a protein that is abundantly secreted in the PV after invasion, and is involved in cholesterol scavenging via the sequestration of host lysosomes within the PV lumen. GRA7-deficient parasites develop poorly as compared with wild-type parasites in the presence of low concentrations of serum in the medium (< 2.5% FBS), as demonstrated by the appearance of small and disordered PVs at 24 h p.i. as shown in Fig. S8. In co-infected cells, the mutant parasites displayed a slower replication rate compared with mono-infection as assessed upon enumeration of parasites per PV (Fig. 8A). On the contrary, C. trachomatis was only minimally affected by the presence of GRA7-deficient parasites, as most of the inclusions were large with normal sized bacteria (inclusion size of 245 ± 35 versus 278 ± 21 μm2 in co-infection and mono-infection respectively). In subcultures, high IFU titres were obtained from co-infections with the mutant parasites, with an approximately fivefold increase over that from co-cultures with wild-type Toxoplasma. Thus, the presence in co-culture of a parasite strain with a defect in nutrient acquisition did not impair the development of the bacterium significantly.
One striking feature of the PV of Toxoplasma is the presence of a network of membranous tubules within the lumen of the vacuole that connects with the vacuolar membrane (Sibley et al., 1995). It has been suggested that the function of this intravacuolar network is to act as a reticulum to increase the surface area of the PV membrane for the transfer of nutrients from the host cytoplasm to the PV. The organization and stability of this membranous network are mediated by the proteins GRA2 and GRA6 (Mercier et al., 2002). Lack of GRA2 and GRA6 results in the disruption of the tubulovesicular network, which may jeopardize the ability of the Δgra2-Δgra6 knockout mutant from retrieving host cell nutrients, although no obvious growth defect has ever been observed for these mutant parasites. We co-infected fibroblasts with Δgra2-Δgra6 knockout parasites and C. trachomatis for 24 h. During co-infection, the mutant parasites had a normal replication rate and the PV, despite lacking their tubulovesicular network, grew similarly to PVs from mono-cultures (Fig. S8). The bacterial inclusions remained small (inclusion size of 112 ± 15 versus 287 ± 33 μm2 in co-infection and mono-infection respectively) and contained many aberrant bodies, regardless of the number of PVs in the cell. No significant difference in chlamydial progeny production was observed in co-infections with Δgra2-Δgra6 knockout or wild-type parasites (Fig. 8A).
Jointly, these results establish an inverse relationship between the multiplication rate of the parasites and the development of C. trachomatis during co-infection, suggesting that the bacteria are able to overcome a slow progressing Toxoplasma infection by diverting sufficient nutrients for themselves.
C. trachomatis development in the presence of Toxoplasma is impaired regardless of the serovar
We next examined the intracellular development of a chlamydial strain of a biovar that is more virulent than the C. trachomatis serovar E strain used in co-infections so far. In vivo, biovar Lymphogranuloma Venereum (LGV/serovars L1–L3) isolates are invasive pathogens that rapidly spread through the submucosa to infect the inguinal lymph nodes and may eventually reach the bloodstream. At the cellular level, the marked difference between serovars L2 and E is the time of inclusion maturation, which is 48 h for the serovar L2 versus 72 h for serovar E in both fibroblasts and epithelial cells (doubling time of serovar L2: ∼ 2.6 h; Schramm and Wyrick, 1995; Wilson et al., 2004). To evaluate the effect of the parasite on the development of the invasive serovar L2, we measured the inclusion size and production of infectious EBs in HFF cells co-infected with serovar L2 and Toxoplasma for 24 h. Smaller, aberrant-shaped inclusions were generally observed in co-infected versus mono-infected cells (Fig. S9). However, the presence of small serovar L2 inclusions was dependent on the number of parasites occupying the host cell in sharp contrast with serovar E for which the presence of a single parasite led to a smaller inclusion size. Indeed, normal actively growing serovar L2 inclusions were observed in host cells containing up to two parasites (Fig. 8B). The inclusion size decreased in host cells with four parasites and was fourfold smaller in host cells with eight parasites (Fig. 8B). Subculture experiments showed that in host cells co-infected for 24 h, about 35% of the L2 bacteria were able to perpetuate the infectious cycle, similar to serovar E. No morphological defect was observed for parasite development.
