https://orcid.org/0000-0003-3444-1069
Figures
Abstract
Anthrax lethal toxin (LT) and edema toxin (ET) are two of the major virulence factors of Bacillus anthracis, the causative pathogen of anthrax disease. While the roles of LT in anthrax pathogenesis have been extensively studied, the pathogenic mechanism of ET remains poorly understood. ET is a calmodulin-dependent adenylate cyclase that elevates intracellular cAMP by converting ATP to cAMP. Thus, it was postulated that the ET-induced in vivo toxicity is mediated by certain cAMP-dependent events. However, mechanisms linking cAMP elevation and ET-induced damage have not been established. Cholera toxin is another bacterial toxin that increases cAMP. This toxin is known to cause severe intestinal fluid secretion and dehydration by cAMP-mediated activation of protein kinase A (PKA), which in turn activates cystic fibrosis transmembrane conductance regulator (CFTR). The cAMP-activated PKA phosphorylation of CFTR on the surface of intestinal epithelial cells leads to an efflux of chloride ions accompanied by secretion of H2O into the intestinal lumen, causing rapid fluid loss, severe diarrhea and dehydration. Due to similar in vivo effects, it was generally believed that ET and cholera toxin would exhibit a similar pathogenic mechanism. Surprisingly, in this work, we found that cAMP-mediated PKA/CFTR activation is not essential for ET to exert its in vivo toxicity. Instead, our data suggest that ET-induced ATP depletion may play an important role in the toxin’s pathogenesis.
Author summary
Anthrax ET is a calmodulin-dependent adenylate cyclase that elevates intracellular cAMP by converting ATP to cAMP. Thus, it was postulated that the ET-induced in vivo toxicity is mediated by certain cAMP-dependent events. PKA and Epac are the two major intracellular targets activated by cAMP. Surprisingly, in this work, we found that cAMP-mediated PKA/CFTR activation as well as Epac activation are not essential for ET to exert its in vivo toxicity. Instead, our data suggest that ET-induced ATP depletion may play an important role in the toxin’s pathogenesis. Our results suggest that ATP depletion might also be an important mechanism underlying pathogenesis of other adenylate cyclase toxins.
Citation: Liu J, Cao Q, Ewing M, Zuo Z, Kennerdell JR, Finkel T, et al. (2025) ATP depletion in anthrax edema toxin pathogenesis. PLoS Pathog 21(4): e1013017. https://doi.org/10.1371/journal.ppat.1013017
Editor: Joseph A. Sorg, Texas A&M University, UNITED STATES OF AMERICA
Received: February 2, 2025; Accepted: March 4, 2025; Published: April 1, 2025
This is an open access article, free of all copyright, and may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose. The work is made available under the Creative Commons CC0 public domain dedication.
Data Availability: All data supporting the findings of this study are available within the paper and its Supplementary Information.
Funding: This research was supported by the institutional seed fund from the Aging Institute of University of Pittsburgh School of Medicine (S.L.) and the grant (R01AI170574) (S.L.) from the National Institute of Allergy and Infectious Diseases (NIAID), NIH, and in part by the Intramural Program of the NIAID (S.H.L), NIH. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Introduction
Bacillus anthracis is a Gram-positive, rod-shaped, spore-forming bacterium and is the causative pathogen of anthrax disease. The B. anthracis spore, the infectious form of the pathogen, has long been considered as a potential warfare agent and was a top bioterrorism concern even before the 2001 anthrax attacks in the USA [1]. B. anthracis contains two large extrachromosomal pathogenic plasmids, pXO1 (182 kb) and pXO2 (96 kb) [2,3]. Plasmid pXO2 encodes proteins that synthesize the unique poly-D-γ-glutamic acid capsule, which disguises B. anthracis from host immune surveillance and confers resistance to phagocytosis by host macrophages [4]. Plasmid pXO1 encodes the three components of anthrax exotoxins: the cellular binding/delivering moiety– protective antigen (PA, 83 kDa), and two enzymatic effector proteins– lethal factor (LF, 89 kDa), and edema factor (EF, 90 kDa) [4]. To target host cells, PA binds to cell surface anthrax toxin receptors CMG2 (capillary morphogenesis protein-2, the major receptor) and TEM8 (tumor endothelium marker-8), leading to proteolytic activation of PA by a cell surface furin protease, yielding the active PA oligomer (heptamer and/or octamer) [4,5]. The PA oligomer then binds and translocates LF and/or EF into the cytosol of target cells to exert their cytotoxic effects. Thus, PA and LF pair to form anthrax lethal toxin (LT), while PA plus EF form anthrax edema toxin (ET), resulting in the two major virulence factors of B. anthracis. LF (the catalytic component of LT) is a zinc-dependent metalloproteinase that cleaves the mitogen-activated protein kinase kinases (MEKs) 1-4 and 6 [6–9], resulting in inactivation of the three key mitogen-activated protein kinase pathways: ERK (through MEK1/2), p38 (through MEK3/6), and JNK (Jun N-terminus kinase) (through MEK4), leading to LT-induced lethality of experimental mice [4,5,10,11]. Since many human cancers rely on ERK pathway for tumorigenesis and growth, LT has been extensively reengineered to selectively target those tumors [12–16].
ET is another major virulence factor of B. anthracis. This is evidenced by the 10-fold reduction in virulence of B. anthracis when the gene encoding EF (the catalytic component of ET) is deleted [17,18]. EF is a calmodulin-dependent adenylyl cyclase that converts ATP to cAMP, the classical second messenger [19]. Thus, it was postulated that ET-induced in vivo toxicity is mediated by certain cAMP-dependent events [4,5]. However, mechanisms linking cAMP elevation and ET-induced damage are poorly understood. cAMP has two major intracellular targets: protein kinase A (PKA) and the exchange protein activated by cAMP (Epac), which is a guanine nucleotide exchange factor/activator for the small G-proteins Rap1 and Rap2 [20–23]. Other bacterial toxins that increase cAMP include cholera toxin, secreted by Vibrio cholera. Cholera toxin causes severe intestinal fluid secretion and dehydration by cAMP-mediated activation of PKA, which in turn activates cystic fibrosis transmembrane conductance regulator (CFTR) [24,25]. CFTR is an ATP-binding cassette transporter-class chloride ion channel found in epithelial cells of many organs, including the lung, liver, digestive tract, and skin. The cAMP-activated PKA phosphorylates CFTR on the surface of intestinal epithelial cells, leading to efflux of chloride ions accompanied by secretion of H2O into the intestinal lumen, causing rapid fluid loss and severe diarrhea and dehydration [25]. Due to the similarity of ET and cholera toxin in their ability to cause fluid loss, it was believed the two toxins had similar mechanisms of action. In this work, we sought to investigate whether PKA-mediated CFTR activation is the cause of ET-induced edema and in vivo toxicity. Surprisingly, we found that cAMP-mediated PKA/CFTR activation is not essential for ET to exert its in vivo toxicity. Instead, our data suggest that ET-induced ATP depletion may play an important role in ET’s pathogenesis.
