Protein tyrosine phosphatase PTP1B is a positive regulator of the intracellular development of Chlamydia trachomatis

ABSTRACT

The intracellular survival and replication of Chlamydia trachomatis rely on the precise manipulation of host signaling pathways. Host kinases are instrumental in the modulation of host signaling during C. trachomatis infection. However, the potential contribution of host phosphatases to chlamydial pathogenesis remains poorly understood. Here, we identified the host tyrosine phosphatase PTP1B as a positive regulator of C. trachomatis intracellular development. Gain-of-function approaches revealed that PTP1B promotes inclusion development and increases the production of infectious elementary bodies, whereas loss-of-function by chemical inhibition or silencing leads to a reduction in both inclusion size and bacterial infectivity. Interestingly, PTP1B inhibition did not affect Chlamydia trachomatis invasion efficiency, suggesting a specific role during the developmental phase of the chlamydial life cycle. To explore the functional relevance of PTP1B and its potential interaction with chlamydial effectors, we focused on the early-secreted effector Tarp, which undergoes tyrosine phosphorylation upon host cell entry. In vitro biochemical assays demonstrated that recombinant PTP1B can dephosphorylate both native and recombinant forms of Tarp. However, PTP1B inhibition during infection did not significantly alter Tarp phosphorylation levels, possibly owing to the overpowering influence of host tyrosine kinases. These findings suggest that while Tarp may not be a major physiological substrate, PTP1B is capable of interacting with phosphorylated chlamydial effectors. Together, these results establish PTP1B as a host factor that supports chlamydial development and underscore the underappreciated role of host phosphatases in bacterial pathogenesis. This study provides a foundation for future work exploring phosphatase-mediated regulation of infection and potential host-directed therapeutic strategies.

INTRODUCTION

Chlamydia trachomatis is the leading cause of bacterial sexually transmitted infections globally and a major contributor to trachoma, the primary infectious cause of preventable blindness in many developing countries (1, 2). This obligate intracellular pathogen undergoes a unique biphasic developmental cycle that alternates between two distinct forms: the infectious elementary body (EB) and the replicative reticulate body (RB) (3). Upon entering host epithelial cells, EBs transition into RBs within a membrane-bound vacuole known as the inclusion, where they replicate (4). To establish and maintain this specialized intracellular niche, C. trachomatis extensively manipulates host cellular machinery, including signaling pathways, membrane trafficking systems, cytoskeletal structures, and metabolic networks (5–9). Its capacity to evade immune responses and persist within host cells contributes to chronic infections that can lead to reproductive complications and inflammatory disease (10–12).

Manipulation of the host cell is largely orchestrated by a suite of chlamydial proteins delivered into the host cytosol or to the inclusion membrane via the bacterial type III secretion system (T3SS) (13–15). These bacterial effectors modulate diverse host pathways and are frequently subject to post-translational modifications, particularly phosphorylation by host kinases (16–18). An early effector, the translocated actin-recruiting phosphoprotein (Tarp), is rapidly phosphorylated upon delivery by host Src family kinases and facilitates actin cytoskeletal rearrangements at the site of Chlamydia invasion (19, 20). While considerable attention has been devoted to the role of host kinases in supporting effector function and promoting infection (18, 21, 22), the role of host phosphatases, key negative regulators of phosphorylation signaling, remains largely unexplored.

Previous studies have shown that bacterial phosphatases contribute to chlamydial development. For instance, a chlamydial-encoded phosphatase was found to be essential for maintaining inclusion membrane integrity and promoting bacterial growth, underscoring the importance of phosphatase activity for physiology (23). However, the roles of host phosphatases in the context of C. trachomatis infection remain poorly understood. This is particularly noteworthy given the dynamic nature of host phosphorylation networks during C. trachomatis infection and the potential for phosphatases to fine-tune key signaling events that influence pathogen survival.

The protein tyrosine phosphatase 1B (PTP1B) is a central tyrosine-specific phosphatase that regulates diverse signaling cascades, including those involved in metabolism, immune responses, and vesicle trafficking (24–27). Notably, several chlamydial effectors, including Tarp, undergo tyrosine phosphorylation, highlighting the potential relevance of a tyrosine phosphatase like PTP1B in modulating C. trachomatis effector activity during infection. PTP1B is anchored to the cytoplasmic face of the endoplasmic reticulum, enabling it to access both cytosolic and membrane-bound substrates (28). Although understanding of its involvement in infection is limited, PTP1B has been shown to influence host responses during Pseudomonas aeruginosa infection (29, 30). Despite this, the contribution of PTP1B to chlamydial infection has not been previously investigated.

In this study, we aimed to investigate whether PTP1B contributes to the intracellular development of C. trachomatis. Using pharmacological inhibition, siRNA-mediated knockdown, and overexpression approaches, we found that PTP1B activity directly correlates with inclusion maturation and the generation of infectious progeny. In vitro assays demonstrated that PTP1B can dephosphorylate the chlamydial effector, Tarp. However, inhibition of PTP1B during infection did not significantly alter Tarp phosphorylation patterns. These findings demonstrate that PTP1B plays a functional role in supporting chlamydial intracellular development and establish it as an important host enzyme in chlamydial infections. It also provides a foundation for future studies to delineate the downstream host pathways influenced by PTP1B during infection, which may offer new avenues for therapeutic targeting of C. trachomatis.