Toxoplasma growth is impaired in cells preinfected with Chlamydia
Our results indicate that Toxoplasma (RH strain) develops normally in co-culture with C. trachomatis, mostly as if nothing was unusual in its environment, and clearly establishing the superior competitive fitness of the parasite over Chlamydia. We next aimed to challenge the parasite's ability to multiply in the presence of chlamydial inclusions by infecting fibroblasts first with Chlamydia and allowing the bacteria to grow for 48 h prior to the addition of Toxoplasma to the culture medium. After 48 h of growth alone in cells, the development of a C. trachomatis inclusion is nearly complete: the bacterial intracellular microcolony contains 500–1000 bacteria, mostly EBs, and occupies most of the free space in the host cytosol (Rockey and Matsumoto, 2000). Fibroblasts were infected with Chlamydia for 48 h, then superinfected with Toxoplasma for an additional 24 h before fixation and staining. Toxoplasma was still able to invade a fibroblast occupied by a very large chlamydial inclusion (Fig. 9A). Under these conditions, most PVs developed abnormally as ∼ 70% still only contained a single parasite. Only ∼ 15% of the PVs had four parasites, i.e. four times less than in PVs from mono-cultures. All the inclusions in superinfected cells were of normal size and aberrant RBs were not detected in these inclusions. This suggests that once the inclusion has nearly completed its development it becomes desensitized to the presence of Toxoplasma in the cell.
Next, cells were first infected with Chlamydia for 48 h, then with Toxoplasma for another 48 h (Fig. 9B). Many of the super-infected cells still contained large inclusions. In the same monolayer, neighbouring cells that were not superinfected with Toxoplasma had lysed and free EBs were visible in the medium. During the ultimate 2 days of co-infection, Toxoplasma continued to grow but at a reduced rate as the majority of the PVs (∼ 70%) harboured only four parasites, whereas 25% and 10% of the PVs had 8 and 16 parasites respectively.
Overall, these results suggest that Toxoplasma is endowed with the extraordinary faculty of adapting to intracellular growth even in the most adverse conditions, such as a cell terminally infected with another pathogen, likely depleted of most of its energy and nutrient stores.
‘The shoe on the other foot’: Toxoplasma cannot replicate normally in cells infected with Chlamydia psittaci
Among Chlamydia spp., Chlamydia psittaci is one of the most virulent and invasive species that infects birds and mammals zoonotically. In cultured epithelial cells, the doubling time of this species is ∼ 1.8 h (Binet and Maurelli, 2005). Like C. trachomatis, C. psittaci develops in a vacuole located in the host perinuclear region in the proximity of the Golgi, and upon nutrient deprivation, enters a persistent state (Moulder et al., 1980; Escalante-Ochoa et al., 1998; Beeckman and Vanrompay, 2010). To further probe the competition between the bacterium and Toxoplasma during a co-infection, we co-infected epithelial cells with the virulent fast-growing RH stain of Toxoplasma and the fast-growing C. psittaci for 24 h and 40 h. Toxopasma was unable to develop properly during a co-infection with C. psittaci, regardless of the number of inclusions in the cell or of the PV location within the cells (Fig. 10A). Some PVs were found in the host perinuclear region, as in mono-infected cells, or at the cell periphery, squeezed between the host plasma membrane and the inclusion, but the parasites managed to remain in their PV (Fig. 10B). In comparison, C. psittaci did not show any noticeable delay in its development: at 24 h p.i., the inclusion sizes were 520 ± 36 versus 489 ± 42 μm2 while at 40 h p.i., they were 1091 ± 120 versus 1203 ± 148 μm2 in mono- and co-cultures respectively. Stress-induced aberrant RBS were not observed (at least detectable by light microcopy) as the host cell cytoplasm was progressively occupied by these fast replicating bacteria as in mono-cultures (Fig. 10A). Interestingly, the inclusions of C. psittaci tightly encircled the parasites similarly to the inclusions of C. trachomatis in pyrimethamine-co-infected cells (Fig. S6). To examine whether the parasite shifted to its cyst form due to the stress of co-infection with C. psittaci, we probed for the expression of the BAG1 protein, a marker for the bradyzoite (cyst) form of the parasite. No BAG1-specific staining was observed on the parasite plasma membrane 40 h p.i. (not shown). Parasites showed defects in cytokinesis as individual parasites were barely discernible in their vacuole. Quantitative assessment of PV dimensions at 40 h p.i. revealed that 35% of the PVs had a size of 5 μm2 (accommodating ∼one parasite), 61% a size of 10 μm2 (∼two parasites) and 4% a size of 25–40 μm2; however, these larger PVs contained an undifferentiated mass of cytoplasm (Fig. 10C, panels a and b). In very rare instance (∼ 0.1% of co-infected cells), host cells contained a small inclusion (size less than 20 μm2), and in these cases, Toxoplasma developed normally with a majority of the PVs harbouring more than 16 parasites (Fig. 10C).
