Cross Talk between ARF1 and RhoA Coordinates the Formation of Cytoskeletal Scaffolds during Chlamydia Infection

ABSTRACT Chlamydia trachomatis is an obligate intracellular bacterium that has developed sophisticated mechanisms to survive inside its infectious compartment, the inclusion. Notably, Chlamydia weaves an extensive network of microtubules (MTs) and actin filaments to enable interactions with host organelles and enhance its stability. Despite the global health and economic burden caused by this sexually transmitted pathogen, little is known about how actin and MT scaffolds are integrated into an increasingly complex virulence system. Previously, we established that the chlamydial effector InaC interacts with ARF1 to stabilize MTs. We now demonstrate that InaC regulates RhoA to control actin scaffolds. InaC relies on cross talk between ARF1 and RhoA to coordinate MTs and actin, where the presence of RhoA downregulates stable MT scaffolds and ARF1 activation inhibits actin scaffolds. Understanding how Chlamydia hijacks complex networks will help elucidate how this clinically significant pathogen parasitizes its host and reveal novel cellular signaling pathways.

Chlamydia trachomatis is the most common cause of bacterial sexually transmitted disease. It is also the most frequent cause of infectious blindness, known as trachoma (5). C. trachomatis displays a unique biphasic lifestyle. The infectious form, called the elementary body (EB), exhibits minimal metabolic activity and promotes its entry into human epithelial cells. Once inside the host cell, Chlamydia EBs reside within a membrane-bound compartment called the inclusion. EBs then rapidly differentiate into a replicative form, called the reticulate body (RB), which is metabolically active and noninfectious. As RBs replicate by binary fission, they synthesize effector proteins that remodel and hijack the host cell to promote their survival (6). In doing so, Chlamydia subverts many aspects of the host cellular environment, including the host cytoskeleton, a critical element of successful infection and dissemination of C. trachomatis within the host.
EBs initially invade the host cell by attaching to the plasma membrane and injecting the chlamydial protein Tarp (translocated actin recruiting protein) into the host cytoplasm via the chlamydial type III secretion system. Phosphorylation of Tarp in the cytosol allows Tarp to recruit the guanine nucleotide exchange factors (GEFs) Sos1 and Vav2, which activate Rac1 (7). Tarp directly binds actin monomers to nucleate new linear filaments (8,9). Subsequently, the Arp2/3 complex is activated, and branched actin filaments are polymerized underneath the bacterial binding site to promote the entry of the EBs into the nascent inclusion (10)(11)(12).
The second wave of cytoskeletal manipulation begins around 16 h postinfection (hpi), when MTs are assembled around the inclusion (13). These scaffolds are subsequently stabilized and undergo posttranslational modification (PTM), including detyrosination and acetylation (14,15). PTMs influence MT structure and depolymerization rates, further increasing their stability (16,17). We have previously established that the small GTPase ARF1 controls the posttranslational modification of MT scaffolds. PTM-MT scaffolds then position Golgi ministacks around the inclusion, which provides lipids to facilitate the rapid expansion of the inclusion (14,15,18). Around 32 hpi, actin scaffolds form around the inclusion. These structures physically reinforce the inclusion membrane to prevent premature lysis, as the inclusion expands to occupy most of the host cytoplasm (19). Using actin-depolymerizing agents and small interfering RNA (siRNA), the formation of F-actin at the inclusion was shown to be RhoA dependent (19).
Finally, Chlamydia exits its host cell either by controlled cell lysis or through extrusion, which also relies on the intervention of the host cytoskeleton. Extrusion is a nonlytic process during which the inclusion protrudes out of the infected cell before pinching off the plasma membrane (20). This event requires the mobilization of RhoA, WASP, and myosin II. Interestingly, MT depolymerization with nocodazole does not affect extrusion, suggesting this event is actin specific (20).
Interference with the host cytoskeleton dramatically impacts the outcome of Chlamydia infection. The inhibition of PTM-MT scaffolds decreases Chlamydia infectivity, whereas their enhancement increases it, demonstrating the importance of this cytoskeletal element in Chlamydia development (14). Global inhibition of actin polymerization with latrunculin also causes early rupture of inclusions, indicating that actin is essential for maintaining inclusion stability (19). Inclusion stability solely depends on actin, as MT depolymerization does not impact inclusion integrity (19).