RESULTS

Chemical inhibition of the tyrosine phosphatase PTP1B impairs Chlamydia trachomatis inclusion development

The phosphoproteome of the infected cell changes during C. trachomatis entry and development, as both host and bacterial proteins are modified by host kinases (31, 32). Phosphoproteomics analysis of the chlamydial developmental cycle indicates that the phosphorylation of many cellular and bacterial targets is temporally controlled with phosphorylation levels both rising and falling (33). While much attention has previously been directed toward the role of kinases during chlamydial development, we sought to investigate the role of host cell phosphatases. To investigate the role of tyrosine phosphatase PTP1B in C. trachomatis development, HeLa cells were treated with increasing concentrations of a commercially available PTP1B chemical inhibitor, followed by infection with C. trachomatis elementary bodies (EBs) (Fig. 1A). Chlamydia-infected cells were fixed 40 h post-infection, and inclusions were visualized by immunofluorescence microscopy using an antibody specific for the chlamydial major outer membrane protein (MOMP). At the highest concentration tested (300  µM), visible signs of cytotoxicity, such as cell rounding and detachment, were observed. To ensure that the observed phenotypes were not due to cytotoxic effects of the inhibitor, we performed an LDH release assay following 27 h of treatment. Absorbance measurements indicated that only the highest concentration (300  µM) of the PTP1B inhibitor caused significant cytotoxicity, whereas 75  µM and 150  µM did not show any significant effect (Fig. 1B). These concentrations were therefore selected for downstream experiments.

Fig 1.

Effect of PTP1B inhibition on Chlamydia trachomatis inclusion formation and treatment-associated cytotoxicity. (A) HeLa cells were pre-treated with increasing concentrations of a PTP1B inhibitor and subsequently infected with C. trachomatis serovar L2 at a multiplicity of infection (MOI) of 5. At 36 h post-infection, cells were fixed with methanol and stained with a mouse monoclonal antibody against the chlamydial major outer membrane protein (MOMP), followed by goat anti-mouse secondary antibody conjugated to Alexa Fluor 488 (green) to visualize inclusions. Fluorescence and phase contrast images were acquired using the BioTek Cytation 5 imaging system. Scale bar, 100 µm. (B) To evaluate cytotoxicity associated with PTP1B inhibitor treatment, HeLa cells were treated with the PTP1B inhibitor for 27 h, and cytotoxicity was assessed using the Invitrogen CyQUANT LDH Cytotoxicity Assay. Absorbance was measured as an indicator of LDH release, with maximum and spontaneous release controls included. Statistical significance was determined using one-way ANOVA followed by Dunnett’s multiple comparisons test. ****P  <  0.0001; ns, not significant. Error bars represent standard deviation (SD).

Compared to the DMSO control, PTP1B inhibition led to a dose-dependent reduction in C. trachomatis inclusion size, with treated cells displaying smaller and less defined inclusions (Fig. 2A). Quantification of captured immunofluorescence images confirmed a significant decrease in both mean inclusion diameter and area in PTP1B-inhibitor treated cells (Fig. 2B and C).

Fig 2.

Chemical inhibition of PTP1B impairs Chlamydia trachomatis development. (A) HeLa cells treated with PTP1B chemical inhibitors (75 or 150 µM) or vehicle control (DMSO) were infected with C. trachomatis EBs. Forty hours post-infection, cells were fixed, and inclusions were visualized using mouse antibodies specific for the chlamydial major outer membrane protein (α MOMP) along with goat anti-mouse antibodies conjugated to Alexa 488 (green). Separate phase contrast (phase) and merged images are provided for each field of view. One representative image (out of 15–20 per condition) is provided for each treatment. Scale bar, 200 µm. (B–C) Quantification of the mean inclusion diameter (µm) and mean inclusion area (µm²) across treatment conditions. Error bars represent SD. (D) Quantification of the chlamydial development cycle by measuring the production of infectious progeny (inclusion-forming units [IFUs/mL]) harvested after 36–42 h of treatment. The number of viable EBs per mL was determined by infecting HeLa cells with serially diluted lysates and quantifying the formation of inclusions after 24 h. Data represent the mean of the three independent experiments. Error bars represent SD. Statistical significance was determined using one-way ANOVA followed by Tukey’s multiple comparisons test. ***P < 0.001; ****P < 0.0001; ns, not significant.

To determine whether impaired inclusion development observed for the PTP1B inhibitor-treated cells affected EB production, infectious progeny were quantified by infecting HeLa cells with EBs harvested 36–42 h post-treatment with and without PTP1B inhibitor. Cells treated with the PTP1B inhibitor exhibited significantly reduced inclusion-forming units (IFUs/mL) relative to the control, indicating diminished bacterial replication (Fig. 2D). These findings demonstrate that chemical inhibition of PTP1B disrupts C. trachomatis inclusion development, underscoring a potential regulatory role for PTP1B in host-pathogen interactions.

PTP1B inhibition does not affect Chlamydia trachomatis invasion

Given our findings that PTP1B inhibition reduces Chlamydia trachomatis inclusion size and progeny, we sought to determine whether this effect was due to impaired bacterial invasion or defects in intracellular development post-entry. To accomplish this, we compared C. trachomatis inclusion numbers and development in HeLa cells treated with the PTP1B inhibitor either 2 h before (pre-treatment) or 2 h after (post-treatment) infection with C. trachomatis. Cells were fixed at 40 h post-infection, and inclusions were visualized by staining with MOMP-specific antibodies (Fig. 3A).

Fig 3.

PTP1B inhibition does not affect Chlamydia trachomatis invasion. (A) HeLa cells were treated with 150 µM PTP1B inhibitor (PTP1Bi) either 2 h before C. trachomatis infection (Pre), 2 h after infection (Post), or with DMSO alone before infection (DMSO). At 40 h post-infection, cells were fixed, and inclusions were visualized using mouse antibodies specific for the chlamydial major outer membrane protein (α MOMP) along with goat anti-mouse antibodies conjugated to Alexa 488 (green). Separate phase contrast (phase) and merged images are provided for each field of view. One representative image is shown for each treatment. Scale bar, 100 µm. (B–D) Quantification of C. trachomatis inclusion size and number across conditions. Inclusion diameter (B), inclusion area (C), and the number of inclusions per field of view (D) were measured. Error bars represent SD. (E) Invasion assay. HeLa cells were pre-treated with DMSO or PTP1B inhibitor (150 µM) for 2 h and then infected with red fluorescent C. trachomatis EBs (pre-labelled with Cell Tracker dye). At 90 min post-infection, cells were fixed with 4% paraformaldehyde for 15 min and stained for extracellular EBs using a mouse anti-Chlamydia LPS antibody followed by Alexa 488-conjugated goat anti-mouse secondary antibody. The percentage of invaded bacteria was calculated by comparing total (red) versus extracellular (green) bacteria. At least 15 fields of view were analyzed per condition. Error bars represent SD. Data represent mean of the three independent experiments. Statistical significance was determined using one-way ANOVA with Tukey’s multiple comparisons test for (B–D) and an unpaired t-test for (E) ****P < 0.0001; ns, not significant.