Collectively, these results indicate the marked competitive edge of C. psittaci over the parasite, possibly due to the faster replication rate of these bacteria and/or to damage inflicted to the host cell, which is incompatible with Toxoplasma growth.
A critical need in the biology of any intravacuolar pathogen immediately after host cell invasion is the transformation of its vacuole into a replication-permissive environment. Variations on the theme of intravacuolar pathogenesis include vacuoles either resisting or permitting fusion with selected host organelles, whereby the vacuole becomes a unique ‘organelle’ within the mammalian cell (Casadevall, 2008; Kumar and Valdivia, 2009). The prokaryote C. trachomatis and the protozoan Toxoplasma, two obligate intracellular pathogens of humans, have evolved similar strategies to exploit host cell organelles and scavenge nutrients thereof (reviewed in Laliberté and Carruthers, 2008; Cocchiaro and Valdivia, 2009; Saka and Valdivia, 2010). Both pathogens reside in a non-acidified vacuole in the cytosol of a wide variety of mammalian cells. There, they multiply, intercept host organelles, eventually filling all free cytoplamic space, then provoke lysis of the host cell at ∼ day 3 p.i. releasing abundant progeny that infect new cells. In this study, we have investigated the interactions of C. trachomatis and Toxoplasma residing simultaneously in the same host cell. Owing to the apparent evolutionary convergence of these two pathogens, we hypothesized that their coexistence in the same cell would influence their respective ability to grow. Consequently, interference with pathways shared by C. trachomatis and T. gondii would provide information on the intracellular lifestyle of either organism, and reveal what these pathogens imperatively need from their host cells. Our results indicate that the PV of Toxoplasma develops normally in cells co-infected with C. trachomatis but the chlamydial inclusion deviates from its typical developmental cycle to enter a stress-induced persistent state. A similar shift usually occurs when chlamydiae are confronted with detrimental growth conditions, such as nutrient starvation or exposure to certain cytokines or antibiotics. Obviously, the presence of live Toxoplasma in a C. trachomatis-infected cell does not represent a ‘normal’ growth environment for these bacteria.
In particular, our work has uncovered new aspects on the intracellular parasitism of C. trachomatis and T. gondii. On the bacterial side, infection of a C. trachomatis-infected cell with Toxoplasma reveals that: (i) C. trachomatis efficiently converts to a stress-induced persistence state independent of the serovar or the type of host cell, confirming that this phenotype is an efficacious and highly adaptable response of the bacterium when faced with a variety of unrelated stresses, (ii) diversion of exogenous cholesterol by the chlamydial inclusion is significantly reduced. The bacterium is unable to compete with Toxoplasma for LDL acquisition. It has been previously demonstrated that chlamydiae can retrieve host cholesterol via a brefeldin A-sensitive pathway (Golgi-dependent) and a brefeldin A-insensitive pathway (lysosome-dependent) but the respective contributions of the two routes for cholesterol supply to the inclusion are unknown (Carabeo et al., 2003). Our data suggest that the Golgi-dependent pathway (which is not targeted by T. gondii) is insufficient to compensate for the lack of cholesterol-LDL availability and that the host Golgi might be a minor source of cholesterol for the bacterium compared with endocytic organelles, and (iii) finally, among nutrients required for chlamydial growth, essential amino acids salvaged from the environment are important as their shortage induces a persistence state.
