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Sepsis is a life-threatening condition resulting from pathogen infection and characterized by organ dysfunction. Programmed cell death (PCD) during sepsis has been associated with the development of multiple organ dysfunction syndrome (MODS), impacting various physiological systems including respiratory, cardiovascular, renal, neurological, hematological, hepatic, and intestinal systems. It is well-established that pathogen infections lead to immune dysregulation, which subsequently contributes to MODS in sepsis. However, recent evidence suggests that sepsis-related opportunistic pathogens can directly induce organ failure by promoting PCD in parenchymal cells of each affected organ. This study provides an overview of PCD in damaged organ and the induction of PCD in host parenchymal cells by opportunistic pathogens, proposing innovative strategies for preventing organ failure in sepsis.
Xunyao Wu and Bin Du have contributed equally to this work.
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Introduction
Sepsis is a life-threatening condition characterized by organ dysfunction due to dysregulated immune responses to infections, and it is associated with high rates of mortality and morbidity [1, 2]. In the intensive care unit (ICU), sepsis is particularly significant, affecting approximately 20–30% of patients, with 25–40% of septic patients dying before hospital discharge [3, 4]. Uncontrolled inflammation in sepsis leads to multiple organ dysfunction syndrome (MODS), which most frequently affects the respiratory, cardiovascular, renal, neurological, hematological, hepatic and intestinal systems [5‐14]. The mortality rate for septic patients diagnosed with MODS can be as high as 28–56% [15‐17].
Uncontrolled cell death during sepsis has been implicated in the development of multiple organ dysfunction syndrome (MODS). Excessive physical or chemical stressors in pathological conditions can induce undesired cellular death, leading to compromised organ functionality [18, 19]. The term "apoptosis" introduced by Kerr in 1972 [20], describes a form of programmed cell death (PCD) that occurs without initiating an immune response. Subsequent research has identified additional forms of cell death, such as apoptosis, necroptosis, autophagy, pyroptosis, and ferroptosis. Initially classified into three distinct types, these forms were redefined under the updated framework established by the Committee on Cell Death Nomenclature in 2018 [21]. PCD can be categorized based on caspase involvement: caspase-dependent cell death, which includes apoptosis and pyroptosis, and non-caspase-dependent cell death, which includes necroptosis, autophagy, and ferroptosis [21]. Various cell death pathways are closely related to organ function impairment in sepsis. Targeting cell death has emerged as a novel therapeutic approach for sepsis.
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Sepsis-induced organ failure is a critical condition often driven by pathogen infections. Bacterial infections, including Staphylococcus aureus (S. aureus), Escherichia coli (E. coli), Klebsiella pneumoniae (K. pneumoniae), and Pseudomonas aeruginosa (P. aeruginosa) [22, 23], as well as viral infections like the novel coronavirus [24] and influenza virus [25], are prevalent in ICU, leading to severe and potentially life-threatening conditions. It is well-established that pathogen infections cause immune dysregulation, which subsequently contributes to MODS in sepsis. However, recent evidence suggests that pathogen can directly induce organ failure by promoting PCD in parenchymal cells of each affected organ. This study summarizes the mechanisms of PCD in damaged organs and the induction of PCD in host cells by opportunistic pathogens. The objective is to propose an innovative approach to protect against organ failure in sepsis.
PCD in sepsis-induced MODS
PCD plays a critical role in sepsis-related MODS, involving pathways such as apoptosis, necroptosis, pyroptosis, autophagy and ferroptosis (Fig. 1) (Table 1).
Fig. 1
In sepsis, cell death mechanisms such as apoptosis, necroptosis, pyroptosis, autophagy and ferroptosis play a direct role in organ dysfunction. Apoptosis and autophagy typically help to remove damaged cells and prevent the spread of inflammation. However, during sepsis, excessive apoptosis and damaged autophagy can result in the loss of many functional and unable to remove damaged cells, thereby weakening organ function. Necroptosis, pyroptosis and ferroptosis may dysregulate inflammatory factors, exacerbating the inflammatory response and causing further tissue and organ damage. Thus, the imbalance of cell death pathways is a critical pathological basis for organ dysfunction in sepsis. The major organ systems that are clinically monitored in patients with sepsis was adapted from Lelubre and Vincent 14]
Apoptosis, a form of PCD, involves two main pathways: intrinsic and extrinsic. The intrinsic pathway, activated by cellular stress like DNA damage or hypoxia, leads to the oligomerization of B-cell lymphoma-2 (BCL-2) family of proteins [26], crucial for mitochondrial outer membrane permeabilization (MOMP) and cytochrome C release. Cytochrome C then binds to apoptotic protease activating factor-1 (APAF1) and caspase-9 to form the apoptosome complex. The extrinsic pathway is initiated by death ligands (such as FasL, TNF, or TRAIL) binding to death receptors (Fas, TNFR1, TNFR2, and TRAIL receptors DR4 and DR5), triggering the recruitment of Fas-associated death domain protein (FADD) and the activation of caspase-8/10 [27, 28]. Both pathways converge to activate executioner caspases 3/6/7, which cleave various intracellular proteins, leading to cell contraction, nuclear fragmentation, and membrane vesicle formation [27, 28].
Increased intestinal epithelial cells (IECs) apoptosis was observed in septic patients compared with non-septic patients by an autopsy study. Mechanistically, increased IECs apoptosis might lead to decreased barrier function, followed by bacterial translocation to systemic circulation or mesenteric lymphatic vessels that contribute to MODS [29]. Overexpression of Bcl-2 in the IECs resulted in a significant reduction of cell apoptosis, giving cecal ligation and puncture (CLP)-induced septic mice a survival advantage [30, 31]. Fas ligand and Fas are highly upregulated in epithelial and endothelial cells of sepsis-related acute respiratory distress syndrome (ARDS) and Fas- and Fas ligand-deficient mice showed marked protection from lung inflammation and apoptosis. Fas-associated death domain (FADD) is an adapter molecule that recruits caspase-8 to Fas. Systemic small interfering RNA (siRNA) targeting FADD can prevent acute lung injury in CLP-induced septic mice [32]. The levels of BCL-2-associated X protein (BAX) in pulmonary microvascular endothelial cells are positively correlated with high mortality in sepsis-induced acute lung injury (S-ALI) patients [33]. What’s more, septic murine pulmonary microvascular dysfunction is mediated through caspase-dependent apoptosis and nitric oxide synthase (iNOS)/NADPH-oxidase dependent signaling [34]. Cardiac dysfunction is an important component of sepsis-related MODS. Enhanced cardiomyocyte apoptosis accompanied with increased reactive oxygen species (ROS) are found in sepsis-related myocardial systolic dysfunction [35]. Lipopolysaccharide (LPS) challenge significantly increased hepatocyte apoptosis, caspase 3 activity, and BAX level while suppressing Bcl-2 expression in liver tissues [36]. Apoptosis in renal tubular and endothelial cells during sepsis-induced acute kidney injury (S-AKI) is notable. Interestingly, plasma from S-AKI patients induced apoptosis in cultured tubular cells and podocytes [36].
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Necroptosis
Necroptosis is a caspase-independent PCD and can be triggered by ligand binding to pathogens, pattern-recognition receptors, and tumor necrosis factor (TNF) family death domain receptors [37]. The formation of the key component, the necrosome, involves FADD, receptor-interacting kinase 1 (RIPK1), receptor-interacting kinase 3 (RIPK3), and caspase-8. In cells stimulated by TNF, inactivated caspase-8 leads to autophosphorylation of RIPK1-RIPK3, which phosphorylates and promotes oligomerization of mixed lineage kinase domain-like protein (MLKL). Phosphorylated MLKL forms oligomers that induce pore swelling and rupture [38].
LPS treatment caused neuronal necroptosis and synaptic damage, which was alleviated by direct inhibition of necroptosis [39]. A previous study demonstrated that genetic deletion of RIPK3 was protective of systemic inflammatory response syndrome and improved survival outcomes in a CLP model. Deletion of RIPK3 or MLKL protected against MODS in septic mice [40]. In sepsis, EGFR activation triggers a significant increase in RIP1 phosphorylation and its binding to receptor-interacting protein 3 (RIP3), leading to NF-κB/MAPK-mediated inflammation and RIPK3-dependent necroptosis in lung epithelial cells [41]. Elevated RIPK3 levels in plasma were associated with ARDS in septic patients, and inhibition of RIPK1 by necrostatin-1 decreased lung injury in neonatal septic mice [42, 43]. RIPK3 and MLKL were implicated in sepsis-associated cardiomyopathy. Peroxisome proliferator-activated receptor (PPAR)-gamma agonists showed therapeutic potential by down-regulating these proteins. RIPK1, RIPK3, and MLKL were upregulated in sepsis-induced hepatic injury, and a chemical inhibitor necrostatin-1 (Nec-1) or RIP1 knockdown accelerated septic liver injury [44‐46]. In a piglet model of LPS-induced sepsis, expressions of necroptosis components (RIP1, RIP3, and MLKL) and mitochondrial proteins (PGAM5 and DRP1) were increased in the liver in a time-dependent manner, followed by hepatic inflammation, morphological damage, and dysfunction [47]. Intestinal injury in sepsis was linked to increased necroptosis and impaired barrier function [48, 49]. Protein expressions of RIP1, RIP3, and MLKL were increased in jejunum of LPS-challenged piglets, which was accompanied by the impairment of jejunal morphology, and digestive and barrier function [50]. RIPK3 knockout in HT-29 cells boosts stimulator of interferon genes (STING) autophagy, reducing STING signaling. Normally, RIPK3 inhibits autophagy during STING activation. MLKL affects STING signaling in two ways: its deficiency enhances signaling, while blocking its pore formation reduces it. When MLKL's necroptotic function is blocked, it binds to STING and is secreted, limiting TBK1 and IRF3 recruitment. Targeting necroptotic signaling ameliorates STING activation during DMXAA-induced intestinal injury and sepsis [51].
Pyroptosis
Pyroptosis is an inflammatory form of PCD marked by cell swelling and the release of inflammatory cytokines, notably interleukin-1β (IL-1β) and IL-18. This process is typically initiated by inflammasomes, such as NLPR1, NLPR3, NLRC4, NLRP6, NLRP12, and AIM2, which act as sensors for a range of threats including bacterial toxins, viruses, parasites, fungi, metabolites, drugs, and ROS. Detection of these dangers leads to the formation of the inflammasome complex and the activation of caspase-1. Caspase-1 then cleaves the pyroptosis executor protein Gasdermin D (GSDMD), releasing its N-terminal domain to form membrane pores. In contrast, the non-canonical pathway circumvents inflammasome assembly and is directly triggered by caspase-4 and caspase-5 in humans, or caspase-11 in mice, in response to intracellular LPS [52‐54].
