The aryl hydrocarbon receptor and FOS mediate cytotoxicity induced by Acinetobacter baumannii

Kew, Chun; Prieto-Garcia, Cristian; Bhattacharya, Anshu; Tietgen, Manuela; MacNair, Craig R.; Carfrae, Lindsey A.; Mello-Vieira, João; Klatt, Stephan; Cheng, Yi-Lin; Rathore, Rajeshwari; Gradhand, Elise; Fleming, Ingrid; Tan, Man-Wah; Göttig, Stephan; Kempf, Volkhard A. J.; Dikic, Ivan

doi:10.1038/s41467-024-52118-7

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Published: 11 September 2024

The aryl hydrocarbon receptor and FOS mediate cytotoxicity induced by Acinetobacter baumannii

Nature Communications volume 15, Article number: 7939 (2024) Cite this article

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Abstract

Acinetobacter baumannii is a pathogenic and multidrug-resistant Gram-negative bacterium that causes severe nosocomial infections. To better understand the mechanism of pathogenesis, we compare the proteomes of uninfected and infected human cells, revealing that transcription factor FOS is the host protein most strongly induced by A. baumannii infection. Pharmacological inhibition of FOS reduces the cytotoxicity of A. baumannii in cell-based models, and similar results are also observed in a mouse infection model. A. baumannii outer membrane vesicles (OMVs) are shown to activate the aryl hydrocarbon receptor (AHR) of host cells by inducing the host enzyme tryptophan-2,3-dioxygenase (TDO), producing the ligand kynurenine, which binds AHR. Following ligand binding, AHR is a direct transcriptional activator of the FOS gene. We propose that A. baumannii infection impacts the host tryptophan metabolism and promotes AHR- and FOS-mediated cytotoxicity of infected cells.

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Introduction

Acinetobacter baumannii is a Gram-negative bacterium that poses an increasing threat to public health based on its unusual ability to acquire multiple antibiotic resistance determinants. The bacterium is extremely versatile and can infect different parts of the human body, causing diseases such as pneumonia, meningitis, wound infections, and sepsis^1,2. It is currently unclear how A. baumannii induces cytotoxicity in host cells. Unlike other pathogenic bacteria, A. baumannii does not rely on classic virulence effectors such as specialized cytotoxins and is often said to have a “persist and resist” pathogenic strategy³. A. baumannii shows an unusual degree of antibiotic resistance, and even pan-drug-resistant strains have been identified⁴. Infection with A. baumannii may, therefore, become untreatable with antibiotics in the future², and the World Health Organization (WHO) considers A. baumannii a “critical priority pathogen”⁵, thus highlighting the urgent need for alternative therapies.

One such alternative approach is host-directed therapy⁶. Rather than targeting bacterial factors directly using antibiotics, host factors are modulated to overcome microbial toxicity or enhance pathogen clearance by augmenting the immune response^6,7. Unlike antibiotics, host-directed interventions impose minimal selective pressure on pathogens, reducing the emergence of resistant strains⁶. Host-directed therapies against the tuberculosis pathogen Mycobacterium tuberculosis have been evaluated successfully in preclinical studies and have entered clinical development⁸. Examples include the modulation of inflammation and the activation of intracellular antimicrobial defenses⁹.

Host-directed therapy requires a clear understanding of the mechanisms underlying pathogen–host interactions. A. baumannii and other Gram-negative bacteria can interact with host cells via the secretion of nanoscale proteoliposomes known as outer membrane vesicles (OMVs) that bud from the bacterial outer membrane¹⁰. These structures carry diverse cargos and are necessary for pathogenesis in many bacteria. For example, Porphyromonas gingivalis^11,12,13 and Escherichia coli^14,15 utilize OMVs to transfer the virulence factors gingipain and hemolysin, respectively, into host cells. OMVs from A. baumannii cause mitochondrial damage, and this toxicity depends on the outer membrane protein OmpA¹⁶.

It is unclear whether A. baumannii OMVs exert other effects on host cells, but one potential mechanism is the modulation of host transcription factors to favor pathogenesis. Indeed, many host transcription factors respond to both internal and external signals. For example, FOS (originally discovered as a proto-oncogene) responds to various growth factors and electrochemical signals as well as pathogens^17,18,19,20. FOS dimerizes with members of the JUN family to form AP-1, which binds directly to DNA as a transcriptional activator. In the context of bacterial infection, the activation of FOS is induced by the MAP kinase signaling pathway and stimulates the production of cytokines^17,18,19,20. Whether FOS activity contributes to other aspects of host-pathogen interactions, such as the cytotoxicity induced by bacteria, is unclear.

Pathogens can also interfere with host metabolism. For example, the aryl hydrocarbon receptor (AHR) is a transcription factor activated by highly diverse ligands such as the environmental pollutant 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD), plant-derived indoles, and the products of tryptophan catabolism such as kynurenine. Activated AHR binds to aryl hydrocarbon receptor nuclear translocator (ARNT) and xenobiotic response elements (XREs) in target gene promoters to regulate transcription²¹. The target genes include those involved in the xenobiotic response, which helps to metabolize AHR ligands. Interestingly, AHR plays a key role in the regulation of immunity, and the activation of AHR is generally regarded as immunosuppressive^22,23. Bacterial pathogens may, therefore, target AHR signaling to manipulate host physiology and immunity, thus favoring pathogenesis.

Here we demonstrate that FOS and AHR play an important role in A. baumannii pathogenesis. We found that a strong induction of FOS is a characteristic feature of A. baumannii infection. The pharmacological inhibition of FOS specifically prevented cell death induced by A. baumannii in cell-based models and led to a reduction of severity in a mouse model of infection. Intriguingly, we found that bacterial OMVs are potent activators of AHR, which directly induces FOS expression. Both the internalization of OMVs and bacterial lipid A synthesis are required for the activation of AHR. Furthermore, the host enzyme tryptophan-2,3-dioxygenase (TDO) is needed for the activation of AHR by OMVs. Exposure to OMVs swiftly activated TDO and the production of endogenous kynurenine in host cells. Together, these findings reveal molecular mechanisms of A. baumannii pathogenesis and suggest potential targets for host-directed therapies.