Quantification of inclusion size revealed a significant reduction in inclusion diameter (Fig. 3B) and inclusion area (Fig. 3C) in both the pre-treatment and post-treatment groups compared to the DMSO controls. However, no significant difference in size was observed between the pre-treatment and post-treatment groups, indicating that PTP1B inhibition impairs inclusion growth regardless of whether it occurs before or after bacterial entry. In contrast, the number of inclusions per field of view remained unchanged across all conditions (Fig. 3D), suggesting that PTP1B inhibition does not affect C. trachomatis invasion. Additionally, an invasion assay using CMTPX-red-labeled EBs confirmed no significant difference in bacterial entry between PTP1B-inhibited and control cells (Fig. 3E). Together, these findings suggest that PTP1B activity is important for the intracellular development of C. trachomatis but is dispensable for bacterial entry into host cells.

PTP1B knockdown impairs Chlamydia trachomatis intracellular development and reduces infectivity

To extend our chemical inhibitor findings and to directly assess the role of PTP1B, we utilized an siRNA-based approach to deplete endogenous PTP1B in HeLa cells. Western blot analysis confirmed efficient and stable depletion of PTP1B protein levels at 24, 48, and 72 h post-transfection with PTP1B-targeting siRNAs, while control siRNA-treated cells retained stable PTP1B expression (Fig. 4A). As PTP1B protein levels were reduced as early as 24 h post-transfection, cells were infected with C. trachomatis 24 h after knockdown and fixed at 40 h post-infection, and inclusions were visualized by immunofluorescence microscopy. PTP1B-depleted cells exhibited smaller inclusions compared to control siRNA and mock-treated cells (Fig. 4B). Quantitative image analysis confirmed a significant reduction in both mean inclusion area and diameter in the knockdown group (Fig. 4C and D). To assess whether the reduction in inclusion size interfered with EB production, we quantified infectious progeny by measuring inclusion-forming units (IFUs/mL). Consistent with reduced inclusion size, PTP1B knockdown resulted in significantly lower IFUs compared to the control groups (Fig. 4E). Together, these findings demonstrate that specific depletion of endogenous PTP1B disrupts C. trachomatis intracellular development and replication, reinforcing the role of PTP1B as a host factor that promotes chlamydial growth.

Fig 4.

PTP1B knockdown impairs C. trachomatis inclusion development and reduces infectivity. (A) HeLa cells were transfected with PTP1B-targeting siRNA or control siRNA, and lysates were collected at 24, 48, and 72 h post-transfection. Samples were resolved by SDS-PAGE and subjected to immunoblotting with antibodies specific for PTP1B (α PTP1B) and actin (α Actin) as a loading control. (B) Forty hours post-infection, cells were fixed, and inclusions were visualized using mouse antibodies specific for the chlamydial major outer membrane protein (α MOMP) and goat anti-mouse antibodies conjugated to Alexa 488 (green). Separate phase contrast (phase) and merged images are provided for each condition. One representative image is shown. Scale bar, 100 µm. (C–E) Quantification of the mean inclusion diameter (µm) (C), mean inclusion area (µm²) (D), and bacterial infectivity based on inclusion-forming units (IFUs/mL) (E). Infectious progeny were harvested at 40 h post-infection, and infectivity was determined by reinfecting fresh HeLa cells with serially diluted lysates and quantifying inclusion formation. Data represent the mean of the three independent experiments. Error bars represent SD. Statistical significance was determined using one-way ANOVA followed by Tukey’s multiple comparisons test. ***P < 0.001; **P  <  0.01; ns, not significant.

PTP1B overexpression promotes Chlamydia trachomatis inclusion development and increases bacterial yield

To assess whether elevated PTP1B levels impact C. trachomatis intracellular development, we cloned the coding sequence of human PTP1B into the mammalian expression vector pEGFPC₃ and transfected HeLa cells. We selected stable cells overexpressing PTP1B with G418 (Geneticin). Stable overexpression of PTP1B in transfected cells (C₃ PTP1B HeLa) was confirmed by immunoblot, which demonstrated increased PTP1B protein levels compared to vector-only control cells (C₃ HeLa) (Fig. 5A). Transfected cells were infected with C. trachomatis, fixed 40 h post-infection, stained with antibodies specific for the chlamydial major outer membrane protein (MOMP), and visualized by fluorescence microscopy. Compared to control cells, PTP1B-overexpressing cells exhibited larger and more prominent inclusions (Fig. 5B). Quantitative analysis revealed a significant increase in both mean C. trachomatis inclusion diameter and area in C₃ PTP1B-transfected cells compared to control-transfected cells (Fig. 5C and D). To evaluate whether PTP1B overexpression enhances bacterial replication, infectious progeny were quantified by measuring inclusion-forming units (IFUs/mL). C₃ PTP1B-transfected cells produced significantly more infectious EBs than vector control-transfected cells (Fig. 5E). Albeit a small but statistically significant increase, these data indicate that elevated levels of PTP1B promote intracellular growth and replication of C. trachomatis, further supporting its positive regulatory role during infection.

Fig 5.