In a rat CLP model, elevated NLRP3, caspase-1, and inflammatory factors were observed [55]. This activation is associated with cognitive dysfunction, blood–brain barrier (BBB) disruption, and brain structural damage. Caspase-1 inhibition has shown to improve cognitive behavior and mitigate brain damage [56]. Lung damage was aggravated through cold-induced RNA binding protein (CIRP)-induced the NLRP3 inflammasome assembly and activation in pulmonary vascular endothelial cells (MLVECs) [57]. GSDMD-NT expression was elevated in sepsis models and GSDMD knockout mice showed higher survival and improved cardiac dysfunction, indicating the involvement of cardiomyocytes pyroptosis in sepsis-related cardiomyopathy [58]. Overexpression of PIK3CG, a well-established target in septic myocardial injury, could exacerbate pyroptosis and worsen myocardial function through modulating the NLRP3/GSDMD signaling pathway. CLP-induced myocardial pyroptosis could be reversed by AS-604850, a PIK3CG inhibitor [59]. Abnormal processing of calcium (Ca2+) in cardiomyocytes is associated with pyroptosis. Inositol 1,4,5-triphosphate receptor type 2 (IP3R2) is a Ca2+ release channel in the ER. IP3R2 promotes NLRP3-mediated pyroptosis by regulating ER Ca2+ release [60]. Liver dysfunction in sepsis is closely linked to caspase-4/5/11 mediated pyroptosis. Hepatocyte release of high mobility group protein 1 (HMGB1) is required for caspase-11-dependent pyroptosis in endotoxemia and bacterial sepsis. HMGB1 to bind LPS and internalize it into the lysosomes of endothelial cells via the receptor for advanced glycosylation end products (RAGE). Subsequently, HMGB1 permeates the phospholipid bilayer in the acidic environment of the lysosome. This leads to leakage of LPS into the cytoplasm and activation of caspase-11 [61, 62]. Caspase-11-induced pyroptosis in renal tubular epithelial cells is a key event in S-AKI. STING was significantly activated in LPS-induced tubule cells. STING triggers ERS, leading to excessive production of mitochondrial ROS (mtROS) and the activation of NLRP3, which causes pyroptosis of renal tubular cells [63]. Tissue inhibitors of metalloproteinase 2 (TIMP2) promotes the ubiquitination and autophagy dependent degradation of NLRP3. TIMP2 knockout mice showed more severe kidney damage and elevated levels of pyroptosis markers than wild-type mice [64]. In disseminated intravascular coagulation (DIC), caspase-1 activation and GSDMD pore formation are critical, with inflammasome activation triggering systemic coagulation [65]. In patients with sepsis, plasma IL-1α and IL-1β concentrations correlate with phosphatidylserine exposure and DIC scores in peripheral leucocytes [66]. Inhibiting the STING-GSDMD-F3 pathway significantly improves survival in sepsis models [67].
Autophagy
Autophagy is a cellular mechanism that maintains homeostasis by degrading damaged or unnecessary components via lysosomes. Endoplasmic reticulum (ER) stress triggers the unfolded protein response (UPR), which in turn induces autophagy to remove misfolded proteins [68]. Autophagosomes form with the assistance of ATG proteins (ATG5, ATG12, and LC3) and subsequently fuse with lysosomes to create autolysosomes [69]. Mitophagy, a specialized form of autophagy, targets damaged mitochondria, which are marked by PINK1 and Parkin, and are similarly degraded through fusion with lysosomes [70]. Additionally, there is the chaperone-mediated autophagy mechanism, in which specific proteins are directly translocated across the lysosomal membrane for degradation via key proteins such as lysosomal-associated membrane protein 2A (Lamp2A) [71].
In septic patients and animal models, dysregulated autophagy was found in MODS. In sepsis-associated murine encephalopathy, reduced expression of Lamp2A in hippocampal neurons is found and is associated with cognitive impairment [72]. Accumulation of microtubule-associated protein light chain 3 (LC3)-II can alleviate S-ALI by enhancing autophagy [73]. Autophagosome formation increases in the left ventricle 24 h post-CLP, but autophagosome degradation decreases. This is consistent with morphological findings showing an increase in autophagosomes in septic hearts, though autolysosomes are rare, indicating incomplete autophagy. Inducing full activation of autophagy with rapamycin restores CLP-induced cardiac dysfunction [74]. Beclin-1 and parkinson disease protein 2 (PARK2) play crucial roles in maintaining heart function and promoting mitophagy during sepsis and their deficiency can impair recovery [75]. Additionally, p21 regulates cardiomyocyte autophagy, with its downregulation exacerbating inflammation and oxidative stress [76]. Hepatic autophagy inactivation during critical illness acutely worsened mitochondrial dysfunction, partially prohibiting acute UPR activation and aggravated liver damage [77]. Increased hepatocellular injury in sepsis was observed when autophagic signaling is abrogated by inhibition of heme oxygenase-1 (HO-1) activity. Aggregated CLP-induced liver injury was found by decreasing the autophagic flux, restoring both autophagosome and autolysosome formation through down-expression of homeodomain-interacting protein kinase 2 (HIPK2) expression [78‐80]. IL-22 pre-administration activates autophagy in hepatocytes and protects mice against septic liver injury partially related to ATF4-ATG7 signaling pathway [81]. Recent studies have confirmed that mitophagy in renal tubular and endothelial cells is known to exert protective effects against S-AKI. The PTEN-induced kinase 1 (PINK1)/PARK2 pathways protect against S-AKI by promoting mitophagy [82]. Upregulated SIRT1, a deacetylase, could deacetylate certain autophagy-related proteins (Beclin1, Atg5, Atg7 and LC3) and protect against S-AKI [83].
Ferroptosis
Ferroptosis is characterized by iron-dependent lipid peroxidation and oxidative stress. Failure of peroxidase (GPX4) inactivation to clear the increase of lipid peroxides or/and the elevated conversion of polyunsaturated fatty acids (PUFAs) to phospholipid hydroperoxides (PLOOHs) can induce iron deposition, leading to iron overload and subsequent production of mitochondrial ROS, ultimately resulting in iron poisoning and ferroptosis [84, 85].
Decreased levels of superoxide dismutase (SOD) were observed in the serum of SAE patients and in the hippocampus of septic mice indicate neuronal ferroptosis [86]. CLP-rats displayed learning and memory impairment, along with heightened ROS, Fe2+, and MDA levels in hippocampal neurons, diminished GSH and SOD levels, and down-regulated ferroptosis-related proteins (GPX4 and SLC7A11) [87]. S-ALI is linked to ferroptosis, which regulated through the Kelch-like ECH-associated protein 1-NF-E2-related factor 2 (KEAP1-NRF2) oxidative stress pathway [88]. Sepsis increases cardiac iron content and lipid peroxidation while reducing glutathione and GPX4 levels, implicating ferroptosis in septic cardiomyopathy and cardiac dysfunction [88‐90]. In addition, ferroptosis has been shown to have a role in an LPS-induced model of septic cardiomyopathy. An LPS-induced increase in sideroflexin 1 expression in the cardiac mitochondrial membrane can increase the production of mtROS, leading to ferroptosis [91]. Hepatic ferroptosis during sepsis was accompanied by G protein-coupled receptor 116 (GPR116) upregulation. GPR116 aggravated mitochondrial damage and lipid peroxidation in hepatocytes by inhibiting system Xc-/GSH/GPX4 [92]. Renal tubular epithelial cell ferroptosis in S-AKI is indicated by increased lipid peroxidation and decreased GPX4 and glutathione (GSH) levels [93]. The activation of lipophagy promoted renal tubular epithelial cell ferroptosis and renal damage in S-AKI were confirmed by constructing S-AKI models in vitro and in vivo [94]. Ferroptosis of kidney epithelial cells may contribute to the severity of AKI and reduce chances for recovery. Oxygenation of polyunsaturated phosphatidylethanolamines increased in AKI patients with or without sepsis and in AKI patients who did not recover renal function [95].
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Regulation of PCD pathways in host parenchymal cells by sepsis-related major opportunistic pathogens
Recent evidence suggests that sepsis-related pathogens can directly induce MODS by promoting PCD in parenchymal cells of each affected organ. Infections by pathogens induce PCD through the disruption of mitochondrial function, the activation of inflammasomes, and the stimulation of receptor-interacting protein kinases, which culminate in the rupture of the cell membrane (Figs. 2 and 3) (Table 2). These molecular mechanisms are involved in the high mortality rate associated with sepsis and MODS.
Fig. 2
Opportunistic pathogenic bacteria elicit the death of organ parenchymal cells by triggering and fine-tuning cell death mechanisms such as apoptosis, necrosis, pyroptosis, autophagy, and ferroptosis, with key molecule interactions in these programmed cell death pathways depicted; this cell death is a critical driver of severe organ dysfunction and the pathological course of sepsis
Pathogenic viruses exploit various cell death mechanisms, including apoptosis, necrosis, pyroptosis, autophagy, and ferroptosis, to induce the demise of organ parenchymal cells, as illustrated by the key molecular interactions within these pathways; this process is a major contributor to the development of severe organ impairment and the progression of sepsis
S. aureus is a gram-positive opportunistic pathogen that normally colonized in human anterior nares. S. aureus is one of the leading causes of bacteremia and case fatality rate for S. aureus has plateaued at 15 to 50% over the past several decades [96‐99]. S. aureus induced MODS is an important cause of high mortality. De Kimpe SJ et al. showed that two cell wall components from S. aureus, peptidoglycan (PepG) and lipoteichoic acid (LTA), work together to cause systemic inflammation and organ failure. For instance, administration of both PepG and LTA in anesthetized rats resulted in MODS, as indicated by a decrease in arterial oxygen pressure (lung) and an increase in plasma concentrations of bilirubin and alanine aminotransferase (liver), creatinine and urea (kidney), lipase (pancreas), and creatine kinase (heart or skeletal muscle) [100, 101]. Wang JE et al. demonstrated that high-dose of peptidoglycan from S. aureus caused increased serum values of aspartate aminotransferase, alanine aminotransferase, gamma-glutamyltransferase, and bilirubin (indicators of liver injury/dysfunction) as well as significantly increased creatinine and urea (indicators of renal dysfunction) in the rat [102, 103]. Proteomics screening singled out the liver as a hotspot for methicillin-resistant S. aureus (MRSA) sepsis pathology, a pattern further confirmed by blood chemistry and histopathological evidence of liver dysfunction, necrosis and thrombosis [104, 105]. Beno DW, et al. found Staphylococcal enterotoxin B and LPS induced a bimodal pattern of hepatic dysfunction and injury. Histological examination of the liver revealed periportal inflammation and fibrosis [106]. Infusion of heat-inactivated S. aureus induced activation of coagulation factor (F) XI that promotes multiorgan failure in rodent models of sepsis. Inhibition of FXII prevented fever, terminal hypotension, respiratory distress, and multiorgan failure [107, 108].