Results

A. baumannii infection strongly induces the expression of FOS

To better understand the host response to A. baumannii infection, we carried out a comparative proteomics analysis of A549 cells infected with A. baumannii (ATCC 19606) and mock-infected controls (Fig. 1a and Supplementary Data 1). A549 cells originate from lung epithelium and are often used in A. baumannii infection experiments^16,24,25. The transcription factor FOS was the most strongly upregulated protein (Fig. 1b). FOSB, another member of the FOS family, was also significantly upregulated (Supplementary Data 1). Gene ontology (GO) analysis revealed an enrichment of proteins involved in mitochondrial functions among the upregulated proteins (Supplementary Fig. 1a), consistent with previous findings showing the damaging effects of A. baumannii on mitochondria¹⁶. In contrast, response to inorganic substances, cell cycle and regulation of cell death were enriched among the downregulated proteins (Supplementary Fig. 1a). The upregulation of FOS was confirmed using different laboratory strains (ATCC 19606 and 17978) and clinical strains (CDC00035, CDC00036, CDC00037²⁶ and FDA-CDC AR-BANK#0280 (#0280)) of A. baumannii (Fig. 1c), and in human monocyte-derived macrophages (MDMs) (Fig. 1d). We also observed a moderate induction of JUN (Supplementary Fig. 1b and Supplementary Data 1). Intriguingly, FOS was induced more strongly by A. baumannii than the other Gram-negative pathogenic bacteria E. coli or Pseudomonas aeruginosa, whereas the activation of NF-κB was comparable for all three bacteria, as indicated by the degradation of IκB (Fig. 1e). The induction of FOS by A. baumannii is therefore more potent than that by E. coli and P. aeruginosa. Induction occurred at the transcriptional level, as shown by the higher FOS transcript levels measured by qRT-PCR (Fig. 1f and Supplementary Fig. 1c). Consistent with the results from cell culture, we observed the strong induction of FOS in the spleen, liver and lung tissues of mice infected with the A. baumannii clinical isolate FDA-CDC AR-BANK#0280 (#0280) (Fig. 1g). These results reveal that the unusual strong induction of FOS is a major host response to A. baumannii infection in vitro and in vivo.

Inhibition of FOS alleviates the cytotoxicity of A. baumannii infection

Given that FOS is the most highly induced protein in A. baumannii-infected cells (Fig. 1), we tested whether FOS contributes to the cytotoxicity induced by A. baumannii infection using pharmacological inhibitors (Fig. 2a). T5224 is a FOS inhibitor with low in vivo toxicity and good bioavailability²⁷ and the only selective FOS inhibitor used in human clinical trials²⁸. Interestingly, T5224 significantly attenuated the cytotoxicity induced by A. baumannii infection in different cells, including A549, lung fibroblast MRC-5, skin fibroblast BJ, keratinocyte HaCaT, and mouse macrophage-like RAW264.7 cells (Fig. 2b) but it did not affect the release of lactate dehydrogenase (LDH) from A549 cells in the absence of infection (Supplementary Fig. 2a, b). We observed the same attenuation of cytotoxicity by T5224 in A549 cells infected with different clinical strains of A. baumannii (Fig. 2c), suggesting this phenomenon is also relevant for clinical strains. SR11302 is a FOS inhibitor with no structural similarity to T5224²⁹. However, like T5224, SR11302 also inhibited cell death induced by A. baumannii (Supplementary Fig. 2c), thus confirming the FOS-specific effect. T5224 did not rescue cell death induced by P. aeruginosa, indicating that the effect is also pathogen-specific (Supplementary Fig. 2d). The FOS inhibitors did not have any effect on the growth of A. baumannii (Supplementary Fig. 2e), indicating that the differences in cytotoxicity were not due to inhibition of the bacteria.

Because FOS is a transcription factor, we next investigated whether transcripts involved in cell death regulation are affected by T5224 treatment. We found that the expression of the anti-apoptotic genes BIRC3, BCL3, and CDK5 was significantly enhanced by T5224 treatment during A. baumannii infection (Fig. 2d). Conversely, T5224 treatment suppressed the expression of the pro-apoptotic genes BNIP3 and BIM in infected cells (Fig. 2e). Based on published data, these cell death regulators are known targets of FOS^30,31. Furthermore, T5224 treatment reduced the cleavage of caspase 9 and PARP triggered by A. baumannii infection in A549 cells (Fig. 2f), suggesting that apoptosis is suppressed. We observed no significant change in cytokine levels during T5224 treatment (Supplementary Fig. 2f), suggesting that inflammation does not play a major role in FOS-mediated cytotoxicity. We therefore concluded that FOS inhibition specifically rescues apoptosis induced by A. baumannii infection.

