Introduction

All organisms that are exposed to oxygen are necessarily subjected to oxidative stress. Specifically, the process of metabolizing substrates in the presence of oxygen can generate reactive oxygen species (ROS), which are toxic at high enough concentrations. Thus, from bacteria to humans, systems have evolved to mitigate the accumulation of ROS. At the same time, host defense mechanisms have evolved to leverage ROS production as a means of limiting pathogen growth and survival. Not surprisingly, pathogens have co-evolved to resist these defense mechanisms. For example, many pathogens possess a variety of antioxidant enzymes, such as catalases, glutathione peroxidases, and peroxiredoxins, that help them both subvert ROS-mediated immune system assaults and mitigate metabolic ROS byproducts (Staerck et al., 2017; Wan et al., 2021).

Chlamydia is an obligate intracellular bacterium that has significantly reduced its genome size and content in adapting to obligate host dependence. Chlamydia trachomatis, the leading cause of bacterial sexually transmitted diseases and preventable infectious blindness, lacks homologs to catalases or glutathione peroxidases but does possess a homolog of Alkyl hydroperoxide reductase subunit C (AhpC). AhpC is a well-known member of the peroxiredoxins family and is widely conserved in prokaryotes (de Oliveira et al., 2021). Peroxiredoxins can scavenge hydrogen peroxide, peroxynitrite, and organic hydroperoxides (Parsonage et al., 2008; Poole & Ellis, 1996; Seaver & Imlay, 2001) and act as the primary scavenger in pathogens that lack both catalase and glutathione peroxidases (Mastronicola et al., 2014; Richard et al., 2011). Several studies from other bacterial systems have shown that AhpC has a significant role in ROS and RNI scavenging, virulence, and persistence (Cosgrove et al., 2007; Kimura et al., 2012; Oh & Jeon, 2014), and deletion of AhpC creates highly oxidized conditions within the bacterium (Zhang et al., 2019). However, the function of AhpC in chlamydial biology has not been characterized.

Chlamydia is unique among pathogens in undergoing a complex developmental cycle that comprises two distinct morphological forms, the elementary body (EB) and reticulate body (RB) (Abdelrahman & Belland, 2005). The EB is the smaller (∼0.3 μm), infectious, and non-dividing form capable of infecting susceptible host cells. Once internalized into a host-derived vacuole termed an inclusion, the EB differentiates into the non-infectious, larger (∼1 μm), and replicating RB - this process is known as primary differentiation and represents the early phase of the developmental cycle (Clifton et al., 2005). RBs replicate within the inclusion in the mid-cycle phase using an asymmetric, MreB-dependent polarized division process (Abdelrahman et al., 2016; Lee et al., 2020; Ouellette et al., 2022; Ouellette et al., 2020). In the late phase of the developmental cycle, RBs asynchronously condense into EBs, and this process is termed secondary differentiation. Despite these well-defined differences between chlamydial morphological forms, how Chlamydia mechanistically differentiates between functional forms remains unclear.

Intriguingly, a recent study evaluated the redox potential of Chlamydia and demonstrated that RBs are reduced, whereas EBs are oxidized (Wang et al., 2014). This is consistent with earlier studies that revealed differences in the crosslinking of outer membrane proteins and type III secretion-related proteins in chlamydial developmental forms (Betts-Hampikian & Fields, 2011; Caldwell et al., 1981; Everett & Hatch, 1995; Wang et al., 2014). Taken together, these observations indicate that, during the developmental cycle, the redox potential of the bacteria is changing. However, whether redox changes in the bacteria directly affect developmental cycle progression is not characterized. Based on the different redox status of the chlamydial developmental forms, we hypothesized that increasing the oxidation of the RB drives secondary differentiation to the EB. We used chlamydial transformants designed to overexpress or reduce AhpC levels to explore the effects of altered redox potential in chlamydial growth and development to test this hypothesis. This study establishes the role of AhpC as an antioxidant in Chlamydia, as demonstrated by its ability to counteract different peroxide stresses when overexpressed. Overexpression of AhpC had no negative effect on bacterial replication but delayed the differentiation of RBs to EBs. In contrast, under conditions of ahpC knockdown, the organism was highly sensitive to oxidizing conditions. Interestingly, this change in redox potential caused earlier expression of EB-associated (late) genes and production of EBs, leading to a shift in developmental cycle progression. This earlier activation of gene expression related to secondary differentiation in ahpC knockdown was also observed when developmental cycle progression was blocked by penicillin treatment. Taken together these data provide mechanistic insight into chlamydial secondary differentiation and are the first to demonstrate redox-regulated differentiation in Chlamydia.

Results

The redox threshold hypothesis and chlamydial secondary differentiation

The chlamydial EB and RB developmental forms differ in size, infectivity, and division capacity. In addition to these characteristics, both forms also differ in their redox status: EBs are oxidized and RBs are reduced (Fig. 1A). Based on this information, we hypothesized that changing redox potential is a critical factor in the process of differentiation from one form to the other. We named this the “redox threshold hypothesis”. In this scenario, as soon as a given RB has crossed an oxidative threshold, the activity of critical proteins is modified to trigger differentiation to the EB (Fig. 1B). AhpC is an antioxidant enzyme, and several reports have indicated that ablation of ahpC creates highly oxidized conditions in bacteria (Feng et al., 2020; Zhang et al., 2019). Therefore, we hypothesize that AhpC, which maintains redox homeostasis in other bacteria, is a critical factor in mediating developmental cycle progression and that altering its activity will impact developmental cycle progression consistent with the redox threshold hypothesis (Fig. 1C).

Fig. 1

Overexpression of ahpC has minimal impact on chlamydial growth

To study the effect of AhpC on chlamydial developmental cycle progression, we first generated an ahpC overexpression (OE) strain using a plasmid encoding an anhydrotetracycline (aTc)-inducible ahpC (untagged) and transformed it into a C. trachomatis L2 strain lacking its endogenous plasmid (-pL2). The same plasmid vector backbone (i.e., encoding mCherry in place of ahpC) was used as an empty vector control (EV). HeLa cells were infected with these transformants, and, at 10 hpi, expression of the construct was induced or not. Lacking an antibody against AhpC, overexpression of ahpC was validated by reverse transcription-quantitative PCR (RT-qPCR). Here, an approximate 1-log increase in transcripts of ahpC was detected at 14 and 24 hpi in the induced strain in comparison to the uninduced strain (Fig. 2A).

Fig. 2

Immunofluorescence analysis (IFA) was performed at 24 hpi to examine the organisms’ morphology and overall inclusion growth. For IFA, individual bacteria were labeled with an anti-ahpC increased the overall inclusion area (Fig. 2B and 2C). We next quantified the total number data support the hypothesis that overexpressing ahpC delays production of infectious EBs.