S. aureus causes apoptosis during infection [109]. The mRNA and protein expressions of death receptor-associated genes (FasL, Fas, FADD and caspase-8) were up-regulated in S. aureus-induced ALI. Ginsenoside Rb1 attenuated S. aureus-induced oxidative damage via the ER stress-mediated pathway and apoptosis through death receptor-mediated pathway [110]. S. aureus-induced cell death was also accompanied by apoptotic alterations, such as DNA fragmentation, but was independent of endogenous FasL and TNF-α death ligand expression. Mesothelial cell death might represent a major mechanism of S. aureus-induced damage of the peritoneum during bacterial peritonitis [111]. Moreover, S. aureus extracellular vesicles (EVs) internalized by bovine mammary epithelial cells (MAC-T cells) reduced the expression of BCL-2 and increased the expression of BAX, causing mitochondrial damage and apoptosis in epithelial cells [112]. S. aureus induced a massive cytosolic Ca2+ increase in epithelial host cells, this was paralleled by a decrease in ER Ca2+ concentration. Additionally, calcium ions from the extracellular space contributed to the cytosolic Ca2+ increase, which led to an increase in mitochondrial Ca2+ concentration, the activation of calpains and caspases, and eventually to cell lysis. It is convinced that intracellular S. aureus disturbs the host cell Ca2+ homeostasis and induces cytoplasmic Ca2+ overload, which results in both apoptotic and necrotic cell death in parallel or succession [113]. Levels of RIP3 and cleaved caspase-1 protein in S. aureus-infected A549 alveolar epithelial cells increased at 12 h post-infection. TNF-α enhances the damage of S. aureus on lung epithelial cells, and its mechanism is associated with RIP3 mediated necroptosis. Decreasing RIP3 levels by siRNA significantly suppressed the release of lactate dehydrogenase as well as A549 cell necrosis induced by TNF-α and S. aureus [114]. Fibrin clots loaded with S. aureus were implanted abdominally into C57BL/6 mice to construct sepsis-induced pulmonary inflammation model. Research indicated that S. aureus could increase both canonical and noncanonical autophagy pathways. Efforts to promote mitophagy may be a useful therapeutic adjunct for S-ALI [115]. Furthermore, a recent investigation revealed that upon internalizing S. aureus, lung epithelial cells generate membrane vesicles (MVs) that not only form but also colocalize with autophagosomes. This discovery offers new understanding into how S. aureus might exploit MVs to facilitate its pathogenic effects within the host [116].
Escherichia coli
Extraintestinal pathogenic E. coli are the most frequent cause of bloodstream infections, mostly related to urinary tract infections and intra-abdominal infections. The burden of disease and mortality associated with bloodstream infections due to E. coli is considerable, in neonatal and older populations. About one-third of patients with E. coli -causing bloodstream infections develop a dysregulated host response (ie, sepsis or septic shock), which is a main driver for mortality [117‐119].
Clinical strains of E. coli isolated from septic patients bind to sheared human aortic endothelial cells under static and hemodynamic shear conditions. This interaction results in disturbances in endothelial barrier integrity, permeability changes, and ultimately apoptosis [120]. E.coli outer membrane vesicles (E.coli-OMVs) increased the apoptosis rate and a decrease in the ratio of Bcl-2/Bax in colon epithelial cells [121]. LPS from E. coli triggers necroptosis in alveolar epithelial cells, leading to ALI. In alveolar epithelial cells treated with LPS from E. coli O111, the accumulation of mitochondrial citrate is attributed to the downregulation of Idh3α and the citrate carrier. This citric acid accumulation induces mitochondrial division and autophagy, promoting hyper-necroptosis through mitochondrial autophagy and initiating ALI [122]. Enterotoxigenic E. coli (ETEC) is a leading cause of intestinal inflammation, damaging intestinal barrier function by modulating necroptosis signaling pathways. ETEC-induced intestinal inflammation and cell damage can be ameliorated by EPA and arachidonic acid (ARA) through inhibition of necroptosis signaling pathways [123]. Pathogenic E. coli harbors a large virulence plasmid encoding HlyF, a cytoplasmic enzyme that leads to the overproduction of outer membrane vesicles (OMVs). These OMVs inhibit macroautophagic/autophagic flux by impairing autophagosome-lysosome fusion, thus preventing the formation of acidic autolysosomes and autophagosome clearance [124]. The Keap1/Nrf2 system is crucial in regulating oxidative stress by controlling antioxidant genes like GPX4 and HO-1. LPS induces ferroptosis in pulmonary microvascular endothelial cells by up-regulating Nrf2 and down-regulating Keap1 and GPX4 [88]. Ferroptosis of cardiomyocytes was observed in a model of septic cardiomyopathy induced by LPS. LPS from E. coli affects cardiac systolic function, evidenced by changes in fractional shortening, peak shortening, and the maximum rate of shortening/re-lengthening on echocardiography, along with increased left ventricular systolic end diameter. LPS induces cardiac systolic abnormalities, oxidative stress and inflammation through NF-κB phosphorylation and cytoplasmic translocation of the transcription factor Nrf2 [125, 126]. LPS induces cytotoxicity by increasing ferroptosis, and Bmal1 inhibits this toxicity via the AKT/p53 pathway [127]. Furthermore, E.coli-OMVs inhibited autophagic flux by impairing the autophagosome-lysosome fusion and activated the non-canonical inflammasome pathways in epithelia cells, highlighting their roles in exacerbating pathogenic properties in the host [124].
Klebsiella pneumoniae
K. pneumoniae primarily resides in the intestines and upper respiratory tract of humans and animals, typically existing as a harmless commensal organism. However, under certain conditions, particularly in immunocompromised individuals, it can become an opportunistic pathogen, causing severe infections such as pneumonia, intestinal infections, and bloodstream infections, which can further lead to sepsis. If the infection is not promptly controlled, sepsis may progress to MODS [128‐131].
Infections caused by K. pneumoniae impact host cell autophagy and ferroptosis through complex mechanisms. During infection, K. pneumoniae releases bacterial extracellular vesicles (bEVs) that interact with lung endothelial cells, leading to endothelial barrier dysfunction. These bEVs significantly reduce RNase1 expression and activate endothelial cells through LPS-dependent toll-like Receptor 4 (TLR4) signaling cascades [132]. Additionally, LPS from K. pneumoniae stimulates caspase-4-dependent GSDMD activation and pyroptosis in cultured human cells, compromising the BBB in caspase-4-expressing mice [133]. Pulmonary infections caused by K. pneumoniae result in insufficient and maladaptive autophagy in the lungs. Therapeutically, activation of Beclin-1, either by forced expression of an active mutant (Becn1F121A) or treatment with TB peptide, enhances autophagy. This significantly reduces sickness scores, systemic infection, and circulating and pulmonary cytokine production, demonstrating benefits such as reduced alveolar congestion, hemorrhage, inflammatory cell infiltration, and alveolar wall thickness. Thus, activation of Beclin-1 alleviates adverse outcomes of K. pneumoniae-induced sepsis, suggesting therapeutic potential [134]. Furthermore, LPS from K. pneumoniae induces ferroptosis in pulmonary microvascular endothelial cells by upregulating Nrf2 and downregulating Keap1 and GPX4. The Keap1/Nrf2 system plays a crucial role in regulating oxidative stress by controlling antioxidant genes like GPX4 and HO-1. This highlights the interplay between autophagy and ferroptosis in the pathogenesis of K. pneumoniae infections [88].
Pseudomonas aeruginosa
P. aeruginosa is widely found in natural environments such as soil, water, and plants, as well as colonizing human skin, the respiratory tract, and the gastrointestinal tract. Driven in part by its metabolic versatility, high intrinsic antibiotic resistance, and a large repertoire of virulence factors, P. aeruginosa is expertly adapted to thrive in a wide variety of environments, and in the process, making it a notorious opportunistic pathogen. In hospital settings, P. aeruginosa is a common pathogen in ICUs, where its strong antibiotic resistance complicates the treatment of sepsis and MODS [135, 136].
A recent study has indicated that P. aeruginosa releases factors that trigger endothelial anoikis through the degradation of matrix proteins, a process akin to the action of many pro-apoptotic proteases [137]. P. aeruginosa utilizes a type III secretion system (T3SS) to inject exoenzyme effectors into the target host’s pulmonary microvascular endothelial cells. Among these effectors, ExoU, disrupts the endothelial cell barrier and initiates caspase-1-mediated inflammation. ExoU is found to target the host cell’s plasma membrane, with its membrane association dependent on C-terminal amino acids between positions 550–687 and its interaction with phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2) on the inner membrane leaflet. The phospholipase A2 (PLA2) activity of ExoU is activated by its interactions with ubiquitinylated proteins and membrane inositol phosphatides, culminating in the destruction of the host cell [138]. Observations in pulmonary microvascular endothelial cells align with previous studies showing ExoU-mediated ROS imbalances in epithelial and endothelial cells, contributing to inflammatory lung injury [134]. Further studies are needed to determine if ExoU targets Nox2- and Nox4-mediated signaling pathways, previously shown to regulate endothelial cell ROS signaling [139]. P. aeruginosa pneumonia-induced sepsis can cause MODS, including ALI and AKI. Molecular analyses revealed that exosomes derived from septic mouse serum could increase levels of cell death biomarkers (cleaved caspase-3, cleaved caspase-1, NLRP3, ASC, and GSDMD) in the lung compared to sham serum exosomes. Additionally, septic serum exosomes increased NGAL (a kidney injury biomarker), cleaved caspase-3, cleaved caspase-1, NLRP3, ASC, and GSDMD levels in the kidney [140]. P. aeruginosa lacks arachidonic acid-phosphatidylethanolamine (AA-PE) but can express lipoxygenase (pLoxA), which oxidizes host AA-PE to 15-hydroperoxide AA-PE, leading to ferroptosis in human bronchial epithelial cells. P. aeruginosa uses pLoxA to convert host AA-PE into the ferrophilic 15-hydroperoxy-arachidonic group, causing ferroptosis. It also degrades host GPX4 defense by activating lysosomal chaperone-mediated autophagy. Pathogens can control or inhibit lipid ROS processes through transcriptional regulation of GPX4-dependent antioxidant pathways or by interfering with lipid ROS via intracellular iron, lipid, and mitochondrial metabolism. They enhance the activity of host lipid-metabolizing enzymes such as acyl-coa synthetase Long Chain Family Member 4 (ACSL4) or ACSL1, promoting the production of PUFA-PL and indirectly leading to lipid ROS synthesis [141, 142]. Massive respiratory epithelial cell death, a hallmark of severe ALI and ARDS, is caused by P. aeruginosa infection. P. aeruginosa induces lung epithelial necroptosis via crosstalk between inflammasome activation and necroptosis. MLKL-dependent necroptosis signaling changes mitochondrial membrane potential, releasing ROS, which triggers functional inflammasome activation. Necroptosis-triggered NLRP3 inflammasome in the epithelium plays a crucial role in P. aeruginosa-mediated injury [143]. Overexpressed RIPK3 continuously attenuates pro-inflammatory cytokine gene expression by inhibiting NF-κB activation, preventing inflammation progression during P. aeruginosa infection [144].