PTP1B overexpression enhances Chlamydia trachomatis development and infectivity. (A) HeLa cells were transfected with either an empty pEGFPC₃ vector (C₃ HeLa) or a PTP1B-expressing construct (C₃ PTP1B HeLa). Forty-eight hours post-transfection, whole-cell lysates were collected and subjected to SDS-PAGE followed by immunoblotting with antibodies specific for PTP1B (α PTP1B) and actin (α Actin) as a loading control. (B) Forty hours post-infection with C. trachomatis, cells were fixed and stained using mouse antibodies specific for the chlamydial major outer membrane protein (α MOMP) and goat anti-mouse antibodies conjugated to Alexa 594 (red). Separate phase contrast (phase) and merged images are shown. One representative field of view is presented for each condition. Scale bar, 100 µm. (C–E) Quantification of the mean inclusion diameter (µm) (C), mean inclusion area (µm²) (D), and bacterial infectivity based on inclusion-forming units (IFUs/mL) (E). Infectious progeny were harvested at 40 h post-infection, and infectivity was determined by reinfecting fresh HeLa cells with serially diluted lysates and quantifying inclusion formation. Data represent the mean of the three independent experiments. Error bars represent SD. Statistical significance was determined using unpaired t-tests. **P < 0.01.

Recombinant PTP1B dephosphorylates the Chlamydia trachomatis early effector Tarp

Early chlamydial effectors such as Tarp are phosphorylated by host tyrosine kinases soon after translocation into a mammalian host cell during EB entry (20). To investigate the phosphorylation kinetics of Tarp during Chlamydia trachomatis infection, we examined Tarp phosphorylation levels at discrete time points between 0 and 12 h post infection in McCoy, HeLa, and J774 macrophage cell lines. As seen previously, immunoblot analysis demonstrated that Tarp is rapidly phosphorylated upon infection, with phospho-Tarp levels peaking early and declining over time depending on the tissue culture cell type employed (Fig. 6A). Interestingly, total Tarp protein levels increased up to 4 h post-infection and then stabilized, indicating that the observed changes in phospho-Tarp levels were due to dynamic phosphorylation rather than new Tarp synthesis. De novo Tarp expression occurs late in the developmental cycle, and therefore, Tarp levels observed early in the infection are EB-derived. The observation that the Tarp effector undergoes active dephosphorylation prompted us to examine whether PTP1B contributes to the regulation of phospho-Tarp levels.

Fig 6.

Tarp phosphorylation kinetics and dephosphorylation by recombinant PTP1B. (A) Time course analysis of Tarp phosphorylation during C. trachomatis infection in different cell lines. McCoy, HeLa, and J774 cells were infected with C. trachomatis L2 (MOI ~ 50–100) and harvested at the indicated time points post-infection. Lysates were resolved by SDS-PAGE and immunoblotted with phosphotyrosine-specific (α YPO₄) and Tarp-specific (α Tarp) antibodies to assess phosphorylation dynamics. Uninfected host cell lysates served as controls. (B) Dephosphorylation of recombinant Tarp by recombinant PTP1B. Purified recombinant Tarp was phosphorylated in the presence of ATP and HeLa lysate and then heat-inactivated to stop kinase activity. Recombinant PTP1B was added for 1 h in the presence or absence of ZnCl₂, and samples were resolved by SDS-PAGE, stained with Coomassie blue, and immunoblotted with phosphotyrosine-specific (αYPO₄) antibodies. (C) Dephosphorylation of native Tarp by recombinant PTP1B. Extracts from C. trachomatis-infected HeLa cells (2 h post-infection) were incubated with recombinant PTP1B for 1 h. Extracts from uninfected HeLa cells served as controls. Proteins were resolved by SDS-PAGE and immunoblotted with phosphotyrosine (α YPO₄) and Tarp-specific (α Tarp) antibodies. Molecular weight markers are shown in kilodaltons (kDa). (D) Tarp phosphorylation kinetics during C. trachomatis infection of HeLa cells treated with either DMSO control (D) or PTP1B inhibitor (I). Cells were harvested at 0, 1, and 6 h post-infection. Lysates were analyzed by SDS-PAGE and immunoblotting with anti-phosphotyrosine (α YPO₄), anti-Tarp (α Tarp), anti-MOMP (α MOMP), and anti-actin (α Actin) antibodies. Actin served as a loading control.

To investigate whether PTP1B is capable of targeting chlamydial effector proteins, we tested whether recombinant PTP1B is able to dephosphorylate Tarp, a key early effector that undergoes tyrosine phosphorylation during Chlamydia trachomatis entry. First, recombinant PTP1B was expressed in E. coli and purified by affinity chromatography. Coomassie-stained SDS-PAGE and immunoblot confirmed the successful purification of soluble recombinant PTP1B protein (~45 kDa) (Fig. S1A). To assess phosphatase activity, purified recombinant Tarp was phosphorylated in vitro using HeLa cell lysate and ATP. Following heat inactivation of the lysate to eliminate residual kinase activity, recombinant PTP1B was added to the reaction (1 h) with or without ZnCl₂. ZnCl₂ was used as an additional negative control for PTP1B activity due to the ability of Zinc ions to inhibit the activity of PTP1B (34). Immunoblot analysis using phosphotyrosine-specific antibodies revealed an absence of detectable phospho-Tarp following treatment with recombinant PTP1B (Fig. 6B), indicating dephosphorylation of Tarp. In contrast, phospho-Tarp was readily detectable in the sample treated with PTP1B in the presence of zinc chloride, suggesting that zinc inhibited PTP1B activity. We next examined the ability of PTP1B to dephosphorylate native Tarp. Lysates from HeLa cells infected with C. trachomatis for 2 h were incubated with recombinant PTP1B for 1 h. Immunoblot revealed an absence of detectable phospho-Tarp in PTP1B-treated samples, while total Tarp levels remained unchanged (Fig. 6C).