Overexpression of ahpC confers resistance to peroxides in Chlamydia

AhpC is an important peroxiredoxin involved in oxidative damage defense. Before determining whether increased ahpC expression impacts chlamydial sensitivity to oxidizing agents, we first sought to determine the response of C. trachomatis to inorganic or organic hydroperoxides such as hydrogen peroxide (H₂O₂), cumene hydroperoxide (CHP), and tert-butyl hydroperoxide (TBHP) as well as peroxynitrite (PN). To do this, we evaluated the sensitivity of HeLa cells, infected or not with the empty vector (EV) control strain, to different concentrations of oxidizing agents using a viability assay. Both uninfected and EV-infected HeLa cells tolerated up to 1 mM oxidizing agents for 30 min (Fig. S1). Next, different concentrations (lower than 1 mM) of these oxidants were tested on wild-type (WT) C. trachomatis L2/434/Bu (Ctr L2). HeLa cells were infected with Ctr L2 and, at 16 hpi, were exposed to different concentrations of exogenous oxidizing agents for 30 min only, before washing out the oxidizing agents and replacing the media. Samples for IFU and IFA assays were then collected at 24 hpi to assess effects of the oxidizing agents on growth of Ctr L2. 62.5 μM concentration of all three inorganic and organic peroxides had no appreciable effect on IFUs, and inclusion size and morphology also remained unaffected at this sublethal concentration. The concentrations of inorganic and organic peroxides that resulted in a decrease in IFUs to ∼50% of the untreated culture were 500 μM (H₂O₂) and 250 μM (CHP and TBHP), respectively, with a concurrent decrease in inclusion size. 1 mM H₂O₂, CHP, and TBHP caused >90% reduction in IFUs and small inclusions. PN at 1 mM concentration was less effective than other oxidizing agents and showed only a ∼30% decline in IFUs compared to the untreated culture (Fig. S2).

To further examine the antioxidant functions of AhpC, we next exploited our ahpC overexpression strain to explore its capability to protect chlamydiae from oxidizing agents. Both ahpC OE and EV strains were used to infect HeLa cells and, at 10 hpi, expression of the constructs was induced or not with 1 nM aTc. At 16 hpi, infected cells were exposed to various concentrations of exogenous oxidizing agents for 30 min. At 24 hpi, IFU and IFA samples were collected to measure chlamydial growth and assess chlamydial morphology, respectively. As shown in Figure 3A, 3B, and S3, the ahpC overexpression strain showed increased resistance to all oxidants tested as the mean IFUs and the size of inclusions were greater than that of the uninduced but treated control. There was no change in bacterial growth and morphology in the case of 62.5 μM concentrations of oxidizing agents, which was anticipated since we determined this was a sublethal concentration. The resistant phenotype was more evident at the higher concentrations of H₂O₂, CHP, and TBHP. Overexpression of ahpC also contributed to higher resistance against PN. Conversely, the empty vector control strain behaved as expected (i.e., like WT) in the presence of oxidizing agents, with similar results observed in the presence or absence of aTc (Fig. 3C, 3D, and S3). Collectively, these data demonstrate that the chlamydial AhpC possesses antioxidant activity, as expected, and that increased ahpC expression is protective for Chlamydia in the presence of increased oxidative stress.

Fig. 3

Knockdown of ahpC negatively impacts chlamydial growth

To further define the function(s) of AhpC in the chlamydial developmental cycle, we used a novel dCas12-based CRISPR interference (CRISPRi) strategy adapted for Chlamydia by our lab (Ouellette, 2018; Ouellette et al., 2021). We generated an ahpC knockdown strain harboring the pBOMBL12CRia with a crRNA targeting the ahpC 5’ intergenic region (plasmid designated as pL12CRia(ahpC)). We used a strain carrying a pL12CRia plasmid with a crRNA with no homology to any chlamydial sequence (i.e., non-targeting [NT]) to serve as a negative control. To confirm the knockdown of ahpC, RT-qPCR was employed. Here, we infected HeLa cells with the ahpC knockdown (ahpC KD) or NT strains and induced dCas12 expression or not at 10 hpi using 1 nM aTc. Nucleic acid samples were harvested at 14 hpi and 24 hpi. RT-qPCR analysis of the ahpC KD revealed approximately 90% reduction of ahpC transcripts compared to the uninduced control, thus confirming the knockdown of ahpC (Fig. 4A). In the case of the NT control, there was no effect on ahpC transcripts. IFA of these strains revealed noticeably smaller inclusions after blocking expression of ahpC as compared to the uninduced sample or the NT conditions (Fig. 4B). We next measured total bacterial counts using genomic DNA. Surprisingly, even though inclusions were smaller during ahpC knockdown, we observed slightly higher gDNA levels at 14 hpi compared to the uninduced control, followed by only a small increase from 14 to 24 hpi (Fig. 4C). In contrast, the uninduced strain showed a logarithmic increase in gDNA levels during this timeframe. These effects were not detected for the NT strain (Fig. 4C). To further explore the effect of reduced ahpC transcripts in Chlamydia, IFU assays were performed. IFU analysis exhibited severely reduced progeny (>90%) in ahpC knockdown compared to its respective uninduced control at both time points assessed (24 and 48 hpi) (Fig. 4D). In contrast, the NT strain showed less than a 50% reduction after inducing dCas12 expression at these timepoints consistent with prior observations that dCas12 expression slightly delays developmental cycle progression (Hatch & Ouellette, 2023; Reuter et al., 2023). Taken together, these analyses indicate that reducing ahpC levels and/or activity severely disrupted chlamydial growth and development.

Fig. 4

To investigate whether ahpC knockdown resulted in increased ROS levels in the bacteria (Zhang et al., 2019), the intracellular ROS levels were measured in infected cells in ahpC knockdown conditions and compared to the uninduced control condition and uninfected cells. We used the cell-permeable, fluorogenic dye CellROX Deep Red to measure ROS levels. This dye remains non-fluorescent in a reduced state and exhibits bright fluorescence once oxidized by ROS. ROS generation was measured in uninfected and ahpC KD-infected HeLa cells at 24 and 48 hpi. As shown in Figure 4E, ahpC knockdown resulted in significantly higher ROS than the uninfected or infected but uninduced samples at both 24 hpi and 48 hpi time points. These results together indicate that AhpC plays an important role in counteracting oxidative stress by reducing ROS in C. trachomatis.