Results
CMG2 receptor-dependent in vivo toxicity of anthrax edema toxin
The in vivo pathogenicity of ET has been carefully assessed since the highly purified EF protein was made available two decades ago [5,26–28]. In addition to the characteristic skin edema when injected subcutaneously, ET is also lethal when administered systemically. In fact, the minimal lethal doses of ET to mice (10-20 µg) are even lower than observed for LT (25-50 µg) [26,29]. Aligning well with this, here, we showed that administration of 20 µg ET (I.P.) resulted in a 60% mortality rate in challenged mice within two days, in a CMG2 receptor-dependent manner (Fig 1A). ET is an adenylate cyclase that quickly increases cellular cAMP levels in cultured cells [19]. To assess this effect in vivo, we measured the cAMP levels in plasma and in tissue lysates of the livers, kidneys, and hearts of mice 18 hours after ET administration. CMG2 receptor-dependent cAMP increases were detected in the livers but not in the kidneys (Fig 1B). The trend to increase cAMP in the hearts was also demonstrated. Surprisingly, significant increases (about 100-fold) of cAMP were also detected in the plasma, revealing that cellular cAMP can efficiently move out of the cells (Fig 1B). However, the mechanism and the significance of this cAMP translocation in ET pathogenesis remain unclear.
A. CMG2-KO mice and their littermate WT control mice were challenged with 20 µg ET (I.P.), and monitored for signs of malaise and survival for 200 hours. Log-rank test for survival comparison. B. ET-induced cAMP increases in mice. Eighteen hours after challenge with 30 µg ET (I.V.), the mice (CMG2-KO and the controls) were euthanized, and blood, liver, heart, kidneys were collected and processed for measuring cAMP levels. Unpaired two-tailed Student’s t-test.
PKA- and CFTR-independent in vivo toxicity of anthrax edema toxin
The in vivo toxicity of ET was initially considered to be mediated by certain cAMP-activated pathological processes. cAMP has at least two distinct intracellular targets: PKA and Epac (Fig 2A) [20–23]. Epac is a guanine nucleotide exchange factor that activates the small G-proteins Rap1 and Rap2, which have positive roles in maintaining the tight junctions between epithelial cells [22,23,30,31]. Thus, activation of Epac by cAMP seems unlikely to mediate the ET’s in vivo toxicity. Consistent with this hypothesis, we found that the Epac specific inhibitor ESI-09 could not protect mice from ET challenges (S1 Fig). PKA exists as an inactive tetramer consisting of two regulatory and two catalytic subunits. Binding of cAMP to the regulatory subunits causes their dissociation from the two catalytic monomers of PKA, which are then activated by ATP binding and are able to phosphorylate/activate many substrate proteins, including the transcription factor cAMP response element-binding protein (CREB) and the cystic fibrosis transmembrane conductance regulator (CFTR).
A. cAMP-increasing bacterial toxins and cAMP-activated pathways. B. Activation of PKA by ET and forskolin. NIH3T3 cells were treated with ET (1 μg/mL) or FSK (forskolin, 10 μM) for various lengths of time as indicated with/without the presence of H89 (0.1 mM). The cell lysates were then analyzed by Western blotting using anti-phospho-CREB and anti-tubulin antibodies. Reprehensive of two independent experiments with similar results. C. Systemic administration of H89 effectively prevents ET-induced activation of CREB in the hearts of the ET-challenged mice. Mice were challenged with 30 µg ET (I.V.) or 30 µg ET (I.V.) plus 500 µg H89 (I.P.). Six 6 hours after administration, mice were euthanized and the heart tissue lysates prepared for Western blotting analysis using the antibodies as indicated (50 µg proteins loaded per lane). Of note, H89 could effectively diminish the ET-induced CREB activation. D. PKA inhibitor could not prevent ET-induced mortality. Mice were administered 30 µg ET (I.V.) or 500 µg H89 (I.P.) plus 30 µg ET (I.V.) and monitored for survival as indicated. E. CFTR-null (CFTR-KO (FABP-CFTR+)) mice and their WT control mice were challenged with 30 µg ET (I.V.) and monitored for survival. F. CFTR inhibitor could not prevent the ET-induced mortality. The mice were administered 30 µg ET (I.V.). 24 h and 1 h prior to ET injection, the mice were injected with PBS or 50 μg CFTRinh-172. (D-F) Log-rank test for survival comparison.
To examine whether ET-induced cAMP could activate PKA, we treated NIH 3T3 cells with ET, and found that phosphorylation (activation) of CREB was enhanced (Fig 2B). The PKA inhibitor H89 completely blocked this ET-induced CREB activation (phosphorylation) (Fig 2B). Further H89 could also completely block CREB activation by forskolin [32], another cAMP-increasing agent that increases cAMP through activating the endogenous adenylyl cyclase (Fig 2B). However, although H89 administration could efficiently diminish ET-induced activation of CREB in ET-challenged mice (Fig 2C), this PKA inhibitor could not provide survival benefit for the ET-challenged mice (Fig 2D).