Previous work investigating Chlamydia has been significantly limited by its reliance on cytoskeleton-modulating drugs. These drugs disrupt the host cytoskeleton in its entirety and often indirectly impact other cytoskeletal elements (21). As a result, elucidating the precise local role of actin and MT scaffolds during Chlamydia infection has been challenging. However, with advances in Chlamydia genetics, targeted disruption of Chlamydia's virulence system and its effector proteins is now possible without indirectly affecting native host processes. Recently, we and others showed that a single chlamydial effector, InaC (also called CT813 or CTL0184), controls the formation of the kinetically and functionally distinct actin and PTM-MT scaffolds (15,19,22). InaC knockout (KO) Chlamydia produces less-infectious progeny and generates smaller inclusions, indicating that InaC-dependent pathways are essential for Chlamydia fitness (15,19). We also demonstrated that InaC directly interacts with the host small GTPases ARF1 and ARF4 to control the formation of PTM-MTs and, consequently, the dispersal of Golgi ministacks around the inclusion during infection (15). This event occurs midcycle at 16 to 24 hpi. In addition to InaC, it has been shown that actin scaffold formation around 32 hpi is mainly dependent on the host GTPase RhoA (19). Whether InaC regulates RhoA and how both GTPases are coordinated during infection to generate their specific scaffolds at the optimal time is unknown.
Here, we report that while RhoA can be recruited to the inclusion in the absence of InaC, its activation, which leads to the formation of actin scaffolds, requires InaC. Furthermore, we discovered that the presence of RhoA downregulates ARF1 activation, while ARF1-GTP inhibits RhoA activation, highlighting cross talk between two small GTPases to coordinate PTM-MT and actin scaffolds during infection. Overall, our findings establish that a single chlamydial effector, InaC, is a master regulator of actin and MT dynamics during Chlamydia infection.

RESULTS
RhoA activation is required for the formation of actin scaffolds around the inclusion. To understand the exact role of RhoA during C. trachomatis infection and to assess whether RhoA is the sole regulator of actin scaffold formation, we generated CRISPR/Cas9 RhoA KO HeLa cells (see Fig. S1A in the supplemental material). HeLa cells transfected with the Cas9 vector without guide RNA were used as a control. RhoA KO cells also express similar amounts of actin, a-tubulin, and acetylated a-tubulin compared to control cells (Fig. S1B). We detected a modest but statistically insignificant increase in detyrosinated a-tubulin in the absence of RhoA. As expected, RhoA KO cells are viable and divide similarly to control cells ( Fig. S1C and D).
Next, we assessed the kinetics of actin scaffold formation in the presence and absence of RhoA during Chlamydia infection. Control and RhoA KO cells were infected with wild-type (WT) C. trachomatis L2 prior to the analysis of actin scaffolds around the inclusion at different times postinfection. In parallel, RhoA KO cells were transfected with a vector encoding myc-RhoA WT (RhoA WT ) at 3 to 4 hpi to restore RhoA expression. In control cells, actin scaffolds are detected at 38 hpi, but not 24 hpi ( Fig. 1A and B, Control), consistent with previous observations that actin scaffolds form late during infection (19). In contrast, actin scaffolds were not detectable at either time point in RhoA KO cells ( Fig. 1A and B, RhoA KO). The expression of myc-RhoA WT rescued the loss of actin scaffolds in RhoA KO cells at 38 hpi (Fig. 1B, RhoA WT ), indicating that the loss of actin scaffolds in RhoA KO cells is not due to an off-target effect of Cas9.
Like all small GTPases, RhoA cycles between GDP-and GTP-bound states, which in turn control its interactions with downstream effectors (23). To assess whether Chlamydia infection influences the activation state of RhoA to promote actin polymerization around the inclusion, we infected HeLa cells with C. trachomatis L2 and generated lysates from mid to late times postinfection (16 to 48 hpi). The levels of RhoA-GTP were then analyzed using the RhoA-binding domain (RBD) of the effector Rhotekin (24). The activation of RhoA reaches a maximal level at 32 hpi (Fig. 1C), which correlates with both the formation of actin scaffolds and the recruitment of endogenous RhoA to the inclusion (Fig. S2). Note that RhoA activation is a transient event, as it is deactivated and returns to noninfected levels at ;40 hpi. These data indicate that C. trachomatis triggers the activation of RhoA at the same time that actin polymerization occurs around the inclusion.