Virus
SARS-CoV-2
Since its emergence in humans at the close of 2019, SARS-CoV-2 has swiftly disseminated across the globe, imposing a substantial mortality toll [145, 146]. Observations have indicated that many critically ill corona virus disease 2019 (COVID-19) patients meet the criteria for sepsis/septic shock and experience impaired liver and kidney function in tandem with severe lung injury [147]. A two-center prospective cohort study enrolled 120 critically ill COVID-19 patients, revealing that patients in various subgroups exhibited distinct systemic markers of pyroptosis, iron metabolism disorders, ferroptosis, or lung cell death, culminating in diverse outcomes in the ICU [148]. An intriguing study noted no differences in apoptosis and ferroptosis between SARS-CoV-2 infected and uninfected lung tissues. However, increased levels of pyroptosis and necroptosis were detected in the infected lungs. Notably, necroptosis showed a strong positive correlation with viral load, implying a virus-mediated necroptotic pathway in the pathogenesis of COVID-19 [149].
The regulation of host cell PCD is pivotal in SARS-CoV-2 induced tissue injury [150]. The SARS-CoV-2 membrane protein can induce mitochondrial apoptosis in lung epithelial cells by stabilizing BCL-2 ovarian killer (BOK) through inhibition of its ubiquitination and promoting its mitochondrial translocation [151]. SARS-CoV-2 infection in cardiomyocytes leads to apoptosis and cell cycle arrest, with upregulated expression of several intrinsic pro-apoptotic genes (BAD, BAX, and BBC3) [152]. In microglia, SARS-CoV-2 infection promotes both intrinsic and extrinsic death receptor-mediated apoptosis, with enhanced expression of extrinsic apoptosis-associated death receptors such as Fas, DR4, DR5, and TNF receptor 2 (TNFR2), and increased activation of BAX-dependent PCD [153]. Using an in vitro culture model of polarized human airway epithelium, a study demonstrated that SARS-CoV-2 infection triggers necroptosis via MLKL, PIPK1, and RIPK3 activation, resulting in epithelial damage and disruption of the airway epithelial barrier function [154]. The SARS-CoV-2 N protein is detectable in damaged tubules of COVID-19 patients and induces tubular cell death through the Smad3-Ripk3/MLKL necroptosis pathway [155]. Necroptosis is predominantly activated in SARS-CoV-2-infected cells, while apoptosis occurs in uninfected neighboring cells. The mechanism involves viral Z-RNA binding to Z-DNA binding protein 1 (ZBP1) in human airway epithelia and lung tissues from COVID-19 patients, interacting with RIPK3 to initiate necroptosis through phosphorylation of MLKL [156]. The non-structural protein 6 (NSP6) of SARS-CoV-2 can induce pyroptosis in lung epithelial cells by triggering NLRP3/ASC-dependent caspase-1 activation [157]. Non-structural protein 5 (NSP5) mediates the cleavage of NLRP1, leading to inflammasome assembly and lung epithelial pyroptosis. The accessory protein Open reading frame 3a (ORF3a) promotes NRF2 degradation by recruiting Keap1, enhancing lipid ROS levels and resulting in lung epithelial ferroptosis [158]. Another accessory protein, ORF7b, induces ferroptosis through upregulation of the transcription regulator c-Myc [159]. NSP6, ORF3a, and ORF10 can initiate ER stress-induced autophagy in HEK293T cells via the cGAS-STING pathway [160‐163]. The ORF7a protein induces autophagosome accumulation across multiple host cell lines by activating the AKT-MTOR signaling pathway [164]. SARS-CoV-2 infection induces host cellular senescence by causing DNA damage and eliciting an altered DNA damage response. Proteins ORF6 and NSP 13 contribute to this process by degrading the DNA damage response kinase CHK1 through proteasomal and autophagic pathways, respectively [165].
Influenza virus
Roughly 30–40% of hospitalized patients with laboratory-confirmed influenza are found to have acute pneumonia. The influenza virus is known to increase the risk of bacterial sepsis, primarily causing severe pneumonia and often coinciding with or leading to secondary bacterial infections, notably by Staphylococcus aureus and Streptococcus pneumoniae [166]. Across all age groups, both primary influenza viral pneumonia and bacterial co-infection can result in respiratory failure, ARDS, septic shock, and MODS [167].
Influenza A virus (IAV) has evolved to produce viral proteins that directly manipulate host cell PCD. The nucleoprotein (NP) of IAV induces apoptosis in infected host cells by directly interacting with and repressing the expression of the host apoptosis inhibitor protein 5 (API5) [168]. IAV exacerbates oxidized low-density lipoprotein-induced apoptosis in human endothelial cells by enhancing p53 signaling, which is associated with increased cytochrome C release and caspase 3 activation [169]. Intriguingly, IAV generates “Z-form” RNA structures (Z-RNAs) that activate the host protein ZBP1, stimulating RIPK3 and leading to the phosphorylation and activation of MLKL, thereby mediating necroptosis in lung epithelial cells [170]. Another study revealed that the NS1 protein of IAV binds to MLKL, promoting its oligomerization and membrane translocation to induce necroptosis in lung epithelial cells [171]. IAV proteins NP and the polymerase subunit polymerase basic protein 1 (PB1) interact with ZBP1 to modulate NLRP3 inflammasome activation and pyroptosis in infected cells, exacerbating epithelial damage [172]. The viral protein HA induces ferroptosis in epithelial cells by interacting with nuclear receptor coactivator 4 (NCOA4) and tax1-binding protein 1 (TAX1BP1), triggering ferritinophagy [173]. The matrix protein M2 of IAV has been found to engage with the host protein LC3B, activating autophagosome formation in human epithelial cells [174]. Recently, it was reported that IAV induces autophagy through HA binding to heat shock protein 90AA1 (HSP90AA1), resulting in an elevated level of LC3-II [175].
Future perspectives
Sepsis leads to unacceptably high morbidity and mortality rates, with MODS being an escalating global issue. Endothelial dysfunction and injury are considered the primary causes of organ dysfunction in sepsis. Sepsis-related organ failure is associated with various PCD modes such as apoptosis, pyroptosis, and autophagy. Mitochondria and ER are pivotal in regulating various cell death pathways, with significant crosstalk between apoptosis, pyroptosis, necroptosis, autophagy, and ferroptosis. These cell death pathways often work together, affecting the same organs, and should not be viewed in isolation. Mitochondria regulates apoptosis through the mitochondrial permeability transition pore, releasing cytochrome C into the cytoplasm, which triggers the intrinsic apoptotic pathway. Mitochondria also contributes to other forms of PCD. For example, GSDMD fragments from pyroptosis can induce MOMP, leading to caspase-3 activation [176]. Caspase-1 can also trigger mitochondrial apoptosis by cleaving BID protein. Mitochondrial apoptosis can, in turn, activate the NLRP3 inflammasome, leading to caspase-1 activity [177]. ROS promotes necroptosis by enhancing RIPK1 autophosphorylation and necrosome formation [178]. RIPK3 activates the pyruvate dehydrogenase complex, increasing aerobic respiration and ROS production [179]. Excess iron in mitochondria causes lipid peroxidation, leading to ferroptosis. Cysteine deprivation also promotes lipid peroxidation by increasing mitochondrial respiration and ROS production [180]. Communication between the ER and mitochondria regulates calcium homeostasis, lipid biosynthesis, and protein interactions, which are critical for cell death regulation. These mechanisms highlight the complexity of cellular responses to severe infections, which might exacerbate the organ dysfunction or failure in sepsis. Understanding these pathways and their common molecular players offers potential targets for therapeutic interventions to mitigate sepsis-induced organ damage.
In recent years, there has been growing interest in pathogen-induced PCD and subsequent organ failure in sepsis. Bacteria and viruses can manipulate inflammatory signaling and cell death pathways in host parenchymal cells either directly or indirectly by releasing toxins or proteins. This manipulation affects the expression of transcription factors and modulates stress responses in mitochondria and the ER. The precise mechanisms linking these organelles to pathogen-activated cell death pathways remain largely elusive. This intricate and vital biological process encompasses a multitude of signaling pathways and molecular regulatory networks, which may vary across different sepsis conditions. Future research is required to understand the specific molecular pathways and interactions that govern various cell death mechanisms in host cells, such as apoptosis, pyroptosis, necroptosis, ferroptosis, and autophagy dysregulation, with particular focus on their organ-specific manifestations and the subsequent inflammatory responses. Strategies targeting pathogen-induced inflammatory and cell death processes could serve as valuable adjuncts in combating infections and preserving organ function. Investigating therapeutic interventions that modulate these cell death pathways, including the development of specific inhibitors or agonists, antioxidant and anti-inflammatory treatments, and gene editing techniques, is pivotal in alleviating sepsis-related inflammation and organ damage, thereby enhancing patient survival rates and quality of life. Future research endeavors will concentrate on advancing personalized and precise sepsis treatments through interdisciplinary collaboration, integrating clinical and laboratory findings. Although numerous proof-of-concept studies exist, the efficacy of treatments targeting cell death pathways during infections is still in its infancy. We are optimistic that a comprehensive understanding of the roles and mechanisms of host cell death in diverse host–pathogen interactions will underpin the rational design of future therapeutic interventions, ultimately improving the prognosis for patients battling life-threatening infections.
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Rhodes A, Evans LE, Alhazzani W, et al. Surviving sepsis campaign: international guidelines for management of sepsis and septic shock: 2016. Intensive Care Med. 2017;43(3):304–77.PubMedCrossRef
2.
Weiss SL, Peters MJ, Alhazzani W, et al. Surviving sepsis campaign international guidelines for the management of septic shock and sepsis-associated organ dysfunction in children. Intensive Care Med. 2020;46(Suppl 1):10–67.PubMedPubMedCentralCrossRef
3.
Kaukonen KM, Bailey M, Suzuki S, et al. Mortality related to severe sepsis and septic shock among critically ill patients in Australia and New Zealand, 2000–2012. JAMA. 2014;311(13):1308–16.PubMedCrossRef
4.
Rudd KE, Johnson SC, Agesa KM, et al. Global, regional, and national sepsis incidence and mortality, 1990–2017: analysis for the global burden of disease study. Lancet. 2020;395(10219):200–11.PubMedPubMedCentralCrossRef
5.
Manabe T, Heneka MT. Cerebral dysfunctions caused by sepsis during ageing. Nat Rev Immunol. 2022;22(7):444–58.PubMedCrossRef
6.
Fowler AA 3rd, Truwit JD, Hite RD, et al. Effect of vitamin C infusion on organ failure and biomarkers of inflammation and vascular injury in patients with sepsis and severe acute respiratory failure: the CITRIS-ALI randomized clinical trial. JAMA. 2019;322(13):1261–70.PubMedPubMedCentralCrossRef
7.