To determine whether the activation of PKA or its downstream target CFTR is involved in ET-induced edema, we tested the effects of H89, as well as CFTRinh-172 [33], a specific CFTR inhibitor, on ET-induced edema using a mouse footpad edema model. ET could rapidly cause edematous lesions in this mouse footpad edema experiment. This pathology could only be marginally ameliorated by local pre-administration of either H89 or CFTRinh-172 (S2A and S2B Fig). However, the footpad edema caused by cholera toxin, another cAMP-increasing bacterial toxin [25], could be more pronouncedly reduced by H89 or CFTRinh-172 (S2C and S2D Fig).
CFTR-null (CFTR-/-) mice die within a month after birth due to severe intestinal occlusion [34]. However, this phenotype of CFTR-/- mice can be corrected when CFTR is specifically expressed in intestines through crossing with the intestinal-specific CFTR transgenic mice (Fabp2-CFTR mice) (CFTR is under the control of an intestinal-specific promotor rat fatty acid binding protein 2 (Fabp2) [35]. As such, CFTR-/-/Fabp2-CFTR mice, which express CFTR only in intestines but not in other tissues, have normal longevity, allowing us to assess the potential role of CFTR in ET pathogenesis. To determine whether the ET-induced in vivo toxicity is CFTR-independent, we generated and challenged the CFTR-/-/Fabp2-CFTR mice and their control mice with ET by intraperitoneal administration. We found that CFTR-/-/Fabp2-CFTR mice and their WT control mice were equally susceptible to ET-induced lethality (Fig 2E). Similarly, systemic administration of CFTRinh-172 could not provide survival benefit for the mice challenged with lethal doses of ET (Fig 2F). Taken together, we conclude that cAMP-mediated activation of PKA, as well as its downstream target CFTR, which appear to be involved in ET-caused edema, are not required for the ET-induced lethality.
Potential role of ATP depletion in ET pathogenesis
One characteristic cytotoxic effect of ET to cultured cells is its ability to rapidly induce morphological change including cell rounding (Fig 3A). This effect is also toxin receptor-dependent, since ET did not morphologically alter anthrax toxin receptor-deficient Chinese hamster ovary (CHO) PR230 cells, but could do so to PR230 cells following reconstitution with the CMG2 receptor (PR230 (CMG2)) (Fig 3A). Interestingly, along with these morphological changes, extracellular cAMP was noted to rapidly accumulate in the conditioned medium of the ET-treated PR230 (CMG2) cells, but not in the conditioned medium of PR230 cells (Fig 3B). To begin to study the potential cytotoxic effect of extracellular cAMP, PR230 cells and PR230 (CMG2) cells grown on different cover slips were co-cultured in a same chamber and were treated with ET (Fig 3C). Although these cells were in the same conditioned medium, morphological change only occurred with the PR230 (CMG2) cells, and not with toxin receptor-deficient PR230 cells (Fig 3C), suggesting that the cell rounding effect of ET was not likely attributable to the accumulation of the extracellular cAMP.
A, B. Toxin receptor-dependent cell rounding effect of ET. PR230 (a PA receptor-deficient CHO mutant cell line) and PR230 (CMG2) (PR230 reconstituted with CMG2 receptor) were incubated with/without ET (0.1 or 1.0 µg/mL) as indicated for 2 h. The ET-induced spherical morphological change was observed on PR230 (CMG2) cells but not on PR230 cells (A). Similarly, the ET-induced extracellular cAMP accumulation was only detected in the conditioned medium of PR230 (CMG2) cells but not that of PR230 cells (B). Means ± SD of three independent biological replicates. C. Co-culture experiment indicates no cell rounding effect of extracellular cAMP. The cover slips grown with PR230 cells and PR230 (CMG2) cells were placed in the same wells of a 6-well plate and co-incubated with/without ET for 2 h. Of note, ET could only induce the characteristic cell rounding morphological change to PR230 (CMG2) cells but not to PR230 cells in the co-culture system.
EF is an extremely efficient bacterial adenylate cyclase, able to convert 1000–2000 molecules of ATP to cAMP per second [36]. Therefore, EF may also deplete cellular ATP, thereby affecting a variety of ATP-depending biological processes, such as vesicle trafficking, tight junction protein delivery. To test whether the ET-induced cellular cAMP increases are accompanied by cellular ATP decreases, NIH3T3 cells were incubated with/without ET for various lengths of time. Remarkably, coincident with the increase in intracellular and extracellular cAMP, we noted a coincident decrease in intracellular ATP levels (Fig 4A–4C). Within 1 hour, cellular ATP had decreased by nearly 80%, followed by a transient elevation (likely due to compensatory increase in ATP production) and then a continuous decline (Fig 4C). Intriguingly, ET was also cytotoxic to the cells, and this was even more pronounced than LT-induced cytotoxicity (Fig 4D). To better understand the mechanism of the cytotoxicity of ET vs. LT, we did annexin V plus propidium iodide (PI) staining of the toxin-treated NIH3T3 cells, followed by flow cytometric analysis of the annexin V positive (early apoptotic cells) and PI positive (late apoptotic cells) cells (S3 Fig). While LT did not significantly kill cells, ET treatment resulted in approximately 50% undergoing apoptosis at 24 h and nearly 70% by 48 h (S3 Fig).
A-C. The cells were incubated with/without ET for various lengths of time, followed by the measurements of cellular (A) and extracellular (B) cAMP levels, along with the cellular ATP levels (C). Means ± SD of three independent biological replicates. D. Cytotoxicity of ET and LT to NIH3T3 cells. Cells were incubated with various concentrations of PA in the presence of EF (1 μg/mL) or LF (1 μg/mL) for 48 h, followed by an MTT assay to assess cell viability. Means ± SD of three independent biological replicates.
We next performed ET’s cytotoxicity analysis on a panel of cells including human breast carcinoma MDA-MB-231 cells, mouse melanoma B16-F10 cells, mouse Glioma 261 cells, mouse primary fibroblasts, and CHO cells, and directly compared the cytotoxicity of ET and LT on these cells. Notably, ET exhibited more pronounced cytotoxicity than LT to all these cells (Fig 5A–5E). In parallel, ET rapidly reduced cellular ATP levels (in a dose-dependent manner) in similar patterns as described for NIH3T3 cells (Fig 4C), whereas LT only exhibited minimal effects (Fig 5F–5I). The ET-induced reduction in ATP levels was also verified in an in vivo study; as we noted hepatic ATP levels of ET-challenged mice were significantly decreased (Fig 5J).