To further explore the role of PTP1B during infection, we examined whether PTP1B inhibition altered native Tarp phosphorylation dynamics. HeLa cells were treated with DMSO or PTP1B inhibitor (150 µM) and infected with C. trachomatis, and cell lysates were collected at 0, 1, and 6 h post-infection. Immunoblot showed that phospho-Tarp levels followed a parallel trajectory in both groups, suggesting that inhibition of PTP1B did not substantially affect Tarp phosphorylation kinetics (Fig. 6D). However, total Tarp levels were transiently decreased at the early infection stage in inhibitor-treated cells and returned to baseline by 6 h. This transient decrease was likely attributable to the chemical stimulation of Chlamydia trachomatis elementary bodies (EBs) by the PTP1B inhibitor, which we found can induce the release of Tarp into the extracellular milieu even in the absence of host cell contact (Fig. S1B). Nevertheless, this does not alter the conclusion that PTP1B inhibition did not significantly impact the phosphorylation kinetics of Tarp during infection. Together, our findings demonstrate that recombinant PTP1B can directly dephosphorylate Tarp and suggest that host phosphatases have the capacity to modulate chlamydial effector function, potentially influencing early host-pathogen interactions.

DISCUSSION

The Chlamydia trachomatis developmental cycle requires microbial control of the infected host cell to subvert host defenses and to promote nutrient acquisition (35). Such control is accomplished in part by a collection of bacterial effectors whose purpose is to commandeer host signaling and metabolic pathways (36–38). While the contribution of host kinases to C. trachomatis pathogenesis has been extensively studied—including their roles in cytoskeletal remodeling, inclusion trafficking, and effector phosphorylation (18, 39)—the role of host phosphatases remains largely uncharacterized. Given the extensive rewiring of host and bacterial phosphorylation networks during infection, understanding how host phosphatases contribute to chlamydial development represents an important and underexplored question. Interestingly, the importance of bacterial phosphatase activity in C. trachomatis development has previously been demonstrated. Claywell et al. identified a chlamydial phosphatase, CppA, whose activity contributed to inclusion stability and development (23). However, the potential regulatory roles of host phosphatases during infection remain unknown.

 In this study, we identify the host protein tyrosine phosphatase PTP1B as a positive regulator of chlamydial intracellular development. Using complementary approaches—chemical inhibition, siRNA-mediated knockdown, and overexpression—we demonstrated that PTP1B plays a significant role in promoting C. trachomatis intracellular development. Treatment of host cells with a specific PTP1B inhibitor resulted in a marked reduction in the size of chlamydial inclusions as well as a substantial decrease in the yield of infectious elementary bodies. These inhibitory effects were observed whether cells were treated with the inhibitor prior to infection or after bacterial entry, suggesting that PTP1B functions primarily during the intracellular stages of the Chlamydia developmental cycle. To determine whether PTP1B also influences bacterial entry, we performed invasion assays under inhibitor-treated conditions. These experiments revealed no measurable effect on bacterial internalization, indicating that PTP1B activity is dispensable for the entry process itself. Silencing of PTP1B by siRNA yielded similar phenotypes, reinforcing the conclusion that PTP1B is required for efficient bacterial replication within host cells. In contrast, ectopic overexpression of wild-type PTP1B enhanced both inclusion expansion and EB production, further validating its role as a host factor that supports chlamydial growth.

To investigate the mechanism by which PTP1B supports chlamydial growth, we examined its interaction with the chlamydial effector Tarp, which is phosphorylated by host tyrosine kinases and contributes to actin remodeling during bacterial entry (19, 20, 40). In vitro assays demonstrated that PTP1B can dephosphorylate both recombinant and native forms of phosphorylated Tarp, suggesting that Tarp is a possible substrate of PTP1B. However, inhibition of PTP1B in infected cells did not significantly alter Tarp phosphorylation kinetics, suggesting that PTP1B is not the primary regulator of Tarp phosphorylation during infection. This may be due to functional redundancy among host phosphatases or spatial compartmentalization of PTP1B. A transient reduction in total Tarp levels was observed early during infection in PTP1B-inhibited cells. This observation was likely due to altered effector secretion during the initial host-pathogen interaction. Cell-free elementary bodies exposed to the PTP1B inhibitor released detectable amounts of Tarp. However, this effect did not persist post-infection or affect downstream phosphorylation, suggesting that it may reflect a minor perturbation in effector release rather than a major regulatory role. Nonetheless, the observation raises the possibility that PTP1B may influence the early dynamics of effector delivery or stabilization in a context-dependent manner.

 Together, these results establish PTP1B as a host factor that supports C. trachomatis intracellular replication. Although prior genome-wide siRNA screens, such as that by Rother et al. (41), included numerous phosphatases—including PTP1B (PTPN1), PPP2CA, PPP1CA, PTPN2, and PTPRJ—none were highlighted as key hits in the published analysis, leaving their potential roles in Chlamydia infection largely unexplored. Our findings suggest that host phosphatases—such as PTP1B—may represent an underappreciated layer of regulation, possibly acting in parallel or compensatory signaling pathways. Our study is the first to functionally validate its role during C. trachomatis infection. Although our biochemical assays demonstrated that PTP1B is capable of dephosphorylating both recombinant and native Tarp, inhibition of PTP1B during infection did not noticeably alter Tarp phosphorylation dynamics. This suggests that Tarp is unlikely to be a major physiological substrate of PTP1B in the context of infection. Nonetheless, these findings highlight PTP1B’s ability to engage with chlamydial effectors, underscoring its potential to influence bacterial–host interactions through other, yet unidentified, targets.

In light of our findings, it is worth considering whether other host phosphatases may similarly contribute to C. trachomatis development. While PTP1B is a member of the non-receptor type protein tyrosine phosphatase (PTPN) family, several other tyrosine phosphatases are expressed in epithelial cells and could potentially influence infection dynamics (42–44). Moreover, serine/threonine phosphatases—though not expected to directly regulate tyrosine-phosphorylated effectors like Tarp—may modulate other bacterial or host targets critical to infection. For example, chlamydial proteins have been shown to undergo diverse post-translational modifications, including serine/threonine phosphorylation (17, 18), which may be influenced by these phosphatases. Future studies aimed at systematically evaluating both tyrosine and serine/threonine phosphatases may reveal broader host phosphatase networks that modulate chlamydial pathogenesis.