T5224 appears to be a suitable candidate of host-targeting therapeutic for A. baumannii infection, so we tested the efficacy of T5224 in a mouse infection model (Fig. 2g). Wild-type BALB/c mice were intraperitoneally (i.p.) infected with the clinical isolate strain FDA-CDC AR-BANK#0280 (#0280), which is highly virulent and extensively drug-resistant³². Intraperitoneal infection in mice causes peritonitis and dissemination of bacteria and leads to systemic disease^33,34,35. We chose to administrate T5224 both before and after infection since this infection model progresses rapidly (Supplementary Fig. 2i). A pre-infection treatment might be necessary for the maximal efficacy in this particular model. At 8 hours post-infection (hpi), we observed high levels of bacteria in the spleen, liver, and lung regardless of the treatment group, confirming bacterial dissemination (Supplementary Fig. 2g). Interestingly, mice treated with T5224 had lower levels of cleaved caspase 9 in their organs (Fig. 2h), which indicates a reduction of cell death, mirroring the phenomenon we observed in cells (Fig. 2f). Intriguingly, histological examination revealed a reduction of neutrophils and fibrin resembling an acute peritonitis on the serosal surface of liver and spleen of infected animals treated with T5224 (Supplementary Fig. 2h). This indicates a less severe peritoneal inflammation. We did not observe any statistically significant increase of survival in infected mice treated with T5224 (Supplementary Fig. 2i). All in all, these results show that FOS inhibition reduces the cytotoxicity due to A. baumannii infection, both in vitro and in vivo.

Bacterial OMVs trigger the induction of FOS

To investigate the molecular mechanism of FOS regulation in infected cells, we tested the effect of heat-inactivated bacteria. This abolished FOS induction but had little effect on NF-κB activation (Fig. 3a and Supplementary Fig. 3a), suggesting that bacterial activity, not just pathogen-associated molecular patterns (PAMPs), is required for FOS induction. Because OMV production is an active process that is required for bacterial pathogenesis^14,16,36,37, we set up transwell plates in which A549 cells were separated from A. baumannii cells by a 0.4-µm filter, preventing direct contact but allowing the exchange of secreted factors, such as OMVs. Under such conditions, A. baumannii was still able to induce FOS in the host cells (Fig. 3b and Supplementary Fig. 3b). Consistently, treating the cells with purified OMVs also resulted in the upregulation of FOS (Fig. 3c). The involvement of OMVs was supported by the use of OMV production modulators. Sub-inhibitory concentrations of colistin destabilize the bacterial outer membrane and promote the production of OMVs³⁸. Accordingly, the presence of colistin during A. baumannii infection further augmented FOS induction (Fig. 3d), whereas the OMV inhibitor BB-Cl-amidine^39,40 had the opposite effect (Fig. 3e).

The outer membrane protein OmpA is a major virulence factor carried by OMVs¹⁶. To determine whether OmpA is required for the induction of FOS, we infected A549 cells with the A. baumannii ΔompA mutant. Unexpectedly, we observed an even stronger induction of FOS in cells infected with this mutant (Supplementary Fig. 3c, d), suggesting that FOS induction does not require OmpA. This was confirmed in the transwell plates setting (Supplementary Fig. 3e), indicating that secreted factors, such as OMVs, are involved. Consistently, FOS inhibition by T5224 also limited cytotoxicity in cells infected with the ΔompA mutant (Supplementary Fig. 3f), confirming that FOS-mediated cytotoxicity does not require OmpA. The ΔompA mutant is known to produce more OMVs than normal laboratory and clinical strains⁴¹, which might explain the stronger induction of FOS during infection.

To gain deeper insight into the effect of OMVs on host cells, we repeated the earlier comparative proteomics analysis, this time using A549 cells treated with OMVs purified from A. baumannii (Fig. 3f). As observed in the infected cells (Fig. 1b), FOS was one of the proteins most strongly induced by OMVs (Fig. 3g and Supplementary Data 2). GO analysis showed that the upregulated proteins were enriched for the establishment of protein localization, ribosome biogenesis, cellular macromolecule localization, and RNA processing, whereas the downregulated proteins were enriched for the regulation of the cell cycle and cell division (Supplementary Fig. 3g).

FOS induction requires AHR

Next, we investigated the downstream steps in the induction of FOS by OMVs. The receptor AHR was downregulated in both infected and OMV-treated cells (Figs. 1b and 3g). AHR is required for the expression of FOS in bone cells during osteoclastogenesis⁴², and AHR is degraded by the proteasomal system following activation (Fig. 4a)⁴³. Therefore, we hypothesized that AHR is required for the induction of FOS by OMVs. First, we confirmed the downregulation of AHR in infected A549 cells (Fig. 4b), infected MDM cells (Fig. 4c), and OMV-treated A549 cells (Fig. 4d). Blocking AHR activity using the chemical inhibitor CH223191 or siRNA significantly reduced the induction of FOS by OMVs (Fig. 4d, e). CH223191 treatment also prevented the degradation of AHR (Fig. 4d), indicating that it inhibits AHR activation. AHR is degraded by the proteasome following OMV exposure, as confirmed by the ability of the proteasomal inhibitor MG132 to prevent the AHR downregulation (Fig. 4f). Although OMV treatment led to rapid degradation of AHR in 3 h, the protein levels of FOS and CYP1A1, a major target of AHR, remained highly upregulated even after 6 h of OMV treatment (Supplementary Fig. 4), suggesting that the effects of AHR activation persist even after its degradation. Furthermore, the activation of AHR using 6-formylindolo(3,2-b)carbazole (FICZ) induced FOS protein expression (Fig. 4g). We concluded that AHR is the mediator of FOS expression induced by OMVs.

AHR is a transcription factor, so we tested whether the FOS gene is directly regulated by AHR. Inhibition of AHR repressed FOS transcript induction by OMVs (Fig. 4h). AHR also physically bound to the FOS promoter region in chromatin immunoprecipitation (ChiP)-qPCR experiments (Fig. 4i). Based on these results, we concluded that AHR is a direct transcriptional regulator of FOS.

