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Hypothesis

Priming, Triggering, Adaptation and Senescence (PTAS): A Hypothesis for a Common Damage Mechanism of Steatohepatitis

Diagnostic & Research Center for Molecular Biomedicine, Diagnostic & Research Institute of Pathology, Medical University of Graz, A-8010 Graz, Austria
*
Author to whom correspondence should be addressed.
Int. J. Mol. Sci. 2021, 22(22), 12545; https://doi.org/10.3390/ijms222212545
Submission received: 30 September 2021 / Revised: 18 November 2021 / Accepted: 19 November 2021 / Published: 21 November 2021
(This article belongs to the Special Issue Recent Advances in Molecular Research of Metabolic Disorders)

Abstract

:
Understanding the pathomechanism of steatohepatitis (SH) is hampered by the difficulty of distinguishing between causes and consequences, by the broad spectrum of aetiologies that can produce the phenotype, and by the long time-span during which SH develops, often without clinical symptoms. We propose that SH develops in four phases with transitions: (i) priming lowers stress defence; (ii) triggering leads to acute damage; (iii) adaptation, possibly associated with cellular senescence, mitigates tissue damage, leads to the phenotype, and preserves liver function at a lower level; (iv) finally, senescence prevents neoplastic transformation but favours fibrosis (cirrhosis) and inflammation and further reduction in liver function. Escape from senescence eventually leads to hepatocellular carcinoma. This hypothesis for a pathomechanism of SH is supported by clinical and experimental observations. It allows organizing the various findings to uncover remaining gaps in our knowledge and, finally, to provide possible diagnostic and intervention strategies for each stage of SH development.

1. Introduction

1.1. What Is Steatohepatitis?

Fatty liver disease is a multifactorial health problem with world-wide increasing prevalence [1,2,3,4]. It is the result of a complex interplay between the liver, adipose tissue, and intestine. According to the aetiology, fatty liver disease is categorized as alcoholic or non-alcoholic. The latter (i.e., non-alcoholic fatty liver disease; NAFLD) arises in the absence of significant alcohol consumption and is the hepatic manifestation of the metabolic syndrome, characterized by visceral obesity, dyslipidaemia, type II diabetes mellitus, insulin resistance (IR), elevated arterial blood pressure, and related cardio-vascular disorders [3,5,6,7,8]. Both disorders may progress from simple steatosis to the necro-inflammatory lesions of alcoholic (ASH) and non-alcoholic (NASH) steatohepatitis (SH), respectively, with cirrhosis and hepatocellular carcinoma as late-onset sequelae [2,3,9].
Fatty liver disease in humans is variable in its morphologic and clinical appearance, depending on the aetiology, severity, and stage [10,11]. It may be caused by a broad spectrum of insults, including alcohol abuse and metabolic disorders related to an imbalance of nutrient intake, energy expenditure, and acute or chronic drug intoxication (drug-induced liver injury, DILI) [12,13,14,15]. Its severity ranges from the relatively benign and reversible fatty liver (termed simple steatosis) to steatohepatitis (SH). SH is characterised by inflammation, hepatocellular damage with necrosis, apoptosis, ballooning, derangement or loss of the keratin intermediate filament cytoskeleton and formation of protein aggregates termed Mallory–Denk bodies (MDBs) [5,16,17], altered energy metabolism [18], oxidative stress, as well as damage to and deficient quality control of mitochondria [19,20,21,22,23,24,25]). It can eventually progress to cirrhosis and hepatocellular carcinoma [2,3,9,26].

1.2. Current State of Knowledge about the Molecular Disease Mechanism

Human SH is characterised by alterations that include morphological changes as key diagnostic criteria [5,16,17], metabolic disturbances [18], oxidative stress [19,23,27,28,29], and inflammation. In addition, it is justified to add cirrhosis and hepatocellular carcinoma (HCC) [9,26] to the phenotypical characteristics of the disease.
A remarkable feature of SH is that it can be the result of a broad aetiological spectrum that nevertheless converges to the same phenotype.
A vast body of literature has accrued over the past decades reporting numerous details regarding contributors to SH, including but not limited to metabolic alterations involving energy metabolism [18], oxidative damage [19,23,27,28,29], mitochondrial dysfunction [19,23,25,27,30], hypoxic response [31,32,33,34,35], and senescence [36,37,38,39], as well as the roles of adipose tissue [40], proinflammatory cytokines [41], and the microbiome [42,43,44]. However, there is presently only limited knowledge about the pathogenic mechanism responsible for the severe forms, particularly the necro-inflammatory lesions in ASH and NASH, respectively. To some extent this is due to the fact that these chronic diseases remain clinically silent for a rather long time (1–5 years for ASH, even longer for NASH, where the onset of insult is often undefined [45]). Moreover, no reliable non-invasive screening procedures are available—the gold standard for the diagnosis of SH is still histopathological examination of liver biopsy. Hence, ASH and NASH are usually detected at a late stage of their development, which explains the insufficient knowledge about their early stages and molecular triggers. Furthermore, while allowing histological assessment, formalin-fixed biopsy material precludes obtaining sufficiently detailed functional data, e.g., about causes and consequences of molecular alterations and mitochondrial function. Animal models have supported the mechanistic research in this respect (see also below) [23,24,25,46] but often give only circumstantial evidence for human SH [16].

1.3. A Mechanistic Framework Is Urgently Needed

Although a considerable amount of data has accumulated over the past decades, the available information has not yet been assembled into a consistent molecular framework that covers all the stages of SH development, including sequelae. Because the processes are highly interconnected, it is difficult to distinguish between causes and consequences, which is, however, indispensable to understand the pathogenesis and to discover and eventually fill gaps in our understanding.
In a complex disease such as NASH, understanding the pathogenesis requires knowledge of the evolution of the disease process from initial damage, via the ‘mature’ phenotype, to sequelae. This is indispensable for the design of screening methods and of efficient therapeutic or preventive strategies. Many of the alterations described in the literature are not drivers but rather consequences of underlying, primary molecular events that affect metabolism, stress response, as well as intra- and intercellular signalling. Identifying drivers, in their chronological sequence, is therefore a useful strategy for the elucidation of the causal chain finally responsible for the fully developed disease.
The evidence for our proposed mechanism of SH development presented below often resulted from human and animal studies without direct reference to SH. However, these studies can explain basic underlying principles that should be confirmed (or refuted) in the context of SH. Moreover, some aspects of the model we present here are substantially underpinned by experimental findings, while other parts, specifically the later phases, are more hypothetical and will need specific research.

1.4. How Can a Mechanistic Framework Be Found?

The working strategy is to uncover events that causally link the primary damage (i.e., immediately resulting from the primary insult) to the final (predominantly morphologic) phenotype, including potential progression to sequelae. This approach allows distinguishing those events that eventually lead to the phenotype from other alterations that occur along the transition from normal to diseased liver. As usual in natural sciences, the mechanism that we describe here is not the only possible one, but it is consistent with a causal chain that is supported by a large body of findings.

2. A Proposed Common Damage Mechanism (CDM) of SH: Priming/Sensitizing, Triggering, Adaptation and Senescence/Sequelae (PTAS)

It is striking that different aetiologies can lead to the SH phenotype. Since each aetiology initially produces a specific type of insult it is likely that the different primary insults converge on a common damage mechanism (CDM). As shown below, the initial damage by the different aetiologies resembles the well-known hypotheses about two or more ‘hits’ [47,48]. Convergence on a CDM can occur early in disease development if the different aetiologies produce similar insults. Alternatively, different primary insults may converge late on a CDM; in this case it may be a secondary mechanism that leads to the SH phenotype which is not a direct consequence of the primary damage.
We favour the second option because the mature phenotype of NASH takes a long time to develop after the initial damage. It was claimed that simple steatosis may be a precursor state for NASH [11,26,49]. We will show in the next section how steatosis fits into a mechanistic model as an initial cause. Moreover, from our previously published work [28,50] and the work of others, we deduce that the development of the SH phenotype requires at least two sequential or parallel insults to hepatocytes prior to the CDM. In more general terms this was already proposed as the ‘two (or multiple)-hit hypothesis’ [47,48], however without the addition of a CDM. Furthermore, from observations in humans and experimental animals we consider the CDM to involve metabolic adaptations following the initial, acute insults that allow tolerating the damage and maintaining basic liver function, in the sense of a chronic disease. Adaptation appears to result in cellular senescence and thus at least temporarily prevents neoplastic transformation that might otherwise be triggered by adaptive metabolic alterations. We propose that adaptation is the committed step of the SH phenotype development; it constitutes a metabolic state of the liver that is essentially functional, but different from normal metabolism, and may be responsible for inflammation through the senescence-associated secretory phenotype (SASP).
It is likely that the altered metabolic state reached during adaptation persists even when the morphological phenotype is resolved after some time of absence of the aetiologic cause. This is observed in ASH patients upon ethanol withdrawal, or in certain toxicity-induced mouse models [51], when the phenotype-inducing toxin is withdrawn; upon re-intoxication, the morphological phenotype will reappear rapidly—actually much more rapidly than upon first-time intoxication. This indicates that the CDM induces an altered metabolic state (the secondary damage mechanism mentioned above), which is responsible for the complex morphological phenotype and the well-known sequelae. Of note, this concept allows that the altered metabolic state will provide susceptibility to any further insult that can induce acute damage.
We propose a four-stage mechanism for SH: (i) a priming event that by itself induces no, or only minor, hepatocellular damage but impairs stress defence capacity and thus sensitizes the cell to further insults; (ii) a trigger that induces the actual (or acute) damage, which is exacerbated by the diminished defence capacity. We posit that in absence of prior priming, the trigger alone can induce some damage, which is, however, mitigated by the defence and repair capacity of the cell. Finally, (iii) irreversible adaptation of the metabolic state initiates a CDM that preserves basal hepatocellular and organ function under sustained conditions of priming and triggering, develops the characteristics of the SH phenotype, and (iv) ends up in cellular senescence that, after escape from this state, favours development of hepatocellular carcinoma (HCC).