Beyond its mechanistic role, PTP1B is a clinically relevant target. Several PTP1B inhibitors have been developed for non-infectious diseases and have reached human clinical trials. Ertiprotafib, developed by Wyeth, was the first PTP1B inhibitor to be tested for treatment of type 2 diabetes and obesity (45). Other inhibitors, including KQ-791, trodusquemine (MSI-1436), and ABBV-CLS-484, are currently being evaluated for efficacy in trials for metabolic disorders and cancer (46, 47). Over eight PTP1B-targeting compounds have entered clinical development (48). Preclinical studies also suggest neuroprotective benefits from PTP1B inhibition, including protection from age-related neurodegeneration (49–51). The existence of this drug pipeline opens the possibility of repurposing PTP1B inhibitors as host-directed therapeutics against intracellular pathogens like C. trachomatis. While further validation is needed, our findings support this concept as a potential avenue for combatting persistent or antibiotic-refractory infections.

These findings presented here contribute to a broader recognition that host phosphatases—long overshadowed by kinases in infection biology—play underappreciated but critical roles in host-pathogen interactions. Our study highlights the value of investigating phosphatase signaling networks in the context of infection. Future research will aim to identify additional PTP1B substrates during C. trachomatis infection and clarify how phosphatase activity is temporally and spatially regulated within infected cells. Such work may reveal novel regulatory circuits that are exploitable for therapeutic intervention and expand our understanding of host-directed strategies to restrict intracellular bacterial growth.

MATERIALS AND METHODS

Organisms and cell culture

Chlamydia trachomatis serovar L2 (LGV 434/Bu) was propagated in HeLa 229 and McCoy cells and purified by density gradient centrifugation using diatrizoate meglumine and diatrizoate sodium as previously described (52). HeLa, McCoy, and J774 cell lines were obtained from the American Type Culture Collection (ATCC) and maintained in Dulbecco’s Modified Eagle Medium (DMEM; Gibco) supplemented with 10% fetal bovine serum (FBS; Gibco) and 1% L-glutamine (Gibco). All cell lines were cultured at 37°C in a humidified incubator with 5% CO₂. HeLa cells were used for the majority of experiments unless otherwise noted.

PTP1B inhibitor treatment

PTP1B inhibition was performed using a selective small-molecule inhibitor (CAS-765317-72-4, Millipore Sigma). An initial dose–response titration was conducted by treating HeLa cells with inhibitor concentrations ranging from 25 to 300 µM for 2 h prior to infection, in order to assess effects on host cell morphology and Chlamydia trachomatis inclusion development.

For pre-treatment protocols, cells were incubated with the inhibitor for 2 h prior to infection, and the inhibitor-containing medium was maintained throughout the 40-hour infection period unless otherwise specified.

In post-infection treatment experiments, HeLa cells were infected with C. trachomatis serovar L2 for 2 h, after which the inoculum was removed and replaced with fresh medium containing either DMSO (vehicle control at the same concentration as inhibitor) or PTP1B inhibitor at the indicated concentrations. Cells were then incubated for the remainder of the 40-hour infection period prior to fixation and analysis.

LDH cytotoxicity assay

LDH release was quantified using the Invitrogen CyQUANT LDH Cytotoxicity Assay Kit according to the manufacturer’s instructions. Briefly, HeLa cells were seeded in 96-well plates (3,000 cells per well) 24 h prior to treatment and incubated with the PTP1B inhibitor (25 µM–300 µM) for 27  h. For controls, maximum LDH release wells were treated with 10× lysis buffer, and spontaneous LDH release wells received sterile water for 45  min before supernatant collection.

At the end of the treatment period, culture supernatants were collected from each well and transferred to fresh assay wells. LDH reaction mixture was added as directed by the kit protocol, incubated per the manufacturer’s instructions at room temperature (protected from light), and the reaction was stopped with the provided stop solution. Absorbance was recorded at 490  nm with 680  nm as the reference wavelength. Absorbance was analyzed as an indicator of LDH release. For each well, background-corrected absorbance was calculated as A490–A680. For data processing, technical replicates were averaged, and treatment groups were compared to vehicle (DMSO) controls.

Chlamydia trachomatis infection and indirect immunofluorescence

HeLa cells were seeded in 6-well plates at a density of 2 × 10⁵ cells per well and pre-treated the following day with PTP1B inhibitor (75–150 µM) or DMSO for 2 h. After pre-treatment, cells were infected with Chlamydia trachomatis serovar L2 at a multiplicity of infection (MOI) ranging from 0.1 to 5 and incubated at 37°C with 5% CO₂ for 40 h. In case of siRNA treatment or PTP1B transfections, cells were infected after a suitable treatment duration (24–72 h).

 At 36–42 h post-infection, cells were fixed with methanol for 15 min at room temperature and washed three times with 1× phosphate-buffered saline (PBS). Fixed cells were blocked with 10% bovine calf serum (Gibco) for 1 h and incubated with a mouse monoclonal antibody against C. trachomatis major outer membrane protein (MOMP) (Invitrogen) at 1:500 dilution for 1 h at room temperature, followed by three to four PBS washes and incubation with an Alexa Fluor–conjugated goat anti-mouse IgG secondary antibodies Alexa 488 or Alexa 594 (Invitrogen) at 1:500 dilution for 1 h in the dark. Fluorescence images were acquired using the Biotek Cytation 5 imaging system using 40× magnification, and inclusion diameters and area were measured using the Gen5 software. Ten to fifteen fields of view were analyzed per condition.