OMVs from A. baumannii activate AHR

Given that AHR is needed for the induction of FOS by OMVs (Fig. 4), it is likely that OMVs activate AHR. To test this hypothesis, we used a luciferase reporter system driven by the xenobiotic response element (XRE), which is the target site for activated AHR⁴³. As expected, purified OMVs from A. baumannii potently activated the reporter in an AHR-dependent manner (Fig. 5a). OMVs facilitate the intracellular delivery of bacterial factors, and endocytosis is believed to be the major route for OMVs to enter mammalian cells¹⁰. To determine whether the activation of AHR by OMVs requires internalization, we repeated the XRE luciferase assay in the presence of cytochalasin B, dynasore, and bafilomycin A1. Cytochalasin B inhibits the assembly of actin filament networks⁴⁴ whereas dynasore inhibits dynamin-mediated membrane scission⁴⁵. Both processes are necessary for the endocytosis of OMVs (Fig. 5b)¹⁰. The treatment of A549 cells with cytochalasin B or dynasore significantly inhibited luciferase reporter gene activation induced by OMVs (Fig. 5c), indicating that OMVs must be taken up to activate AHR. The release of cargo from internalized OMVs can be triggered by endolysosomal acidification (Fig. 5b), and this process is inhibited by bafilomycin A1^14,46. The addition of bafilomycin A1 also slightly inhibited luciferase reporter gene activation induced by OMVs (Fig. 5d). Endolysosomal acidification might play a secondary role in the activation of AHR as the effect of bafilomycin A1 was relatively small. In addition, AHR target genes such as CYP1A1, CYP1B1, AHRR, TIPARP, and STC2 were strongly induced by OMV treatment, as determined by qRT-PCR, and the induction was partially abolished by the AHR inhibitor CH223191 (Fig. 5e). The induction of AHR target genes was also observed in cells infected with A. baumannii (Supplementary Fig. 5a, b). Accordingly, organs from mice infected with A. baumannii contained elevated levels of CYP1A1 protein (Fig. 5f), indicating that AHR is activated in vivo following infection with A. baumannii. Based on these observations, we concluded that internalized OMVs activate AHR.

We also tested whether the FOS inhibitors T5224 and SR11302 are inducers of AHR since AHR is known to be promiscuous²¹. The XRE luciferase was potently activated by FICZ, a known activator of AHR, while there was no significant induction by T5224 or SR11302 (Supplementary Fig. 5c). FICZ also potently induced the transcription of CYP1A1. On the other hand, T5224 resulted in a very small induction of CYP1A1 while SR13302 did not have an effect (Supplementary Fig. 5d). All in all, these data indicate that T5224 and SR11302 are not relevant activators of AHR.

Lipopolysaccharide (LPS) is an abundant component of OMVs. LPS isolated from E. coli can activate AHR²². A. baumannii does not synthesize LPS but produces a similar molecule known as lipooligosaccharide (LOS)³. Lipid A is an obligatory component of both LPS and LOS and is the main PAMP sensed by host cells⁴⁷. Although lipid A synthesis is essential for the survival of most bacteria, A. baumannii mutants lacking lipid A remain viable^48,49. We exploited this unique property to investigate the role of lipid A in the activation of AHR. Interestingly, while OMVs from wild-type bacteria efficiently activated AHR, equal amounts of OMVs isolated from the lipid A-deficient mutant ΔlpxA (Fig. 5g) were unable to activate AHR (Fig. 5h), demonstrating that lipid A synthesis is required for OMVs to activate AHR. To determine whether host receptors for lipid A are also required, we repeated the AHR reporter gene experiments using HEK 293T cells, which lack functional receptors for lipid A (including TLR4 and caspase 4/5) and are therefore unresponsive to lipid A stimulation^50,51. Surprisingly, OMVs still activated AHR in these cells in a lipid A-dependent manner (Fig. 5i). Neither purified lipid A nor cell lysate from A. baumannii was sufficient to activate AHR, highlighting the importance of OMVs as a bacterial factor (Supplementary Fig. 5e). Collectively, these results suggest a novel AHR activation mechanism which is dependent on OMVs and bacterial lipid A synthesis.

OMVs activate AHR by modulating host tryptophan metabolism

One way that OMVs could activate AHR is to stimulate the endogenous production of AHR ligands. The major AHR ligands produced in human cells are products of tryptophan catabolism, and this pathway is controlled by three different enzymes (IDO1, IDO2 and TDO), each responsible for catalyzing the same rate-limiting step (Fig. 6a)⁵². IDO1 and IDO2 are unlikely to be involved because they are not expressed at detectable levels in A549 and Huh-7 cells (Supplementary Fig. 6a–d). In contrast, we readily detected TDO (Fig. 6b and Supplementary Fig. 6e), suggesting this is the main enzyme that breaks down tryptophan in A549 and Huh-7 cells. Consistently, the specific TDO chemical inhibitor 680C91 blocked the activation of AHR in response to OMVs (Fig. 6c) whereas 1-methyl-d-tryptophan (1-M-D-T), a specific inhibitor of IDOs, had no significant effect (Fig. 6d). TDO levels were not altered by A. baumannii infection or OMV treatment (Fig. 6b) although the transcript levels were reduced slightly (Supplementary Fig. 6f). TDO is strongly expressed in the liver⁵², so we tested whether OMVs have the same effect on hepatocyte-derived Huh-7 cells⁵³. These responded to OMVs in the same way as A549 cells (Supplementary Fig. 5d, e, g), suggesting the whole pathway is conserved in liver cells.

Finally, we measured the levels of tryptophan metabolites in cells treated with OMVs by liquid chromatography-mass spectrometry (LC-MS). We found that kynurenine, the downstream product of TDO and the main endogenous AHR ligand, was elevated significantly following OMV exposure for just 1 h. Furthermore, the addition of 680C91 largely suppressed the induction of kynurenine caused by OMV treatment (Fig. 6e). Taken together, these results suggest that A. baumannii OMVs do not induce the expression of the host tryptophan catabolic enzymes, but specifically activate TDO post-translationally, which in turn generates ligands that activate AHR.