3. Experimental Evidence Supporting PTAS

3.1. Priming Leads to Compromised Stress Defence

Experimental evidence for the existence of the first two stages, namely priming and triggering, has been obtained with a mouse model based on prolonged dietary administration of the porphyrinogenic substance 3,5-diethoxycarbonyl-1,4-dihydrocollidine (DDC; 0.1% (wt/wt) for 8–10 weeks) leading to inhibition of ferrochelatase [52,53]. In this model, acute hepatocellular damage with focal apoptotic and necrotic cell death occurs early (during the first 2–5 weeks of DDC administration), whereas development of the SH phenotype requires 8–10 weeks of continuous DDC treatment [50,51]. A similar situation was observed in ferrochelatase-deficient mice [29]. In the acute intoxication stage, DDC primes the liver by suppression of the Nrf2-dependent stress response [50]. In this context, a significant downregulation of Pparα and PPARα-controlled genes [50] occurred, affecting not only genes involved in energy and lipid metabolism but also typical stress-response genes, such as Gclc, Sod1, Sod2, and Hmox1. The effects of priming persisted for several months after cessation of DDC administration (P.M.A., et al., unpublished data), probably as a result of an irreversible metabolic alteration due to the interplay of PPARα-mediated lowering of stress response [50] in combination with c-Myc [54,55], succinate efflux from hepatocytes [56] leading to persistent hypoxic signalling [57,58]. This may explain the rapid reappearance of the SH phenotype during re-intoxication [59]. The trigger may be based on the dysfunction of mitochondrial complex II [28], probably caused by inhibition of haem production and resulting in impaired formation of haem-containing proteins. Table 1 summarizes the priming factors.
Of note, downregulation of Ppara and PPARα-dependent genes by itself does not lead to damage and does not (yet) produce the SH phenotype but only lowers resilience towards stress. That is the reason why Ppara-deficient mice do not spontaneously develop SH unless there is an additional trigger, such as ethanol [60].
Table 1. Factors that prime the liver for extensive damage after triggering.
Table 1. Factors that prime the liver for extensive damage after triggering.
FactorMechanismModel
Porphyrinogens (DDC, griseofulvin); liver toxinsAhR ↑ → PPARα ↓ & c-Myc ↓ → Nrf2-dependent genes ↓Intoxication mouse models [16,28,50,51,52,53,61,62]; DILI [12,13,14]
High-fat diet 1AhR ↑ → PPARα ↓ & c-Myc ↓ → Nrf2-dependent genes ↓HFD mouse model; human NASH [16,63,64,65,66,67,68]
High-fat diet 1Palmitoyl-CoA → NNT inhibition → NADPHmito ↓ → GSHmitoHFD mouse model; human NASH [69,70,71,72,73]
Excess keratin 8Impaired mitochondrial QC via Pirh2Keratin 18−/− mouse model, human NASH [16,59,74,75,76,77,78,79,80,81]
1 HFD in normal experiments does not lead to the SH phenotype [16], most likely because HFD only primes but needs a separate trigger.

3.1.1. Mechanistic Implications of PPARα Downregulation

PPARα is predominantly regarded as a regulator of energy (lipid) metabolism in the liver. However, PPARα also activates the Nrf2-dependent stress response [50], very likely via c-Myc [54,55]. It is connected with hypoxic signalling and mitochondrial QC [57,82,83].
The initial downregulation of PPARα may be caused by activation of the aryl hydrocarbon receptor [67] by oxidative stress [84], or (e.g., in the DDC model) by porphyrin degradation products, such as bilirubin [68]; this notion is corroborated by observations in ferrochelatase-deficient mice that also develop porphyria and the SH phenotype [29]. Another possibility is that oxidative stress directly leads to stabilization of hypoxia-inducible factors [57,85], which then contribute to persistent PPARα downregulation at later stages of the disease. A similar pattern of downregulation of stress response genes was described for mice fed a high-fat diet (HFD) [86], although these mice did not develop the SH phenotype after prolonged HFD feeding. This may be due to a lack of a suitable trigger, which could be, e.g., an imbalance of the keratin 8/18 content of the cell [65,74,75,79] (see below), or due to insufficient time for phenotype development. Similarly, it was shown recently that hepatocyte-specific ablation of PPARα induces steatosis, but not SH [66].
Cross-talk between PPARα and the Nrf2-controlled stress response has been shown to affect the expression of a wide variety of genes [87,88,89]. Since suppression of stress response also affects the redox state of the hepatocyte and its mitochondria, in particular the glutathione (GSH) redox buffer, this is probably a key element of the reduced resilience to subsequent insults, revealed by impaired mitochondrial function and increased oxidative damage [28].
Recent studies showed that expression of PPARα target genes may be amplified by c-Myc [90], which enables the full stress response via Nrf2 [54,55]. Conversely, downregulation of PPARα reduces this amplification.
Another factor that determines stress response is the mitochondrial NADPH pool, which is required to keep the sulfhydryl cofactors, specifically GSH, in a reduced state. Mitochondrial NADPH is regenerated from NADP+ in the tricarboxylic acid (TCA) cycle and, also, with NADH as the electron donor, through nicotinamide nucleotide transhydrogenase (NNT), a proton-translocating enzyme located in the inner mitochondrial membrane [70,91]. In addition to the well-known genetic NNT defect in the C57BL/6J mouse substrain [72,73,92,93] that severely affects their mitochondrial stress-resilience, the function of NNT may be compromised by a variety of biogenic inhibitors [69]; one of them is palmitoyl-CoA, which may be relevant in HFD-fed animals and human steatosis [94].

3.1.2. Priming in Human SH

Since priming occurs early in the development of SH in humans, persistent markers are likely to serve as indicators. The persistent downregulation of PPARα found in mice [50] is also found in patients with SH [61] but not in patients with simple steatosis [31,62,95]. Agonists for PPARα and other isotypes showed promising therapeutic effects [96,97] similar to the observations in mice, A recent review and meta-analysis of the efficacy of the PPARα/δ agonist elafibranor in clinical trials reported that it improved most metabolic parameters, particularly serum lipid profiles and liver enzymes [98].

3.1.3. Priming Is a Factor Determining Sexual Dimorphism

Recently it was shown that the downregulation of PPARα exhibits pronounced sexual dimorphism in diet-induced NAFLD in mice, as female mice were protected from the adverse effects of a hypercaloric diet [99]. A similar sexual dimorphism of PPARα and PPARα-controlled genes has been shown in humans with simple steatosis, moderate fibrosis, and absence of ballooning and, therefore, did not relate to SH [99].

3.2. Triggering Induces Severe Hepatocellular Injury

The factors that prime and trigger may differ with regard to aetiology. In many cases, the triggering event cannot be clearly separated from priming. For example, in the DDC-intoxication mouse model the induction of porphyria may be responsible both for priming, by lowering PPARα expression, as well as for triggering, since the production of ferrohaem is inhibited leading to mitochondrial dysfunction. This is most likely due to impaired synthesis of haem-proteins of the electron transport chain, resulting in impairment of the tricarboxylic acid cycle and increased formation of reactive oxygen species (ROS) [28,100,101,102]. Particularly prominent was the inhibition of succinate:quinone oxidoreductase (SQR; also termed succinate dehydrogenase, SDH; or complex II) in the DDC mouse model within about two weeks [28,50], accompanied by substantial weight loss and 40% lower hepatic ATP content. Consequences of mitochondrial dysfunction, such as reduced ATP content, were also observed in human ASH [103] and NASH [22,30]. Another cause of SQR inhibition may be superoxide production by mitochondrial respiratory complex I [104]. Succinate produced in the TCA cycle is oxidized by SQR with concomitant reduction of ubiquinone to ubiquinol in the mitochondrial electron transport chain (TCA). Besides contributing directly to oxidative damage by production of ROS [71,102,105], SQR dysfunction can impair both ETC and TCA cycles, leading to increased succinate and succinyl-CoA concentrations. Succinyl-CoA is an inhibitor of the upstream TCA cycle enzymes, α-ketoglutarate dehydrogenase and citrate synthase, which is the likely cause of the reduced ATP synthesis after inhibition of SQR activity [28]. On the other hand, succinate induces hypoxic signalling by stabilizing hypoxia inducible factor 1α (HIF-1α) and Hif-2α through (product) inhibition of the HIFα prolyl hydroxylases (PHD) [34].
In line with observations in other models and in human SH, elevated succinate is a signal of liver damage. It can bind to a G-protein-coupled succinate receptor, SUCNR1 [106], that activates hepatic stellate cells [107,108,109], triggers an immune response [56,110,111], and impairs the mitochondrial quality control machinery [112]. Moreover, it has been demonstrated in an acetaminophen-toxicity model that, by uncoupling the succinate dehydrogenase and quinone reductase activities of SQR with methylene blue, the adverse effects of SQR inhibition can be relieved [113].
Interestingly, it was recently described that uncoupling protein 1 (UCP1) in adipocytes [40], which is also expressed in the kidney [114] and in hepatocytes [115], may influence the systemic and hepatic succinate pools [40], which is a possible link between obesity and liver disease.
The impairment of the TCA cycle eventually lowers formation of NADPH, required for the regeneration of (mitochondrial) glutathione (GSH). Together with the unbalanced utilization of TCA cycle substrates, which favours ROS formation, the reduced antioxidant capacity of the GSH pool may promote damage of key enzymes of the TCA cycle and ETC.

3.2.1. Triggering in Human SH

Similar to some animal models, the identification of a triggering insult in human SH is hampered by the difficulty to choose the right time, i.e., before adaptive processes start; early events in the development of SH can usually not be detected in isolation. However, several consequences persist for a longer time period, such as lower hepatic ATP levels [30,116], succinate efflux [101,106], and inflammatory processes [56], accompanied by oxidative and mitochondrial damage. Damage is not a result of priming alone, and it persists during adaptation to some extent since it cannot be completely mitigated, e.g., in NASH patients, the activities of all respiratory electron transfer complexes were found reduced, most pronouncedly complex II [20,22,71]. Table 2 summarizes triggers for mitochondrial damage and Table 3 the outcome of both priming and triggering.