Merx MW, Weber C. Sepsis and the heart. Circulation. 2007;116(7):793–802.PubMedCrossRef
8.
Lv X, Wang H. Pathophysiology of sepsis-induced myocardial dysfunction. Mil Med Res. 2016;27(3):30.
9.
Singer G, Houghton J, Rivera CA, et al. Role of LPS in the hepatic microvascular dysfunction elicited by cecal ligation and puncture in mice. J Hepatol. 2007;47(6):799–806.PubMedPubMedCentralCrossRef
10.
Gimson AE. Hepatic dysfunction during bacterial sepsis. Intensive Care Med. 1987;13(3):162–6.PubMedCrossRef
11.
Zarbock A, Nadim MK, Pickkers P, et al. Sepsis-associated acute kidney injury: consensus report of the 28th acute disease quality initiative workgroup. Nat Rev Nephrol. 2023;19(6):401–17.PubMedCrossRef
12.
Mittal R, Coopersmith CM. Redefining the gut as the motor of critical illness. Trends Mol Med. 2014;20(4):214–23.PubMedCrossRef
13.
Vincent JL, Francois B, Zabolotskikh I, et al. Effect of a recombinant human soluble thrombomodulin on mortality in patients with sepsis-associated coagulopathy: the SCARLET randomized clinical trial. JAMA. 2019;321(20):1993–2002.PubMedPubMedCentralCrossRef
Singer M, Deutschman CS, Seymour CW, et al. The third international consensus definitions for sepsis and septic shock (Sepsis-3). JAMA. 2016;315(8):801–10.PubMedPubMedCentralCrossRef
16.
Graetz TJ, Hotchkiss RS. Sepsis: preventing organ failure in sepsis - the search continues. Nat Rev Nephrol. 2017;13(1):5–6.PubMedCrossRef
17.
Gotts JE, Matthay MA. Sepsis: pathophysiology and clinical management. BMJ. 2016;23(353):i1585.CrossRef
18.
Linkermann A, Stockwell BR, Krautwald S, et al. Regulated cell death and inflammation: an auto-amplification loop causes organ failure. Nat Rev Immunol. 2014;14(11):759–67.PubMedCrossRef
19.
Yuan J, Ofengeim D. A guide to cell death pathways. Nat Rev Mol Cell Biol. 2024;25(5):379–95.PubMedCrossRef
20.
Kerr JF, Wyllie AH, Currie AR. Apoptosis: a basic biological phenomenon with wide-ranging implications in tissue kinetics. Br J Cancer. 1972;26(4):239–57.PubMedPubMedCentralCrossRef
21.
Tang D, Kang R, Berghe TV, et al. The molecular machinery of regulated cell death. Cell Res. 2019;29(5):347–64.PubMedPubMedCentralCrossRef
22.
Markwart R, Saito H, Harder T, et al. Epidemiology and burden of sepsis acquired in hospitals and intensive care units: a systematic review and meta-analysis. Intensive Care Med. 2020;46(8):1536–51.PubMedPubMedCentralCrossRef
23.
Timsit JF, Ruppé E, Barbier F, et al. Bloodstream infections in critically ill patients: an expert statement. Intensive Care Med. 2020;46(2):266–84.PubMedPubMedCentralCrossRef
24.
Perkins GD, Ji C, Connolly BA, et al. Effect of noninvasive respiratory strategies on intubation or mortality among patients with acute hypoxemic respiratory failure and COVID-19: the RECOVERY-RS randomized clinical trial. JAMA. 2022;327(6):546–58.PubMedPubMedCentralCrossRef
25.
Sumner KM, Masalovich S, O’Halloran A, et al. Severity of influenza-associated hospitalisations by influenza virus type and subtype in the USA, 2010–19: a repeated cross-sectional study. Lancet Microbe. 2023;4(11):e903–12.PubMedPubMedCentralCrossRef
26.
Ashkenazi A, Fairbrother WJ, Leverson JD, et al. From basic apoptosis discoveries to advanced selective BCL-2 family inhibitors. Nat Rev Drug Discov. 2017;16(4):273–84.PubMedCrossRef
27.
Wieder T, Essmann F, Prokop A, et al. Activation of caspase-8 in drug-induced apoptosis of B-lymphoid cells is independent of CD95/Fas receptor-ligand interaction and occurs downstream of caspase-3. Blood. 2001;97(5):1378–87.PubMedCrossRef
28.
Bitomsky N, Conrad E, Moritz C, et al. Autophosphorylation and Pin1 binding coordinate DNA damage-induced HIPK2 activation and cell death. Proc Natl Acad Sci USA. 2013;110(45):E4203–12.PubMedPubMedCentralCrossRef
29.
Hotchkiss RS, Swanson PE, Freeman BD, et al. Apoptotic cell death in patients with sepsis, shock, and multiple organ dysfunction. Crit Care Med. 1999;27(7):1230–51.PubMedCrossRef
30.
Coopersmith CM, Chang KC, Swanson PE, et al. Overexpression of Bcl-2 in the intestinal epithelium improves survival in septic mice. Crit Care Med. 2002;30(1):195–201.PubMedCrossRef
31.
Coopersmith CM, Stromberg PE, Dunne WM, et al. Inhibition of intestinal epithelial apoptosis and survival in a murine model of pneumonia-induced sepsis. JAMA. 2002;287(13):1716–21.PubMedCrossRef
32.
Perl M, Chung CS, Lomas-Neira J, et al. Silencing of Fas, but not caspase-8, in lung epithelial cells ameliorates pulmonary apoptosis, inflammation, and neutrophil influx after hemorrhagic shock and sepsis. Am J Pathol. 2005;167(6):1545–59.PubMedPubMedCentralCrossRef
33.
Wang Z, Guo Z, Wang X, et al. Reduning alleviates sepsis-induced acute lung injury by reducing apoptosis of pulmonary microvascular endothelial cells. Front Immunol. 2023;3(14):1196350.CrossRef
34.
Gill SE, Rohan M, Mehta S. Role of pulmonary microvascular endothelial cell apoptosis in murine sepsis-induced lung injury in vivo. Respir Res. 2015;16(1):109.PubMedPubMedCentralCrossRef
35.
Zhu H, Zhang L, Jia H, et al. Tetrahydrocurcumin improves lipopolysaccharide-induced myocardial dysfunction by inhibiting oxidative stress and inflammation via JNK/ERK signaling pathway regulation. Phytomedicine. 2022;104:154283.PubMedCrossRef
36.
Chen SN, Tan Y, Xiao XC, et al. Deletion of TLR4 attenuates lipopolysaccharide-induced acute liver injury by inhibiting inflammation and apoptosis. Acta Pharmacol Sin. 2021;42(10):1610–9.PubMedPubMedCentralCrossRef
37.
Zhang H, Zeng L, Xie M, et al. TMEM173 drives lethal coagulation in sepsis. Cell Host Microbe. 2020;27(4):556-570.e6.PubMedPubMedCentralCrossRef
38.
Cai Z, Jitkaew S, Zhao J, et al. Plasma membrane translocation of trimerized MLKL protein is required for TNF-induced necroptosis. Nat Cell Biol. 2014;16(1):55–65.PubMedCrossRef
39.
Duprez L, Takahashi N, Van Hauwermeiren F, et al. RIP kinase-dependent necrosis drives lethal systemic inflammatory response syndrome. Immunity. 2011;35(6):908–18.PubMedCrossRef
40.
Murphy JM, Czabotar PE, Hildebrand JM, et al. The pseudokinase MLKL mediates necroptosis via a molecular switch mechanism. Immunity. 2013;39(3):443–53.PubMedCrossRef
41.
Zhang H, Zhang X, Ling C, et al. EGFR-TNFR1 pathway in endothelial cell facilitates acute lung injury by NF-κB/MAPK-mediated inflammation and RIP3-dependent necroptosis. Int Immunopharmacol. 2023;117:109902.PubMedCrossRef
42.
Liao Z, Ou X, Zhou C, et al. Xenon attenuated neonatal lipopolysaccharide exposure induced neuronal necroptosis and subsequently improved cognition in juvenile rats. Front Pharmacol. 2022;2(13):1002920.CrossRef
43.
Bolognese AC, Yang WL, Hansen LW, et al. Inhibition of necroptosis attenuates lung injury and improves survival in neonatal sepsis. Surgery. 2018;S0039–6060(18):30096–105.
44.
Hansen LW, Jacob A, Yang WL, et al. Deficiency of receptor-interacting protein kinase 3 (RIPK3) attenuates inflammation and organ injury in neonatal sepsis. J Pediatr Surg. 2018;53(9):1699–705.PubMedCrossRef
45.
Zechendorf E, Vaßen P, Zhang J, et al. Heparan sulfate induces necroptosis in murine cardiomyocytes: a medical-in silico approach combining in vitro experiments and machine learning. Front Immunol. 2018;20(9):393.CrossRef
46.
Zhang Q, Wei S, Lu J, et al. Necrostatin-1 accelerates time to death in a rat model of cecal ligation and puncture and massively increases hepatocyte caspase-3 cleavage. Am J Physiol Gastrointest Liver Physiol. 2019;316(4):G551–61.PubMedCrossRef
47.
Xu Q, Guo J, Li X, et al. Necroptosis underlies hepatic damage in a piglet model of lipopolysaccharide-induced sepsis. Front Immunol. 2021;12(12):633830.PubMedPubMedCentralCrossRef
48.
Zelic M, Roderick JE, O’Donnell JA, et al. RIP kinase 1-dependent endothelial necroptosis underlies systemic inflammatory response syndrome. J Clin Invest. 2018;128(5):2064–75.PubMedPubMedCentralCrossRef
49.
Zhang X, Wu J, Liu Q, et al. mtDNA-STING pathway promotes necroptosis-dependent enterocyte injury in intestinal ischemia reperfusion. Cell Death Dis. 2020;11(12):1050.PubMedPubMedCentralCrossRef
50.
Liu Y, Xu Q, Wang Y, et al. Necroptosis is active and contributes to intestinal injury in a piglet model with lipopolysaccharide challenge. Cell Death Dis. 2021;12(1):62.PubMedPubMedCentralCrossRef
51.
Zhang X, Wu J, Liu Q, et al. RIPK3-MLKL necroptotic signalling amplifies STING pathway and exacerbates lethal sepsis. Clin Transl Med. 2023;13(7):e1334.PubMedPubMedCentralCrossRef
52.
Mariano F, Cantaluppi V, Stella M, et al. Circulating plasma factors induce tubular and glomerular alterations in septic burns patients. Crit Care. 2008;12:R42.PubMedPubMedCentralCrossRef
53.
Wang Y, Gao W, Shi X, et al. Chemotherapy drugs induce pyroptosis through caspase-3 cleavage of a gasdermin. Nature. 2017;547(7661):99–103.PubMedCrossRef
54.