Discussion

We discovered a mechanism of cytotoxicity induced by A. baumannii infection which involves the interplay between invading bacteria, the host transcription factor FOS, and host tryptophan metabolism (Supplementary Fig. 7). The potential contribution of host transcription factors to the cytotoxicity induced by bacteria has been largely neglected. A. baumannii infections result in a stronger induction of FOS when compared to the two other similar bacterial pathogens that we tested, and FOS activity is required for full virulence of A. baumannii in vitro and in vivo. The induction of FOS is mediated at least in part by the take-up of secreted bacterial OMVs, which increase the activity of the host enzyme TDO to promote tryptophan catabolism and the accumulation of kynurenine. This metabolite then activates AHR, a direct transcriptional activator of FOS. Importantly, this pathway is easy to block with available drug candidates, making it an attractive target for host-directed therapeutics.

FOS is a regulator of cell death⁵⁴. The involvement of FOS in innate immunity and neoplasia has also been documented^17,20,55. However, whether FOS plays a role in the cell death caused by bacterial infections remains unknown. We found that A. baumannii infections cause a high upregulation of FOS, whose activity results in host cell death (Fig. 2). FOS inhibition prevented cell death induced by A. baumannii but not P. aeruginosa (Fig. 2 and Supplementary Fig. 2d). It should be noted that in the current study, we compared only a small number of bacterial strains. In the future, it would be interesting to investigate more bacteria with regard to the induction of FOS and the potential prevention of cytotoxic effects by FOS inhibitors. The FOS inhibitor T5224 has already been used in human clinical trials for other diseases, such as arthritis²⁸. The repurposing of T5224 for bacterial infections would be more convenient than de novo drug development.

We found that OMVs are potent activators of AHR, which can directly induce the expression of FOS (Figs. 4 and 5). The role of OMVs in bacterial pathogenesis has recently gained more prominence⁵⁶. Our findings represent an important addition to the arsenal of ways in which OMVs usurp host physiology. AHR activation directs T-cell maturation and differentiation, which leads to the suppression of immunity and inflammation⁵⁷. AHR is involved in diverse pathogenic conditions, including graft vs host disease, autoimmune diseases, neurodegenerative diseases, and cancer²¹, which are potentially influenced by bacterial OMVs. This is important because OMVs are stable in vivo and can penetrate biological barriers that are non-permissive to intact bacterial cells, such as the blood–brain barrier⁵⁸. This would enable the bacteria to influence the physiology of distant organs.

Our results indicate that OMVs activate AHR indirectly via host tryptophan metabolism, specifically the catabolic enzyme TDO but not IDOs (Fig. 6). Tryptophan degradation is enhanced during inflammation to generate immunoregulatory ligands for AHR, such as kynurenine²², in a process mediated by IDOs⁵². IDOs are normally found at basal levels but their expression can be induced in a wide range of tissues by IFNγ during inflammation⁵². In contrast, TDO is believed to be responsible for the maintenance of homeostatic levels of tryptophan and kynurenine⁵². TDO is expressed at stable levels mainly in the liver⁵², but also in the lung⁵⁹, kidney^60,61, and brain^62,63. Consistently, we did not detect IDO expression in A549 or Huh7 cells where TDO was readily detectable (Fig. 6b and Supplementary Fig. 6a–f). In agreement with these observations, AHR activation by OMVs is also dependent on TDO but not IDOs (Fig. 6c, d). Contrary to the current beliefs of the predominate role of IDOs during inflammation, the expression or activity of IDOs are not altered by OMVs. Rather, OMVs activate TDO without affecting its expression level (Fig. 6b and Supplementary Fig. 6e), indicating a novel mechanism for enhanced kynurenine production via a post-translational activation of TDO by OMVs. The proposed regulation of the liver-enriched enzyme TDO by bacterial OMVs is fascinating because OMVs accumulate in the liver in vivo^12,64,65. We hypothesized that high levels of OMVs in the liver might result in the activation of TDO and AHR in hepatocytes, thus influencing hepatic functions. An increase of TDO activity can lead to the depletion of tryptophan, which triggers the activation of GCN2⁶⁶ and thus the suppression of immune responses^67,68,69, which could assist A. baumannii infections. The precise molecular mechanism used by OMVs to induce TDO activity should be investigated in future studies.

Lipid A is the main PAMP of Gram-negative bacteria and their OMVs⁷⁰. We found that the activation of AHR by OMVs requires bacterial lipid A synthesis and internalization (Fig. 5), suggesting that intracellular lipid A may be responsible for the OMV-mediated activation of the TDO–AHR signaling pathway. Alternatively, the full activation of AHR might require other components of OMVs. Cellular recognition of lipid A is mediated by TLR4 and caspase 4/5 (caspase 11 in mice)⁷⁰. Lipid A and OMVs have been shown to induce potent host responses in cells expressing TLR4 and/or caspase 4/5/11 whereas cells devoid of functional TLR4 and caspase 4/5/11, such as HEK293T cells, are unresponsive to lipid A^50,51. Surprisingly, OMVs still induce robust lipid A-dependent AHR activation in HEK 293T cells (Fig. 5i). This suggests the lipid A-dependent activation of AHR is independent of TLR4 and caspase 4/5/11. Alternatively, a lack of LpxA may alter the structures or components of OMVs in addition to lipid A, which might also contribute to the loss of AHR activation. Given the swiftly increasing kynurenine production in host cells (Fig. 6e, samples harvested 1 hour after OMV treatment), we speculate that TDO or its interacting partners might directly interact with OMVs.

This study provides the first promising results regarding the reduction of the severity of intraperitoneal A. baumamnnii infection in a mouse model using FOS inhibitors. However, a more thorough investigation is needed to further consolidate the potential of FOS inhibitors as host-directed-therapeutics against bacterial infections. For example, it would be useful to include pneumonia models in future studies, which might be more representative for human infections.