Knockdown of ahpC sensitizes Chlamydia to oxidizing agents

As overexpression of ahpC resulted in increased resistance to oxidizing agents, we next explored whether knockdown of ahpC resulted in increased sensitivity to such agents. As described above, ahpC KD and NT strains were used to infect HeLa cells and, at 10 hpi, expression of dCas12 was induced or not with 1 nM aTc. Considering that the ahpC KD already demonstrated reduced IFUs and inclusion sizes at 24 hpi, only sublethal concentrations (62.5 μM) of exogenous oxidizing agents were applied. At 24 hpi, IFA and IFU analyses were performed to quantify the effect of the oxidants on chlamydial growth in the absence of ahpC activity. Under conditions of ahpC knockdown, even sublethal concentrations (62.5 μM) of H₂O₂, CHP, or TBHP further reduced the inclusion size, and the IFU data revealed a decrease from >90% to <30% in comparison to the untreated control (Fig. 5A, 5B, and S4). Reduced expression of ahpC also affected the survival of Chlamydia against peroxynitrite. Notably, there was no significant change in the NT control strain in the uninduced and induced samples in response to oxidizing agents as assessed by IFU and IFA (Fig. 5C, 5D, and S4). These data further support that the chlamydial AhpC is a critical antioxidant enzyme in these bacteria.

Fig. 5

Complementation restores the growth and resistance to low levels of peroxide stress of the ahpC knockdown strain

To validate that the impaired growth, altered inclusion morphology, and enhanced sensitivity to peroxides were due to the decreased level/activity of AhpC, we generated a complemented strain of ahpC knockdown. For the construction of the ahpC complementing plasmid, the ahpC gene was cloned and transcriptionally fused 3’ to the dCas12 in the pL12CRia(ahpC) knockdown plasmid. Here, the complementing ahpC allele is also under the control of the aTc-induced Ptet promoter and is co-expressed with the aTc-inducible dCas12. Consequently, the ahpC knockdown effect is ablated, and the observed phenotypes should be restored. After inducing dCas12-ahpC expression with aTc, the resultant strain was verified by RT-qPCR. Increased transcripts for ahpC were quantified under these conditions, indicating successful complementation of the knockdown effect (Fig. 6A). Of note, the ahpC transcript levels remained elevated in comparison to the uninduced control at the 24hpi time point. After confirming this strain, we measured genomic DNA to quantify total bacteria and performed IFA and IFU assays to examine if complementation restored the phenotypes observed during ahpC knockdown. These assays revealed normal inclusion morphology, gDNA levels, and EB progeny production in the complemented strain, indicating successful complementation of the knockdown phenotype (Fig. 6B, C, and D). Of note, a C-terminal 6xHis tagged AhpC was not capable of complementing the knockdown phenotype (data not shown), indicating a requirement for a free C-terminus in the function of AhpC in Chlamydia. Previous studies in other bacteria have revealed that the C-terminal residues in AhpC play a crucial role in the structural stability and enzymatic activity of AhpC (Dip et al., 2014; Feng et al., 2020; Wan et al., 2021).

Fig. 6

We next assessed whether complementation of the knockdown phenotype could also restore the resistance to low levels of peroxide stress. As in previous experiments, a sublethal concentration of oxidizing agents was added for 30’ at 16 hpi after having induced dCas12-ahpC expression at 10 hpi. Consistent with the growth parameters (Fig. 6B-D), the complemented strain showed wild-type responses in these conditions (Fig. 6E, 6F, and S4). These data indicate that the enhanced susceptibility of the ahpC knockdown strain to oxidizing agents was due to the reduced levels of ahpC and not an indirect effect of knockdown.

We predicted that the growth defects resulting from higher production of ROS during ahpC knockdown could be rescued by treating the ahpC KD with ROS scavengers. To test this prediction, we utilized two characterized ROS scavengers, DMTU (N,N’-dimethylthiourea) and α-tocopherol, which scavenge H₂O₂ and peroxyl radical, respectively (Hollander-Czytko et al., 2005; Kiffin et al., 2006; Walch et al., 2015). First, the effect of ROS scavengers on uninfected and infected HeLa cells was investigated using a viability assay. This assay revealed that 10 mM DMTU and 100 µM α-tocopherol had no adverse effects on uninfected or EV-infected HeLa cells (data not shown). These same concentrations were tested on WT Ctr L2 infected HeLa cells in untreated and 500 µM H₂O₂ treated conditions to test the potency of these scavengers to rescue growth defects associated with this concentration of oxidizing agent. Scavengers were added at 9.5 hpi and washed away at 16 hpi - the time of addition of H₂O₂ in the respective samples. At 16.5 hpi, after 3 wash steps, scavengers were added again with fresh media, and, at 24 hpi, bacterial growth and morphology were assessed using IFU and IFA assays. Neither scavenger had a significant impact on chlamydial morphology or infectious progeny. In the case of peroxide (500 µM H₂O₂) treated samples, scavengers restored IFUs from ∼50% to ∼100%, and inclusion size was also recovered (Fig. S5A and B).

Next, the effect of the ROS scavengers under ahpC knockdown conditions was assessed. In this experiment, scavengers were added at 9.5 hpi to provide the protective effect of scavengers before reducing the activity of ahpC. At 10 hpi, knockdown was induced or not with 1 nM aTc, and, at 14 hpi, scavengers were added again. At 24 hpi, IFU and IFA samples were collected to examine bacterial growth and morphology to assess the effect of ROS scavenging on the ahpC knockdown phenotype. As shown in Fig. 6G and 6H, both ROS scavengers had a positive impact on restoring growth of the ahpC knockdown strain; inclusions were larger, and IFUs were increased from <10% to >90% in the presence of scavengers in the induced samples. These data provide compelling evidence that the adverse effects of ahpC knockdown are due to increased ROS accumulation in this strain.

Chlamydial developmental cycle progression is altered by ahpC knockdown/ overexpression

Whereas the overexpression of AhpC delayed overall developmental progression and was consistent with our hypothesis, the results with the ahpC knockdown strain appeared to refute our hypothesis given the apparent decrease in IFUs and smaller inclusion sizes after knockdown. To investigate the effects of changes in redox potential on chlamydial developmental cycle progression, we performed a transcriptional analysis of well-characterized late-cycle genes associated with secondary differentiation (hctA, hctB, glgA, tsp, and omcB) (Belland et al., 2003; Gehre et al., 2016; Newhall, 1987; Ouellette et al., 2006; Swoboda et al., 2023). In the uninduced samples, transcript levels of all the tested late genes were higher at 24 hpi compared to 14 hpi, indicating their normal expression during the late stage of the developmental cycle. In comparison to the uninduced control, under conditions of ahpC knockdown, significantly higher expression of hctA, hctB, omcB, tsp, and glgA was observed at the mid-developmental cycle timepoint of 14 hpi (Fig. 7A). At 24 hpi, expression of these genes was comparable between the uninduced and induced conditions in the ahpC knockdown strain. In contrast, the complementation strain showed no increase in the expression of these tested late genes at 14 hpi, and tsp and glgA transcripts were reduced (Fig. 7C). Consistent with our model, the overexpression of ahpC resulted in significantly lower expression of hctA, hctB, omcB, tsp, and glgA at 14 and 24 hpi (Fig. 7D), suggesting a delayed transition to EBs.