To establish the role of RhoA activation in the formation of actin scaffolds, we exploited the well-characterized G14V (myc-RhoA G14V , GTP locked) and T19N (myc-RhoA T19N , dominant negative) RhoA mutants, which have been used extensively to explore the biological activities of RhoA (25)(26)(27). At 3 to 4 hpi with C. trachomatis L2, we transfected RhoA KO cells with either myc-RhoA G14V or myc-RhoA T19N and compared the formation of actin scaffolds to that of cells transfected with myc-RhoA WT . We found that myc-RhoA G14V , but not myc-RhoA T19N , supports actin scaffold formation (Fig. 1D).
Compared to infected myc-RhoA WT -expressing cells, the expression of myc-RhoA G14V maximizes actin scaffold formation to 126% 6 3.7%. In comparison, only 9% 6 2.1% of inclusions displayed actin scaffolds in RhoA T19N -expressing cells, demonstrating that active RhoA is required to promote actin scaffold formation. Interestingly, actin scaffolds become detectable at 24 hpi in cells expressing constitutively active myc-RhoA G14V , while they are absent in cells expressing myc-RhoA WT and myc-RhoA T19N (see Fig. S3 in the supplemental material). In this case, we observe that 86% 6 4.2% of inclusions have actin scaffolds as early as 24 hpi. This is not due to a shift in RhoA recruitment since endogenous RhoA is already present on the inclusion at 24 hpi (Fig.  S2) despite the absence of actin scaffolds (Fig. 1A). Altogether, these data indicate that the temporal activation of RhoA is critical and dictates the kinetics of actin scaffold formation during infection.
The chlamydial effector InaC is required for the activation of RhoA. InaC is a chlamydial effector that is required for the formation of actin scaffolds during Chlamydia infection (15,19). To establish whether InaC regulates the activity of RhoA during infection, we made use of a C. trachomatis L2 mutant in which the InaC gene was inactivated using Targetron (InaC KO Chlamydia) (15). We then infected HeLa cells with InaC KO C. trachomatis L2 and quantified the amount of RhoA-GTP at 32 hpi. Infection with WT C. trachomatis L2 increased RhoA-GTP levels 2.9 6 0.54-fold compared to uninfected control cells ( Fig. 2A). In contrast, InaC KO-infected cells exhibited RhoA-GTP levels similar to control cells (0.983 6 0.34-fold change), indicating that InaC is required in RhoA activation during C. trachomatis infection.
One way that proteins regulate each other is through direct interactions. We thus tested whether InaC forms a complex with RhoA to control its activation locally. To do so, we conducted a series of immunoprecipitation experiments in the presence and absence of infection in different configurations. However, we could not detect an interaction (see Fig. S4 in the supplemental material), indicating that either InaC and RhoA do not bind or the interaction is too weak or transient to detect.
Even though InaC and RhoA do not form a detectable complex, the recruitment of RhoA to the inclusion could still be indirectly impacted by the absence of InaC, which would explain the reduction in RhoA-GTP in cells infected with InaC KO C. trachomatis L2 ( Fig. 2A). To assess whether RhoA recruitment to the inclusion depends on InaC, we used RhoA KO cells complemented with myc-RhoA WT and infected with WT or InaC KO C. trachomatis L2. At 32 to 38 hpi, infected cells were fixed and stained with anti-myc and anti-InaC antibodies to label RhoA and the inclusion membrane, respectively. As expected, myc-RhoA WT was present on the inclusion membrane in WT C. trachomatis L2-infected cells (Fig. 2B, WT L2). Interestingly, myc-RhoA WT was also detected on the inclusion membrane of InaC KO C. trachomatis L2-infected cells (Fig. 2B, InaC KO), indicating that RhoA recruitment is InaC independent. Next, we assessed whether both mutant forms of RhoA are differentially recruited during infection with InaC KO Chlamydia. As shown in Fig. 2C and D, myc-RhoA G14V (Fig. 2C) and myc-RhoA T19N (Fig. 2D) are also recruited to the inclusion in both WT and InaC KO C. trachomatis L2infected cells. These data further indicate that RhoA is recruited to the inclusion independently of its activation state.
We then determined whether the loss of actin scaffolds in InaC KO C. trachomatis L2-infected cells is due to a defect in RhoA activation. To do so, we infected RhoA KO HeLa cells with WT or InaC KO C. trachomatis L2, followed by transfection with myc-RhoA WT , myc-RhoA G14V , or myc-RhoA T19N DNA. The cells were fixed at 32 to 38 hpi and labeled with anti-myc and anti-InaC antibodies. As observed earlier, myc-RhoA WT and myc-RhoA G14V , but not myc-RhoA T19N , promote the formation of actin scaffolds during WT C. trachomatis infection ( Fig. 2B to E, WT L2). Strikingly, the expression of myc-RhoA G14V , but not myc-RhoA WT or myc-RhoA T19N , rescues the InaC KO phenotype and fully restores the formation of actin scaffolds ( Fig. 2B to E, InaC KO), indicating that the loss of actin scaffolds in InaC KO-infected cells is due to an inability to activate RhoA. Altogether, these results demonstrate that InaC is not required to recruit RhoA but rather to locally activate it and promote actin scaffold formation around the inclusion. Since RhoA does not interact with InaC, it is likely that an additional host or bacterial factor that remains to be identified is involved.