In conclusion, our study facilitates the future exploration of the intriguing connection between bacterial OMVs, host metabolism, and cell death pathways. Given the current antibiotic resistance crisis, we propose further in vivo investigation of FOS inhibitors in bacterial infections. This represents a direction for developing therapies alternative to traditional antibiotics.

Methods

Ethical regulation

The animal studies were reviewed and approved by the ethic committee PDST IACUC (Pharmacology Discovery Services Taiwan Institutional Animal Care and Use Committee). We confirm that the animal experiments complied with all relevant ethical regulations.

Mammalian cell culture

A549, RAW264.7, MRC-5, BJ, and HEK 293 T cells were obtained from the American Type Culture Collection (ATCC). Human monocyte-derived macrophages (MDMs) were differentiated from peripheral blood mononuclear cells (PBMCs) collected from a local blood donation center. PBMCs were washed with PBS and incubated in RPMI 1640 medium supplemented with 2.5% heat-inactivated human serum for ~7 days, allowing them to differentiate. All other cells were cultivated in RPMI 1640 (A549 cells) or Dulbecco’s modified Eagle’s medium (DMEM) (RAW264.7, MRC-5, BJ, HaCaT, Huh7, and HEK 293T cells) supplemented with 10% fetal calf serum. All cells were cultured at 37 °C in a humid 5% carbon dioxide atmosphere. All cells used in this study were tested negative for mycoplasma contamination.

Bacterial culture

The A. buamannii ΔlpxA mutant²⁶ and the corresponding ATCC 19606 wild-type strain were cultured in tryptic soy broth (TSB) and on tryptic soy agar (TSA). The following strains were cultured in lysogeny broth (LB) and on LB agar: E. coli O127:H6, P. aeruginosa ATCC 21853, A. baumannii ATCC 17978, 19606 (when not serving as the wild-type strain for the ΔlpxA mutant), FDA-CDC AR-BANK#0280, CDC00035, CDC00036 and CDC00037. FDA-CDC AR-BANK#0280 was cultivated in brain heart infusion (BHI) broth for the mouse experiments. Bacteria were cultivated at 37 °C following standard procedures.

Mouse housing and husbandry

Experiments on live animals were carried out by Pharmacology Discovery Services Taiwan, Ltd. (partner of Eurofins Discovery) as an experimental paid service (SOW # ODS112-01006934). All aspects of animal experiments, including housing, treatments, and disposal, adhered to the “Guide for the Care and Use of Laboratory Animals: Eight Edition”⁷¹ and were carried out in an Association for Assessment and Accreditation of Laboratory Animal Care (AAALAC)-accredited animal facility.

All animal experiments involved BALB/c immunocompetent female mice at 5-6 weeks of age, which were closely monitored for health on a daily basis. The animals were maintained in a controlled temperature (20 − 24 °C) and humidity (30% − 70%) environment with 12-h light/dark cycles. Following infection with A. baumannii, animals were monitored for mortality and health at least twice per day. When they reached early humane endpoints, showed signs of pain, or became moribund, they were immediately euthanized. The responsible veterinarian had full authority to remove animals from the study and/or perform euthanasia prior to the experimental endpoint in order to prevent unacceptable levels of pain or distress.

In vitro model of bacterial infection

Mammalian cells were seeded on the day before infection to generate confluent monolayers for infection experiments. Saturated bacterial cultures were diluted 1:10 in fresh LB or TSB (for experiments involving the ΔlpxA mutant). The diluted cultures were then grown at 37 °C for ~2 h shaking at 180 rpm to reach an OD₆₀₀ of ~1.0. Bacterial cells were then pelleted by centrifugation and washed twice with RPMI or DMEM. After resuspending the bacteria in cell culture medium supplemented with 10% FBS, the suspension was added to the cells at a MOI of 200 unless otherwise stated. Chemicals were added to the bacterial suspension if applicable. For transwell infection experiments, bacteria were placed in a TC-insert (Sarstedt) with a pore size of 0.4 µm.

Mass spectrometry

A549 cell pellets were heated at 95 °C for 10 min in lysis buffer containing 2% SDS, 50 mM Tris-HCl (pH 8), 1 mM tris(2-carboxyethyl)phosphine (TCEP) and 4 mM chloroacetamide. The protein content was precipitated with chloroform/methanol. The protein pellets were then solubilized in 8 M urea and digested with Lys-C and trypsin at 37 °C overnight. The resulting peptides were desalted using Sep-Pak cartridges or C-18 stage tips and labeled with TMT reagents for MS analysis in an Orbitrap Fusion Lumos mass spectrometer. MS data were analyzed with Proteome Discoverer v2.4 (Thermo Fisher Scientific) and subsequently with Perseus.

Gene ontology enrichment analysis

GO enrichment analysis was carried out using ShinyGO 0.76.2 according to the authors’ instructions⁷² (http://bioinformatics.sdstate.edu/go/).

In vivo model of bacterial infection

Groups of 15 BALB/c immunocompetent female mice (body weight 20 ± 2 g) were inoculated i.p. with 3 × 10³ CFU A. baumannii (FDA-CDC AR-BANK#0280) suspended in 0.5 mL BHI broth containing 5% mucin. T5224 was dissolved in a vehicle solution at a concentration of 50 mg/mL as suggested in the corresponding patent⁷³. Briefly, 500 mg of T5224 was dissolved in a mixture composed of 3.9 mL 0.5 mol/L NaOH and 5.1 mL pure H₂O. We then added 1.5 g of Plasdone K29/32 to obtain a 50 mg/mL solution at pH 8.3. Vehicle solution (15 animals) and T5224 (15 animals) were administered orally 23 h before and 1 h after A. baumannii infection at a dose of 250 mg/kg. Five mice per group were euthanized 8 hpi, and the lung, spleen, and kidney were harvested for analysis.