A-D. Cytotoxicity of ET and LT to multiple cell lines. MDA-MB-231 (A), B16-F10 (B), Glioma 261 (C) cells, and mouse primary fibroblasts (D) were incubated with various concentrations of PA in the presence of EF (1 μg/mL) or LF (1 μg/mL) for 48 h, followed by an MTT assay to assess the cell viability. E. Cytotoxicity of ET to PR230 (CMG2) cells. The cells were incubated with various concentrations of PA in the presence of EF (1 μg/mL) or LF (1 μg/mL) for 48 h, followed by an MTT assay to assess the cell viability. F-I. Cellular ATP depletion by ET vs. LT in multiple cell lines. MDA-MB-231 (F), B16-F10 (G), Glioma 261 (H) cells, and mouse primary fibroblasts (I) were incubated with ET (0.5 μg/mL or 5 μg/mL) or LT (5 μg/mL) for various lengths of time as indicated, followed by measuring cellular ATP levels. (A-I) Means ± SD of three independent biological replicates. J. The significant reduction in cellular ATP levels in livers from the mice treated with ET. Eighteen hours after challenge with 30 µg ET (I.V.), the mice were euthanized, and livers collected and processed for measuring tissue ATP levels. Means ± SD. Unpaired two-tailed Student’s t-test.
Cellular ATP depletion by ET but not by cholera toxin
Cholera toxin is another bacterial toxin that indirectly increases cellular cAMP through activating the endogenous adenylyl cyclases. Thus, the cAMP-increasing ability of cholera toxin would be limited by the availability of endogenous adenylyl cyclases. As such, in contrast to ET, which itself is a potent adenylate cyclase, cholera toxin could only gradually raise cAMP levels, in much slower pace and lesser extent than did ET (Fig 6A and 6B). As expected, while ET efficiently depleted cellular ATP, cholera toxin could not do so but instead induced up-regulation of ATP levels likely via a compensatory mechanism. Interestingly, forskolin, which also indirectly increases cAMP levels through activating endogenous adenylyl cyclases, could only temporarily decrease cellular ATP, following by compensatory up-regulation of ATP production (Fig 6C).
A. CHO cells were incubated with ET (PA/EF, each 2.5 µg/mL) for various lengths of time as indicated. Then cAMP (from conditioned medium, left panel) and cellular ATP (from cell lysates, middle panel) levels were assessed. Right panel, cell rounding morphological changes (at 24 h) were shown. ***, p < 0.01, when compared to the “0” groups. Unpaired two-tailed Student’s t-test. B, C. CHO cells were incubated with cholera toxin (5 µg/mL) (B) or forskolin (12.5 µM) (C) for various lengths of time as indicated. Then cAMP (from conditioned medium, left panels) and cellular ATP (from cell lysates, middle panels) levels were assessed. Right panels, cell rounding morphological changes (at 24 h) were shown. **, p < 0.05, ***, p < 0.01, when compared to the “0” groups. Unpaired two-tailed Student’s t-test. (A-C) Means ± SD of three independent biological replicates.
Taken together, the ET-induced ATP depletion appears to be an important mechanism underlying ET pathogenesis.
Discussion
Certain protein toxins produced by diverse bacterial pathogens dramatically increase intracellular concentrations of cAMP, suggesting that over-production of this second messenger represents a strategy for bacterial pathogenesis. These toxins include adenylate cyclase toxins, such as anthrax edema toxin, Bordetella pertussis CyaA, and Pseudomonas aeruginosa ExoY, which rapidly convert ATP to cAMP [4,5,37,38]. Although cholera toxin, secreted by Vibrio cholerae, is not an adenylate cyclase, it can ADP-ribosylate the α subunit (Gαs) of trimeric G proteins, and locks Gαs in its GTP-bound, activated form, allowing Gαs to bind and activate endogenous adenylyl cyclases to produce cAMP [39]. Because cholera toxin is the direct mediator of the severe diarrheal symptoms seen in life-threatening cholera, the pathogenic mechanism of this toxin have been well studied [25]. It is widely believed that cholera toxin triggers cAMP-dependent, PKA activation in intestinal epithelial cells. The active PKA then phosphorylates the CFTR chloride channel protein on the surface of intestinal epithelial cells, leading to ATP-mediated efflux of chloride ions, accompanied by secretion of H2O, Na+, K+, and HCO3− into the intestinal lumen. This causes rapid fluid loss from the intestine, resulting in severe dehydration of cholera patients that can culminate in death. As such, a small molecule inhibitor of CFTR was shown to be able to protect mice from acute diarrhea caused by toxigenic V. cholerae [40], attesting to the importance of CFTR in cholera toxin’s pathogenesis.
Since anthrax ET is an adenylate cyclase and can cause intestinal fluid accumulation and tissue edema [26,29], it has been postulated that the cAMP-PKA-CFTR pathway may also mediate the in vivo toxicity of ET [4,5]. Here, indeed, we showed that ET can rapidly increase cellular cAMP and activate PKA. However, although the PKA inhibitor H89 could efficiently block PKA activation, surprisingly, we found that H89 could not protect mice from ET-induced mortality following a systemic challenge. Consistent with these observations, CFTR-null mice were found to be equally susceptible as WT control mice to ET challenges. Further, the potent CFTR inhibitor (CFTRInh-172) also could not protect mice from ET-induced mortality. Therefore, we conclude that ET-induced in vivo toxicity appears to be independent from the cAMP-PKA-CFTR pathway.