On the parasite's side, infection of a Toxoplasma-infected cell with C. trachomatis reveals that: (i) although metabolic, biosynthetic and regulatory pathways are predictably profoundly altered in mammalian cells infected by C. trachomatis, the parasite can still proficiently invade and form a PV within these cells. This indicates that the parasite is actively promoting its invasion and has a minimal requirement for host cell molecules, (ii) the parasite shifts to a slow growing mode similar to that of the bradyzoite form in cells that have been previously precolonized by the bacterium. This indicates that Toxoplasma is undergoing stress in response to Chlamydia co-infecting the host cell, (iii) among the known parasite proteins that contribute to nutrient acquisition, GRA7 seems important since GRA7-deficient parasites cannot outcompete Chlamydia. Our data did not support a direct function for GRA2 or GRA6, and therefore for the tubulo-vesicular network, in promoting nutrient scavenging activities since double GRA2- and GRA6-deficient parasites are as efficient as the wild-type parasite at multiplying in co-infected cells, and (iv) co-infection of T. gondii and C. psittaci results in an arrest in parasite growth, likely due to the fast replication rate of this chlamydial species, which is then able to rapidly colonize the host cell and consume most of the host resources, thereby outcompeting the parasite.
Our results demonstrate that Toxoplasma co-infection alters C. trachomatis development similar to other known stressors. Inclusions in co-infected cells are phenotypically undistinguishable from previously described stress-induced in vitro persistent chlamydial inclusions. Among the common characteristic features of the in vitro persistence phenotype that were reproduced in our study are: the altered ultrastructural morphology of the inclusion that is smaller than normal and contains markedly enlarged aberrant RBs, the frequent observation of intra-periplasmic blebs in the outer envelope of RBs, reduced levels of late chlamydial antigens, decreased production of infectious EBs, and the reversion to normal growth upon stressor removal. We demonstrate here that the stressed behaviour of the C. trachomatis inclusions in co-infected cells may be the direct result of decreased nutrient and energy availability owing to overwhelming competition by Toxoplasma. Interestingly, co-infection of host cells with C. trachomatis and Mycoplasma does not lead to similar alterations of chlamydial development, indicating that the growth requirements of these two bacteria may be substantially different (Baczynska et al., 2006). It is however not surprising that a Toxoplasma and C. trachomatis co-infection is particularly noxious for the bacterium because unlike mycoplasmas, the parasite diverts host organelles whose function is known to be essential for C. trachomatis.
Among the needed nutrients, essential amino acids play a central role for C. trachomatis growth. In adapting inside of a mammalian cell, chlamydiae and Toxoplasma have lost the ability to synthesize a number of amino acids and vitamins essential for their multiplication, putting them in competition with each other and with the host cell for specific amino acids. Hence, blockade of host protein synthesis in chlamydiae-infected cells results in the stimulation of bacterial growth due to reduced competition by the host cell for nutrient pools. Studies of C. trachomatis serovar L2 cultured in medium containing 0–100% amino acid levels revealed that chlamydial inclusions at 48 h p.i. were filled with aberrant bodies upon reduction of amino acid pools in the medium to 10% (Coles et al., 1993). A return to productive infection and maturation of RBs to EBs could be accomplished by the addition of cysteine or isoleucine. Degradation of tryptophan by indoleamine-2,3-dioxygenase activated by IFN-γ has been reported to trigger chlamydial persistence, and addition of exogenous indole in the presence of IFN-γ results in a return to the normal chlamydial development, making tryptophan an important amino acid for the bacteria (Caldwell et al., 2003). Toxoplasma is a natural tryptophan auxotrophic organism as it dies in cells depleted from this amino acid (Sibley et al., 1994), making it a strong competitor for host tryptophan. Other studies emphasize the essential requirement for phenylalanine, histidine and two branched amino acids: leucine and valine, for the intracellular development of many C. trachomatis serovars. The need for leucine for these bacteria is not surprising since this amino acid has the property to stimulate protein synthesis and inhibit protein breakdown in both mammalian cells and E. coli under conditions of nutrient limitation (Tischler et al., 1982).