Li Y, Huang H, Liu B, et al. Inflammasomes as therapeutic targets in human diseases. Signal Transduct Target Ther. 2021;6(1):247.PubMedPubMedCentralCrossRef
55.
Xia S, Zhang Z, Magupalli VG, et al. Gasdermin D pore structure reveals preferential release of mature interleukin-1. Nature. 2021;593(7860):607–11.PubMedPubMedCentralCrossRef
56.
Zhou R, Yang X, Li X, et al. Recombinant CC16 inhibits NLRP3/caspase-1-induced Pyroptosis through p38 MAPK and ERK signaling pathways in the brain of a neonatal rat model with sepsis. J Neuroinflammation. 2019;16(1):239.PubMedPubMedCentralCrossRef
57.
Cheng KT, Xiong S, Ye Z, et al. Caspase-11-mediated endothelial pyroptosis underlies endotoxemia-induced lung injury. J Clin Invest. 2017;127(11):4124–35.PubMedPubMedCentralCrossRef
58.
Dai S, Ye B, Zhong L, et al. GSDMD mediates LPS-induced septic myocardial dysfunction by regulating ROS-dependent NLRP3 inflammasome activation. Front Cell Dev Biol. 2021;8(9):779432.CrossRef
59.
Lu C, Liu J, Escames G, et al. PIK3CG regulates NLRP3/GSDMD-mediated pyroptosis in septic myocardial injury. Inflammation. 2023;46(6):2416–32.PubMedCrossRef
60.
Wu QR, Yang H, Zhang HD, et al. IP3R2-mediated Ca2+ release promotes LPS-induced cardiomyocyte pyroptosis via the activation of NLRP3/Caspase-1/GSDMD pathway. Cell Death Discov. 2024;10(1):91.PubMedPubMedCentralCrossRef
61.
Deng M, Tang Y, Li W, et al. The endotoxin delivery protein HMGB1 mediates caspase-11-dependent lethality in sepsis. Immunity. 2018;49(4):740-753.e7.PubMedPubMedCentralCrossRef
62.
Tang Y, Wang X, Li Z, et al. Heparin prevents caspase-11-dependent septic lethality independent of anticoagulant properties. Immunity. 2021;54(3):454-467.e6.PubMedCrossRef
63.
Cao Y, Chen X, Zhu Z, et al. STING contributes to lipopolysaccharide-induced tubular cell inflammation and pyroptosis by activating endoplasmic reticulum stress in acute kidney injury. Cell Death Dis. 2024;15(3):217.PubMedPubMedCentralCrossRef
64.
Xu D, Jiang J, Liu Y, et al. TIMP2 protects against sepsis-associated acute kidney injury by cAMP/NLRP3 axis-mediated pyroptosis. Am J Physiol Cell Physiol. 2024;326(5):C1353–66.PubMedCrossRef
65.
Liu J, Du S, Kong Q, et al. HSPA12A attenuates lipopolysaccharide-induced liver injury through inhibiting caspase-11-mediated hepatocyte pyroptosis via PGC-1α-dependent acyloxyacyl hydrolase expression. Cell Death Differ. 2020;27(9):2651–67.PubMedPubMedCentralCrossRef
66.
Yang X, Cheng X, Tang Y, et al. Bacterial endotoxin activates the coagulation cascade through gasdermin D-dependent phosphatidylserine exposure. Immunity. 2019;51(6):983-996.e6.PubMedCrossRef
67.
Wu C, Lu W, Zhang Y, et al. Inflammasome activation triggers blood clotting and host death through pyroptosis. Immunity. 2019;50(6):1401-1411.e4.PubMedPubMedCentralCrossRef
68.
Chen X, Shi C, He M, et al. Endoplasmic reticulum stress: molecular mechanism and therapeutic targets. Signal Transduct Target Ther. 2023;8(1):352.PubMedPubMedCentralCrossRef
69.
Cybulsky AV. Endoplasmic reticulum stress, the unfolded protein response and autophagy in kidney diseases. Nat Rev Nephrol. 2017;13(11):681–96.PubMedCrossRef
70.
Pi S, Mao L, Chen J, et al. The P2RY12 receptor promotes VSMC-derived foam cell formation by inhibiting autophagy in advanced atherosclerosis. Autophagy. 2021;17(4):980–1000.PubMedCrossRef
71.
Wang S, Long H, Hou L, et al. The mitophagy pathway and its implications in human diseases. Signal Transduct Target Ther. 2023;8(1):304.PubMedPubMedCentralCrossRef
72.
Issa AR, Sun J, Petitgas C, et al. The lysosomal membrane protein LAMP2A promotes autophagic flux and prevents SNCA-induced Parkinson disease-like symptoms in the Drosophila brain. Autophagy. 2018;14(11):1898–910.PubMedPubMedCentralCrossRef
73.
Li Y, Fan Z, Jia Q, et al. Chaperone-mediated autophagy (CMA) alleviates cognitive impairment by reducing neuronal death in sepsis-associated encephalopathy (SAE). Exp Neurol. 2023;365:114417.PubMedCrossRef
74.
Lo S, Yuan SS, Hsu C, et al. Lc3 over-expression improves survival and attenuates lung injury through increasing autophagosomal clearance in septic mice. Ann Surg. 2013;257(2):352–63.PubMedCrossRef
75.
Wang Y, Jasper H, Toan S, et al. Mitophagy coordinates the mitochondrial unfolded protein response to attenuate inflammation-mediated myocardial injury. Redox Biol. 2021;45:102049.PubMedPubMedCentralCrossRef
76.
Piquereau J, Godin R, Deschênes S, et al. Protective role of PARK2/Parkin in sepsis-induced cardiac contractile and mitochondrial dysfunction. Autophagy. 2013;9(11):1837–51.PubMedCrossRef
77.
Thiessen SE, Derese I, Derde S, et al. The role of autophagy in critical illness-induced liver damage. Sci Rep. 2017;7(1):14150.PubMedPubMedCentralCrossRef
78.
Huang S, Xu M, Liu L, et al. Autophagy is involved in the protective effect of p21 on LPS-induced cardiac dysfunction. Cell Death Dis. 2020;11(7):554.PubMedPubMedCentralCrossRef
79.
Li Y, Xu J, Chen W, et al. Hepatocyte CD36 modulates UBQLN1-mediated proteasomal degradation of autophagic SNARE proteins contributing to septic liver injury. Autophagy. 2023;19(9):2504–19.PubMedPubMedCentralCrossRef
80.
Carchman EH, Rao J, Loughran PA, et al. Heme oxygenase-1-mediated autophagy protects against hepatocyte cell death and hepatic injury from infection/sepsis in mice. Hepatology. 2011;53(6):2053–62.PubMedCrossRef
81.
Shao L, Xiong X, Zhang Y, et al. IL-22 ameliorates LPS-induced acute liver injury by autophagy activation through ATF4-ATG7 signaling. Cell Death Dis. 2020;11(11):970.PubMedPubMedCentralCrossRef
82.
Wang Y, Zhu J, Liu Z, et al. The PINK1/PARK2/optineurin pathway of mitophagy is activated for protection in septic acute kidney injury. Redox Biol. 2021;38:101767.PubMedCrossRef
83.
Deng Z, Sun M, Wu J, et al. SIRT1 attenuates sepsis-induced acute kidney injury via Beclin1 deacetylation-mediated autophagy activation. Cell Death Dis. 2021;12(2):217.PubMedPubMedCentralCrossRef
84.
Bersuker K, Hendricks JM, Li Z, et al. The CoQ oxidoreductase FSP1 acts parallel to GPX4 to inhibit ferroptosis. Nature. 2019;575(7784):688–92.PubMedPubMedCentralCrossRef
85.
Mao C, Liu X, Zhang Y, et al. DHODH-mediated ferroptosis defence is a targetable vulnerability in cancer. Nature. 2021;593(7860):586–90.PubMedPubMedCentralCrossRef
86.
Wu H, Li N, Peng S, et al. Maresin1 improves hippocampal neuroinflammation and cognitive function in septic rats by activating the SLC7A11 / GPX4 ferroptosis signaling pathway. Int Immunopharmacol. 2024;20(131):111792.CrossRef
87.
Wang H, Xu L, Tang X, et al. Lipid peroxidation-induced ferroptosis as a therapeutic target for mitigating neuronal injury and inflammation in sepsis-associated encephalopathy: insights into the hippocampal PEBP-1/15-LOX/GPX4 pathway. Lipids Health Dis. 2024;23(1):128.PubMedPubMedCentralCrossRef
88.
Shen K, Wang X, Wang Y, et al. miR-125b-5p in adipose derived stem cells exosome alleviates pulmonary microvascular endothelial cells ferroptosis via Keap1/Nrf2/GPX4 in sepsis lung injury. Redox Biol. 2023;62:102655.PubMedPubMedCentralCrossRef
89.
Wang C, Yuan W, Hu A, et al. Dexmedetomidine alleviated sepsis-induced myocardial ferroptosis and septic heart injury. Mol Med Rep. 2020;22(1):175–84.PubMedPubMedCentralCrossRef
90.
Liu C, Zou Q, Tang H, et al. Melanin nanoparticles alleviate sepsis-induced myocardial injury by suppressing ferroptosis and inflammation. Bioact Mater. 2022;27(24):313–21.
91.
Li N, Wang W, Zhou H, et al. Ferritinophagy-mediated ferroptosis is involved in sepsis-induced cardiac injury. Free Radic Biol Med. 2020;20(160):303–18.CrossRef
92.
Wang Y, Wang T, Xiang Q, et al. GPR116 promotes ferroptosis in sepsis-induced liver injury by suppressing system Xc-/GSH/GPX4. Cell Biol Toxicol. 2023;39(6):3015–30.PubMedCrossRef
93.
Yang Y, Wang A, Zhou J, et al. LncRNA SNHG11 induces ferroptosis in liver injury cells through miR-324–3p/GPX4 axis-mediated sepsis. Cell Mol Biol. 2023;69(12):163–9.PubMedCrossRef
94.
Yang Y, Lin Q, Zhu X, et al. Activation of lipophagy is required for RAB7 to regulate ferroptosis in sepsis-induced acute kidney injury. Free Radic Biol Med. 2024;218:120–31.PubMedCrossRef
95.
Wenzel SE, Tyurina YY, Zhao J, et al. PEBP1 wardens ferroptosis by enabling lipoxygenase generation of lipid death signals. Cell. 2017;171(3):628-641.e26.PubMedPubMedCentralCrossRef
96.
Menéndez R, Montull B, Reyes S, et al. Pneumonia presenting with organ dysfunctions: causative microorganisms, host factors and outcome. J Infect. 2016;73(5):419–26.PubMedCrossRef
97.