Fig. 7

Given the increase in late gene expression during ahpC knockdown at an earlier time point in the developmental cycle than normal, we reasoned that the increase in oxidizing conditions might prematurely trigger secondary differentiation and EB production. However, complicating such an analysis is that fewer RBs are present to convert to EBs, which would result in overall lower IFU yields as we had measured at 24 and 48 hpi (Fig. 4D). Nonetheless, to assess EB production more rigorously at earlier time points in the developmental cycle, we performed a growth curve analysis to precisely measure IFUs at two-hour intervals from 16 to 24 hpi. HeLa cells were infected, and knockdown was induced or not at 10 hpi with 1 nM aTc. At the indicated time points, IFU samples were collected and quantified. Consistent with our prediction, this experiment revealed higher IFUs (i.e., EBs) at 16 and 18 hpi during ahpC knockdown compared to the uninduced samples (Fig. 7B). This difference was statistically significant at 18 hpi. However, further EB production was stalled, with the uninduced strain continuing to produce EBs such that, by 24 hpi, there were significantly higher EB yields under these conditions (as noted in Fig. 4D). These data show that the phenotypic consequence of higher expression at 14 hpi of genes functionally related to EBs is the concomitant earlier production of EBs. These data underscore that reduced activity of ahpC causes earlier secondary differentiation in C. trachomatis.

ahpC knockdown activates transcription of late-cycle genes when bacterial replication is blocked

We detected earlier expression of EB-related genes during ahpC knockdown and were curious if the ahpC knockdown condition could activate these genes under conditions when the chlamydial developmental cycle is blocked. Pathogenic Chlamydia species undergo a polarized cell division process, in which peptidoglycan is transiently synthesized only at the division septum (Abdelrahman et al., 2016; Cox et al., 2020; Ouellette et al., 2020). As a result, during penicillin treatment, division of RBs is blocked (Moulder, 1993; Moulder et al., 1956; Ouellette et al., 2020). However, the bacteria continue to grow in size resulting in aberrantly enlarged RBs (Barbour et al., 1982; Matsumoto & Manire, 1970; Ouellette et al., 2012), in which EB-related genes are not transcribed and production of EBs is inhibited (Ouellette et al., 2006; Panzetta et al., 2018).

To examine this, we generated an ahpC KD strain with spectinomycin resistance (ahpC KD-spec) and validated its phenotype as being the same as the penicillin-resistant ahpC KD strain. This new strain, ahpC KD-spec, was used to infect HeLa cells, and knockdown was induced or not at 10 hpi with 1 nM aTc. At the same time, samples were treated or not with 1 unit per mL of penicillin (Pen). RNA samples were harvested at 16 and 24 hpi and processed for RT-qPCR (Fig. S6B-D). IFA controls from these different conditions demonstrated the expected phenotypes (Fig. S6A). For example, Pen treatment caused aberrantly enlarged RBs, irrespective of ahpC KD, whereas ahpC KD itself caused smaller inclusions. Similarly, ahpC KD resulted in reduced ahpC transcripts in the induced conditions irrespective of the presence of Pen (Fig. S6B). We next quantified transcript levels for the late genes hctA and hctB in these different conditions (Fig. S6C and D). As expected, in the uninduced and untreated condition, both transcripts had higher expression at 24 hpi, indicating their regular developmental expression during the late stage of the developmental cycle. Consistent with prior observations (Ouellette et al., 2006), transcript levels of these genes in uninduced+Pen conditions was low. As we previously noted (Fig. 7A), the transcripts of hctA and hctB are higher in the induced than uninduced samples at 16 hpi in the absence of Pen. Interestingly, in the induced+ Pen condition, these genes had higher expression at 24 hpi than the uninduced+Pen control. At 16 hpi, their expression was either higher or similar between both samples (i.e., induced+Pen and uninduced+Pen). Collectively, these data support our observation that ahpC knockdown activates the transcription of late-cycle genes – even under conditions where developmental cycle progression is blocked.

Discussion

Secondary differentiation (differentiation from RBs to EBs) is an essential step for chlamydial growth and survival, but there is a dearth of information regarding the mechanisms of its regulation. EBs and RBs have significantly different proteomic repertoires, and our group has identified critical functions for the cytoplasmic ClpXP and ClpCP and periplasmic Tsp proteases during secondary differentiation (Pan, Jensen et al., 2023; Swoboda et al., 2023; Wood et al., 2022). The tsp gene, in addition to another late-cycle gene, hctB, is transcriptionally regulated by sigma factor 28 (σ²⁸) (Hatch & Ouellette, 2023). Another sigma factor, σ⁵⁴ has been linked to regulation of outer membrane components, type III secretion system components, and other genes typically expressed late in development (Hatch & Ouellette, 2023; Soules, LaBrie, et al., 2020) . In addition to the two minor sigma factors, σ²⁸ and σ⁵⁴, Chlamydia encodes one major sigma factor, σ⁶⁶, which is transcriptionally regulated by the relative levels of RsbV1 (antagonist) and RsbW (anti-sigma factor) (Thompson et al., 2015). This Rsb system senses ATP availability and has been proposed to regulate σ⁶⁶ (Thompson et al., 2015). A related study from Soules et al. (Soules, Dmitriev, et al., 2020) showed that TCA intermediates act as ligands for RsbU, thereby linking the Rsb system to the TCA cycle and ATP synthesis. Collectively, these data show that post-translational mechanisms drive secondary differentiation in Chlamydia. However, no definitive “switch” that triggers this step has been identified.

Aside from the clear morphological and functional differences between the EB and RB, they also differ in their redox status: EBs are more oxidized whereas RBs are more reduced (Wang et al., 2014). This same study noted that the overall redox status of the host cell was relatively unchanged throughout the developmental cycle, suggesting that changes in the bacteria were driven by endogenous processes (i.e., metabolism). Though Chlamydia is dependent on its host for most of its energy requirements, it has some metabolic activities, such as a partial TCA cycle and oxidative phosphorylation (Gerard et al., 2002; Iliffe-Lee & McClarty, 1999) that can be possible sources of intracellular ROS. We hypothesize that accumulating redox stress from its metabolic activities serves as a signal to trigger secondary differentiation from RBs to EBs. There is a paucity of knowledge on how C. trachomatis modulates oxidative stress during disease pathogenesis. However, some studies explored the effects of redox changes on Chlamydia growth. Boncompain et al. reported that infection of C. trachomatis induced the transient production of ROS by the host cell at a moderate level for the initial few hours of infection only (Boncompain et al., 2010). Another study also found a similar observation about ROS during infection, reporting that Chlamydia requires host-derived ROS for its growth (Abdul-Sater et al., 2010). Given that no studies have examined the role of chlamydial proteins related to redox and how these proteins impact the growth of the pathogen, we initiated experiments to investigate the function of a key antioxidant enzyme, AhpC, as a means of testing our hypothesis.