All cells have an absolute requirement for iron or related translational metals for survival; intracellular bacterial pathogens require iron for entry and replication in mammalian cells, and chlamydiae are no exception. Addition of iron-chelating chemicals, e.g. desferal leads to a distinct form of persistent infection. Reversal to productive infection and recovery of infectious EB could be achieved with the removal of desferal and supplementation of the culture with iron-saturated transferrin or ferric chloride (Raulston, 1997; Thompson and Carabeo, 2011). The source of iron for Toxoplama is still unknown but the parasite would need to retrieve this metal from its host cell. However, addition of large quantities of iron in the co-infection medium only partially restores chlamydial growth in comparison with exogenous amino acids. The modest effect on C. trachomatis development also observed following the addition of selected lipids suggests that lipids like iron are in sufficient amounts in the host cell for the three species (the host cell, the parasite and the bacterium) in competition but then poorly salvaged by the bacteria under stress conditions. Alternatively, the bacterial needs for lipids and iron may be minimal, and C. trachomatis' persistent state in co-infected cells is instead induced by the lack of other host molecules that are more important for chlamydial replication than lipids or iron. C. trachomatis is auxotrophic for lipoic acid and needs to scavenge host lipoic for protein lipoylation to sustain its growth (Ramaswamy and Maurelli, 2010). Toxoplasma scavenges host-derived lipoate from mitochondria but has also the capacity of synthesizing lipoic acid (Crawford et al., 2006). The respective contributions of biosynthesis versus scavenging of lipoate to parasite growth are not known; however, the addition of excess lipoate in the medium did not benefit chlamydial growth, suggesting that the bacteria are able to salvage enough lipoate to persist in co-infected cells. In summary, under conditions in which the host's pool of nutrients becomes limiting, C. trachomatis may fail to successfully compete with the host cell and the parasite for specific metabolic precursors, and hence enters a state of arrested development. Alterations in the extracellular environment in order to increase the host intracellular pools of needed nutrients reactivate persistent chlamydiae to a state of productive developmental growth. In addition, co-infection with less competitive forms of Toxoplasma, like the Prugniaud strain, which has a slower metabolism than the RH strain and forms dormant cysts in vivo, or the GRA7 knockout Toxoplasma, which is impaired in its ability to re-route host lysosomes to the PV and hence has poor access to nutrients supplied from the endocytic network (Coppens et al., 2006), confirms our hypothesis that the replication capability of C. trachomatis largely depends on host cell resources available for the bacteria. This is further strengthened by our observations that during co-infection, the rapidly growing C. psittaci is able to outcompete Toxoplasma and develops normally despite the presence of the parasite.
It is known that Toxoplasma secretes many proteins that may directly target the chlamydiae and impact chlamydial development. We observed that the distance between the PV and the inclusion within a co-infected cell had a major impact on the expression of the persistent phenotype, whereby inclusions in close proximity to a PV would tend to contain very large aberrant bodies, whereas low but detectable production of PmpB was noted in inclusions distant from PVs. However, it seems unlikely that parasite-induced chlamydial persistence may have been indirectly due to host cell mediators activated by T. gondii based on observations that other Toxoplasma strains or mutants derived from the RH strain did not negatively impact bacterial growth. Regardless of the signal or processes that trigger the entry into a persistent state of C. trachomatis in PV-containing cells, it is worth noting that all inclusions established a state of persistence in our co-infection system. In comparison, upon intracellular depletion of tryptophan or exposure to IFN-γ, about 20% of the chlamydial inclusions fail to enter into a persistent state and instead achieve normal development to productive growth. Possible reasons for these heterogeneous responses may be the pre-existing resistance to starvation or phenotypic heterogeneity among bacteria with respect to cytokine signalling (Kokab et al., 2010). In this regard, Toxoplasma-induced persistence may then constitute a more accurate, and perhaps less complex, model for analysing the expression of chlamydial genes specifically regulated during persistent versus productive chlamydial infection.