Tong SY, Davis JS, Eichenberger E, et al. Staphylococcus aureus infections: epidemiology, pathophysiology, clinical manifestations, and management. Clin Microbiol Rev. 2015;28(3):603–61.PubMedPubMedCentralCrossRef
98.
Skinner D, Keefer CS. Significance of bacteremia caused by Staphylococcus aureus. Arch Intern Med. 1941;68:851–75.CrossRef
99.
Spink WW, Hall WH. Penicillin therapy at the University of Minnesota hospitals: 1942–1944. Ann Intern Med. 1945;22:510–25.CrossRef
100.
De Kimpe SJ, Kengatharan M, Thiemermann C, et al. The cell wall components peptidoglycan and lipoteichoic acid from Staphylococcus aureus act in synergy to cause shock and multiple organ failure. Proc Natl Acad Sci USA. 1995;92(22):10359–63.PubMedPubMedCentralCrossRef
101.
Kengatharan KM, De Kimpe S, Robson C, et al. Mechanism of gram-positive shock: identification of peptidoglycan and lipoteichoic acid moieties essential in the induction of nitric oxide synthase, shock, and multiple organ failure. J Exp Med. 1998;188(2):305–15.PubMedPubMedCentralCrossRef
102.
Wang JE, Dahle MK, Yndestad A, et al. Peptidoglycan of Staphylococcus aureus causes inflammation and organ injury in the rat. Crit Care Med. 2004;32(2):546–52.PubMedCrossRef
103.
Zacharowski K, Berkels R, Olbrich A, et al. The selective guanylate cyclase inhibitor ODQ reduces multiple organ injury in rodent models of gram-positive and gram-negative shock. Crit Care Med. 2001;29(8):1599–608.PubMedCrossRef
104.
Toledo AG, Golden G, Campos AR, et al. Proteomic atlas of organ vasculopathies triggered by Staphylococcus aureus sepsis. Nat Commun. 2019;10(1):4656.PubMedPubMedCentralCrossRef
105.
Kim HK, Missiakas D, Schneewind O. Mouse models for infectious diseases caused by Staphylococcus aureus. J Immunol Methods. 2014;410:88–99.PubMedPubMedCentralCrossRef
106.
Beno DW, Uhing MR, Goto M, et al. Chronic Staphylococcal enterotoxin B and lipopolysaccharide induce a bimodal pattern of hepatic dysfunction and injury. Crit Care Med. 2003;31(4):1154–9.PubMedCrossRef
107.
Silasi R, Keshari RS, Regmi G, et al. Factor XII plays a pathogenic role in organ failure and death in baboons challenged with Staphylococcus aureus. Blood. 2021;138(2):178–89.PubMedPubMedCentralCrossRef
108.
Silasi R, Keshari RS, Lupu C, et al. Inhibition of contact-mediated activation of factor XI protects baboons against S aureus-induced organ damage and death. Blood Adv. 2019;3(4):658–69.PubMedPubMedCentralCrossRef
109.
Zhang X, Hu X, Rao X. Apoptosis induced by Staphylococcus aureus toxins. Microbiol Res. 2017;205:19–24.PubMedCrossRef
110.
Shaukat A, Shaukat I, Rajput SA, et al. Ginsenoside Rb1 protects from Staphylococcus aureus-induced oxidative damage and apoptosis through endoplasmic reticulum-stress and death receptor-mediated pathways. Ecotoxicol Environ Saf. 2021;219:112353.PubMedCrossRef
111.
Haslinger-Löffler B, Wagner B, Brück M, et al. Staphylococcus aureus induces caspase-independent cell death in human peritoneal mesothelial cells. Kidney Int. 2006;70(6):1089–98.PubMedCrossRef
112.
Wang X, Li H, Wang J, et al. Staphylococcus aureus extracellular vesicles induce apoptosis and restrain mitophagy-mediated degradation of damaged mitochondria. Microbiol Res. 2023;273:127421.PubMedCrossRef
113.
Stelzner K, Winkler AC, Liang C, et al. Intracellular Staphylococcus aureus perturbs the host cell Ca2+ homeostasis to promote cell death. mBio. 2020;11(6):e02250-20.PubMedPubMedCentralCrossRef
114.
Wen SH, Lin LN, Wu HJ, et al. TNF-α increases Staphylococcus aureus-induced death of human alveolar epithelial cell line A549 associated with RIP3-mediated necroptosis. Life Sci. 2018;15(195):81–6.CrossRef
115.
Chang AL, Ulrich A, Suliman HB, et al. Redox regulation of mitophagy in the lung during murine Staphylococcus aureus sepsis. Free Radic Biol Med. 2015;78:179–89.PubMedCrossRef
116.
Bitto NJ, Cheng L, Johnston EL, et al. Staphylococcus aureus membrane vesicles contain immunostimulatory DNA, RNA and peptidoglycan that activate innate immune receptors and induce autophagy. J Extracell Vesicles. 2021;10(6):e12080.PubMedPubMedCentralCrossRef
117.
Maldonado N, López-Hernández I, García-Montaner A, et al. Whole-genome characterisation of Escherichia coli isolates from patients with bacteraemia presenting with sepsis or septic shock in Spain: a multicentre cross-sectional study. Lancet Microbe. 2024;5(4):e390–9.PubMedCrossRef
118.
Stoll BJ, Puopolo KM, Hansen NI, et al. Early-onset neonatal sepsis 2015 to 2017, the rise of Escherichia coli, and the need for novel prevention strategies. JAMA Pediatr. 2020;174(7):e200593.PubMedPubMedCentralCrossRef
119.
Tabah A, Buetti N, Staiquly Q, et al. Epidemiology and outcomes of hospital-acquired bloodstream infections in intensive care unit patients: the EUROBACT-2 international cohort study. Intensive Care Med. 2023;49(2):178–90.PubMedPubMedCentralCrossRef
120.
McHale TM, Garciarena CD, Fagan RP, et al. Inhibition of vascular endothelial cell leak following Escherichia coli attachment in an experimental model of sepsis. Crit Care Med. 2018;46(8):e805–10.PubMedCrossRef
121.
Jiang Y, Ma J, Long Y, et al. Extracellular membrane vesicles of Escherichia coli induce apoptosis of CT26 colon carcinoma cells. Microorganisms. 2024;12(7):1446.PubMedPubMedCentralCrossRef
122.
Yang HH, Jiang HL, Tao JH, et al. Mitochondrial citrate accumulation drives alveolar epithelial cell necroptosis in lipopolysaccharide-induced acute lung injury. Exp Mol Med. 2022;54(11):2077–91.PubMedPubMedCentralCrossRef
123.
Xiao K, Yang Y, Zhang Y, et al. Long-chain PUFA ameliorate enterotoxigenic Escherichia coli-induced intestinal inflammation and cell injury by modulating pyroptosis and necroptosis signaling pathways in porcine intestinal epithelial cells. Br J Nutr. 2022;128(5):835–50.PubMedCrossRef
124.
David L, Taieb F, Pénary M, et al. Outer membrane vesicles produced by pathogenic strains of Escherichia coli block autophagic flux and exacerbate inflammasome activation. Autophagy. 2022;18(12):2913–25.PubMedPubMedCentralCrossRef
125.
Tan Y, Wan HH, Sun MM, et al. Cardamonin protects against lipopolysaccharide-induced myocardial contractile dysfunction in mice through Nrf2-regulated mechanism. Acta Pharmacol Sin. 2021;42(3):404–13.PubMedCrossRef
126.
Pang J, Peng H, Wang S, et al. Mitochondrial ALDH2 protects against lipopolysaccharide-induced myocardial contractile dysfunction by suppression of ER stress and autophagy. Biochim Biophys Acta Mol Basis Dis. 2019;1865:1627–41.PubMedCrossRef
127.
Lin H, Ji F, Lin KQ, et al. LPS-aggravated ferroptosis via disrupting circadian rhythm by Bmal1/AKT/p53 in sepsis-induced myocardial injury. Inflammation. 2023;46(4):1133–43.PubMedCrossRef
128.
Roach DJ, Sridhar S, Oliver E, et al. Clinical and genomic characterization of a cohort of patients with Klebsiella pneumoniae bloodstream infection. Clin Infect Dis. 2024;78(1):31–9.PubMedCrossRef
129.
Kumar CK, Sands K, Walsh TR, et al. Global, regional, and national estimates of the impact of a maternal Klebsiella pneumoniae vaccine: a bayesian modeling analysis. PLoS Med. 2023;20(5):e1004239.PubMedPubMedCentralCrossRef
130.
Fostervold A, Raffelsberger N, Hetland MAK, et al. Risk of death in Klebsiella pneumoniae bloodstream infections is associated with specific phylogenetic lineages. J Infect. 2024;88(5):106155.PubMedCrossRef
131.
Antimicrobial Resistance Collaborators. Global burden of bacterial antimicrobial resistance in 2019: a systematic analysis. Lancet. 2022;399(10325):629–55.CrossRef
132.
Laakmann K, Eckersberg JM, Hapke M, et al. Bacterial extracellular vesicles repress the vascular protective factor RNase1 in human lung endothelial cells. Cell Commun Signal. 2023;21(1):111.PubMedPubMedCentralCrossRef
Nikouee A, Kim M, Ding X, et al. Beclin-1-dependent autophagy improves outcomes of pneumonia-induced sepsis. Front Cell Infect Microbiol. 2021;15(11):706637.CrossRef
135.
Singh VK, Almpani M, Maura D, et al. Tackling recalcitrant Pseudomonas aeruginosa infections in critical illness via anti-virulence monotherapy. Nat Commun. 2022;13(1):5103.PubMedPubMedCentralCrossRef
136.
Metais A, Torregrosa Diaz JM, Gallego Hernanz MP, et al. Efficacy of antibiotic short course for bloodstream infections in acute myeloid leukemia patients with febrile neutropenia: a retrospective comparative study. J Infect. 2022;84(1):1–7.PubMedCrossRef
137.
Beaufort N, Corvazier E, Mlanaoindrou S, et al. Disruption of the endothelial barrier by proteases from the bacterial pathogen Pseudomonas aeruginosa: implication of matrilysis and receptor cleavage. PLoS ONE. 2013;8(9):e75708.PubMedPubMedCentralCrossRef
138.
Hardy KS, Tuckey AN, Renema P, et al. ExoU induces lung endothelial cell damage and activates pro-inflammatory caspase-1 during Pseudomonas aeruginosa infection. Toxins (Basel). 2022;14(2):152.PubMedCrossRef
139.
Pendyala S, Gorshkova IA, Usatyuk PV, et al. Role of Nox4 and Nox2 in hyperoxia-induced reactive oxygen species generation and migration of human lung endothelial cells. Antioxid Redox Signal. 2009;11(4):747–64.PubMedPubMedCentralCrossRef
140.
Chen X, Guo J, Mahmoud S, et al. Regulatory roles of SP-A and exosomes in pneumonia-induced acute lung and kidney injuries. Front Immunol. 2023;15(14):1188023.CrossRef
141.