Inclusion forming unit assay

HeLa cells seeded at 2 × 10⁵ cells per well were infected with C. trachomatis serovar L2 at a multiplicity of infection (MOI) ranging from 0.1 to 5 and incubated at 37°C for 36–42 h. Cells were scraped, and the entire contents of each well (cells plus media) were collected. Samples were subjected to brief sonication to release elementary bodies. The resulting lysates were serially diluted in Hanks’ Balanced Salt Solution (HBSS) and used to infect fresh monolayers of HeLa cells seeded in 24-well plates. After a 24-hour secondary infection period, cells were fixed with methanol for 15 min, washed with PBS, blocked with 10% bovine calf serum, and stained using a mouse monoclonal antibody against C. trachomatis MOMP (1:500 dilution), followed by an Alexa Fluor 488–conjugated goat anti-mouse secondary antibody (1:500 dilution). Fluorescence images were acquired using the Biotek Cytation 5 imaging system, and inclusion-forming units (IFUs) were quantified using the Gen5 software by analyzing 15 random fields per well at 20× magnification.

C. trachomatis invasion assay

Chlamydia trachomatis elementary bodies (EBs) were labeled with CellTracker Red CMTPX Dye (Invitrogen) during propagation in HeLa cells, as previously described (53). For invasion assays, CMTPX-labeled EBs were harvested and used to infect HeLa cells seeded in 24-well tissue culture plates. Cells were grown in Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 1% L-glutamine for 24 h prior to infection. To synchronize the infections, plates were pre-chilled on ice for 5 min. CMTPX-labeled EBs were added to the wells at a multiplicity of infection (MOI) of ~10 and allowed to attach for 20 min while the plates remained on ice. After attachment, pre-warmed DMEM containing 10% FBS was added, and plates were transferred to 37°C for 90 min to allow internalization. Following incubation, media was removed, and cells were fixed with 4% paraformaldehyde (PFA) for 15 min at room temperature. Cells were not permeabilized. To label extracellular EBs, a mouse monoclonal anti-Chlamydia LPS antibody (Invitrogen) was used at a 1:100 dilution for 1 h at room temperature. After four PBS washes, Alexa Fluor 488-conjugated anti-mouse secondary antibody was applied (dilution 1:500) for 1 h. After final washes, fluorescence imaging was performed directly in the 24-well plates using the Biotek Cytation 5 imaging system. At least 15 fields of view per well were acquired under identical exposure settings. Quantification of invasion efficiency was conducted using Gen5 software (Agilent). Red fluorescence (CMTPX) marked total EBs, while green fluorescence indicated extracellular EBs. The percentage of internalized EBs was calculated using the formula: [((red − green)/red) × 100] = % Invasion.

siRNA-mediated knockdown of PTP1B

PTP1B knockdown was performed using small interfering RNAs (siRNAs) synthesized by Integrated DNA Technologies (IDT). Cells were seeded in antibiotic-free DMEM supplemented with 10% FBS and allowed to reach 50–60% confluency prior to transfection. Transfections were carried out using Lipofectamine 3000 (Thermo Scientific) according to the manufacturer’s protocol. Briefly, cells were transfected with 20–60 nM siRNAs targeting human PTPN1 or a non-targeting negative control siRNA (Table S1). To assess knockdown efficiency, cells were lysed directly in 5× SDS sample buffer at 24, 48, and 72 h post-transfection.

Cloning and protein expression

The in-frame amino-terminal glutathione-S-transferase (GST) fusion protein for PTP1B was generated by PCR amplifying the PTP1B gene from cDNA from human dermal fibroblast cells (ATCC). PCR was performed with synthesized oligonucleotide primers (Integrated DNA Technologies) engineered with EcoRI and NotI linkers ( Table S1). PCR products were purified, digested with restriction enzymes (New England BioLabs), and subcloned into linearized pGEX-6p-1 to generate translational fusions with GST. The PTP1B gene was also cloned into mammalian expression vector pEGFP-C3 (Clontech) to allow for ectopic expression of PTP1B in HeLa cells. PCR was performed with DNA primers engineered with NheI and BamHI linkers (Table S1). Protein expression and purification were performed according to the procedures outlined for glutathione Sepharose 4B in the Bulk GST Purification Module (Cytiva). In some experiments, the GST tag was removed prior to kinase and phosphatase assays by treatment with PreScission Protease according to the manufacturer’s recommendations (Cytiva).

PTP1B overexpression by transient and stable transfection

HeLa cells were transfected (transient or stable) with PTP1B construct cloned into the pEGFPC₃ vector (PTP1BC₃) or with the empty pEGFPC₃ vector (Clontech). Transfections were performed in 6-well plates (Costar) at ~50–60% confluency using Lipofectamine 3000 (Invitrogen), following the manufacturer’s protocol. For each well, 5  µg of plasmid DNA was mixed with Lipofectamine 3000 and P3000 reagent in Opti-MEM and added directly to the cells. For transient transfection, cells were incubated for 48 h post-transfection before being processed for downstream experiments. For stable transfection, G418 (geneticin) was added at 150 µg/mL 24 h post-transfection, and the selection medium was replaced every 24–36 h. Stably transfected cells were expanded and used for downstream applications once resistant populations were established.

Tarp phosphorylation kinetics

To evaluate the temporal phosphorylation of the Chlamydia trachomatis effector protein Tarp during early infection, HeLa, McCoy, and J774 cells (2 × 10⁵ cells) were seeded in 6-well plates and infected with C. trachomatis serovar L2 at a multiplicity of infection (MOI) of 50–100. A synchronized infection was performed to ensure uniform bacterial entry: prior to infection, cells were placed on ice and infected with cold bacterial suspensions for 30 min, followed by the addition of warm DMEM to initiate synchronous internalization. Plates were then transferred to a 37°C incubator, and infections were allowed to proceed for the indicated durations. Cells were harvested at defined time points between 0 and 12 h post-infection (hpi) to capture dynamic changes in Tarp phosphorylation. At each time point, media was removed, and cells were lysed directly in 5× SDS sample buffer (62.5 mM Tris-HCl [pH 6.8], 10% glycerol, 2% SDS, 5% β-mercaptoethanol, 0.01% bromophenol blue). Lysates were boiled for 5 minutes and stored at −20°C until further analysis.