3.2.2. Triggering by Mitochondrial Dysfunction May Involve the Intermediate Filament Cytoskeleton

Mice fed a high-fat diet (HFD) show reduced expression of PPARα [63,64,86,118,119], but normally do not produce the SH phenotype, probably due to lack of a suitable trigger or too short duration of treatment. Mitochondrial dysfunction, prominent in human SH [19,27,120], was identified as the triggering event in the DDC-model as a consequence of haem deficiency, leading predominantly to inhibited function of SQR [28]. A first indication of how cytoskeletal elements could lead to mitochondrial dysfunction came from early experiments where griseofulvin-treated mice recovered for one month on a normal diet, upon which the SH phenotype disappeared. Re-challenge of these mice with either griseofulvin or colchicine (but not lumicolchicine or cytochalasin B) led to rapid reappearance of the SH phenotype [121]. In contrast to DDC, both griseofulvin and colchicine [122] interact with the tubulin cytoskeleton, and microtubules were described to be required for MDB formation [123]. Re-challenging DDC- or griseofulvin-fed mice after recovery with DDC achieves the same effect, and an excess of keratin 8 over keratin 18 was required to precipitate the SH phenotype [51,59,75,81,124,125].
The implication of keratin 8 (or the ratio keratin 8/18) as a trigger was corroborated by the finding that HFD-fed mice overexpressing keratin 8 developed the SH phenotype [65], which may be related to their propensity to be deposited as β-pleated sheet aggregates under these conditions [124]. Moreover, keratin 18-deficient mice (which only express keratin 8) spontaneously develop the SH phenotype (MDBs containing only keratin 8 are formed) [79]. Aggregation and deposition of misfolded proteins do probably not contribute to a CDM [125]. However, it is difficult to assess whether they are causally involved or merely represent a marker of the underlying processes. Increased levels of misfolded protein oligomers are known to be the toxic principle in protein misfolding diseases; therefore, they may contribute to maintaining the disease process, e.g., by sustaining mitochondrial dysfunction and ROS production (reviewed in [125,126]).
It is as yet unclear how the imbalance of keratin 8 and 18 triggers mitochondrial dysfunction; however, one possibility is the disturbance of intracellular mitochondrial distribution after the disruption of the interaction of the E3 ubiquitin ligase Pirh2 with the keratin intermediate filament, a process that may alter mitochondrial quality control [74]. Since this interaction is affected by phosphorylation of either Pirh2 [74] or the keratins, this may explain why the genetic background (in mice and humans) [75,78,80] plays an important role for the susceptibility for developing the SH phenotype. Pirh2 regulates stability and degradation of p53 and c-Myc [76] and could, thus, contribute to the persistent downregulation of the stress response [55,90] and to persistent mitochondrial damage [127,128,129].
Interestingly, Pirh2 also targets HuR [130], which is part of an RNA binding protein network that is activated during the HIF-2α-mediated hypoxic response [131]. HuR was reported to activate p62/SQSTM1-dependent selective autophagy in several cell types [132,133,134] and prevent steatosis in high-fat diet fed mice [135], possibly by increasing the stability of Pten mRNA. Moreover, HuR is also centrally involved in cell cycle regulation and might therefore link the immediate damage to adaptation and senescence [77]. This hypothesis might be tested, e.g., by using the keratin 18-deficient mouse model for SH.

3.3. Adaptation—Mitigation of Acute Hepatocyte Damage and Involvement of Other Cell Types

Adaptation, leading to the SH phenotype and probably transitioning to senescence, is the most complex part of the PTAS model since several different processes are deployed both in parallel and in succession: the hypoxic response, mediated by Hif-1 and Hif-2, the wound-healing response, inflammation, autophagy and mitophagy, among others. Therefore, different aspects of this phase may be observable at different time-points, and may partially obscure each other, in particular in human SH, but also in the better controlled environment of animal models. Therefore, more evidence specifically for this phase is needed for better understanding.
As indicated above, mitochondrial damage leads to impairment of the capacity of the TCA cycle to produce ATP, probably due to defective SDH, and succinate efflux into the cytosol. Elevated levels of succinate were described as a paracrine liver damage signal [106] and elicit a (pseudo)hypoxic response, similar to ischemia [136], by relieving the inhibition of Hif-1/2α through product inhibition of PHD. It allows survival of the hepatocyte but is also proto-oncogenic: succinate links metabolic deregulation to initiation of carcinogenesis, by transformation of cell function [137] and suppression of DNA repair [138]. These events, in addition to oxidative stress, trigger the development of a senescent state, especially the DNA-damage response (DDR) that is known to induce cell-cycle arrest, possibly as a means to prevent neoplastic hyperproliferation. Moreover, besides the activation of stellate cells [101] and triggering neoplastic transformation [139,140], succinate is involved in cross-talk of adipocytes with hepatocytes [40] and activation of immune cells [56,110].
In fibroblasts, both Hif-1 and Hif-2 induction leads to cell cycle arrest, without hypoxia, which may also represent senescence [141]. Interestingly, while the activation of hypoxia-dependent gene products occurs via Hif-1α, Hif-2α seems to involve translational modulation that increases formation of specific hypoxia-related proteins [142]. It was found that Hif-2α initially leads to a shutdown of transcription, while chronic hypoxia induces the Hif-2α-mediated remodelling of the central carbon metabolism by activating a specific translation initiation factor, eIF5B [143]; this process may also be operative during the chronic pseudohypoxic state described above. Recently, a network of RNA-binding proteins activated by Hif-2α was described [131], one of which, HuR, appears to be a target of the E3 ubiquitin ligase Pirh2 [130], which is involved in mediating mitochondrial localization [74]. This constitutes another link between hypoxia and cell cycle arrest as a consequence of the DDR [77].
Of note, Hif-2α was also described to activate the mTORC1 complex and to override the inhibition by Hif-1α in the liver and lung [144]. Interestingly, in peripheral blood mononuclear cells, upregulated expression of hypoxia-inducible genes was found in patients with chronic liver disease [145]. Hypoxic signalling might also be triggered focally by reduced hepatic microcirculation [145,146].

Mitigation of Damage and Restoration of Stress Response

Mouse models have shown that the initially suppressed stress response is restored [28], and typical stress proteins, such as haem oxygenase 1 or SQSTM1/p62 [51], are upregulated. Upregulation of SQSTM1/p62 is an important factor in the formation of MDBs [59,125,147,148,149] and is involved in autophagy and stress defence [150,151,152,153]. SQSTM1/p62 plays a role in the activation of stress response, by inducing selective autophagy and mitophagy; both may be independent [154] or dependent on a PINK1/PARKIN-related mechanism [155,156]. Mitophagy is apparently enhanced by ULK1 [155,157,158,159,160] that enhances binding to PINK1. On the other hand, SQSTM1/p62 directly activates the Nrf2-dependent stress response by interacting with Keap1, leading to its proteasomal degradation [153,161,162].
Moreover, ULK1 (re-)activates the stress response by enhancing the binding of SQSTM1/p62 to KEAP1 [157] followed by its autophagic clearance [162,163] and activation of the Nrf2-dependent induction of stress response proteins [164,165]. Interestingly, however, a splice-variant of SQSTM1/p62 lacking the Keap1-interacting region suppressed the stress response by increasing the amount of Keap1 [166].
Adaptation appears also to involve the activation of developmental pathways that are primarily responding to liver injury (excellently reviewed in [167]), such as Notch [168], possibly induced in stellate cells by osteopontin released from hedgehog-activated hepatocytes [169] as a response to injury [170].
Although sirtuins (Sirts) are probably not drivers of the CDM, they may play an important role in linking priming and triggering to the adaptive process and ageing. Therefore, and since the implications of NAD+ metabolism and Sirts with the CDM is beyond the scope of this paper, we provide only a few examples for this link; the dependence of Sirts on NAD+ implicates energy metabolism [171]. The deacetylase and desuccinylase activities, specifically of Sirt3 (acting exclusively on mitochondrial proteins), regulate mitochondrial function and the stress response [172]. Activation of the aryl hydrocarbon receptor (e.g., by oxidized fatty acids) deactivates Sirt3, which in turn lowers SOD2 activity and contributes to reduced mitochondrial stress defence [173]. This also results in altered energy homeostasis, which eventually favours sequelae such as HCC [172]. Sirt1 [174] and Sirt7 [175] have been implicated in the regulation of the hypoxic response via Hif-1 and thus constitute a potential link to ageing: Sirt1 is downregulated by autophagy in senescence [176], and there are links between overexpression of the cell-cycle regulator p16INK4a, Sirt1, and PPARα.

3.4. Senescence Prevents Neoplastic Transformation

Cellular senescence is a known tumour suppressor mechanism. Chronic liver disease, specifically SH, is often accompanied by senescence [39], in particular of hepatocytes [177,178,179,180,181] and cholangiocytes [182,183]. There is strong indication that many of the determinants for adaptation eventually culminate in senescence: mitochondrial dysfunction [100,184,185,186,187], metabolic alterations such as hypoxic signalling [129], oxidative stress [91,188,189], or downregulation of PPARα [190]. Importantly, there are potential routes to some of the hallmarks of the SH phenotype involving senescence, such as the formation of Mallory–Denk bodies [191].

3.5. Sequelae of SH—Cirrhosis and HCC

Hypoxic signalling has been found to be associated with hedgehog signalling, at least in paediatric NASH [35]. The hedgehog pathway plays a role in wound-healing and the response of the liver to injury [170]. Aberrant activation of hedgehog signalling was reported to be involved in development of HCC [192] by inducing metabolic changes in myofibroblasts that provide lactate for transformed hepatocytes [193]. The hedgehog pathway was found to be involved in progression of SH and development of fibrosis by inducing osteopontin [169]. Moreover, hedgehog/YAP signalling activates hepatic stellate cells in patients carrying the PNPLA3 I148M variant that is associated with risk of NASH progression [194,195], accompanied by metabolic changes, such as anaerobic glycolysis, favouring HCC development.