InaC-RhoA-mediated cytoskeletal scaffolds contribute to inclusion stability. InaC is required for the formation of both PTM-MT and actin scaffolds (15,22). However, their direct contribution to inclusion stability is not well understood. To test the importance of these scaffolds in Chlamydia fitness, we infected HeLa cells with InaC KO C. trachomatis L2 and assessed inclusion stability over time. In WT C. trachomatis L2-infected cells, we observed that ;40% of inclusions were ruptured at 48 hpi. In contrast, ;50% of the InaC KO inclusions were lysed by 32 hpi (Fig. 3A), indicating a substantial shift in inclusion stability in the absence of InaC. This early release of Chlamydia into the cytosol correlated with a 2.4-fold increase in cell death measured by caspase 3/7 activation. While only 5% 6 0.98% of WT C. trachomatis L2-infected cells were positive for caspase 3/7 activation, 12% 6 0.86% of InaC KO C. trachomatis L2-infected cells were positive at 72 hpi (Fig. 3B). InaC KO inclusions were stabilized when InaC-FLAG was reintroduced on a plasmid (see Fig. S5A and C in the supplemental material). As expected, the complementation of InaC expression also rescued the loss of actin scaffolds (Fig. S5B). Overall, this indicates that InaC is critical for the maintenance of inclusion stability.
To specifically test the impact of the loss of actin scaffolds on inclusion stability, RhoA KO cells were infected with WT C. trachomatis L2, and inclusion lysis was measured at 72 hpi as it corresponds to maximal caspase 3/7 activation (Fig. 3B). As controls, we infected RhoA KO cells with InaC KO C. trachomatis L2 and treated WT C. trachomatis L2-infected cells with latrunculin B to depolymerize all host cell actin. Consistent with previous reports (19), latrunculin B treatment increased levels of inclusion lysis The presence of RhoA inhibits the formation of PTM-MT scaffolds during infection. PTM-MT scaffolds around the inclusion are most evident around 24 hpi, whereas the formation of actin scaffolds around the inclusion begins later, around 32 hpi (15,19). Furthermore, actin scaffolds do not form in the absence of RhoA, and PTM-MT scaffolds are disorganized in the absence of ARF1. These data suggest that these two cytoskeletal pathways are independent of each other and are controlled by their corresponding small GTPase. This concept is supported by the fact that each GTPase is activated at specific times during infection: 16 hpi for ARF1 (see Fig. S6 in the supplemental material) and 32 hpi for RhoA (Fig. 1C). Yet, the requirement for the chlamydial protein InaC to control both cytoskeletal events indicates that some degree of cross talk likely exists between these two pathways. Since the inclusion is a fundamental component of Chlamydia pathogenicity, this cross talk may allow InaC to finely control the activation/inactivation of each pathway to appropriately time Golgi repositioning and inclusion stability. In this case, we would expect that altering the expression level and/or activity level of either GTPase would impact the formation of the other cytoskeletal element.