Histology

Formalin-fixed samples were embedded in paraffin and 5-μm sections were prepared using a MICROM HM 355S rotational microtome (Thermo Fisher Scientific). The slides were deparaffinized and rehydrated by sequential immersion in xylene, ethanol and water before staining with hematoxylin and eosin using an H&E staining kit (Abcam).

Bacterial burden

Mice were sacrificed at 8 hpi. The organs were weighed and homogenized to release the bacteria. CFUs were determined using the dilution plating technique and were divided by the weight of the organs for normalization.

Heat inactivation of bacteria

Bacteria were washed twice with RPMI and then resuspended in RPMI supplemented with 10% FBS. The suspension was then heated to 95 °C for 10 min. Complete inactivation was confirmed by the absence of growth on LB agar plates after overnight incubation at 37 °C. Equal amounts of live and inactivated bacteria (MOI 200) were used for infection experiments.

OMV isolation

One liter of overnight bacterial culture was used for OMV isolation. Bacterial cells were removed by centrifugation (4969 g, 30 min, 4 °C) and passing the supernatant through a 0.22-µm PVDF filter. The filtered medium was centrifuged (369548.3 g, 1 h, 4 °C) to pellet the OMVs, which were washed with PBS. The pelleting and washing steps were repeated twice. The final preparation of washed OMVs was then resuspended in 1 mL PBS and sterilized by passing through a 0.22-µm syringe filter. The concentration was determined using a bicinchoninic acid (BCA) assay.

Western blot

Cell pellets were mixed with SDS lysis buffer (1% SDS in 50 mM Tris-HCl, pH 8) and immediately heated to 95 °C for 10 min to ensure lysis and protein solubilization. Mouse tissues were lysed in RIPA lysis buffer containing proteinase inhibitor by sonication in a Branson Sonifier 250. The protein concentration was determined using the BCA method. For each sample, 10–15 µg protein was loaded on an SDS polyacrylamide gel and fractionated by electrophoresis. Separated proteins were transferred to PVDF membranes, which were then blocked with 5% non-fat milk powder in PBS containing 0.1% Tween-20 (PBST). The membranes were then incubated sequentially with the primary antibodies and horseradish peroxidase (HRP)-conjugated secondary antibodies, with intervening washes in PBST. Proteins were detected using the WesternBright chemiluminescence reagent (Advansta). Please refer to the “reporting summary” for the list of antibodies and the dilution used.

LDH release assay

Cell culture supernatant was collected at the indicated time points, and LDH activity was determined using a Cytotoxicity Detection Kit (Roche).

RNA isolation and qRT-PCR

Total RNA was extracted using the RNeasy Mini kit (Qiagen), and cDNA was prepared using the RevertAid First Strand cDNA Synthesis Kit (Thermo Fisher Scientific). The cDNA was amplified by qPCR using Takyon No ROX SYBR 2X MasterMix blue dTTP (Eurogentec). The relative abundance of the target sequences was calculated using the ∆∆Ct method. GAPDH was used for internal normalization. For the sequences of the primers please refer to the supplementary “Oligonucleotides” file.

Chromatin immunoprecipitation assay

We used the Pierce Magnetic ChIP Kit (Thermo Fisher Scientific) according to the manufacturer’s protocols. Eluted DNA was analyzed by qPCR to determine the relative abundance of the targets as described above.

Transfection of mammalian cells

Plasmids were introduced into A549 and HEK 293T cells using polyethyleneimine as the transfection reagent, whereas Lipofectamine 2000 was used for the transfection of Huh-7 cells. Lipofectamine RNAiMAX was used for the introduction of siRNA into A549 cells. Pre-designed siRNAs (ON-TARGETplus siRNA SMARTPool) were purchased from Horizon Discovery.

Luciferase reporter assay

On the day before the experiments, cells were transfected with three plasmids: XRE-Luc (pGL4.43[luc2P/XRE/Hygro], Promega), a Renilla luciferase expression vector driven by the thymidine kinase promoter, and pEGFP-C1 (to monitor transfection efficiency). On the day of the experiments, cells were treated with the indicated chemicals and/or OMVs for 3 h before lysis, and luciferase activity was measured using the Dual-Luciferase Reporter Assay System (Promega). Firefly luciferase activity was normalized to the activity of Renilla luciferase.

Lipid A and bacterial lysate treatment

A549 cells were treated with purified lipid A from Salmonella enterica (sigma Aldrich) at a concentration of 15 μg/mL or A. baumannii lysate (1:150 diluted) for 3 hours before measuring luciferase activity. To generate cell lysate from A. baumannii, strain 19606 was grown in LB medium till OD600 1. Bacteria from 1 mL of the culture were pelleted, washed, and resuspended in 50 μl of PBS. The suspension was then heated at 95 °C for 10 min. Insoluble material was removed using centrifugation.

Measurement of tryptophan metabolites

Cell pellets (~1 × 10⁶ cells per sample) were lysed in 200 μl methanol extraction buffer (1 mM TCEP, 1 mM ascorbic acid, 0.1% formic acid in 85:15 methanol:water) containing the internal standards 20 µM homotaurine and 20 µM serotonin-d4. Protein precipitates were removed by centrifugation and the supernatant was freeze-dried overnight. For each sample, 70 μl of 200 mM boric acid buffer was added for reconstitution. We then added 10 μl of 6-aminoquinolyl-N-hydroxysuccinimidyl carbamate (AQC) to each sample and incubated at 55 °C for 10 min to derivatize amines. Samples were then centrifuged to remove lipids and remaining particulates, and the supernatant was transferred to MS glass vials.