4. Discussion: Open Questions

We have presented the PTAS (priming, triggering, adaptation, senescence) model for SH that reflects our interpretation of the causal chain from aetiology to phenotype on the basis of experimental studies in correlation with observations in humans.
However, at present no single animal model can account for all elements of SH in humans with regard to aetiology and phenotype. One important future goal will therefore be the creation of an animal model that represents the PTAS mechanism whenever exposed to an aetiology that is relevant for human SH and exhibits the mature SH phenotype, including sequelae. This model will then serve as a strong support for the PTAS model and allow further studies on the development of SH.
In Figure 1 the PTAS model is summarized: a two-pronged initial, acute insult leads to liver (hepatocyte) damage, by simultaneous or sequential priming and triggering. Neither of these alone can produce damage that is severe or long-lasting enough to lead to the CDM and eventually to the SH phenotype. We suppose that after priming and triggering, the liver damage may be pronounced but reversible (after cessation of priming and triggering events); however, after adaptation, which is essential for survival and sustained basic liver function of the hepatocyte under persisting conditions of priming and triggering, a new steady-state is reached. To avoid metabolic breakdown (caused by impaired function of the ETC and reduced ATP production) the cell adapts by switching to an alternative metabolism, similar to that in hypoxia, which is, on the one hand, less efficient and results in reduced hepatocyte function but, on the other, allows survival. Whereas acute hypoxia causes transcriptional arrest, during chronic hypoxia Hif-2 activates a translational program that enables anaerobic metabolism [131,142,143]—a process that is known as the ‘Warburg-effect’ and is also found in cancer cells. Increased succinate efflux triggers activation of stellate cells that leads to increased fibrosis/cirrhosis and inflammation through activated macrophages, both mediated by the SUCNR1 receptor. Hepatocellular pseudohypoxia may be regarded as a precancerous state that can lead to formation of HCC; however, the events during and after adaptation may also enter the senescence state as an alternative route, which prevents neoplastic transformation. The disadvantage of senescence is that through the SASP, hepatocytes persist in a metabolically minimally active state and can no longer contribute to normal liver function. Due to the regenerative capacity of the liver, this loss may be temporarily compensated by cell proliferation and increase of liver mass; however, liver failure may be the final consequence.
There are similarities of the mechanism described here with processes occurring during ischemia and reperfusion injury, with the difference that under these conditions, instead of a pseudohypoxic state, true hypoxic signalling develops; however, there is clear evidence that succinate acts as a danger signal also in this case.
Moreover, although evidence for adaptation and senescence is rather widespread in the literature, there are still gaps in establishing mechanistic links between late and early parts of the mechanism.

4.1. Open Questions Regarding Priming and Triggering Events

It was already mentioned above that in the literature the function of PPARα is almost exclusively related to lipid metabolism rather than stress response. There is, however, no doubt about the causal involvement of PPARα in human SH [61,63,64,82,83,118,196,197]. Since the causes for downregulation of PPARα and stress response are not fully elucidated, it is necessary to also revisit the metabolism-related approach of SH pathogenesis [18,31,86,198,199,200], in addition to more work on the proposed mechanism by activation of the AhR [67].
The role of mitochondria in triggering human SH is still insufficiently elucidated, although mitochondrial involvement in SH pathogenesis has been known for a long time [19,20,23,24,25,27,46,113,120]. The crucial factor of mitochondrial dysfunction in the PTAS model is that it is aggravated by impaired stress response and leads to disruption of oxidative phosphorylation and the tricarboxylic acid cycle, followed by succinate release and hypoxic signalling.
Hence, the effect of succinate, which is clearly involved in SH, and is known as part of a damage-associated molecular pattern, is another topic that merits more attention [40,56,106,107,136,201,202]. It is probably closely associated with the (pseudo-)hypoxic state that is proposed to play a role at the transition to adaptation [32,35,57,141,145,203,204]. Their causal connection, specifically in the context of SH, is still insufficiently studied both in human SH and in animal models. It is quite interesting that succinate may also link the metabolic state of adipose tissue to that of the liver [40], and future studies might be facilitated by a PTAS-relevant animal model.

4.2. Open Questions Regarding Adaptation

In the PTAS model, adaptation mitigates the damage incurred during priming and triggering. During adaptation the hepatocyte reaches a new metabolic state that is stable but different from that of the ‘naïve’ hepatocyte. It allows mitigation of the damage at the expense of functionality and reduced resilience towards further insults. It eventually leads to the formation of the SH phenotype and constitutes a transition state towards neoplastic development or senescence.
Adaptation is characterized by a (pseudo-)hypoxic state, and hypoxic signalling has been described for human SH in the literature [32,33,35,57,144,145,203,204,205,206,207,208]. Adaptation is the committed step of SH development; however, this stage is quite complex, e.g., it is made persistent through several feedback loops to priming [57,205] and triggering [117,209,210], and many of these processes overlap and make their analysis difficult. Therefore, in human SH direct evidence is scarce, linking, e.g., succinate to hypoxic signalling, or the consequences of pseudohypoxia on cell cycle arrest and senescence. During adaptation, hepatocytes communicate their status to other cell types, such as stellate cells, e.g., by release of succinate [40,56,101,106,136,211]. Here, too, a specific PTAS-aligned animal model would be helpful.

4.3. Open Questions Regarding Senescence

Besides the fully adapted state, senescence is the second ‘endpoint’ in the PTAS model. We think that fully developed SH consists of both adapted hepatocytes and senescent cells (hepatocytes, cholangiocytes, and stellate cells). While there is a substantial body of literature that indicates senescence as a cellular state in SH [36,37,38,39,177,178,179,180,181,212,213,214,215,216,217,218], there are scarcely any functional data about induction of senescence and the fate of senescent cells, the potential for spreading of senescence via the SASP and escape from senescence with the potential for neoplastic transformation.

5. Conclusions: What We Presently Know and Do Not Know, and Some Diagnostic and Therapeutic Options

We think that the PTAS mechanism described above represents a good ‘working’ model of SH since it accommodates a broad spectrum of experimental findings (morphology, metabolic alterations, senescence, and sequelae) in animal models and humans [16]. We have certainly not included all phenomena associated with SH, either because we think that they are a result and not part of PTAS, or because they accrued from models that do neither produce the phenotype nor have the aetiology in common with human SH [16]. The PTAS model is intended as a starting point and guideline for focused research on this complex disease mechanism.

Diagnostic and Therapeutic Options

Armed with better understanding of the mechanistic underpinnings of the cascade that eventually leads to irreversible liver damage in SH, we provide a few ideas for diagnostic and therapeutic intervention. It should be noted, however, that besides the CDM, different initiating factors of different aetiologies also represent specific therapeutic targets [15].
A first diagnostic indicator may be serum succinate, which was also found elevated in perfusate of transplant livers that undergo ischemia reperfusion injury [219]; succinate may thus be a discriminant between steatosis and SH. It may also be of interest to use the expression profiles of peripheral blood mononuclear cells to detect the signature of hepatic (pseudo)hypoxic zones in the liver [145]. Finally, at later stages we may aim at detecting circulating senescence markers, such as soluble urokinase plasminogen activator receptor [218] or specific signatures in circulating extracellular vesicles [220,221].
The important therapeutic action at the transition from steatosis to SH will be to revert mitochondrial damage (e.g., by administration of mitochondrially targeted antioxidants/SDH uncouplers [222,223,224,225], which have already been used in clinical trials [46]), improve mitochondrial quality control [226], and prevent pseudohypoxia (e.g., by Hif-1 and Hif-2 inhibitors). Late-intervention therapy might be supported by senolytics [39,227].
There are still gaps in explaining the mechanism of MDB formation, the cause of ballooning, and similar morphological hallmarks. A functional role for keratins has been implied in this process [51,59,81,125], and apoptosis-triggered release of keratin 18 [228] and keratin 18-related tissue polypeptide-specific antigen into the circulation has been reported to be a diagnostic marker for SH [229]. However, with the aid of a framework model, it is possible that these gaps can be closed more rapidly and with greater focus.

Author Contributions

Conceptualization: P.M.A., H.D. and K.Z., Investigation: P.M.A., Original draft preparation: P.M.A., Review and editing: P.M.A., H.D. and K.Z. All authors have read and agreed to the published version of the manuscript.

Funding

No external funding was received for this work.