Unlike cholera toxin, EF itself is a direct and extremely efficient adenylate cyclase, able to convert 1000–2000 molecules/second of ATP to cAMP [36]. Therefore, in addition to increasing cellular cAMP, we found EF can also rapidly deplete cellular ATP, thereby potentially affecting a variety of biological processes dependent on ATP, such as vesicle trafficking (such as cell junction protein delivery) [41,42]. Unlike ET, cholera toxin increases cAMP through activation of the endogenous adenylyl cyclases with their activities regulated through negative feedback loops [43]. Therefore, the capacity of cholera toxin to convert ATP to cAMP is limited by the availability of endogenous adenylyl cyclases, and that cholera toxin is unlikely to be able to deplete ATP. In fact, CFTR’s action is also ATP-dependent [25]. It is estimated that vesicle trafficking and active transport processes would consume approximately 50% of cellular ATP [41,42]. One characteristic cytotoxic effect of ET to cultured cells is its ability to rapidly induce the cell rounding morphological change, suggesting that the integrity of cell junctions is compromised in ET-treated cells. This is consistent with a study indicated that in the larval wing of Drosophila melanogaster, Rab11-mediated endocytic recycling was disrupted following exogenous expression of EF, resulting in cargo proteins such as cadherins failing to reach intercellular junctions [44]. Therefore, the ET-induced cell rounding, compromised cell junctions, and tissue edema are more likely due to the cellular ATP depletion rather than the cAMP over-production and the cAMP-mediated events. In fact, numerous studies have demonstrated the positive role of induction of cAMP in establishment and stabilization of intercellular junctions, through both the cAMP-activated PKA and Epac pathways [22,30,31,45,46]. Since sufficient levels of cellular ATP are key to various biological processes, here, we did consistently observe more pronounced cytotoxic effects of ET than LT to the cultured cells. Recently, we found that through downregulating the MEK-ERK-cMyc-bioenergetics axis, LT also has inhibitory effects on the cellular ATP production [47]. Therefore, B. anthracis has evolved to use these two exotoxins to synergistically reduce ATP levels via ATP depletion (ET) as well as its production (LT). Indeed, anthrax LT and ET demonstrate potent synergistic in vivo toxicity in challenged experimental animals [48,49].
In summary, our results support the notion that the ET-induced cellular ATP depletion rather than the cAMP overproduction contributes to ET lethality. Our results further suggest that ATP depletion might be an important mechanism underlying pathogenesis of other related adenylate cyclase toxins. However, the present work used a reductionist approach to investigate the role of ET in anthrax pathogenesis, and the results need to be further confirmed in studies using virulent B. anthracis strains.
Methods and materials
Ethics statement
All studies were caried out in with the protocols approved by the Institutional Biosafety Committee (IBC 201800198), the Institutional Animal Care and Use Committee of the University of Pittsburgh (#22030855), and by the National Institute of Allergy and Infectious Diseases, NIH (LPD 1E).
Proteins and reagents
Recombinant PA, EF (with original N-terminal sequence MNE…), and LF (with original N-terminal sequence AGG…) proteins were purified from supernatants of BH500, an avirulent, sporulation-defective, protease-deficient B. anthracis strain, as described previously [50,51]. Cholera toxin (from Vibrio cholera, C8052-.5MG) was obtained from Sigma. H89 (S1582), CFTRInh-172 (S7139), forskolin (S2449), and ESI-09 (S7499) were purchased from Selleckchem. MTT (3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide, 475989-1GM) was obtained from Sigma.
Cells and cytotoxicity assay
All cultured cells were grown at 37oC in a 5% CO2 atmosphere. NIH3T3, B16F10, MDA-MB-231, and Glioma 261 cells were from ATCC, and were cultured in DMEM supplemented with 10% FBS. CHO PR230 and PR230 (CMG2) cells were described in [52], and were cultured in AMEM supplemented with 10% FBS. Primary fibroblasts were isolated from skins of newborn mice as described [53], cultured in DMEM supplemented with 10% FBS. For cytotoxicity assays, cells grown in 96-well plates were treated with various concentrations of the toxins as indicated for 48 h. Cell viabilities were then assessed by MTT as described previously [54], expressed as % of MTT signals of untreated cells. In co-culture experiments, PR230 cells and PR230 (CMG2) cells grown on different cover slips were placed into the same well of a 6-well plate and co-incubated with/without ET for 2 h. Cellular ATP levels were measured using ATPlite 1step kit (PerkinElmer, Boston, MA), following the manufacture’s manual and are expressed as % of ATP levels of untreated cells. cAMP levels in cell lysates and in conditioned medium were measured with the HTRF cAMP HiRange Kit (Cisbio) according to the manufacturer’s protocol.
For assessing the effect of H89 on PKA activation, NIH3T3 cells were serum-starved for 3 h. The cells were then incubated with ET (1 μg/mL) or forskolin (10 μM) with/without H89 (0.1 mM) for various lengths of time as indicated. Then cell lysates were prepared in the modified RIPA lysis buffer containing protease inhibitors as described [54]. Cell lysates were separated on SDS-PAGE gels, transferred onto nitrocellulose membranes, and analyzed by Western blotting using anti-phospho-CREB (Proteintech, 81871-1-RR) and anti-tubulin (Proteintech, 66031) antibodies.
For PI/annexin V staining, NIH3T3 cells treated with or without ET or LT were collected (including the detached cells in medium) and resuspended in 1× binding buffer (BD Biosciences) at a concentration of 1×106 cells/mL. Then, 0.1 mL of the solution was stained with 5 μL each of annexin V (BD Biosciences) and 50 μg/mL PI (Invitrogen) at room temperature for 15 min. The cells were analyzed using a BD FACSCanto Flow Cytometer and percentages of each cell population were obtained.
Animal studies
CFTR-/-/Fabp2-CFTR mice were from the Jackson Laboratory (Strain #: 002364). In ET challenge experiments, 8-10 week old male and female mice with various genotypes were injected with ET (20-50 μg) in 0.5 ml PBS intraperitoneally or in 0.2 ml PBS intravenously. We used the following criteria to score mouse disease progression induced by ET: 0: healthy mouse; 1: slight ruffled coat but no problem in running around cage; 2: ruffled coat and decrease in activity; 3: ruffled coat, hunched posture and little movement; 3.5: moribund; 4: found dead. All toxin-challenged or infected mice were monitored twice daily for two weeks post challenge for signs of malaise or mortality. For testing PKA inhibitor H89, mice were administered with 30 µg ET (I.V.) or 30 µg ET (I.V.) plus 0.5 mg H89 (in 0.5 mL PBS, I.P.). For CFTR inhibitor CFTRinh-172 experiment, mice were administered (I.P.) twice at 24 h and 1 h with PBS or 50 μg CFTRinh-172 prior to ET injection (30 µg in 0.2 mL PBS, I.V.).