Reactive oxygen species (ROS) generation is an inevitable condition for pathogens growing in aerobic conditions, and resistance against oxidative stress (imbalanced level of ROS) is a key survival mechanism (Fang, 2011). Hence, pathogens have evolved detoxifying proteins, such as peroxiredoxins, to eliminate ROS (Dip et al., 2014; Wang et al., 2004; Yang et al., 2002). AhpC is a 2-cys peroxiredoxin and has been reported to have a crucial role in bacterial physiology, survival, and virulence by scavenging ROS and RNI (Cosgrove et al., 2007; Kimura et al., 2012; Loprasert et al., 2003; Oh & Jeon, 2014). In the process of detoxification of peroxides and peroxynitrite, AhpC is converted into an oxidized dimer, requiring alkyl hydroperoxide reductase AhpF or AhpD to regenerate its activity (Koshkin et al., 2004; Poole & Ellis, 1996; Wong et al., 2017). C. trachomatis encodes AhpC (Ct603) but lacks any annotated homologs of AhpF or AhpD. Therefore, it remains an open question how AhpC activity is regulated. Some studies have mentioned ahpC as an iron-responsive gene in Chlamydia (Brinkworth et al., 2018; Pokorzynski et al., 2019), but there is no detailed investigation to date about the role of this crucial antioxidant in the growth and development of Chlamydia.

Using a broad set of methodologies, we created and characterized overexpression and knockdown conditions of ahpC. Higher expression of ahpC resulted in lower EB production at 24 hpi coupled with a higher number of RBs and larger inclusion size (Fig. 2), suggesting differentiation from RBs to EBs is delayed. The number of EBs is further decreased at 48 hpi, but the difference is not statistically significant. This may be due to the highly oxidized conditions in Chlamydia at 48 hpi (Wang et al., 2014) or the inability of ahpC overexpression to generate sufficient reducing conditions to neutralize it completely. In our studies, ahpC expression was induced at 10hpi, and adequate enzyme levels may not have been sustained later during the developmental cycle. Consistent with studies in other bacteria (Sherman et al., 1999; Zuo et al., 2014), overexpression of ahpC created resistance to different oxidants (Fig. 3 and S3), indicating that AhpC is the principal defense mechanism against oxidative stress in Chlamydia. During ahpC knockdown, the attenuated activity of AhpC severely affected the growth of bacteria (Fig. 4). Notably, the morphology of inclusions during ahpC knockdown is strikingly similar to Chlamydia treated with a high concentration (1 mM) of oxidizing agents (Fig. 4 and Fig S2), indicating similar scenarios in both conditions. This was further supported by the increased sensitivity of ahpC knockdown to sublethal concentrations of oxidants (Fig. 5 and S4). Again, this observation is consistent with previous data from other bacteria (Cosgrove et al., 2007; Seaver & Imlay, 2001; Storz et al., 1989; Zhang et al., 2019).

An essential aspect of the scavenging activity of AhpC for many bacterial pathogens in which it has been studied is that it works best with endogenous (i.e., low) levels of H₂O₂. These bacteria, such as Staphylococcus aureus and Yersinia pseudotuberculosis, encode catalases, which have high Km for hydrogen peroxide and serve a predominant role in scavenging exogenous H₂O₂ (Cosgrove et al., 2007; Wan et al., 2021). Catalases can detoxify H₂O₂ at high levels (millimolar levels) and are crucial in responding to external H₂O₂ stress (Mishra & Imlay, 2012). The ahpC mutants in these bacteria became sensitive to organic peroxides but resistant to H₂O₂ due to higher catalase activity as a compensatory response to the lack of AhpC (Antelmann et al., 1996; Mongkolsuk et al., 2000; Ochsner et al., 2000). Further, simultaneous mutations in both catalase and ahpC displayed drastically enhanced sensitivity to all oxidizing agents tested (Cosgrove et al., 2007; Ezraty et al., 2017; Seaver & Imlay, 2001). C. trachomatis does not encode a catalase gene (Boncompain et al., 2014; Rusconi & Greub, 2013), and hypersensitivity of ahpC knockdown to both inorganic and organic peroxides indicates the absence of catalase and establishes that AhpC is the primary scavenger of ROS in C. trachomatis. We did not observe significant effects of PN with AhpC expression. One possible explanation may be our study’s concentration (1 mM) of PN. Higher concentrations of PN could not be used due to toxic effects on the host cell. The other possibility may be the presence of some other unknown mechanism(s) to detoxify PN. For example, Chlamydia also encodes a superoxide dismutase (SOD) that may prevent the accumulation of the necessary precursors that are needed for PN production. We are currently investigating the function of the chlamydial SOD enzyme.

Previous studies in other bacterial systems indicated that ROS production significantly increased after the deletion of ahpC (Zhang et al., 2019). Similarly, in Chlamydia during ahpC knockdown, the ROS level was dramatically higher. Its ability to dissipate ROS produced by exogenous oxidative stress was severely compromised (Fig. 4E). Further, the addition of ROS scavengers to the culture medium rescued the negative phenotypes associated with ahpC knockdown (Fig. 6G and H). This strongly suggests that, in the absence of AhpC, higher amounts of reactive oxygen species accumulate in Chlamydia and that the negative impact on growth during ahpC knockdown is due to highly oxidized conditions in the organism. Moreover, for the complete restoration of inclusion size, multiple doses of scavengers were required throughout the experiment, further emphasizing that the internal chlamydial environment is oxidized and/or susceptible to oxidation in the absence of AhpC. These findings suggest that AhpC in Chlamydia is indispensable, having a functional role even more extensive than previously reported in other bacteria.