In contrast to C. trachomatis, Toxoplasma does not show any overt signs of stress in the presence of C. trachomatis and appears to carry on ‘business as usual’ in co-infected cells. However, in cells that were infected first with C. trachomatis, Toxoplasma growth was slower than in cells that were simultaneously infected with both pathogens. After 48 h of C. trachomatis mono-culture, a time at which most of the host cytosol was occupied by the inclusion, and host organelles were typically squeezed between the inclusion membrane and the host plasmalemma, newly infecting parasites were at last challenged. Remarkably, however, internalized parasites still managed to divide and gradually fill the PV with new daughter cells. In the presence of C. psittaci, Toxoplasma does not develop since it forms very small PVs containing atrophic parasites up to 40 h p.i. The fast intracellular development of C. psittaci is in part due to its capacity to acquire the host adenine nucleotides through a carrier-mediated transport system, which hydrolyses host ATP to establish energized membranes (Hatch et al., 1982). Additionally, a reduction of host DNA and RNA syntheses has been observed in C. psittaci-infected cells. This results in a decrease in metabolic activities of the host cell, and therefore allows the bacteria to benefit from the pool of intracellular nutrients, which promotes inclusion growth. Nonetheless, this situation is disadvantageous for the parasite to achieve its programme of infectivity. Lack of parasite growth may happens either because Toxoplasma is starving and cannot adeptly attract host organelles to its PV and have therefore enough nutrients to multiply, or because the reduced transcriptional activities in the host cell are detrimental for the parasite that need to activate the expression of specific host genes to achieve pathogenicity.
In humans, C. trachomatis is the leading cause of infectious blindness (Schachter, 1999). Chlamydial genital infection is also the most frequently reported infectious disease in the USA according to CDC (see http://www.cdc.gov/std/stats09/default.htm). T. gondii causes life-threatening disease in immunocompromised individuals such as HIV/AIDS patients, and is responsible for retinochoroiditis and lethal encephalitis in these patients (Luft and Remington, 1992). Improved characterization of the specific interactions of the chlamydial inclusion or the PV with their respective host cell will expand our knowledge on the pathogenic strategies evolved by C. trachomatis and T. gondii, and may also inspire novel therapeutic avenues by exposing new vulnerabilities.
Reagents and antibodies
All chemicals were obtained from Sigma Chem. (St. Louis, MO) or Fisher (Waltham, MA) unless indicated otherwise. The nitrobenzoxadiazole (NBD) cholesterol was from Invitrogen (Carlsbad, CA) Molecular Probes (Seattle, WA). Lipids including cholesterol, sphingomyelin and triglycerides (tripalmitin, trilaurin, 1,2-dipalmitoyl-3-lauroyl-sn-glycerol and 1,2-dipalmitoyl-3-myristoyl-sn-glycerol) were purchased from Avanti Polar Lipids (Alabaster, AL). The antibodies used for immunofluorescence assays (IFA) included rabbit or rat polyclonal anti-TgGRA7 (Coppens et al., 2006); mouse monoclonal anti-EF-Tu (Zhang et al., 1994); guinea pig polyclonal anti-PmpB (Tan et al., 2010); rabbit polyclonal TgSAG1 and TgBAG1 (gift from J.-F. Dubremetz, Université de Montpellier, France). All primary antibodies were used at the dilution of 1:100 except for the anti-TgGRA7 and anti-TgSAG1 antibodies, used at the dilution of 1:200. The secondary antibodies were goat anti-IgG conjugated to either Alexa 488 or Alexa 594 (Invitrogen) before dilution at 1:2000.