For ET footpad skin edema model, mice were injected intradermally in their right and left rear footpads with H89 (20 µg in 20 µl PBS) or CFTRinh-172 (12 µg in 20 µl 20% DMSO), and rear right footpads were injected with same volume of vehicles. Two hours later, the mice were injected intradermally in their right and left rear footpads with 0.2 μg ET or 0.2 μg CT in 20 μL PBS. The thicknesses of footpads (dorsal/plantar) were measured with a digital caliper before treatment, and at the indicated time points after ET or CT injection, expressed as % of footpad sizes before any injection.
To extract and measure cAMP levels of the tissues/organs from ET challenged mice, 18 h after challenge with 30 µg ET (I.V.), mice were euthanized, and blood, liver, heart, kidneys collected. cAMP levels from sera prepared from blood samples were measured with the HTRF cAMP HiRange Kit (Cisbio) according to the manufacturer’s protocol. To prepare tissue extracts for cAMP and ATP measurements, we followed the procedure previously described [55]. Briefly, ~ 0.3 g each tissue sample was homogenized in 3 mL of ice-cold phenol-TE on ice. In 1 mL homogenate, 200 μL of chloroform and 150 μL of deionized water were added. The samples were then mixed well and centrifuged at 10,000 x g for 5 min, after which the upper aqueous phase (50 μL) was collected. The tissue extracts were then diluted for cAMP measurements using with the HTRF cAMP HiRange Kit (Cisbio). The liver extracts were also used to measure the ATP levels using ATPliteTM 1step (PerkinElmer, 6016731). ATP levels of the livers were normalized by the tissue wet weight, presented as percentages of heart ATP levels from the untreated mice.
Supporting information
S1 Fig. Epac-independent in vivo toxicity of anthrax edema toxin.
A. Mice were injected with Epac inhibitor ESI-09 (200 µg, I.P.) 2 h prior to challenge with 50 µg ET (I.P.), and monitored for survival as indicated. Log-rank test for survival comparison. B. Mice were injected with Epac inhibitor ESI-09 (200 µg, I.P.) 2 h prior to injection with 0.25 μg ET (in 20 µl PBS) into their rear footpads. The thicknesses (dorsal/plantar) of footpads were measured with a digital caliper at the time points indicated after ET injection. Edema was expressed as % of footpad sizes before ET injection. Of note, ESI-09 could not diminish the ET-induced in vivo toxicity. Means ± SD. Unpaired two-tailed Student’s t-test.
https://doi.org/10.1371/journal.ppat.1013017.s001
(EPS)
S2 Fig. Amelioration of ET- and CT-induced footpad edema by PAK or CFTR inhibition.
A, B. ET-induced footpad edema in mice. Mice were injected with 0.2 μg ET (in 20 µl PBS) into their rear right and left footpads, two hours after the rear right footpads were pre-injected with H89 (20 µg in 20 µl PBS) (C) or CFTRinh-172 (12 µg in 20 µl 20% DMSO) (D), with their rear left footpads were injected with same volume of vehicle. The thicknesses of footpads were measured with a digital caliper before injection and at the time points indicated after ET injection, expressed as % of footpad sizes before injection. C, D. CT-induced footpad edema in mice. Mice were injected with 0.2 μg CT (in 20 µl PBS) into their rear right and left footpads two hours after their rear right footpads were injected with H89 (20 µg in 20 µl PBS) (E) or CFTRinh-172 (12 µg in 20 µl 20% DMSO) (F), and their left rear footpads were injected with same volume of vehicle. The thicknesses of footpads were measured before injection and at the indicated time points after CT injection, expressed as % of footpad sizes before injection. Of note, H89 and CFTRinh-172 could ameliorate the CT-induced footpad edema. (A-D) Means ± SD of independent biological repeats (each symbol represents one mouse). Unpaired two-tailed Student’s t-test.
https://doi.org/10.1371/journal.ppat.1013017.s002
(EPS)
S3 Fig. Apoptotic cell death induced by ET.
A. NIH 3T3 cells were incubated with or without ET (5 µg/mL), or LT (5 µg/mL) for various lengths of time, followed by flow cytometry analyses of the cells after staining with propidium iodide (PI) and annexin V. B. Images of the NIH 3T3 cells treated with/without of ET (5 µg/mL), LT (5 µg/mL), or PA + FP59 (0.1 µg/mL PA + 0.1 µg/mL FP59) for 48 h.
https://doi.org/10.1371/journal.ppat.1013017.s003
(EPS)
References
- 1. Hudson MJ, Beyer W, Bohm R, Fasanella A, Garofolo G, Golinski R, et al. Bacillus anthracis: balancing innocent research with dual-use potential. Int J Med Microbiol. 2008;298:345–64.
- 2.
Leppla SH. Chapter 281 - anthrax lethal factor. In: Rawlings ND, Salvesen GS, editors. Handbook of proteolytic enzymes; 2013. p. 1257–61. Academic Press.
- 3. Fouet A, Mock M Regulatory networks for virulence and persistence of Bacillus anthracis. Curr Opin Microbiol. 2006.
- 4. Moayeri M, Leppla SH, Vrentas C, Pomerantsev AP, Liu S. Anthrax Pathogenesis. Annu Rev Microbiol. 2015;69:185–208. pmid:26195305
- 5. Liu S, Moayeri M, Leppla SH. Anthrax lethal and edema toxins in anthrax pathogenesis. Trends Microbiol. 2014;22(6):317–25. pmid:24684968
- 6. Duesbery NS, Webb CP, Leppla SH, Gordon VM, Klimpel KR, Copeland TD, et al. Proteolytic inactivation of MAP-kinase-kinase by anthrax lethal factor. Science. 1998;280(5364):734–7. pmid:9563949
- 7. Lee C-S, Dykema KJ, Hawkins DM, Cherba DM, Webb CP, Furge KA, et al. MEK2 is sufficient but not necessary for proliferation and anchorage-independent growth of SK-MEL-28 melanoma cells. PLoS One. 2011;6(2):e17165. pmid:21365009
- 8. Vitale G, Pellizzari R, Recchi C, Napolitani G, Mock M, Montecucco C. Anthrax lethal factor cleaves the N-terminus of MAPKKs and induces tyrosine/threonine phosphorylation of MAPKs in cultured macrophages. Biochem Biophys Res Commun. 1998;248:706–11.