SDS-PAGE and immunoblot analysis

Protein samples were prepared in 5× SDS sample buffer, heated at 95°C for 5 min, and resolved on 4–12% gradient polyacrylamide gels (Invitrogen). Following electrophoresis, gels were either stained with Imperial Protein Stain (Thermo Scientific) or used for protein transfer onto nitrocellulose membranes via a wet transfer system (Bio-Rad). Membranes were blocked in 5% non-fat dry milk prepared in 1× phosphate-buffered saline with 0.001% Tween-20 for 1 h at room temperature. Primary antibody incubations were performed overnight at 4°C in 1× PBST containing 5% non-fat dry milk. The following primary antibodies were used (dilution 1:1000): monoclonal anti-PTP1B (Santa Cruz Biotechnology), monoclonal anti-actin C4 (Millipore Sigma), monoclonal anti-pan actin clone 7A8.2.1 (Cytoskeleton), monoclonal anti-phosphotyrosine 4G10 (Millipore Sigma), and monoclonal Chlamydia trachomatis anti-MOMP (Invitrogen). Polyclonal rabbit antibodies against Chlamydia trachomatis L2 LGV 434 Tarp (CT456) were generated at Rocky Mountain Laboratories, as previously described (16). After three washes with 1× PBST, membranes were incubated with horseradish peroxidase (HRP)-conjugated goat anti-mouse or goat anti-rabbit secondary antibodies (Invitrogen) for 1 h at room temperature, followed by three to four additional washes with 1× PBST. Blots were developed using Protein Biology chemiluminescent substrate (Thermo Scientific, Pierce), and signals were visualized using a Bio-Rad imaging system or by exposure to autoradiographic film.

Recombinant Tarp dephosphorylation assay

Recombinant N-terminal domain of Tarp was incubated with HeLa cell lysate and ATP at 37°C for 1 h to allow phosphorylation as previously described (20). An aliquot was collected as a pre-heat inactivation control. The remaining reaction was heat-inactivated at 75°C for 6–8 min. Heat-inactivated samples were then divided into three groups: untreated control, PTP1B-treated, and PTP1B + ZnCl₂-treated. All groups were incubated at 37°C for 1 h. Reactions were terminated by adding 5× SDS sample buffer and heating at 95°C for 5 min. Samples were resolved by SDS-PAGE, and Tarp phosphorylation was assessed by immunoblot using a monoclonal anti-phosphotyrosine antibody. Coomassie staining was used as a loading control.

Native Tarp dephosphorylation assay

HeLa cells (2 × 10⁵ cells per well in a 6-well plate seeded one day prior to infection) were infected with Chlamydia trachomatis serovar L2 at a multiplicity of infection of 100 (MOI = 100) and incubated at 37°C for 2 h. Following infection, media was removed, and cells were scraped and collected in Xenopus buffer (100 mM KCl, 2 mM MgCl₂, 10 mM HEPES, pH 7.7). Cell lysates were prepared by brief sonication. Lysates were then divided into two groups: one incubated with recombinant PTP1B and the other left untreated. Both reactions were carried out at 37°C for 1 h and terminated by the addition of 5× SDS sample buffer followed by heating at 95°C for 5 min. Samples were resolved by SDS-PAGE, and Tarp phosphorylation was assessed by immunoblotting using a monoclonal anti-phosphotyrosine antibody. Total Tarp levels were detected using a polyclonal anti-Tarp antibody.

Cell-free type III secretion system (T3SS) induction assay

Cell-free induction of type III secretion system was performed as previously described (54). C. trachomatis L2 elementary bodies (EBs), stored at −80°C in sucrose-phosphate-glutamate (SPG) buffer, were thawed on ice. EBs were pelleted by centrifugation at 13,000  RPM for 5 min at 4°C. Supernatants were discarded, and pellets were resuspended in 50  mM potassium acetate buffer (KAC, pH 4.8). EBs were pelleted again under the same conditions, and the supernatant was removed. Pellets were then resuspended in the appropriate experimental solution. Experimental solutions included PTP1B inhibitor (at the indicated concentrations), DMSO (vehicle control at the same concentration as inhibitor), or 3% fetal bovine serum (FBS) as a positive control for T3SS induction. Samples were incubated at 37°C for 30 min. After incubation, EBs were pelleted again at 13,000 RPM for 5 min at 4°C, and the supernatants were transferred to fresh tubes. Pellets were discarded. Supernatants were centrifuged once more at 13,000 RPM for 5 min at 4 °C to remove residual debris. Clarified supernatant was mixed with 5× SDS sample buffer to obtain samples. Samples were loaded onto SDS-PAGE gels for immunoblot analysis. Secreted chlamydial proteins were detected using antibodies against Tarp and MOMP.

Statistical analysis

All quantitative experiments were performed in at least three independent biological replicates, each with the indicated number of technical replicates. Data are presented as mean + standard deviation (SD), unless otherwise noted. Statistical comparisons between two groups were made using unpaired, two-tailed Student’s t-test. For comparisons involving more than two groups, one-way ANOVA followed by Tukey’s post hoc test or Dunnett’s multiple comparison test was applied. P-values less than 0.05 were considered statistically significant. Graphing and statistical analyses were conducted using GraphPad Prism version 10.

SUPPLEMENTAL MATERIAL

The following material is available online at https://doi.org/10.1128/iai.00373-25.

Fig. S1. iai.00373-25-s0001.docx.

Purification of recombinant PTP1B and T3SS induction in response to PTP1B inhibition.

Table S1. iai.00373-25-s0002.xlsx.

Cloning and SiRNA primers.

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