Dar HH, Tyurina YY, Mikulska-Ruminska K, et al. Pseudomonas aeruginosa utilizes host polyunsaturated phosphatidylethanolamines to trigger theft-ferroptosis in bronchial epithelium. J Clin Invest. 2018;128(10):4639–53.PubMedPubMedCentralCrossRef
142.
Dar HH, Anthonymuthu TS, Ponomareva LA, et al. A new thiol-independent mechanism of epithelial host defense against Pseudomonas aeruginosa: iNOS/NO• sabotage of theft-ferroptosis. Redox Biol. 2021;45:102045.PubMedPubMedCentralCrossRef
143.
Li H, Guan J, Chen J, et al. Necroptosis signaling and NLRP3 inflammasome cross-talking in epithelium facilitate Pseudomonas aeruginosa mediated lung injury. Biochim Biophys Acta Mol Basis Dis. 2023;1869(3):166613.PubMedCrossRef
144.
Yun M, Park SH, Kang DH, et al. Inhibition of Pseudomonas aeruginosa LPS-Induced airway inflammation by RIPK3 in human airway. J Cell Mol Med. 2022;26(21):5506–16.PubMedPubMedCentralCrossRef
145.
Planès R, Pinilla M, Santoni K, et al. Human NLRP1 is a sensor of pathogenic coronavirus 3CL proteases in lung epithelial cells. Mol Cell. 2022;82(13):2385-2400.e9.PubMedPubMedCentralCrossRef
146.
Li F, Li J, Wang PH, et al. SARS-CoV-2 spike promotes inflammation and apoptosis through autophagy by ROS-suppressed PI3K/AKT/mTOR signaling. Biochim Biophys Acta Mol Basis Dis. 2021;1867(12):166260.PubMedPubMedCentralCrossRef
147.
Li H, Liu L, Zhang D, et al. SARS-CoV-2 and viral sepsis: observations and hypotheses. Lancet. 2020;395(10235):1517–20.PubMedPubMedCentralCrossRef
148.
Peleman C, Van Coillie S, Ligthart S, et al. Ferroptosis and pyroptosis signatures in critical COVID-19 patients. Cell Death Differ. 2023;30(9):2066–77.PubMedPubMedCentralCrossRef
149.
Riegler AN, Benson P, Long K, et al. Differential activation of programmed cell death in patients with severe SARS-CoV-2 infection. Cell Death Discov. 2023;9(1):420.PubMedPubMedCentralCrossRef
150.
Ivanisenko NV, Seyrek K, Kolchanov NA, et al. The role of death domain proteins in host response upon SARS-CoV-2 infection: modulation of programmed cell death and translational applications. Cell Death Discov. 2020;6(1):101.PubMedPubMedCentralCrossRef
151.
Yang Y, Wu Y, Meng X, et al. SARS-CoV-2 membrane protein causes the mitochondrial apoptosis and pulmonary edema via targeting BOK. Cell Death Differ. 2022;29(7):1395–408.PubMedPubMedCentralCrossRef
152.
Huang X, Fan W, Sun J, et al. SARS-CoV-2 induces cardiomyocyte apoptosis and inflammation but can be ameliorated by ACE inhibitor captopril. Antiviral Res. 2023;215:105636.PubMedPubMedCentralCrossRef
153.
Jeong GU, Lyu J, Kim KD, et al. SARS-CoV-2 infection of microglia elicits proinflammatory activation and apoptotic cell death. Microbiol Spectr. 2022;10(3):e0109122.PubMedCrossRef
154.
Hao S, Ning K, Kuz CA, et al. SARS-CoV-2 infection of polarized human airway epithelium induces necroptosis that causes airway epithelial barrier dysfunction. J Med Virol. 2023;95(9):e29076.PubMedPubMedCentralCrossRef
155.
Liang L, Wang W, Chen J, et al. SARS-CoV-2 N protein induces acute kidney injury in diabetic mice via the Smad3-Ripk3/MLKL necroptosis pathway. Signal Transduct Target Ther. 2023;8(1):147.PubMedPubMedCentralCrossRef
156.
Liang K, Barnett KC, Hsu M, et al. Initiator cell death event induced by SARS-CoV-2 in the human airway epithelium. Sci Immunol. 2024;9(97):eadn0178.PubMedCrossRef
157.
Sun X, Liu Y, Huang Z, et al. SARS-CoV-2 non-structural protein 6 triggers NLRP3-dependent pyroptosis by targeting ATP6AP1. Cell Death Differ. 2022;29(6):1240–54.PubMedPubMedCentralCrossRef
158.
Liu L, Du J, Yang S, et al. SARS-CoV-2 ORF3a sensitizes cells to ferroptosis via Keap1-NRF2 axis. Redox Biol. 2023;63:102752.PubMedPubMedCentralCrossRef
159.
Deshpande R, Li W, Li T, et al. SARS-CoV-2 accessory protein Orf7b induces lung injury via c-Myc mediated apoptosis and ferroptosis. Int J Mol Sci. 2024;25(2):1157.PubMedPubMedCentralCrossRef
160.
Su J, Shen S, Hu Y, et al. SARS-CoV-2 ORF3a inhibits cGAS-STING-mediated autophagy flux and antiviral function. J Med Virol. 2023;95(1):e28175.PubMedCrossRef
161.
Jiao P, Fan W, Ma X, et al. SARS-CoV-2 nonstructural protein 6 triggers endoplasmic reticulum stress-induced autophagy to degrade STING1. Autophagy. 2023;19(12):3113–31.PubMedPubMedCentralCrossRef
162.
Han L, Zheng Y, Deng J, et al. SARS-CoV-2 ORF10 antagonizes STING-dependent interferon activation and autophagy. J Med Virol. 2022;94(11):5174–88.PubMedPubMedCentralCrossRef
163.
Zhang X, Yang Z, Pan T, et al. SARS-CoV-2 ORF3a induces RETREG1/FAM134B-dependent reticulophagy and triggers sequential ER stress and inflammatory responses during SARS-CoV-2 infection. Autophagy. 2022;18(11):2576–92.PubMedPubMedCentralCrossRef
164.
Hou P, Wang X, Wang H, et al. The ORF7a protein of SARS-CoV-2 initiates autophagy and limits autophagosome-lysosome fusion via degradation of SNAP29 to promote virus replication. Autophagy. 2023;19(2):551–69.PubMedCrossRef
165.
Gioia U, Tavella S, Martínez-Orellana P, et al. SARS-CoV-2 infection induces DNA damage, through CHK1 degradation and impaired 53BP1 recruitment, and cellular senescence. Nat Cell Biol. 2023;25(4):550–64.PubMedPubMedCentralCrossRef
166.
Kalil AC, Thomas PG. Influenza virus-related critical illness: pathophysiology and epidemiology. Crit Care. 2019;23(1):258.PubMedPubMedCentralCrossRef
Mayank AK, Sharma S, Nailwal H, et al. Nucleoprotein of influenza A virus negatively impacts antiapoptotic protein API5 to enhance E2F1-dependent apoptosis and virus replication. Cell Death Dis. 2015;6(12):e2018.PubMedPubMedCentralCrossRef
169.
Suo J, Zhao L, Wang J, et al. Influenza virus aggravates the ox-LDL-induced apoptosis of human endothelial cells via promoting p53 signaling. J Med Virol. 2015;87(7):1113–23.PubMedCrossRef
Gaba A, Xu F, Lu Y, et al. The NS1 protein of influenza a virus participates in necroptosis by interacting with MLKL and increasing its oligomerization and membrane translocation. J Virol. 2019;93(2):e01835-e1918.PubMedPubMedCentralCrossRef
172.
Kuriakose T, Man SM, Malireddi RK, et al. ZBP1/DAI is an innate sensor of influenza virus triggering the NLRP3 inflammasome and programmed cell death pathways. Sci Immunol. 2016;1(2):2045.CrossRef
173.
Ouyang A, Chen T, Feng Y, et al. The hemagglutinin of influenza A virus induces ferroptosis to facilitate viral replication. Adv Sci. 2024;11(39):e2404365.CrossRef
174.
Bell TAS, Velappan N, Gleasner CD, et al. Nonclassical autophagy activation pathways are essential for production of infectious Influenza A virus in vitro. Mol Microbiol. 2022;117(2):508–24.PubMedPubMedCentralCrossRef
175.
Wang X, Zheng T, Lin L, et al. Influenza A virus induces autophagy by its hemagglutinin binding to cell surface heat shock protein 90AA1. Front Microbiol. 2020;7(11):566348.CrossRef
176.
Bock FJ, Tait SWG. Mitochondria as multifaceted regulators of cell death. Nat Rev Mol Cell Biol. 2020;21(2):85–100.PubMedCrossRef
177.
Tsuchiya K, Nakajima S, Hosojima S, et al. Caspase-1 initiates apoptosis in the absence of gasdermin D. Nat Commun. 2019;10(1):2091.PubMedPubMedCentralCrossRef
178.
Zhang Y, Su SS, Zhao S, et al. RIP1 autophosphorylation is promoted by mitochondrial ROS and is essential for RIP3 recruitment into necrosome. Nat Commun. 2017;8(8):14329.PubMedPubMedCentralCrossRef
179.
Yang Z, Wang Y, Zhang Y, et al. RIP3 targets pyruvate dehydrogenase complex to increase aerobic respiration in TNF-induced necroptosis. Nat Cell Biol. 2018;20(2):186–97.PubMedCrossRef
180.
Gao M, Yi J, Zhu J, Minikes AM, Monian P, Thompson CB, Jiang X. Role of mitochondria in ferroptosis. Mol Cell. 2019;73(2):354-363.e3.PubMedCrossRef
Immer mehr Ärztinnen und Ärzte arbeiten angestellt in Praxen bzw. MVZ. Was im Arbeitsvertrag geklärt werden kann und sollte und wo Risiken liegen, erklärt Medizin- und Arbeitsrechtlerin Gabriele Leucht.
Medizinischen Rat von Chatbots auf der Basis sogenannter künstlicher Intelligenz haben laut Umfragen bereits knapp die Hälfte aller Erwachsenen schon einmal eingeholt. Welche Chancen und Risiken birgt das?
Bei erwachsenen, mechanisch beatmeten kritisch Kranken führt der Verzicht auf eine dauerhafte körperliche Fixierung im Vergleich zur routinemäßigen Anwendung von Handgelenkriemen nicht zu mehr Tagen mit Delirium oder Koma. Zu diesem Ergebnis kommt eine französische Studie.
Bei Kopfschmerzen unklarer Ursache kann eine okuläre Sonografie früh Hinweise auf einen erhöhten Hirndruck liefern und damit die Entscheidung für eine Lumbalpunktion erleichtern. Bei einer 37-jährigen Frau führte diese zur Diagnose einer Varizella-zoster-Meningitis.