- 9. Vitale G, Bernardi L, Napolitani G, Mock M, Montecucco C. Susceptibility of mitogen-activated protein kinase kinase family members to proteolysis by anthrax lethal factor. Biochem J. 2000;352 Pt 3(Pt 3):739–45. pmid:11104681
- 10. Van Hauwermeiren F, Van Opdenbosch N, Van Gorp H, de Vasconcelos N, van Loo G, Vandenabeele P, et al. Bacillus anthracis induces NLRP3 inflammasome activation and caspase-8-mediated apoptosis of macrophages to promote lethal anthrax. Proc Natl Acad Sci U S A. 2022;119(2):e2116415119. pmid:34996874
- 11. Liu W, Nestorovich EM. Anthrax toxin channel: What we know based on over 30 years of research. Biochim Biophys Acta Biomembr. 2021;1863:183715
- 12. Liu S, Liu J, Ma Q, Cao L, Fattah RJ, Yu Z, et al. Solid tumor therapy by selectively targeting stromal endothelial cells. Proc Natl Acad Sci U S A. 2016;113(28):E4079-87. pmid:27357689
- 13. Schafer JM, Peters DE, Morley T, Liu S, Molinolo AA, Leppla SH, et al. Efficient targeting of head and neck squamous cell carcinoma by systemic administration of a dual uPA and MMP-activated engineered anthrax toxin. PLoS One. 2011;6(5):e20532. pmid:21655226
- 14. Alfano RW, Leppla SH, Liu S, Bugge TH, Ortiz JM, Lairmore TC, et al. Inhibition of tumor angiogenesis by the matrix metalloproteinase-activated anthrax lethal toxin in an orthotopic model of anaplastic thyroid carcinoma. Mol Cancer Ther. 2010;9(1):190–201. pmid:20053778
- 15. Zuo Z, Liu J, Sun Z, Silverstein R, Zou M, Finkel T, et al. A potent tumor-selective ERK pathway inactivator with high therapeutic index. PNAS Nexus. 2022;1(3):pgac104. pmid:35899070
- 16. Zuo Z, Liu J, Sun Z, Cheng Y-W, Ewing M, Bugge TH, et al. ERK and c-Myc signaling in host-derived tumor endothelial cells is essential for solid tumor growth. Proc Natl Acad Sci U S A. 2023;120(1):e2211927120. pmid:36574698
- 17. Pezard C, Berche P, Mock M. Contribution of individual toxin components to virulence of Bacillus anthracis. Infect Immun. 1991;59(10):3472–7. pmid:1910002
- 18. Pezard C, Duflot E, Mock M. Construction of Bacillus anthracis mutant strains producing a single toxin component. J Gen Microbiol. 1993;139(10):2459–63. pmid:8254316
- 19. Leppla SH. Anthrax toxin edema factor: a bacterial adenylate cyclase that increases cyclic AMP concentrations of eukaryotic cells. Proc Natl Acad Sci U S A. 1982;79(10):3162–6. pmid:6285339
- 20. Steinberg GR, Carling D. AMP-activated protein kinase: the current landscape for drug development. Nat Rev Drug Discov. 2019;18(7):527–51. pmid:30867601
- 21. Murray AJ. Pharmacological PKA inhibition: all may not be what it seems. Sci Signal. 2008;1(22):re4. pmid:18523239
- 22. Kooistra MRH, Corada M, Dejana E, Bos JL. Epac1 regulates integrity of endothelial cell junctions through VE-cadherin. FEBS Lett. 2005;579(22):4966–72. pmid:16115630
- 23. Fukuhara S, Sakurai A, Sano H, Yamagishi A, Somekawa S, Takakura N, et al. Cyclic AMP potentiates vascular endothelial cadherin-mediated cell-cell contact to enhance endothelial barrier function through an Epac-Rap1 signaling pathway. Mol Cell Biol. 2005;25(1):136–46. pmid:15601837
- 24. Riordan JR, Rommens JM, Kerem B, Alon N, Rozmahel R, Grzelczak Z, et al. Identification of the cystic fibrosis gene: cloning and characterization of complementary DNA. Science. 1989;245(4922):1066–73. pmid:2475911
- 25. Thiagarajah JR, Donowitz M, Verkman AS. Secretory diarrhoea: mechanisms and emerging therapies. Nat Rev Gastroenterol Hepatol. 2015;12(8):446–57. pmid:26122478
- 26. Firoved AM, Miller GF, Moayeri M, Kakkar R, Shen Y, Wiggins JF, et al. Bacillus anthracis edema toxin causes extensive tissue lesions and rapid lethality in mice. Am J Pathol. 2005;167(5):1309–20. pmid:16251415
- 27. Kuo SR, Willingham MC, Bour SH, Andreas EA, Park SK, Jackson C, et al. Anthrax toxin-induced shock in rats is associated with pulmonary edema and hemorrhage. Microb Pathog. 2008;44:467–72.
- 28. Leysath CE, Phillips DD, Crown D, Fattah RJ, Moayeri M, Leppla SH. Anthrax edema factor toxicity is strongly mediated by the N-end rule. PLoS One. 2013;8(8):e74474. pmid:24015319
- 29. Liu S, Zhang Y, Moayeri M, Liu J, Crown D, Fattah RJ, et al. Key tissue targets responsible for anthrax-toxin-induced lethality. Nature. 2013;501(7465):63–8. pmid:23995686
- 30. Lorenowicz MJ, Fernandez-Borja M, Kooistra MRH, Bos JL, Hordijk PL. PKA and Epac1 regulate endothelial integrity and migration through parallel and independent pathways. Eur J Cell Biol. 2008;87(10):779–92. pmid:18635287
- 31. Pannekoek W-J, Vliem MJ, Bos JL. Multiple Rap1 effectors control Epac1-mediated tightening of endothelial junctions. Small GTPases. 2020;11(5):346–53. pmid:29388865
- 32. Alasbahi RH, Melzig MF. Forskolin and derivatives as tools for studying the role of cAMP. Pharmazie. 2012;67(1):5–13. pmid:22393824
- 33. Young PG, Levring J, Fiedorczuk K, Blanchard SC, Chen J. Structural basis for CFTR inhibition by CFTR(inh)-172. Proc Natl Acad Sci U S A. 2024;121:e2316675121.