Acknowledgments

The authors thank Penelope Kungl for critical reading of the manuscript.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Geier, A.; Tiniakos, D.; Denk, H.; Trauner, M. From the origin of NASH to the future of metabolic fatty liver disease. Gut 2021, 70, 1570–1579. [Google Scholar] [CrossRef]
  2. Angulo, P. Nonalcoholic fatty liver disease. N. Engl. J. Med. 2002, 346, 1221–1231. [Google Scholar] [CrossRef] [Green Version]
  3. Brunt, E.M.; Neuschwander-Tetri, B.A.; Burt, A.D. Fatty liver disease: Alcoholic and non-alcoholic. In MacSween’s Pathology of the Liver, 6th ed.; Burt, A.D., Portmann, B.D.F.L., Eds.; Churchill Livingstone/Elsevier: London, UK, 2012; pp. 293–359. [Google Scholar]
  4. Younossi, Z.M.; Koenig, A.B.; Abdelatif, D.; Fazel, Y.; Henry, L.; Wymer, M. Global epidemiology of nonalcoholic fatty liver disease-Meta-analytic assessment of prevalence, incidence, and outcomes. Hepatology 2016, 64, 73–84. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  5. Kleiner, D.E. Histopathology, grading and staging of nonalcoholic fatty liver disease. Minerva Gastroenterol. Dietol. 2017, 64, 28–38. [Google Scholar] [CrossRef]
  6. Nalbantoglu, I.L.; Brunt, E.M. Role of liver biopsy in nonalcoholic fatty liver disease. World J. Gastroenterol. 2014, 20, 9026–9037. [Google Scholar] [CrossRef] [PubMed]
  7. Neuschwander-Tetri, B.A. Hepatic lipotoxicity and the pathogenesis of nonalcoholic steatohepatitis: The central role of nontriglyceride fatty acid metabolites. Hepatology 2010, 52, 774–788. [Google Scholar] [CrossRef] [PubMed]
  8. Marchesini, G.; Bugianesi, E.; Forlani, G.; Cerrelli, F.; Lenzi, M.; Manini, R.; Natale, S.; Vanni, E.; Villanova, N.; Melchionda, N.; et al. Nonalcoholic fatty liver, steatohepatitis, and the metabolic syndrome. Hepatology 2003, 37, 917–923. [Google Scholar] [CrossRef] [PubMed]
  9. Anstee, Q.M.; Reeves, H.L.; Kotsiliti, E.; Govaere, O.; Heikenwalder, M. From NASH to HCC: Current concepts and future challenges. Nat. Rev. Gastroenterol. Hepatol. 2019, 16, 411–428. [Google Scholar] [CrossRef] [PubMed]
  10. Working Group; Association of Pathologists; The Japan Society of Hepatology. Pathological Findings of NASH and NAFLD: For Guidebook of NASH and NAFLD, 2015: The Japan Society of Hepatology. Hepatol. Res. 2017, 47, 3–10. [Google Scholar] [CrossRef]
  11. Diehl, A.M.; Day, C. Cause, Pathogenesis, and Treatment of Nonalcoholic Steatohepatitis. N. Engl. J. Med. 2017, 377, 2063–2072. [Google Scholar] [CrossRef]
  12. McGill, M.R.; Jaeschke, H. Animal models of drug-induced liver injury. Biochim. Biophys. Acta Mol. Basis Dis. 2019, 1865, 1031–1039. [Google Scholar] [CrossRef]
  13. McGill, M.R.; Jaeschke, H. Biomarkers of drug-induced liver injury: Progress and utility in research, medicine, and regulation. Expert Rev. Mol. Diagn. 2018, 18, 797–807. [Google Scholar] [CrossRef]
  14. Woolbright, B.L.; Jaeschke, H. Mechanisms of Inflammatory Liver Injury and Drug-Induced Hepatotoxicity. Curr. Pharmacol. Rep. 2018, 4, 346–357. [Google Scholar] [CrossRef] [PubMed]
  15. Joshi-Barve, S.; Kirpich, I.; Cave, M.C.; Marsano, L.S.; McClain, C.J. Alcoholic, Nonalcoholic, and Toxicant-Associated Steatohepatitis: Mechanistic Similarities and Differences. Cell Mol. Gastroenterol. Hepatol. 2015, 1, 356–367. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  16. Denk, H.; Abuja, P.M.; Zatloukal, K. Animal models of NAFLD from the pathologist’s point of view. Biochim. Biophys. Acta Mol. Basis Dis. 2019, 1865, 929–942. [Google Scholar] [CrossRef] [PubMed]
  17. Machado, M.V.; Diehl, A.M. Pathogenesis of Nonalcoholic Steatohepatitis. Gastroenterology 2016, 150, 1769–1777. [Google Scholar] [CrossRef] [Green Version]
  18. Larter, C.Z.; Chitturi, S.; Heydet, D.; Farrell, G.C. A fresh look at NASH pathogenesis. Part 1: The metabolic movers. J. Gastroenterol. Hepatol. 2010, 25, 672–690. [Google Scholar] [CrossRef]
  19. Begriche, K.; Massart, J.; Robin, M.A.; Bonnet, F.; Fromenty, B. Mitochondrial adaptations and dysfunctions in nonalcoholic fatty liver disease. Hepatology 2013, 58, 1497–1507. [Google Scholar] [CrossRef] [PubMed]
  20. Caldwell, S.H.; Swerdlow, R.H.; Khan, E.M.; Iezzoni, J.C.; Hespenheide, E.E.; Parks, J.K.; Parker, W.D., Jr. Mitochondrial abnormalities in non-alcoholic steatohepatitis. J. Hepatol. 1999, 31, 430–434. [Google Scholar] [CrossRef]
  21. Galloway, C.A.; Lee, H.; Brookes, P.S.; Yoon, Y. Decreasing mitochondrial fission alleviates hepatic steatosis in a murine model of nonalcoholic fatty liver disease. Am. J. Physiol. Gastrointest. Liver Physiol. 2014, 307, G632–G641. [Google Scholar] [CrossRef] [Green Version]
  22. Perez-Carreras, M.; del Hoyo, P.; Martin, M.A.; Rubio, J.C.; Martin, A.; Castellano, G.; Colina, F.; Arenas, J.; Solis-Herruzo, J.A. Defective hepatic mitochondrial respiratory chain in patients with nonalcoholic steatohepatitis. Hepatology 2003, 38, 999–1007. [Google Scholar] [CrossRef]
  23. Pessayre, D. Role of mitochondria in non-alcoholic fatty liver disease. J. Gastroenterol. Hepatol. 2007, 22 (Suppl. 1), S20–S27. [Google Scholar] [CrossRef]
  24. Pessayre, D.; Berson, A.; Fromenty, B.; Mansouri, A. Mitochondria in steatohepatitis. Semin. Liver Dis. 2001, 21, 57–69. [Google Scholar] [CrossRef] [PubMed]
  25. Pessayre, D.; Fromenty, B. NASH: A mitochondrial disease. J. Hepatol. 2005, 42, 928–940. [Google Scholar] [CrossRef] [PubMed]
  26. Farrell, G.C.; Larter, C.Z. Nonalcoholic fatty liver disease: From steatosis to cirrhosis. Hepatology 2006, 43, S99–S112. [Google Scholar] [CrossRef]
  27. Begriche, K.; Igoudjil, A.; Pessayre, D.; Fromenty, B. Mitochondrial dysfunction in NASH: Causes, consequences and possible means to prevent it. Mitochondrion 2006, 6, 1–28. [Google Scholar] [CrossRef] [PubMed]
  28. Nikam, A.; Patankar, J.V.; Lackner, C.; Schock, E.; Kratky, D.; Zatloukal, K.; Abuja, P.M. Transition between Acute and Chronic Hepatotoxicity in Mice Is Associated with Impaired Energy Metabolism and Induction of Mitochondrial Heme Oxygenase-1. PLoS ONE 2013, 8, e66094. [Google Scholar] [CrossRef] [Green Version]
  29. Singla, A.; Moons, D.S.; Snider, N.T.; Wagenmaker, E.R.; Jayasundera, V.B.; Omary, M.B. Oxidative stress, Nrf2 and keratin up-regulation associate with Mallory-Denk body formation in mouse erythropoietic protoporphyria. Hepatology 2012, 56, 322–331. [Google Scholar] [CrossRef] [Green Version]
  30. Cortez-Pinto, H.; Chatham, J.; Chacko, V.P.; Arnold, C.; Rashid, A.; Diehl, A.M. Alterations in liver ATP homeostasis in human nonalcoholic steatohepatitis: A pilot study. JAMA 1999, 282, 1659–1664. [Google Scholar] [CrossRef] [Green Version]
  31. Fromenty, B.; Pessayre, D. Inhibition of mitochondrial beta-oxidation as a mechanism of hepatotoxicity. Pharmacol. Ther. 1995, 67, 101–154. [Google Scholar] [CrossRef]
  32. Tirosh, O. Hypoxic Signaling and Cholesterol Lipotoxicity in Fatty Liver Disease Progression. Oxid. Med. Cell Longev. 2018, 2018, 2548154. [Google Scholar] [CrossRef] [PubMed]
  33. Polotsky, V.Y.; Patil, S.P.; Savransky, V.; Laffan, A.; Fonti, S.; Frame, L.A.; Steele, K.E.; Schweizter, M.A.; Clark, J.M.; Torbenson, M.S.; et al. Obstructive sleep apnea, insulin resistance, and steatohepatitis in severe obesity. Am. J. Respir. Crit. Care Med. 2009, 179, 228–234. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  34. Selak, M.A.; Armour, S.M.; MacKenzie, E.D.; Boulahbel, H.; Watson, D.G.; Mansfield, K.D.; Pan, Y.; Simon, M.C.; Thompson, C.B.; Gottlieb, E. Succinate links TCA cycle dysfunction to oncogenesis by inhibiting HIF-alpha prolyl hydroxylase. Cancer Cell 2005, 7, 77–85. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  35. Sundaram, S.S.; Swiderska-Syn, M.; Sokol, R.J.; Halbower, A.C.; Capocelli, K.E.; Pan, Z.; Robbins, K.; Graham, B.; Diehl, A.M. Nocturnal Hypoxia Activation of the Hedgehog Signaling Pathway Affects Pediatric Nonalcoholic Fatty Liver Disease Severity. Hepatol. Commun. 2019, 3, 883–893. [Google Scholar] [CrossRef]
  36. Schafer, M.J.; Miller, J.D.; LeBrasseur, N.K. Cellular senescence: Implications for metabolic disease. Mol. Cell Endocrinol. 2017, 455, 93–102. [Google Scholar] [CrossRef]