Interestingly, in our RT-qPCR study, late-cycle genes such as hctA, hctB, glgA, omcB, and tsp are expressed at a higher level at an earlier time (14 hpi) in the chlamydial developmental cycle as a result of reduced AhpC activity (Fig. 7). Notably, in the developmental cycle of C. trachomatis, only RBs are present at 14 hpi, and secondary differentiation starts after 16 hpi (Abdelrahman & Belland, 2005). Expression of most of the late-cycle genes occurs after this time in the developmental cycle (Belland et al., 2003). Among the tested late-cycle genes, hctA and hctB encode histone-like proteins responsible for chromosomal condensation during the differentiation of RBs into EBs. The hctA gene is among the first to be transcribed in the late-stage (Chiarelli et al., 2020). The other three late genes, tsp, omcB, and glgA, are all well-characterized late genes associated with secondary differentiation. Tsp is a periplasmic protease thought to be crucial in the degradation of RB-specific periplasmic proteins during secondary differentiation in C. trachomatis (Swoboda et al., 2023). OmcB is responsible for the rigid cell wall and osmotic stability of the EBs (Mygind et al., 1998; Newhall, 1987). GlgA is a non-essential secretory protein involved in glycogen metabolism and is responsible for glycogen accumulation in the inclusion lumen at late stages of the developmental cycle (Gehre et al., 2016; Sun et al., 2020). One crucial point to consider is the redox status of Chlamydia. 14 hpi is when reduced RBs predominate with virtually no oxidized EBs detectable. However, we established that ahpC knockdown shifts the redox status of the bacteria towards oxidation. Hence, the earlier and significantly higher detection of transcripts for these late-cycle genes indicate earlier secondary differentiation in the ahpC knockdown. Importantly, ahpC complementation during knockdown restored the regular developmental expression of these late-cycle genes, further supporting this phenotype was due to the highly oxidized conditions created by reduced AhpC activity.

The higher expression of genes functionally related to EBs and EB production at an early stage of the chlamydial developmental cycle indicates earlier secondary differentiation as an outcome of diminished activity of ahpC in C. trachomatis. Consequently, increased late gene transcription should have the phenotypic effect of causing earlier production of EBs. Indeed, we quantified more IFUs (proxy for EBs) in the ahpC knockdown at 16 and 18 hpi compared to the uninduced control condition. In contrast, for the AhpC overexpression strain, the expression of these late-cycle genes in the induced conditions compared to the uninduced control at 24 hpi is significantly lower, indicating reduced production of EBs. These data support our hypothesis that the developmental cycle is delayed in the reduced environment. We propose a simple model to explain these scenarios (Fig. 8). During ahpC KD, higher amounts of ROS accumulation in the bacterium create highly oxidized conditions. As secondary differentiation is asynchronous and RBs divide through an asymmetric budding mechanism (Ouellette et al., 2020), ROS levels are unevenly distributed between mother and daughter cell. This difference will lead some RBs to breach an oxidative threshold sooner, allowing activation of late genes and secondary differentiation earlier than other RBs. In contrast, ahpC overexpression scavenges ROS at a greater level leading to a more reducing environment. This results in a delay in achieving the oxidative threshold, thus allowing RBs to continue to divide before committing to secondary differentiation. Taken together, our data directly link oxidation state and secondary differentiation in Chlamydia. Ongoing studies are focused on characterizing redox-sensitive chlamydial proteins to understand which specific factors drive the shift from RBs to EBs (or EBs to RBs) in C. trachomatis.

Fig. 8

Materials and methods

Strains and cell culture

For chlamydial transformation, McCoy mouse fibroblast cells (kind gift of Dr. Harlan Caldwell (NIH/NIAID)) were used. Human cervix adenocarcinoma epithelial HeLa cells (kind gift of Dr. Harlan Caldwell (NIH/NIAID)) were used for RT-qPCR, immunofluorescence assays (IFA), inclusion forming unit assays (IFU), viability assays, oxidative stress, ROS measurements, and ROS scavenger assays. Both cell types were routinely grown and passaged in Dulbecco’s modified Eagle’s medium (DMEM; Gibco, Waltham, MA) supplemented with 10% fetal bovine serum (FBS; Sigma, St. Louis, MO) and 10 µg/mL gentamicin (Gibco, Waltham, MA) at 37⁰C and 5% CO2. All strains were verified to be Mycoplasma-negative using LookOut mycoplasma PCR detection kit (Sigma). For chlamydial transformations, Chlamydia trachomatis serovar L2 EBs lacking the endogenous pL2 plasmid (kind gift of Dr. Ian Clarke, University of Southampton) were used (Wang et al., 2011). Wild-type, density gradient-purified Chlamydia trachomatis 434/Bu (ATCC VR902B) EBs were used for sensitivity to oxidizing agents and ROS scavenger assays. Molecular biology reagents, oxidizing agents, scavengers, and CellROX Deep Red dye were purchased from Thermo Fisher unless otherwise noted.

Plasmid construction

The primers, gBlock gene fragments, plasmids, and bacterial strains used for molecular cloning are listed in Table S1 in the supplemental material. Constructs for chlamydial transformation were cloned using high-fidelity (HiFi) cloning system from New England BioLabs (NEB). Primers were designed using the NEBuilder online primer generation tool (https://nebuilderv1.neb.com). For the overexpression strain, the ahpC gene was amplified by PCR with Phusion DNA polymerase (NEB) using C. trachomatis serovar L2 434/Bu genomic DNA as a template. The PCR product was purified using a PCR purification kit (Qiagen, Hilden, Germany). The HiFi assembly reaction was performed as per the manufacturer’s instructions in conjunction with the pBOMBDC plasmid digested with Fast Digest EagI and KpnI enzymes and dephosphorylated with FastAP (Thermo Fisher, Waltham, MA). The HiFi reaction mix was transformed into E. coli 10β (NEB). Plasmids were first confirmed by restriction enzyme digestion, and final verification of insert was performed using Sanger sequencing. A dCas12-based CRISPRi approach was used for ahpC knockdown generation (Ouellette et al., 2021). The pBOMBL12CRia plasmid having anhydrotetracycline (aTc) inducible catalytically dead dCas12 protein was used as vector. A crRNA targeting the 5’ intergenic region of ahpC was designed and ordered as a presynthesized DNA fragment. For ahpC knockdown construct, 2 ng of the gBlock (Integrated DNA Technologies [IDT], Coralville, IA) listed in Table S1 was combined with 25 ng of BamHI-digested, alkaline phosphatase-treated pBOMBL12CRia(e.v.)::L2 in a HiFi reaction according to the manufacturer’s instructions (NEB). The plasmid was transformed into NEB 10β cells and verified by Sanger sequencing prior to transformation into Chlamydia trachomatis. To generate the complementation strain, the ahpC gene was amplified using primers listed in Table S1 and fused with the ahpC knockdown construct (i.e., pBOMBL12CRia (ahpC)) digested with the Fast Digest restriction enzyme SalI (Thermo Fisher) and alkaline phosphatase-treated, using the HiFi reaction as mentioned above.