- 34. Snouwaert JN, Brigman KK, Latour AM, Malouf NN, Boucher RC, Smithies O, et al. An animal model for cystic fibrosis made by gene targeting. Science. 1992;257(5073):1083–8. pmid:1380723
- 35. Zhou L, Dey CR, Wert SE, DuVall MD, Frizzell RA, Whitsett JA. Correction of lethal intestinal defect in a mouse model of cystic fibrosis by human CFTR. Science. 1994;266(5191):1705–8. pmid:7527588
- 36. Shen Y, Lee Y-S, Soelaiman S, Bergson P, Lu D, Chen A, et al. Physiological calcium concentrations regulate calmodulin binding and catalysis of adenylyl cyclase exotoxins. EMBO J. 2002;21(24):6721–32. pmid:12485993
- 37. Melvin JA, Scheller EV, Miller JF, Cotter PA. Bordetella pertussis pathogenesis: current and future challenges. Nat Rev Microbiol. 2014;12(4):274–88. pmid:24608338
- 38. Yahr TL, Vallis AJ, Hancock MK, Barbieri JT, Frank DW. ExoY, an adenylate cyclase secreted by the Pseudomonas aeruginosa type III system. Proc Natl Acad Sci U S A. 1998;95(23):13899–904. pmid:9811898
- 39. White C, Bader C, Teter K. The manipulation of cell signaling and host cell biology by cholera toxin. Cell Signal. 2022;100:110489. pmid:36216164
- 40. Rivera-Chávez F, Meader BT, Akosman S, Koprivica V, Mekalanos JJ. A potent inhibitor of the cystic fibrosis transmembrane conductance regulator blocks disease and morbidity due to toxigenic Vibrio cholerae. Toxins (Basel). 2022;14(3):225. pmid:35324722
- 41. Puurand M, Tepp K, Klepinin A, Klepinina L, Shevchuk I, Kaambre T. Intracellular energy-transfer networks and high-resolution respirometry: a convenient approach for studying their function. Int J Mol Sci. 2018;19.
- 42. Cui L, Li H, Xi Y, Hu Q, Liu H, Fan J, et al. Vesicle trafficking and vesicle fusion: mechanisms, biological functions, and their implications for potential disease therapy. Mol Biomed. 2022;3(1):29. pmid:36129576
- 43. Hanoune J, Pouille Y, Tzavara E, Shen T, Lipskaya L, Miyamoto N, et al. Adenylyl cyclases: structure, regulation and function in an enzyme superfamily. Mol Cell Endocrinol. 1997;128(1–2):179–94. pmid:9140089
- 44. Guichard A, McGillivray SM, Cruz-Moreno B, van Sorge NM, Nizet V, Bier E. Anthrax toxins cooperatively inhibit endocytic recycling by the Rab11/Sec15 exocyst. Nature. 2010;467(7317):854–8. pmid:20944747
- 45. Sugawara K, Shibasaki T, Mizoguchi A, Saito T, Seino S. Rab11 and its effector Rip11 participate in regulation of insulin granule exocytosis. Genes Cells. 2009;14(4):445–56. pmid:19335615
- 46. Park SW, Schonhoff CM, Webster CRL, Anwer MS. Rab11, but not Rab4, facilitates cyclic AMP- and tauroursodeoxycholate-induced MRP2 translocation to the plasma membrane. Am J Physiol Gastrointest Liver Physiol. 2014;307(8):G863-70. pmid:25190474
- 47. Liu J, Zuo Z, Fattah R, Finkel T, Leppla SH, Liu S. Anthrax lethal toxin exerts potent metabolic inhibition of the cardiovascular system. mBio. 2024:e0216024.
- 48. Cui X, Li Y, Li X, Laird MW, Subramanian M, Moayeri M, et al. Bacillus anthracis edema and lethal toxin have different hemodynamic effects but function together to worsen shock and outcome in a rat model. J Infect Dis. 2007;195(4):572–80. pmid:17230417
- 49. Firoved AM, Moayeri M, Wiggins JF, Shen Y, Tang WJ, Leppla SH. Anthrax edema toxin sensitizes DBA/2J mice to lethal toxin. Infect Immun. 2007;75:2120–5.
- 50. Gupta PK, Moayeri M, Crown D, Fattah RJ, Leppla SH. Role of N-terminal amino acids in the potency of anthrax lethal factor. PLoS One. 2008;3(9):e3130. pmid:18769623
- 51. Pomerantsev AP, Pomerantseva OM, Moayeri M, Fattah R, Tallant C, Leppla SH. A Bacillus anthracis strain deleted for six proteases serves as an effective host for production of recombinant proteins. Protein Expr Purif. 2011;80(1):80–90. pmid:21827967
- 52. Liu S, Leung HJ, Leppla SH. Characterization of the interaction between anthrax toxin and its cellular receptors. Cell Microbiol. 2007;9(4):977–87. pmid:17381430
- 53. Reiisi S, Esmaeili F, Shirazi A. Isolation, culture and identification of epidermal stem cells from newborn mouse skin. In Vitro Cell Dev Biol Anim. 2010;46(1):54–9. pmid:19915936
- 54. Liu S, Leppla SH. Cell surface tumor endothelium marker 8 cytoplasmic tail-independent anthrax toxin binding, proteolytic processing, oligomer formation, and internalization. J Biol Chem. 2003;278(7):5227–34. pmid:12468536
- 55. Chida J, Kido H. Extraction and quantification of adenosine triphosphate in mammalian tissues and cells. Methods Mol Biol. 2014;1098:21–32. pmid:24166365