  37. Hunt, N.J.; Kang, S.W.S.; Lockwood, G.P.; le Couteur, D.G.; Cogger, V.C. Hallmarks of Aging in the Liver. Comput. Struct. Biotechnol. J. 2019, 17, 1151–1161. [Google Scholar] [CrossRef]
  38. Huda, N.; Liu, G.; Hong, H.; Yan, S.; Khambu, B.; Yin, X.M. Hepatic senescence, the good and the bad. World J. Gastroenterol. 2019, 25, 5069–5081. [Google Scholar] [CrossRef]
  39. Papatheodoridi, A.M.; Chrysavgis, L.; Koutsilieris, M.; Chatzigeorgiou, A. The Role of Senescence in the Development of Nonalcoholic Fatty Liver Disease and Progression to Nonalcoholic Steatohepatitis. Hepatology 2020, 71, 363–374. [Google Scholar] [CrossRef]
  40. Mills, E.L.; Harmon, C.; Jedrychowski, M.P.; Xiao, H.; Garrity, R.; Tran, N.V.; Bradshaw, G.A.; Fu, A.; Szpyt, J.; Reddy, A.; et al. UCP1 governs liver extracellular succinate and inflammatory pathogenesis. Nat. Metab. 2021, 3, 604–617. [Google Scholar] [CrossRef]
  41. Diehl, A.M.; Li, Z.P.; Lin, H.Z.; Yang, S.Q. Cytokines and the pathogenesis of non-alcoholic steatohepatitis. Gut 2005, 54, 303–306. [Google Scholar] [CrossRef] [Green Version]
  42. Moschen, A.R.; Kaser, S.; Tilg, H. Non-alcoholic steatohepatitis: A microbiota-driven disease. Trends Endocrinol. Metab. 2013, 24, 537–545. [Google Scholar] [CrossRef]
  43. Mouzaki, M.; Comelli, E.M.; Arendt, B.M.; Bonengel, J.; Fung, S.K.; Fischer, S.E.; McGilvray, I.D.; Allard, J.P. Intestinal microbiota in patients with nonalcoholic fatty liver disease. Hepatology 2013, 58, 120–127. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  44. Mouzaki, M.; Wang, A.Y.; Bandsma, R.; Comelli, E.M.; Arendt, B.M.; Zhang, L.; Fung, S.; Fischer, S.E.; McGilvray, I.G.; Allard, J.P. Bile Acids and Dysbiosis in Non-Alcoholic Fatty Liver Disease. PLoS ONE 2016, 11, e0151829. [Google Scholar] [CrossRef] [Green Version]
  45. Neuman, M.G.; French, S.W.; French, B.A.; Seitz, H.K.; Cohen, L.B.; Mueller, S.; Osna, N.A.; Kharbanda, K.K.; Seth, D.; Bautista, A.; et al. Alcoholic and non-alcoholic steatohepatitis. Exp. Mol. Pathol. 2014, 97, 492–510. [Google Scholar] [CrossRef] [Green Version]
  46. Smith, R.A.; Murphy, M.P. Animal and human studies with the mitochondria-targeted antioxidant MitoQ. Ann. N. Y. Acad. Sci. 2010, 1201, 96–103. [Google Scholar] [CrossRef] [PubMed]
  47. Day, C.P.; James, O.F. Steatohepatitis: A tale of two “hits”? Gastroenterology 1998, 114, 842–845. [Google Scholar] [CrossRef]
  48. Tilg, H.; Moschen, A.R. Evolution of inflammation in nonalcoholic fatty liver disease: The multiple parallel hits hypothesis. Hepatology 2010, 52, 1836–1846. [Google Scholar] [CrossRef]
  49. Browning, J.D.; Horton, J.D. Molecular mediators of hepatic steatosis and liver injury. J. Clin. Investig. 2004, 114, 147–152. [Google Scholar] [CrossRef] [Green Version]
  50. Nikam, A.; Patankar, J.V.; Somlapura, M.; Lahiri, P.; Sachdev, V.; Kratky, D.; Denk, H.; Zatloukal, K.; Abuja, P.M. The PPARalpha Agonist Fenofibrate Prevents Formation of Protein Aggregates (Mallory-Denk bodies) in a Murine Model of Steatohepatitis-like Hepatotoxicity. Sci Rep. 2018, 8, 12964. [Google Scholar] [CrossRef] [PubMed]
  51. Stumptner, C.; Fuchsbichler, A.; Lehner, M.; Zatloukal, K.; Denk, H. Sequence of events in the assembly of Mallory body components in mouse liver: Clues to the pathogenesis and significance of Mallory body formation. J. Hepatol. 2001, 34, 665–675. [Google Scholar] [CrossRef]
  52. McCluskey, S.A.; Marks, G.S.; Sutherland, E.P.; Jacobsen, N.; Ortiz de Montellano, P.R. Ferrochelatase-inhibitory activity and N-alkylprotoporphyrin formation with analogues of 3,5-diethoxycarbonyl-1,4-dihydro-2,4,6-trimethylpyridine (DDC) containing extended 4-alkyl groups: Implications for the active site of ferrochelatase. Mol. Pharmacol. 1986, 30, 352–357. [Google Scholar] [PubMed]
  53. McCluskey, S.A.; Marks, G.S.; Whitney, R.A.; Ortiz de Montellano, P.R. Differential inhibition of hepatic ferrochelatase by regioisomers of N-butyl-, N-pentyl-, N-hexyl-, and N-isobutylprotoporphyrin IX. Mol. Pharmacol. 1988, 34, 80–86. [Google Scholar]
  54. Benassi, B.; Fanciulli, M.; Fiorentino, F.; Porrello, A.; Chiorino, G.; Loda, M.; Zupi, G.; Biroccio, A. c-Myc phosphorylation is required for cellular response to oxidative stress. Mol. Cell. 2006, 21, 509–519. [Google Scholar] [CrossRef]
  55. Levy, S.; Forman, H.J. C-Myc is a Nrf2-interacting protein that negatively regulates phase II genes through their electrophile responsive elements. IUBMB Life 2010, 62, 237–246. [Google Scholar] [CrossRef]
  56. Mills, E.; O’Neill, L.A. Succinate: A metabolic signal in inflammation. Trends Cell Biol. 2014, 24, 313–320. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  57. Li, D.; Du, Y.; Yuan, X.; Han, X.; Dong, Z.; Chen, X.; Wu, H.; Zhang, J.; Xu, L.; Han, C.; et al. Hepatic hypoxia-inducible factors inhibit PPARalpha expression to exacerbate acetaminophen induced oxidative stress and hepatotoxicity. Free Radic. Biol. Med. 2017, 110, 102–116. [Google Scholar] [CrossRef] [PubMed]
  58. Li, J.; Yang, Y.L.; Li, L.Z.; Zhang, L.; Liu, Q.; Liu, K.; Li, P.; Liu, B.; Qi, L.W. Succinate accumulation impairs cardiac pyruvate dehydrogenase activity through GRP91-dependent and independent signaling pathways: Therapeutic effects of ginsenoside Rb1. Biochim. Biophys. Acta Mol. Basis Dis. 2017, 1863, 2835–2847. [Google Scholar] [CrossRef]
  59. Stumptner, C.; Fuchsbichler, A.; Zatloukal, K.; Denk, H. In vitro production of Mallory bodies and intracellular hyaline bodies: The central role of sequestosome 1/p62. Hepatology 2007, 46, 851–860. [Google Scholar] [CrossRef]
  60. Nakajima, T.; Kamijo, Y.; Tanaka, N.; Sugiyama, E.; Tanaka, E.; Kiyosawa, K.; Fukushima, Y.; Peters, J.M.; Gonzalez, F.J.; Aoyama, T. Peroxisome proliferator-activated receptor alpha protects against alcohol-induced liver damage. Hepatology 2004, 40, 972–980. [Google Scholar] [CrossRef]
  61. Francque, S.; Verrijken, A.; Caron, S.; Prawitt, J.; Paumelle, R.; Derudas, B.; Lefebvre, P.; Taskinen, M.R.; van Hul, W.; Mertens, I.; et al. PPARalpha gene expression correlates with severity and histological treatment response in patients with non-alcoholic steatohepatitis. J. Hepatol. 2015, 63, 164–173. [Google Scholar] [CrossRef]
  62. Kersten, S.; Stienstra, R. The role and regulation of the peroxisome proliferator activated receptor alpha in human liver. Biochimie 2017, 136, 75–84. [Google Scholar] [CrossRef] [PubMed]
  63. Abdelmegeed, M.A.; Yoo, S.H.; Henderson, L.E.; Gonzalez, F.J.; Woodcroft, K.J.; Song, B.J. PPARalpha expression protects male mice from high fat-induced nonalcoholic fatty liver. J. Nutr. 2011, 141, 603–610. [Google Scholar] [CrossRef] [PubMed]
  64. Deng, Q.G.; She, H.; Cheng, J.H.; French, S.W.; Koop, D.R.; Xiong, S.; Tsukamoto, H. Steatohepatitis induced by intragastric overfeeding in mice. Hepatology 2005, 42, 905–914. [Google Scholar] [CrossRef] [PubMed]
  65. Kucukoglu, O.; Guldiken, N.; Chen, Y.; Usachov, V.; El-Heliebi, A.; Haybaeck, J.; Denk, H.; Trautwein, C.; Strnad, P. High-fat diet triggers Mallory-Denk body formation through misfolding and crosslinking of excess keratin 8. Hepatology 2014, 60, 169–178. [Google Scholar] [CrossRef]
  66. Regnier, M.; Polizzi, A.; Smati, S.; Lukowicz, C.; Fougerat, A.; Lippi, Y.; Fouche, E.; Lasserre, F.; Naylies, C.; Betoulieres, C.; et al. Hepatocyte-specific deletion of Pparalpha promotes NAFLD in the context of obesity. Sci. Rep. 2020, 10, 6489. [Google Scholar] [CrossRef]
  67. Shaban, Z.; El-Shazly, S.; Abdelhady, S.; Fattouh, I.; Muzandu, K.; Ishizuka, M.; Kimura, K.; Kazusaka, A.; Fujita, S. Down regulation of hepatic PPARalpha function by AhR ligand. J. Vet. Med. Sci. 2004, 66, 1377–1386. [Google Scholar] [CrossRef] [Green Version]
  68. Sinal, C.J.; Bend, J.R. Aryl hydrocarbon receptor-dependent induction of cyp1a1 by bilirubin in mouse hepatoma hepa 1c1c7 cells. Mol. Pharmacol. 1997, 52, 590–599. [Google Scholar] [CrossRef]
  69. Bicego, R.; Francisco, A.; Ruas, J.S.; Siqueira-Santos, E.S.; Castilho, R.F. Undesirable effects of chemical inhibitors of NAD(P)(+) transhydrogenase on mitochondrial respiratory function. Arch. Biochem. Biophys. 2020, 692, 108535. [Google Scholar] [CrossRef]