To test this hypothesis, we first investigated the impact of RhoA depletion on the formation of PTM-MTs during infection. RhoA KO cells were infected with WT C. trachomatis L2, and the PTM-MT scaffolds around the inclusion at 24 hpi were monitored using immunofluorescence microscopy. As expected, PTM-MT scaffolds still assemble around the inclusion in the absence of RhoA, demonstrating that RhoA is not required for their formation ( Fig. 4A and B). However, we observe increases of 48% 6 8.8% and 28% 6 7.4% of detyrosinated and acetylated a-tubulin scaffolds, respectively, compared to control cells ( Fig. 4A and B, graphs). This enhanced formation of PTM-MT scaffolds is suppressed to WT levels when myc-RhoA WT is expressed in RhoA KO cells, confirming that the phenotype is specifically due to the loss of RhoA ( Fig. 4A and B, RhoA WT ). Furthermore, the expression of either active or inactive RhoA mutants in RhoA KO cells suppresses the enhanced formation of PTM-MTs similarly to myc-RhoA WT (see Fig. S7 in the supplemental material). These results indicate that this phenomenon is due to the presence of RhoA rather than a specific nucleotide-bound state. Altogether, these data suggest that the absence of RhoA enhances the formation of PTM-MTs.  Since ARF1 is a critical factor in forming PTM-MT scaffolds (15), the enhancement of PTM-MT scaffolds in the RhoA KO could be due to increased complex formation between InaC and ARF1 and/or increased ARF1 activation. To test both possibilities, we infected RhoA KO cells with C. trachomatis L2 expressing InaC-FLAG. We then assessed the interactions between InaC-FLAG and endogenous ARF1 compared to the complex formed in control cells. As shown in Fig. 4C, we observe a striking increase in ARF1 binding to InaC-FLAG in the absence of RhoA at 24 hpi. Next, we tested whether the enhanced interaction between ARF1 and InaC is with active or inactive ARF1. Control and RhoA KO cells were transfected with either GDP-bound (ARF1 T31N -HA [hemagglutinin]) or GTP-bound (ARF1 Q71L -HA) ARF1 (15, 28) at 3 to 4 hpi with C. trachomatis L2 expressing InaC-FLAG. InaC was immunoprecipitated using an anti-FLAG antibody, and the immunoprecipitates were analyzed by Western blotting to identify which ARF1 mutant was bound. As shown in Fig. 4D, only ARF1 Q71L -HA binds to InaC. The interaction with ARF1-Q71L and InaC is enhanced in the absence of RhoA, demonstrating that the lack of RhoA results in an increase in InaC:ARF1-GTP binding and consequently PTM-MT scaffold formation.
Activation of ARF1 impairs RhoA-GTP-dependent actin scaffold formation. The recruitment of RhoA to the inclusion begins around 24 hpi (Fig. S2). During this time, ARF1 is already activated, and PTM-MT scaffolds are forming. Since (i) the absence of RhoA concomitantly leads to the loss of actin scaffolds and enhances the interaction between ARF1-GTP and InaC (Fig. 4D) and (ii) ARF1 is not required for the formation of actin scaffolds (see Fig. S8 in the supplemental material), we hypothesized that the deactivation of ARF1 might be required for the subsequent downstream activation of RhoA and the formation of actin scaffolds. To test this hypothesis, we expressed the constitutively active ARF1 Q71L -HA and dominant-negative ARF1 T31N -HA mutants at 3 to 4 hpi with WT C. trachomatis L2 and assessed the formation of actin scaffolds at 32 hpi. Interestingly, the constitutive activation of ARF1 drastically reduces actin scaffold formation by 45% ( Fig. 5A and B, ARF1 Q71L -HA). This effect is specific to the active form of ARF1, as the expression of GDP-bound ARF1 does not influence actin scaffold formation ( Fig. 5A and B, ARF1 T31N -HA).
Next, we determined whether the constitutive activation of ARF1 blocks the activation of RhoA. As shown in Fig. 5C, the activation of RhoA is inhibited by 68% 6 22% in cells expressing ARF1 Q71L -HA compared to cells transfected with ARF1 WT -HA. Collectively, these data demonstrate that cross talk between ARF1 and RhoA is critical for the coordination of PTM-MT and actin scaffolds during C. trachomatis infection.
Together, our data support a model in which Chlamydia uses a single effector to coordinate two small GTPases and regulate critical cytoskeletal pathways necessary for its survival (Fig. 6). Specifically, ARF1 is activated midcycle (;16 hpi) in an InaC-dependent manner. At this time, RhoA is absent from the inclusion, leaving ARF1 activation unimpeded. As a result, ARF1-GTP actively drives the formation of PTM-MTs. In turn, PTM-MT scaffolds coordinate Golgi rearrangement around the inclusion, which contributes to inclusion growth (15). Around 24 hpi, RhoA starts to relocate to the inclusion membrane. The presence of RhoA blocks ARF1 activation and the generation of additional PTM-MTs. Simultaneously, ARF1-GTP inhibits actin scaffold formation by interfering with RhoA activation. As the infection progresses, the concentration of RhoA on the inclusion membrane increases, further inhibiting ARF1 activation. As ARF1 returns to its inactive GDP-bound state, it can no longer block RhoA activation. The InaC-dependent activation of RhoA fully engages, peaking at 32 hpi and maximizing actin polymerization around the inclusion. As a result, Chlamydia's infectious niche, which has grown significantly due to the continuous supply of lipids from Golgi ministacks, is now fortified against premature lysis and ultimately provides a safe space for RBs to differentiate into infectious EBs and perpetuate the infectious cycle.