Chlamydial transformation

Chlamydial transformations were performed using a protocol described previously, with some modifications (Mueller et al., 2017). One day before transformation, 1 x 10⁶ McCoy cells were seeded in one 6-well plate, and two wells were used per plasmid transformation. Briefly, for each well of the 6-well plate, 2 µg of sequenced verified plasmids were incubated with 2.5 x 10⁶ C. trachomatis serovar L2 without plasmid (-pL2) EBs in 50 µL Tris-CaCl₂ (10 mM Tris, 50 mM CaCl2, pH 7.4) at room temperature for 30 min. McCoy cells were washed with 2 mL Hank’s Balanced Salt Solution (HBSS; Gibco), and 1 mL HBSS was added back into each well. 1 mL of HBSS was added to each transformant mixture, and one well of a 6-well plate was infected using this transformation solution. Cells were centrifuged at 400 × g for 15 min at room temperature followed by 15 min incubation at 37°C. HBSS was aspirated and replaced with antibiotic-free DMEM. At 8 hpi, 1 µg/mL of cycloheximide and 1 or 2 U/mL of penicillin G or 500 μg/mL spectinomycin were added to the culture media. The infection was passaged every 48 h until a population of penicillin or spectinomycin resistant, green fluorescent protein (GFP) positive C. trachomatis was established. The chlamydial transformants were then serially diluted to isolate clonal populations. These isolated populations were further expanded and frozen at - 80⁰C in a sucrose phosphate solution (2SP). To verify plasmid sequences, DNA was harvested from infected cultures using the DNeasy kit (Qiagen) and transformed into NEB 10β for plasmid propagation. Isolated plasmids were then verified by restriction digest and Sanger sequencing.

Inclusion forming unit assay

Inclusion forming unit assay was performed to determine the infectious progeny (number of EBs) from a primary infection based on inclusions formed in a secondary infection. C. trachomatis transformants were infected into HeLa cells and induced or not with 1 nM aTc at 10 hpi. At 24 and 48 hpi, samples were harvested by scraping three wells of a 24-well plate in 2 sucrose-phosphate (2SP) solution and lysed via a single freeze-thaw cycle, serially diluted, and used to infect a fresh HeLa cell monolayer and allowed to grow for 24 h. Samples were fixed with methanol, stained with a goat antibody specific to C. trachomatis major outer membrane protein (MOMP; Meridian Biosciences, Memphis TN) followed by staining with donkey antigoat Alexa Fluor 594-conjugated secondary antibody (Invitrogen) and titers were enumerated using a 20x lens objective. All experiments were performed three times for three biological replicates. Induced values were expressed as a percentage of the uninduced values, which was considered as 100%.

Immunofluorescence assay

HeLa cells were cultured on glass coverslips in 24-well tissue cultures plates at 2 x 10⁵ cells/well, infected with the relevant strains, and, at 10 hpi, samples were induced or not with 1 nM aTc. At 24 hpi, samples were fixed and permeabilized using 100% methanol. Organisms were stained with anti-MOMP (Meridian Biosciences) primary antibody for all the strains and primary mouse anti-Cpf1 (dCas12) (Sigma-Millipore) for knockdown or complementation samples. Donkey anti-goat Alexa Fluor 594-conjugated secondary antibody (Invitrogen) was used to visualize Chlamydia in all the samples, and donkey anti-mouse Alexa Fluor 488-conjugated secondary antibody (Invitrogen) was used for dCas12 expression. DAPI (Invitrogen) was used for visualization of host and bacterial cell DNA. These stained coverslips were mounted on glass slides using ProLong glass antifade mounting media (Invitrogen) and imaged using a 100x lens objective on a Zeiss Axioimager Z.2 equipped with Apotome.2 optical sectioning hardware and X-Cite Series 120PC illumination lamp using a 2MP Axiocam 506 monochrome camera.

Nucleic acid extraction and RT-qPCR

HeLa cells were seeded in 6-well tissue culture plates, infected with the C. trachomatis transformants, and induced or not at 10 hpi with 1 nM aTc. For each condition, triplicate wells were used for simultaneous harvest of RNA (for transcript analysis by RT-qPCR), gDNA (to normalize RT-qPCR data and quantification of gDNA), and IFA (to verify the morphological changes in samples in the tested conditions). For RNA extraction, cells were rinsed with DPBS twice and lysed with 1 mL TRIzol (Invitrogen/Thermo Fisher) per well as per manufacturer’s instructions. 200 μL of chloroform was added to extract the aqueous layer containing total RNA, which was precipitated with isopropanol. A total of 10 µg of purified RNA was treated with TURBO DNase (Invitrogen/Thermo Fisher) according to the manufacturer’s instructions to remove DNA contamination. DNA-free RNA was used for cDNA synthesis using random nonamers (N9; NEB) and SuperScript III reverse transcriptase (Invitrogen/Thermo Fisher) following the manufacturer’s instructions. cDNA samples were diluted 10-fold with molecular biology-grade water and stored at -80°C. A total of 2.5 µL of each diluted cDNA sample was used per well of a 96-well qPCR plate. For each of three biological replicates, each sample was analyzed in triplicate on a QuantStudio 3 system (Applied Biosystems/Thermo Fisher) using the standard amplification cycle with melting curve analysis. For gDNA, one well of a 6-well plate per condition was scraped in 500 μL DPBS, split in half (i.e., 250 μL), and frozen at -80°C. Each sample was then thawed and frozen twice more for a total of three freeze/thaw cycles, and gDNA was extracted using the DNeasy DNA extraction kit (Qiagen) according to the manufacturer’s guidelines. The isolated gDNA was quantified and diluted down to 5 ng/mL prior to use in quantitative PCR (qPCR). A total of 2.5 µL of each diluted gDNA sample was mixed with 10 µL of PowerUp SYBR green master mix in a 96-well qPCR plate and was analyzed on a QuantStudio 3 system (Applied Biosystems). Each sample from each biological replicate was tested in triplicate. For each primer set used, a standard curve of gDNA was generated against purified C. trachomatis L2 genomic DNA, and the cDNA levels were normalized to gDNA levels or 16S rRNA for analysis. All experiments were performed three times for three biological replicates. At the same time, morphological differences were monitored in the IFA control, with samples fixed with methanol at the time of harvesting of RNA/gDNA. Staining and imaging were performed as described above.

Viability assay

Cell viability assays were performed using PrestoBlue (Invitrogen/Thermo Fisher). HeLa cells were seeded in 96 well plates and were infected or not with the indicated chlamydial strains (e.g., pBOMBDC.ev). Samples were treated or not as per the experimental parameters (e.g., different concentrations of oxidizing agents or scavengers). Medium without cells was used as the blank control. At the end point, 10% PrestoBlue (v/v) was added in the wells and incubated at 37⁰ C for 30 min while protecting the plate from light. Fluorescence was read at excitation 560 nm and emission 590 nm using an Infinite M200 Pro (Tecan). The treated values were expressed as a percentage of the untreated values, which was considered as 100%. All experiments were performed three times for three biological replicates.