Propagation of mammalian cells and pathogens
The cell lines used in this study and obtained from the American Type Culture Collection (Manassas, VA) include: primary HFF, HCT-8, a human ileocaecal adenocarcinoma cell line and HeLa 299 cells, a human cervical adenocarcinoma epithelial cell line. Cells were cultured in DMEM supplemented with 10% FBS, penicillin/streptomycin (100 U ml−1 per 100 μg ml−1) and 2 mM l-glutamine. The tachyzoite RH strain (type I) of T. gondii were used throughout this study. In one experiment, the Prugniaud and 17 K-GFP strains (type II) of T. gondii was used. Red fluorescent protein (RFP)-expressing stable transgenics derived from the RH strain was generously provided by F. Dzierszinski (McGill University, Canada). We also used the double knockout T. gondii for gra2 and gra6, a gift from M.-F. Cesbron-Delauw (Université Fourier, Grenoble, France; Mercier et al., 2002) and the single knockout parasite for gra7 kindly provided by J.C. Boothroyd (Stanford University; Coppens et al., 2006); both mutants were derived from the RH strain. All parasites were propagated in vitro by serial passage in monolayers of HFF in DMEM plus 10% FBS (Roos et al., 1994). The C. trachomatis lab reference serovar E/UW5-CX was used throughout this study. In other experiments, the C. trachomatis lab reference serovar L2 or the pathogenic avian type strain C. psittaci Cal-10 were used. All chlamydiae were propagated in HeLa cells at 37°C with 5% CO2 in DMEM supplemented with 10% FBS, 25μg ml−1 gentamicin and 1.25 μg ml−1 fungizone as described (Tan et al., 2010).
Mammalian cells were seeded onto coverslips in 24-well plates for IFA or in six-well tissue culture dishes for EM and grown at 37°C in a CO2 incubator until 70% confluence. Prior to infection experiments, cells were washed with PBS and incubated 24 h in antibiotic- and fungizone-free medium. Two infection protocols were established to examine the interactions of T. gondii and C. trachomatis or C. psittaci. Co-infection protocol: Mammalian cells in DMEM with 10% FBS (‘infection medium’) were infected with 2 × 105Toxoplasma (moi of ∼ 1) harvested from culture supernatant and washed in PBS and with a titrated inoculum of Chlamydia species seed stock (5–25 μl per dish, yielding an 80% infection rate) in SPG (0.25 M sucrose, 10 mM sodium phosphate and 5 mM l-glutamic acid). Infected monolayers were then spun at 700 g in a Beckman Allegra centrifuge for 30 min at 37°C before addition of fresh ‘infection medium’ to dishes which marks the time of infection (T = 0 h p.i.) and incubation at 37°C with 5% CO2 for the various times. Superinfection protocol: HFF cultivated in the ‘infection medium’ were first infected with bacteria as described above. After 48 h of bacterial growth, the cells were superinfected with 2 × 105 and 2 × 106Toxoplasma respectively, and incubated for an additional 24 h or 48 h prior to processing for IFA. In some experiments, the antiparasitic drug pyrimethamine was added in the culture medium during the co-infection or superinfection protocols.
Chlamydial infectivity assays
After 24 h, 48 h or 72 h of co-infection, cell monolayers were washed, scraped, pelleted and resuspended in 1 ml of cold growth medium. Infected cells were then lysed by freeze/thaw and sonication, followed by centrifugation at 500 g for 5 min to discard host cell and parasite debris in the pellet. Supernatants were collected and centrifuged twice at 8000 g for 30 min to recover chlamydial bodies in the pellet. Serial dilutions of the pellets in SPG were applied to HeLa cells plated on glass coverslips in triplicates. After infection, the HeLa monolayers were re-fed with bacterial growth medium containing 1 μM pyrimethamine. After 36 h, the cells were fixed and chlamydial inclusions were stained with DAPI. The average number of cells containing inclusions within 10 optical fields, as viewed through the 40× objective of a Nikon Eclipse E800 microscope was used to calculate the number of infectious units per ml for each condition. To score the size of chlamydial inclusions, random fields of EF-Tu-labelled bacteria grown in the indicated cell lines in the presence or the absence of Toxoplasma were selected and images of 0.8 μm thick sections were collected. Determination of the cross-sectional area in square micrometers was performed using the LSM510 software.