DISCUSSION
Here, we have identified a virulence pathway in which the bacterial pathogen Chlamydia trachomatis uses a single chlamydial effector to integrate ARF1 and RhoA cross talk to precisely time the formation of PTM-MT and actin scaffolds (Fig. 6). Many other intracellular pathogens also target GTPases (29) and integrate GTPase cross talk into their virulence systems. An example is the effector protein EspG expressed by enteropathogenic E. coli OH157:H7, which disrupts ER-to-Golgi trafficking by linking ARF1-specific vesicular trafficking events to Rab1 inactivation (30,31). EspG also blocks phagocytosis by interacting with ARF1 to prevent Rac1 activation and consequently the polymerization of actin at the plasma membrane (32). In contrast, Salmonella enterica co-opts an ARF6-ARF1-Rac1 axis to drive plasma membrane ruffling required for entry (33). The Chlamydia virulence pathway is most similar to that utilized by Salmonella since Chlamydia activates rather than inhibits the ARF1-RhoA network to promote successful infection. In this pathway, both RhoA and ARF1 elegantly fine-tune their activity by generating simultaneous feedback loops. The activation of ARF1 prevents the activation of RhoA, while the presence of RhoA downregulates ARF1 activation. While GTPases are well known to regulate each other (34)(35)(36)(37), how RhoA and ARF1 influence the function of each other is unknown. Furthermore, whether this process requires other bacterial factors or is an inherent property of these GTPases and their respective effector proteins is under investigation.
Since GTPases are critical components of host signaling networks, manipulating their function or activity can have detrimental consequences for the host. In particular, dysregulation of GTPases involved in coordinating the host cytoskeleton has been linked to cancer hallmarks, such as uncontrolled cell proliferation, migration, and Coordination of Actin and Microtubules by Chlamydia ® differentiation (38,39). ARF1 and RhoA have been implicated in some aspects of cancer development, including tumor cell invasion and proliferation (40)(41)(42)(43)(44)(45). Given that Chlamydia infection has been associated with an increased risk for developing cervical and ovarian cancers (46)(47)(48), it would be interesting to determine whether the activation of the ARF1-RhoA signaling axis contributes to the potentiation of cancer development in those with a history of chronic Chlamydia infection.
As an absolute intracellular bacterium, Chlamydia must ensure the integrity of its intracellular niche to establish a successful infection. As Chlamydia replicates inside the inclusion, this bacterial compartment matures and expands to eventually occupy most of the host cytoplasm (49). We have previously shown that InaC KO Chlamydia displays smaller inclusions and is replication deficient (15). These defects may be due to a combination of factors. (i) They may be a consequence of reduced PTM-MT scaffold formation, which impairs the recruitment of Golgi ministacks around the inclusion that play a role in lipid acquisition and inclusion development (50). (ii) Premature lysis could result in the release of Chlamydia into the cytoplasm before the cells have successfully differentiated from RBs into infectious EBs. We show here that InaC KO inclusions lyse significantly earlier than their WT counterparts (Fig. 3A). Since the infected cells then undergo more cell death (Fig. 3B) due to cytoplasmic Chlamydia, the bacterium reaches an impasse. While both possibilities likely contribute to the survival of Chlamydia, our data demonstrate that the chlamydial effector protein InaC is critical for the protection of Chlamydia's intracellular niche and successful infection by controlling two cytoskeletal networks.
InaC is required to activate both ARF1 and RhoA, which is tightly regulated during Chlamydia infection (Fig. 2) (15). This suggests that InaC may possess intrinsic GEF activity. However, we have shown that InaC does not exhibit GEF activity in vitro (15), indicating that other factors likely activate ARF1 and RhoA. How these GTPases are being inactivated and how this inactivation is coordinated are also unknown. Due to the significant number of ARF1 and RhoA GEFs and GTPase-activating proteins (GAPs) (51,52), it is challenging to postulate what specific regulators may be involved. Nevertheless, GEFs containing GAP domains (52), or GAPs that have both ARF and RhoA GAP domains (53), may be critical for coupling the activities of ARF1 and RhoA.