Oxidizing agents’ susceptibility testing

Susceptibility of host cells and chlamydiae to different oxidizing agents was tested in HeLa cells infected with density gradient-purified Chlamydia trachomatis 434/Bu EBs or C. trachomatis transformants. For C. trachomatis transformants, expression of the construct was induced or not with 1 nM aTc at 10 hpi. At 16 hpi, different concentrations of oxidizing agents were added, and samples were incubated at 37⁰C for 30 min. These samples were washed three times with HBSS, fresh DMEM media was added in the wells, and cultures were allowed to grow until 24 hpi. At 24 hpi, inclusion forming unit assays and immunofluorescence analysis were performed as described above.

ROS scavenging by chemical compounds

The scavenging capacity of different ROS scavengers was tested using HeLa cells infected with Chlamydia trachomatis 434/Bu EBs or the ahpC KD strain. To examine the protective effect of scavengers in the absence or presence of oxidative stress, Chlamydia trachomatis 434/Bu EBs infected HeLa cells were used. In these samples, scavengers were added or not at 10 hpi in respective wells. At 16 hpi, wells were washed three times with HBSS, and fresh DMEM containing 500 μM H₂O₂ or not was added. After 30 min, wells were washed three times with HBSS and fresh DMEM containing scavengers or not was added back in the respective wells, which were incubated until 24 hpi prior to collecting samples for IFU assay and IFA analysis. The effect of these scavengers was further tested in ahpC KD. HeLa cells were infected with ahpC KD, scavengers were added or not at 9.5 hpi before induction at 10 hpi with 1nM aTc. At 14 hpi, scavengers were added again in the respective wells, and, at 24 hpi, samples were collected for IFU assay and IFA.

ROS detection by CellROX Deep Red

Intracellular ROS levels were measured in uninfected and ahpC KD-infected HeLa cells using CellROX Deep Red dye (Invitrogen/Thermo Fisher). Samples were induced or not at 10 hpi with 1 nM aTc. Medium without cells was used as the blank control. At 16 hpi, samples were treated or not with 62.5 μM or 1 mM TBHP for 30 min. At the end point, media was removed, samples were washed three times with DPBS and incubated with CellROX Deep Red dye for 30 min in the dark at 37⁰C. Fluorescence was read at excitation 644 nm and emission 665 nm using an Infinite M200 Pro (Tecan). All experiments were performed three times for three biological replicates.

Acknowledgements

We thank Dr. H. Caldwell (NIH/NIAID) for providing eukaryotic cell lines and Dr. I. Clarke (University of Southampton) for providing the plasmid less strain of C. trachomatis serovar L2. We thank Dr. Elizabeth A. Rucks for critical feedback. We thank the members of the Rucks/Ouellette research group for thoughtful discussion of the material presented. Funding for this work was provided by the National Institutes of Health (NIH/NIAID) grants R01AI170688 and R21AI178150 to SPO.

Supplementary Information

Table S1 List of plasmids, strains, and primers used in this study.

Fig. S1 Viability assay of uninfected or infected HeLa cells treated with oxidizing agents H₂O₂ (A), CHP (B), TBHP (C), and PN (D). HeLa cells were infected or not with empty vector control, pBOMBDC.ev (EV), and treated or not with different concentrations of oxidizing agents at 16 hpi for 30 min. At 24 hpi, an end point viability assay was performed using PrestoBlue as mentioned in materials and methods. The treated values were expressed as a percentage of the untreated values, which were considered as 100%. Data represent three biological replicates.

Fig. S2 Response of C. trachomatis L2 against oxidizing agents. (A) IFA of wild-type Ctr L2 exposed to oxidizing agents. Oxidizing agents’ treatment, staining, and imaging were performed as mentioned in materials and methods. Representative images from three biological replicates are shown. (B) IFU analysis of Ctr L2 post-exposure with oxidizing agents. Conditions were same as in section (A), and IFUs were harvested at 24 hpi. IFUs were calculated as percentage of untreated samples. ***p< 0.0001 vs untreated sample by using one way ANOVA. Data represent three biological replicates.

Fig. S3 Overexpression of ahpC provides resistance to peroxides in Chlamydia. IFA of ahpC (A) or EV (B) exposed to oxidizing agents (CHP, TBHP, and PN). Experiments were performed as mentioned in the legend of Fig. 3A. Representative images from three biological replicates are shown.

Fig. S4 Chlamydia is hypersensitive to oxidizing agents as a result of reduced levels of ahpC. IFA of ahpC KD (A), NT (B), comp (C) treated with CHP, TBHP, and PN. Experimental conditions were the same as mentioned in the legend of Fig. 5A. Representative images from three biological replicates are shown.

Fig. S5 Rescue of oxidative stress phenotype by ROS scavengers in C. trachomatis L2. (A) IFA of wild-type Ctr L2 incubated or not with scavengers, α-Tocopherol (100 µM) and DMTU (10 mM), and treated or not with 500 µM H₂O₂ as mentioned in materials and methods. Representative images from three biological replicates are shown. (B) IFU analysis of Ctr L2 grown under the same conditions as mentioned in the legend of Fig. 6G. IFUs were calculated as percentage of untreated samples. ***p< 0.0001, statistical analysis was performed using ordinary one-way ANOVA. H₂O₂ treated sample and samples incubated with scavengers only were compared to the untreated control. Samples treated with H₂O₂ and incubated with scavengers were compared with sample treated with H₂O₂. Data represent three biological replicates. Only significant differences are noted.

Fig. S6 Effect of penicillin treatment during ahpC knockdown. (A) IFA was performed to assess inclusion size and morphology of ahpC KD-spec following induction (1 nM aTc) and penicillin treatment (1 U/mL) at 10 hpi. At 24 hpi, cells were fixed with methanol and stained using primary antibodies to major outer membrane protein (MOMP), Cpf1 (dCas12), and DAPI. All images were acquired on Zeiss Axio Imager Z.2 with Apotome2 at 100x magnification. Bars, 2 μm. Representative images of three biological replicates are shown. Transcriptional analysis of (B) ahpC, (C) hctA, and (D) hctB in ahpC KD-spec using RT-qPCR using the same conditions as in section (A). RNA samples were harvested at 16 and 24 hpi. Quantified cDNA was normalized to 16SrRNA, and values were plotted on a log scale. ***p< 0.0001, **p< 0.001, *p< 0.01 vs uninduced sample by using two-way ANOVA. Data represent three biological replicates.