  70. Gameiro, P.A.; Laviolette, L.A.; Kelleher, J.K.; Iliopoulos, O.; Stephanopoulos, G. Cofactor Balance by Nicotinamide Nucleotide Transhydrogenase (NNT) Coordinates Reductive Carboxylation and Glucose Catabolism in the Tricarboxylic Acid (TCA) Cycle. J. Biol. Chem. 2013, 288, 12967–12977. [Google Scholar] [CrossRef] [Green Version]
  71. McLennan, H.R.; Degli Esposti, M. The contribution of mitochondrial respiratory complexes to the production of reactive oxygen species. J. Bioenerg. Biomembr. 2000, 32, 153–162. [Google Scholar] [CrossRef]
  72. Ronchi, J.A.; Figueira, T.R.; Ravagnani, F.G.; Oliveira, H.C.F.; Vercesi, A.E.; Castilho, R.F. A spontaneous mutation in the nicotinamide nucleotide transhydrogenase gene of C57BL/6J mice results in mitochondrial redox abnormalities. Free Radic. Biol. Med. 2013, 63, 446–456. [Google Scholar] [CrossRef] [Green Version]
  73. Simon, M.M.; Greenaway, S.; White, J.K.; Fuchs, H.; Gailus-Durner, V.; Wells, S.; Sorg, T.; Wong, K.; Bedu, E.; Cartwright, E.J.; et al. A comparative phenotypic and genomic analysis of C57BL/6J and C57BL/6N mouse strains. Genome Biol. 2013, 14, R82. [Google Scholar] [CrossRef]
  74. Duan, S.; Yao, Z.; Zhu, Y.; Wang, G.; Hou, D.; Wen, L.; Wu, M. The Pirh2-keratin 8/18 interaction modulates the cellular distribution of mitochondria and UV-induced apoptosis. Cell Death Differ. 2009, 16, 826–837. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  75. Guldiken, N.; Zhou, Q.; Kucukoglu, O.; Rehm, M.; Levada, K.; Gross, A.; Kwan, R.; James, L.P.; Trautwein, C.; Omary, M.B.; et al. Human keratin 8 variants promote mouse acetaminophen hepatotoxicity coupled with c-jun amino-terminal kinase activation and protein adduct formation. Hepatology 2015, 62, 876–886. [Google Scholar] [CrossRef] [Green Version]
  76. Hakem, A.; Bohgaki, M.; Lemmers, B.; Tai, E.; Salmena, L.; Matysiak-Zablocki, E.; Jung, Y.S.; Karaskova, J.; Kaustov, L.; Duan, S.; et al. Role of Pirh2 in mediating the regulation of p53 and c-Myc. PLoS Genet. 2011, 7, e1002360. [Google Scholar] [CrossRef] [PubMed]
  77. Halaby, M.J.; Hakem, R.; Hakem, A. Pirh2: An E3 ligase with central roles in the regulation of cell cycle, DNA damage response, and differentiation. Cell Cycle 2013, 12, 2733–2737. [Google Scholar] [CrossRef] [Green Version]
  78. Kwan, R.; Hanada, S.; Harada, M.; Strnad, P.; Li, D.H.; Omary, M.B. Keratin 8 phosphorylation regulates its transamidation and hepatocyte Mallory-Denk body formation. FASEB J. 2012, 26, 2318–2326. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  79. Magin, T.M.; Schroder, R.; Leitgeb, S.; Wanninger, F.; Zatloukal, K.; Grund, C.; Melton, D.W. Lessons from keratin 18 knockout mice: Formation of novel keratin filaments, secondary loss of keratin 7 and accumulation of liver-specific keratin 8-positive aggregates. J. Cell Biol. 1998, 140, 1441–1451. [Google Scholar] [CrossRef]
  80. Toivola, D.M.; Ku, N.O.; Resurreccion, E.Z.; Nelson, D.R.; Wright, T.L.; Omary, M.B. Keratin 8 and 18 hyperphosphorylation is a marker of progression of human liver disease. Hepatology 2004, 40, 459–466. [Google Scholar] [CrossRef]
  81. Zatloukal, K.; Stumptner, C.; Lehner, M.; Denk, H.; Baribault, H.; Eshkind, L.G.; Franke, W.W. Cytokeratin 8 protects from hepatotoxicity, and its ratio to cytokeratin 18 determines the ability of hepatocytes to form Mallory bodies. Am. J. Pathol. 2000, 156, 1263–1274. [Google Scholar] [CrossRef] [Green Version]
  82. Rakhshandehroo, M.; Hooiveld, G.; Muller, M.; Kersten, S. Comparative analysis of gene regulation by the transcription factor PPARalpha between mouse and human. PLoS ONE 2009, 4, e6796. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  83. Rakhshandehroo, M.; Knoch, B.; Muller, M.; Kersten, S. Peroxisome proliferator-activated receptor alpha target genes. PPAR Res. 2010, 2010, 612089. [Google Scholar] [CrossRef] [Green Version]
  84. Opitz, C.A.; Litzenburger, U.M.; Sahm, F.; Ott, M.; Tritschler, I.; Trump, S.; Schumacher, T.; Jestaedt, L.; Schrenk, D.; Weller, M.; et al. An endogenous tumour-promoting ligand of the human aryl hydrocarbon receptor. Nature 2011, 478, 197–203. [Google Scholar] [CrossRef]
  85. Bruick, R.K. Oxygen sensing in the hypoxic response pathway: Regulation of the hypoxia-inducible transcription factor. Genes Dev. 2003, 17, 2614–2623. [Google Scholar] [CrossRef] [Green Version]
  86. Hernandez-Rodas, M.C.; Valenzuela, R.; Echeverria, F.; Rincon-Cervera, M.A.; Espinosa, A.; Illesca, P.; Munoz, P.; Corbari, A.; Romero, N.; Gonzalez-Manan, D.; et al. Supplementation with Docosahexaenoic Acid and Extra Virgin Olive Oil Prevents Liver Steatosis Induced by a High-Fat Diet in Mice through PPAR-alpha and Nrf2 Upregulation with Concomitant SREBP-1c and NF-kB Downregulation. Mol. Nutr. Food Res. 2017, 61, 1700479. [Google Scholar] [CrossRef]
  87. Maher, J.M.; Aleksunes, L.M.; Dieter, M.Z.; Tanaka, Y.; Peters, J.M.; Manautou, J.E.; Klaassen, C.D. Nrf2- and PPAR alpha-mediated regulation of hepatic Mrp transporters after exposure to perfluorooctanoic acid and perfluorodecanoic acid. Toxicol. Sci. 2008, 106, 319–328. [Google Scholar] [CrossRef] [Green Version]
  88. Xu, J.; Donepudi, A.C.; Moscovitz, J.E.; Slitt, A.L. Keap1-knockdown decreases fasting-induced fatty liver via altered lipid metabolism and decreased fatty acid mobilization from adipose tissue. PLoS ONE 2013, 8, e79841. [Google Scholar] [CrossRef]
  89. Aleksunes, L.M.; Klaassen, C.D. Coordinated regulation of hepatic phase I and II drug-metabolizing genes and transporters using AhR-, CAR-, PXR-, PPARalpha-, and Nrf2-null mice. Drug Metab. Dispos. 2012, 40, 1366–1379. [Google Scholar] [CrossRef] [Green Version]
  90. Kim, D.; Brocker, C.N.; Takahashi, S.; Yagai, T.; Kim, T.; Xie, G.; Wang, H.; Qu, A.; Gonzalez, F.J. Keratin 23 Is a Peroxisome Proliferator-Activated Receptor Alpha-Dependent, MYC-Amplified Oncogene That Promotes Hepatocyte Proliferation. Hepatology 2019, 70, 154–167. [Google Scholar] [CrossRef] [PubMed]
  91. Nesci, S.; Trombetti, F.; Pagliarani, A. Nicotinamide Nucleotide Transhydrogenase as a Sensor of Mitochondrial Biology. Trends Cell Biol. 2020, 30, 1–3. [Google Scholar] [CrossRef] [PubMed]
  92. Navarro, C.D.C.; Figueira, T.R.; Francisco, A.; Dal’Bo, G.A.; Ronchi, J.A.; Rovani, J.C.; Escanhoela, C.A.F.; Oliveira, H.C.F.; Castilho, R.F.; Vercesi, A.E. Redox imbalance due to the loss of mitochondrial NAD(P)-transhydrogenase markedly aggravates high fat diet-induced fatty liver disease in mice. Free Radic. Biol. Med. 2017, 113, 190–202. [Google Scholar] [CrossRef]
  93. Nakamura, A.; Terauchi, Y. Lessons from mouse models of high-fat diet-induced NAFLD. Int. J. Mol. Sci. 2013, 14, 21240–21257. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  94. Evangelou, E.; Warren, H.R.; Mosen-Ansorena, D.; Mifsud, B.; Pazoki, R.; Gao, H.; Ntritsos, G.; Dimou, N.; Cabrera, C.P.; Karaman, I.; et al. Publisher Correction: Genetic analysis of over 1 million people identifies 535 new loci associated with blood pressure traits. Nat. Genet. 2018, 50, 1755. [Google Scholar] [CrossRef] [PubMed]
  95. Ahrens, M.; Ammerpohl, O.; von Schonfels, W.; Kolarova, J.; Bens, S.; Itzel, T.; Teufel, A.; Herrmann, A.; Brosch, M.; Hinrichsen, H.; et al. DNA methylation analysis in nonalcoholic fatty liver disease suggests distinct disease-specific and remodeling signatures after bariatric surgery. Cell Metab. 2013, 18, 296–302. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  96. Ratziu, V.; Harrison, S.A.; Francque, S.; Bedossa, P.; Lehert, P.; Serfaty, L.; Romero-Gomez, M.; Boursier, J.; Abdelmalek, M.; Caldwell, S.; et al. Elafibranor, an Agonist of the Peroxisome Proliferator-Activated Receptor-alpha and -delta, Induces Resolution of Nonalcoholic Steatohepatitis Without Fibrosis Worsening. Gastroenterology 2016, 150, 1147–1159. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  97. Li, M.H.; Chen, W.; Wang, L.L.; Sun, J.L.; Zhou, L.; Shi, Y.C.; Wang, C.H.; Zhong, B.H.; Shi, W.G.; Guo, Z.W. RLA8-A New and Highly Effective Quadruple PPAR-alpha/gamma/delta and GPR40 Agonist to Reverse Nonalcoholic Steatohepatitis and Fibrosis. J. Pharmacol. Exp. Ther. 2019, 369, 67–77. [Google Scholar] [CrossRef] [Green Version]