Our data demonstrate that PTM-MT and actin scaffold formation occurs with distinct kinetics and that disruption of one of these pathways impacts the other. For example, we show that the constitutive activation of ARF1 blocks the formation of actin scaffolds (Fig. 5). Why can't actin scaffolds form in the presence of PTM-MT scaffolds? One possibility is steric hindrance. Since PTM-MTs woven around the inclusion possess enhanced stability and rigidity, the presence of these cytoskeletal elements might limit access to the inclusion membrane either for actin effectors to interact with chlamydial effectors or for the polymerization of actin itself. Additionally, PTM-MT and actin scaffolds may share critical components for their formation. We have already identified InaC as such a regulator; however, other host effectors may also be shared between these two pathways. For example, host formins control the generation of PTM-MTs and the polymerization of actin filaments (54). Thus, the activation of ARF1dependent PTM-MTs may sequester critical components away from the actin pathway until ARF1 is inactivated, and these effectors can then participate in RhoA-dependent actin scaffold formation. Identification of these components will provide insight into how Chlamydia coordinates the specific kinetics of each of these cytoskeletal scaffolds.
While the activation of ARF1 blocks actin scaffold formation, the loss of RhoA results in the enhanced interaction between ARF1 and InaC and increased PTM-MT scaffold formation (Fig. 4). This phenotype can be reversed by the expression of RhoA in any of its nucleotide-bound states ( Fig. 4; Fig. S7). The nucleotide independence of this complementation suggests that (i) the presence of RhoA on the inclusion is sufficient to downregulate ARF1 or (ii) RhoA may interact with effectors irrespective of its nucleotidebound state to shut off ARF1. Several reports have documented that RhoA and other GTPases interact with other proteins in a nucleotide-independent manner to control their localization and activation (55)(56)(57). Thus, not only has Chlamydia co-opted two host GTPases to coordinate the formation of two cytoskeletal scaffolds, but it has also taken advantage of both nucleotide-dependent and -independent functions of these GTPases.
In summary, our findings have revealed a mechanism by which Chlamydia utilizes a single effector protein to control two cytoskeletal pathways. By employing GTPase cross talk, Chlamydia temporally regulates the assembly of PTM-MT and actin scaffolds around its inclusion to maintain its parasitic niche. Ultimately, identifying the molecular players participating in these pathways will uncover new targets to combat Chlamydia trachomatis infection.

MATERIALS AND METHODS
Cell culture and transfections. HeLa cells (CCL-2; ATCC) and Nuclight Red HeLa cells (Essen Biosciences) were cultured as described previously in Dulbecco's modified Eagle's medium (DMEM) containing 10% fetal bovine serum (FBS), 10 mg/ml gentamicin, and nonessential amino acids (15). Cells were transfected with Continuum transfection reagent (Gemini Bioproducts) using 25 to 50 ng of DNA according to the manufacturer's instructions. For siRNA transfections, HeLa cells were transfected using 25 nM siRNA and DharmaFect 1 reagent (Horizon) according to the manufacturer's instructions 48 h before infection. SmartPool ON-TARGETplus human ARF1 and nontargeting control siRNA were purchased from Dharmacon (15).

Coordination of Actin and Microtubules by Chlamydia
Generation and characterization of the RhoA KO CRISPR/Cas9 cell line. Guide RNAs targeting human RhoA was designed using the sequences available in GenBank and analyzed for their efficiency and off-target effects, using the CRISPOR (61) and the Broad Institute single guide RNA (sgRNA) designer (62) databases with default cutoffs. The optimal guide RNA was prepared by annealing primers FO1033 (59-CAC CGG AAC TAT GTG GCA GAT ATC G-39) and FO1034 (59-AAA CCG ATA TCT GCC ACA TAG TTC C-39). The guide RNA was then inserted into the BsbI site of the pSpCas9(BB)-2A-GFP (PX458) vector (S. Kim, Thomas Jefferson University) and confirmed by Sanger sequencing (Genewiz). HeLa cells were then transfected with pSpCas9(BB)-2A-GFP containing the guide RNA as well as the empty vector (no guide RNA) as described previously (63). Individual clones were isolated by limited dilution. Successful knockout of RhoA was confirmed by Western blotting. Cell division analysis using carboxyfluorescein succinimidyl ester (CFSE) labeling was conducted as described previously (63).
Preparation of cell lysates for Western blot analysis. Cells were washed with PBS and lysed with ice-cold lithium dodecyl sulfate (LDS)-PAGE sample buffer (Invitrogen) containing 250 U/ml benzonase (Accelagen), 1 mg/ml pepstatin A, 5 mg/ml leupeptin, 1 mM phenylmethylsulfonyl fluoride, 10 mM sodium fluoride, and 5.4 mM sodium orthovanadate for 10 min on ice. b-Mercaptoethanol was then added to a final concentration of 0.36 M at 95°C for 5 min, followed by centrifugation at 20,000 Â g at 4°C for 10 min. Protein concentrations were determined using the Pierce 660-nm protein assay reagent containing ionic detergent compatibility reagent and read at 660 nm in a SpectraMax M2 plate reader.