We injected 5 μl of each sample into an Infinity II Bio liquid chromatography system coupled to a 6495 C triple quadrupole mass spectrometer (Agilent Technologies). Tryptophan metabolites were separated on a Zorbax Extend C18 column (RRHD 2.1 ×150 mm, 1.8 µm; Agilent Technologies) at a flow rate of 0.3 mL/min. Metabolites were eluted in a gradient of solvent A (0.1% formic acid in water) and solvent B (0.1% formic acid in acetonitrile) over a period of 19 min: 0 min 1% B, 0–2 min 1% B, 2–9 min 15% B, 9–14 min 30% B, 14–16 min 60% B, 16–17 min 65% B, and 17–19 min 1% B, followed by a 2-min post-time run. Eluted standards were detected in positive ionization dynamic MRM AJS-ESI mode. The gas temperature was set to 290 °C and the gas flow to 20 L/min. The nebulizer was set to 45 psi. The sheath gas flow was set to 11 L/min, with a temperature of 400 °C. The capillary voltage was set at 3800 V with a nozzle voltage of 500 V. The voltages of the High-Pressure RF and Low-Pressure RF were set to 150 and 60 V, respectively.

Quantification and statistical analysis

GraphPad Prism v8 was used for statistical analysis. Pairs of variables were compared using unpaired and two-tailed Student’s t tests. Data are represented as means ± standard deviations.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

Source data are provided with this paper. The proteomics data generated in this study have been deposited in the ProteomeXchange Consortium via the PRIDE partner repository⁷⁴ partner repository with the dataset identifier PXD053768. Source data are provided with this paper.

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Acknowledgements

The authors thank Dr. Andreas Weigert from Goethe University of Frankfurt, Germany for the generous support with the generation of the MDM cells. The mouse infection experiments were carried out by the commercial service provider Pharmacology Discovery Services Taiwan, Ltd. We would also like to thank Julia Bein from Goethe University of Frankfurt, Germany for the technical assistance with the processing of the histological samples. This work was supported by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) – Project-ID 259130777 – SFB 1177, the Cluster Project ENABLE funded by the Hessian Ministry for Science and the Leistungszentrum Innovative Therapeutics (TheraNova) funded by the Fraunhofer Society and the Hessian Ministry of Science and Art.

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Authors and Affiliations

Institute of Biochemistry II, Faculty of Medicine, Goethe University, Frankfurt, Germany
Chun Kew, Cristian Prieto-Garcia, Anshu Bhattacharya, João Mello-Vieira, Yi-Lin Cheng, Rajeshwari Rathore & Ivan Dikic
Buchmann Institute for Molecular Life Sciences, Goethe University, Frankfurt, Germany
Chun Kew, Anshu Bhattacharya, João Mello-Vieira, Yi-Lin Cheng & Ivan Dikic
Institute for Medical Microbiology and Infection Control, Hospital of the Goethe University, Frankfurt, Germany
Manuela Tietgen, Stephan Göttig & Volkhard A. J. Kempf
University Center of Competence for Infection Control of the State of Hesse, Frankfurt, Germany
Manuela Tietgen
Department of Infectious Diseases, Genentech Inc., 1 DNA Way, South San Francisco, CA, USA
Craig R. MacNair, Lindsey A. Carfrae & Man-Wah Tan
Institute for Vascular Signalling, Department of Molecular Medicine, CPI, Goethe University, Frankfurt, Germany
Stephan Klatt & Ingrid Fleming
Institute of Basic Medical Sciences, College of Medicine, National Cheng Kung University, Tainan, Taiwan
Yi-Lin Cheng
Department of Pathology, Dr. Senckenberg Institute of Pathology, Goethe University, Frankfurt, Germany
Elise Gradhand
Fraunhofer Institute for Molecular Biology and Applied Ecology, Branch Translational Medicine and Pharmacology, Frankfurt, Germany
Ivan Dikic
Max Planck Institute of Biophysics, Frankfurt, Germany
Ivan Dikic

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Chun Kew
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Cristian Prieto-Garcia
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Anshu Bhattacharya
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Contributions

Conceptualization, C.K. and I.D.; Methodology, C.K., C.P., A.B., M.T., C.M., L.C., S.K., Y.C., R.R., E.G., I.F., and S.G.; Investigation, C.K., C.P., A.B., C.M., L.C., J.M., S.K., and E.G.; Writing – original draft, C.K.; Writing – review & editing, C.K., C.P., M.T., Y.C., M-W.T., S.G., V.K. and I.D.; Funding acquisition, I.D.; Resources, I.F., M-W.T., S.G., V.K. and I.D.; Supervision, I.D.

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Kew, C., Prieto-Garcia, C., Bhattacharya, A. et al. The aryl hydrocarbon receptor and FOS mediate cytotoxicity induced by Acinetobacter baumannii. Nat Commun 15, 7939 (2024). https://doi.org/10.1038/s41467-024-52118-7

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Received: 07 March 2023
Accepted: 27 August 2024
Published: 11 September 2024
DOI: https://doi.org/10.1038/s41467-024-52118-7

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Introduction

Results

A. baumannii infection strongly induces the expression of FOS

Inhibition of FOS alleviates the cytotoxicity of A. baumannii infection

Bacterial OMVs trigger the induction of FOS

FOS induction requires AHR

OMVs from A. baumannii activate AHR

OMVs activate AHR by modulating host tryptophan metabolism

Discussion

Methods

Ethical regulation

Mammalian cell culture

Bacterial culture

Mouse housing and husbandry

In vitro model of bacterial infection

Mass spectrometry

Gene ontology enrichment analysis

In vivo model of bacterial infection

Histology

Bacterial burden

Heat inactivation of bacteria

OMV isolation

Western blot

LDH release assay

RNA isolation and qRT-PCR

Chromatin immunoprecipitation assay

Transfection of mammalian cells

Luciferase reporter assay

Lipid A and bacterial lysate treatment

Measurement of tryptophan metabolites

Quantification and statistical analysis

Reporting summary

Data availability

References

Acknowledgements

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Supplementary information

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