Cholesterol uptake assays
Human LDL (density: 1.019–1.063 g ml−1) were isolated from fresh serum by zonal density gradient ultracentrifugation as described (Poumay and Ronveaux-Dupal, 1985). The fluorescent lipid NBD-cholesterol was incorporated into LDL by mixing 50 μl of the lipid stock solution with 20 ml of filtered fresh human plasma. To visualize fluorescent cholesterol associated with T. gondii and C. trachomatis, infected HFF were incubated with 0.1 mg ml−1 LDL containing NBD-cholesterol in lipoprotein-free medium for 2 h and observed as live by fluorescence microscopy. In some assays, the cells were infected with the parasites or the bacteria for 24 h, then exposed to 10 μM pyrimethamine and 100 units ml−1 of penicillin G, respectively, for an additional day before the pulse.
Assays of nutrient enrichment on chlamydial growth
Human foreskin fibroblasts were co-infected with C. trachomatis serovar E and T. gondii RH strain for 24 h in normal conditions (‘infection medium’), or in medium enriched in iron (ferric chloride, holo-transferrin), lipoate, lipids (cholesterol, sphingomyelin, mixture of triacylgycerols), and/or essential amino acids (histidine, isoleucine, leucine, lysine, methionine, phenylalanine, phreonine, tryptophan, valine) and non-essential amino acids (glycine, alanine, asparagine, aspartic acid, glutamic acid, proline, serine) before determining the inclusion size and the number of IFU.
For IFA, cells were fixed in a solution consisting of 4% PFA (Polysciences, Warrington, PA) and 0.02% glutaraldehyde in PBS for 15 min and permeabilized with 0.3% Triton X-100 for 5 min. Samples were then blocked with 3% BSA dissolved in PBS for 45 min and probed with primary antibodies diluted in blocking buffer for 1–2 h. Samples were then washed three times and probed with secondary antibodies diluted in the blocking buffer for 45 min. Coverslips were mounted onto glass microscope slides using ProLong Gold anti-fade mounting solution with or without DAPI (Invitrogen). The cyst wall of Toxoplasma was visualized by staining with tetramethyl rhodamine isothiocyanate-conjugated Dolichos biflorus lectin used at a dilution of 1:300 in 3% bovine serum albumin/PBS. For all fluorescence assays, images were acquired on a Nikon Eclipse E800 microscope equipped with a Spot RT CCD Camera and processed using Image-Pro-Plus software (Media Cybernetics, Silver Spring, MD) before assembly using Adobe Photoshop (Adobe Systems, Mountain View, CA). Some images were also viewed with a Nikon Plan Apo 100× objective using a Nikon 90i microscope and pictures were taken using a Hammatsu ORCA-ER camera and Volocity software. Images (a z-stack per field) were processed using iterative restoration (confidence limit 98% and iteration limit 25) and further processing was done using Adobe Photoshop software. Protocols for filipin labelling and visualization were performed as described (Coppens and Joiner, 2003). To quantify the level of filipin accumulation, images were collected using sequential scanning, processed and merged using Volocity software. For quantification of fluorescence intensity, the total intensity in the UV channel was determined for each vacuole and compared with the total fluorescence intensity of the whole same cell using the following equation: [sum intensity (pathogenic vacuole centre)/sum intensity (entire host cell)] × 100. Same quantitative measurement was applied to monitor NBD-cholesterol levels associated to the vacuole in the green (537) channel. For each vacuole, data on NBD-cholesterol were compared with the total fluorescence intensity of the entire host cell.
For thin-section transmission EM, cells were fixed in 2.5% glutaraldehyde (Electron Microscopy Sciences, Hatfield, PA) in 0.1 M sodium cacodylate buffer (pH 7.4) for 1 h at RT, and processed as described (Coppens and Joiner, 2003) before examination with a Philips CM120 Electron Microscope (Eindhoven, the Netherlands) under 80 kV.
For the comparison of means, P-values were determined by the analysis of variance against control (anova 2).
The authors are grateful to the members of the Coppens and Bavoil laboratories, and Dr Ru-ching Hsia for their helpful discussions during the course of this work. We are grateful to the generous providers of non-commercial antibodies. We also have a special gratitude for the work of H. Zhang in co-infecting cells and performing the IFA. We would like to thank the competent technical staff from the Microscopy Facility at Johns Hopkins University. This work was supported by National Institutes of Health Grant RO1 AI06767 to I.C. and U19 AI084044 to P.M.B.