Western blotting. Samples were separated on 4-to 12% bis-Tris SDS-PAGE gels (Invitrogen) and transferred to polyvinylidene difluoride membranes for 1 h at 90 V and 4°C in transfer buffer (25 mM Tris, 192 mM glycine, 10% methanol). Membranes were washed in TBS (25 mM Tris base, 150 mM NaCl [pH 7.5]) and then dried at room temperature for 1 h. Membranes were rehydrated with methanol and washed with TBS and TBST (TBS containing 0.1% Tween 20). Membranes were then blocked for 1 h at room temperature with blocking buffer (3% bovine serum albumin and 0.05% sodium azide in TBST). After blocking, membranes were incubated in primary antibody overnight at 4°C in blocking buffer. Membranes were washed with TBST before incubation with HRP-conjugated secondary antibody for 1 h at room temperature in 0.5% milk diluted in TBST. Membranes were then washed several times with TBST and TBS and revealed with SuperSignal West Dura extended duration substrate (Thermo Scientific). Membranes were imaged on a FluorChem R system (ProteinSimple). Band intensities were quantified using AlphaView software (ProteinSimple).
Inclusion lysis analysis. Cells were infected with WT or InaC KO C. trachomatis L2 at a multiplicity of infection (MOI) of 0.5 and fixed at the indicated times. When noted, WT C. trachomatis L2-infected cells were treated with either 0.5 mM latrunculin B or ethanol (vehicle control) in cell culture medium for 30 min at 37°C 5 h prior to fixation. Latrunculin B and ethanol were washed out with cell culture medium, and cells were fixed at the indicated times. For complementation of inclusion stability, HeLa cells were infected with WT or InaC-FLAG-expressing InaC KO C. trachomatis L2 at an MOI of 0.5 and fixed at 48 hpi. Inclusion integrity was determined by labeling inclusions with IncA and identifying discontinuity in the labeling of IncA on the inclusion membrane. An inclusion was considered broken or lysed when one or more substantial gaps in the incidence of IncA labeling were observed using wide-field fluorescence microscopy. Most of the broken inclusions were missing large pieces of the inclusion membrane. When inclusion integrity was less clear, the presence of Chlamydia in the cytosol was used to identify a broken inclusion.
Immunofluorescence microscopy. HeLa cells were fixed with either (i) 4% paraformaldehyde in PEMS buffer (80 mM PIPES, 5 mM EDTA, 2 mM MgCl 2 , 50 mM sucrose [pH 6.8]) for 15 min at room temperature or (ii) ice-cold methanol for 10 min at room temperature. Cells were then washed in IF-G buffer (25 mM HEPES, 150 mM NaCl, 900 nM CaCl 2 , 500 nM MgCl 2 , 100 mM glycine [pH 7.5]) for paraformaldehyde fixation or IF buffer (25 mM HEPES, 150 mM NaCl, 900 nM CaCl 2 , 500 nM MgCl 2 [pH 7.5]) for methanol fixation. Permeabilization was performed with 0.2% Triton X-100 in IF buffer for 10 min, followed by washes with 0.1% Triton X-100 in IF buffer. Cells were then blocked for 1 h with either (i) goat serum (10% goat serum, 0.05% sodium azide, 0.1% Triton X-100 in IF buffer) or (ii) donkey serum (10% donkey serum, 0.05% sodium azide, 0.1% Triton X-100 in IF buffer) blocking buffer. After blocking, cells were treated with the primary antibodies diluted in the appropriate blocking buffer for 1 h at room temperature. Cells were washed with 0.1% Triton X-100 in IF buffer and subsequently incubated with Alexa Fluor-conjugated secondary antibodies and Hoechst, diluted in the appropriate blocking buffer for 1 h at room temperature. Cells were washed 0.1% Triton X-100 in IF buffer, followed by multiple washes with IF buffer, and mounted on coverslips with ProLong glass antifade mounting medium (Invitrogen). For inclusion lysis experiments, permeabilization, blocking, and subsequent washes were performed in 0.1% saponin instead of Triton X-100. A minimum of 100 cells per condition were observed and imaged with a 60Â oil immersion lens on a Nikon TiE inverted fluorescence microscope and Elements software (Nikon). Images were processed using ImageJ (NIH).