  98. Malik, A.; Nadeem, M.; Malik, M.I. Efficacy of elafibranor in patients with liver abnormalities especially non-alcoholic steatohepatitis: A systematic review and meta-analysis. Clin. J. Gastroenterol. 2021, 14, 1–8. [Google Scholar] [CrossRef]
  99. Smati, S.; Polizzi, A.; Fougerat, A.; Ellero-Simatos, S.; Blum, Y.; Lippi, Y.; Regnier, M.; Laroyenne, A.; Huillet, M.; Arif, M.; et al. Integrative study of diet-induced mouse models of NAFLD identifies PPARalpha as a sexually dimorphic drug target. Gut 2021. [Google Scholar] [CrossRef]
  100. Atamna, H.; Liu, J.; Ames, B.N. Heme deficiency selectively interrupts assembly of mitochondrial complex IV in human fibroblasts: Revelance to aging. J. Biol. Chem. 2001, 276, 48410–48416. [Google Scholar] [CrossRef] [Green Version]
  101. Cho, E.H. Succinate as a Regulator of Hepatic Stellate Cells in Liver Fibrosis. Front. Endocrinol. 2018, 9, 455. [Google Scholar] [CrossRef] [Green Version]
  102. Grivennikova, V.G.; Kozlovsky, V.S.; Vinogradov, A.D. Respiratory complex II: ROS production and the kinetics of ubiquinone reduction. Biochim. Biophys. Acta Bioenerg. 2017, 1858, 109–117. [Google Scholar] [CrossRef]
  103. Cunningham, C.C.; Coleman, W.B.; Spach, P.I. The effects of chronic ethanol consumption on hepatic mitochondrial energy metabolism. Alcohol Alcohol. 1990, 25, 127–136. [Google Scholar] [CrossRef] [PubMed]
  104. Wong, H.S.; Mezera, V.; Dighe, P.; Melov, S.; Gerencser, A.A.; Sweis, R.F.; Pliushchev, M.; Wang, Z.; Esbenshade, T.; McKibben, B.; et al. Superoxide produced by mitochondrial site IQ inactivates cardiac succinate dehydrogenase and induces hepatic steatosis in Sod2 knockout mice. Free Radic. Biol. Med. 2021, 164, 223–232. [Google Scholar] [CrossRef] [PubMed]
  105. Quinlan, C.L.; Orr, A.L.; Perevoshchikova, I.V.; Treberg, J.R.; Ackrell, B.A.; Brand, M.D. Mitochondrial complex II can generate reactive oxygen species at high rates in both the forward and reverse reactions. J. Biol. Chem. 2012, 287, 27255–27264. [Google Scholar] [CrossRef] [Green Version]
  106. Correa, P.R.; Kruglov, E.A.; Thompson, M.; Leite, M.F.; Dranoff, J.A.; Nathanson, M.H. Succinate is a paracrine signal for liver damage. J. Hepatol. 2007, 47, 262–269. [Google Scholar] [CrossRef] [Green Version]
  107. Park, S.Y.; Le, C.T.; Sung, K.Y.; Choi, D.H.; Cho, E.H. Succinate induces hepatic fibrogenesis by promoting activation, proliferation, and migration, and inhibiting apoptosis of hepatic stellate cells. Biochem. Biophys. Res. Commun. 2018, 496, 673–678. [Google Scholar] [CrossRef]
  108. De Castro Fonseca, M.; Aguiar, C.J.; da Rocha Franco, J.A.; Gingold, R.N.; Leite, M.F. GPR91: Expanding the frontiers of Krebs cycle intermediates. Cell Commun. Signal. 2016, 14, 3. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  109. Li, Y.H.; Woo, S.H.; Choi, D.H.; Cho, E.H. Succinate causes alpha-SMA production through GPR91 activation in hepatic stellate cells. Biochem. Biophys. Res. Commun. 2015, 463, 853–858. [Google Scholar] [CrossRef]
  110. Harber, K.J.; de Goede, K.E.; Verberk, S.G.S.; Meinster, E.; de Vries, H.E.; van Weeghel, M.; de Winther, M.P.J.; van den Bossche, J. Succinate Is an Inflammation-Induced Immunoregulatory Metabolite in Macrophages. Metabolites 2020, 10, 372. [Google Scholar] [CrossRef]
  111. Macias-Ceja, D.C.; Ortiz-Masia, D.; Salvador, P.; Gisbert-Ferrandiz, L.; Hernandez, C.; Hausmann, M.; Rogler, G.; Esplugues, J.V.; Hinojosa, J.; Alos, R.; et al. Succinate receptor mediates intestinal inflammation and fibrosis. Mucosal. Immunol. 2019, 12, 178–187. [Google Scholar] [CrossRef] [Green Version]
  112. Lu, Y.T.; Li, L.Z.; Yang, Y.L.; Yin, X.; Liu, Q.; Zhang, L.; Liu, K.; Liu, B.; Li, J.; Qi, L.W. Succinate induces aberrant mitochondrial fission in cardiomyocytes through GPR91 signaling. Cell Death Dis. 2018, 9, 672. [Google Scholar] [CrossRef] [Green Version]
  113. Lee, K.K.; Imaizumi, N.; Chamberland, S.R.; Alder, N.N.; Boelsterli, U.A. Targeting mitochondria with methylene blue protects mice against acetaminophen-induced liver injury. Hepatology 2015, 61, 326–336. [Google Scholar] [CrossRef]
  114. Jia, P.; Wu, X.; Pan, T.; Xu, S.; Hu, J.; Ding, X. Uncoupling protein 1 inhibits mitochondrial reactive oxygen species generation and alleviates acute kidney injury. EBioMedicine 2019, 49, 331–340. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  115. Lian, A.; Li, X.; Jiang, Q. Irisin inhibition of growth hormone secretion in cultured tilapia pituitary cells. Mol. Cell Endocrinol. 2017, 439, 395–406. [Google Scholar] [CrossRef]
  116. Serviddio, G.; Bellanti, F.; Tamborra, R.; Rollo, T.; Romano, A.D.; Giudetti, A.M.; Capitanio, N.; Petrella, A.; Vendemiale, G.; Altomare, E. Alterations of hepatic ATP homeostasis and respiratory chain during development of non-alcoholic steatohepatitis in a rodent model. Eur. J. Clin. Investig. 2008, 38, 245–252. [Google Scholar] [CrossRef] [PubMed]
  117. Mantena, S.K.; Vaughn, D.P.; Andringa, K.K.; Eccleston, H.B.; King, A.L.; Abrams, G.A.; Doeller, J.E.; Kraus, D.W.; Darley-Usmar, V.M.; Bailey, S.M. High fat diet induces dysregulation of hepatic oxygen gradients and mitochondrial function in vivo. Biochem. J. 2009, 417, 183–193. [Google Scholar] [CrossRef] [Green Version]
  118. Stienstra, R.; Mandard, S.; Patsouris, D.; Maass, C.; Kersten, S.; Muller, M. Peroxisome proliferator-activated receptor alpha protects against obesity-induced hepatic inflammation. Endocrinology 2007, 148, 2753–2763. [Google Scholar] [CrossRef] [Green Version]
  119. Cong, W.N.; Tao, R.Y.; Tian, J.Y.; Liu, G.T.; Ye, F. The establishment of a novel non-alcoholic steatohepatitis model accompanied with obesity and insulin resistance in mice. Life Sci. 2008, 82, 983–990. [Google Scholar] [CrossRef]
  120. Berson, A.; de Beco, V.; Letteron, P.; Robin, M.A.; Moreau, C.; El Kahwaji, J.; Verthier, N.; Feldmann, G.; Fromenty, B.; Pessayre, D. Steatohepatitis-inducing drugs cause mitochondrial dysfunction and lipid peroxidation in rat hepatocytes. Gastroenterology 1998, 114, 764–774. [Google Scholar] [CrossRef]
  121. Denk, H.; Eckerstorfer, R. Colchicine-induced Mallory body formation in the mouse. Lab. Investig. 1977, 36, 563–565. [Google Scholar] [PubMed]
  122. Leung, Y.Y.; Yao Hui, L.L.; Kraus, V.B. Colchicine—Update on mechanisms of action and therapeutic uses. Semin. Arthritis Rheum. 2015, 45, 341–350. [Google Scholar] [CrossRef] [Green Version]
  123. Riley, N.E.; Bardag-Gorce, F.; Montgomery, R.O.; Li, J.; Lungo, W.; Lue, Y.H.; French, S.W. Microtubules are required for cytokeratin aggresome (Mallory body) formation in hepatocytes: An in vitro study. Exp. Mol. Pathol. 2003, 74, 173–179. [Google Scholar] [CrossRef]
  124. Mahajan, V.; Klingstedt, T.; Simon, R.; Nilsson, K.P.; Thueringer, A.; Kashofer, K.; Haybaeck, J.; Denk, H.; Abuja, P.M.; Zatloukal, K. Cross beta-sheet conformation of keratin 8 is a specific feature of Mallory-Denk bodies compared with other hepatocyte inclusions. Gastroenterology 2011, 141, 1080–1090. [Google Scholar] [CrossRef] [PubMed]
  125. Strnad, P.; Zatloukal, K.; Stumptner, C.; Kulaksiz, H.; Denk, H. Mallory-Denk-bodies: Lessons from keratin-containing hepatic inclusion bodies. Biochim. Biophys. Acta 2008, 1782, 764–774. [Google Scholar] [CrossRef] [Green Version]
  126. Gonzalez-Garcia, M.; Fusco, G.; de Simone, A. Membrane Interactions and Toxicity by Misfolded Protein Oligomers. Front. Cell Dev. Biol. 2021, 9, 642623. [Google Scholar] [CrossRef] [PubMed]
  127. Agarwal, E.; Altman, B.J.; Seo, J.H.; Ghosh, J.C.; Kossenkov, A.V.; Tang, H.Y.; Krishn, S.R.; Languino, L.R.; Gabrilovich, D.I.; Speicher, D.W.; et al. Myc-mediated transcriptional regulation of the mitochondrial chaperone TRAP1 controls primary and metastatic tumor growth. J. Biol. Chem. 2019, 294, 10407–10414. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  128. Sarin, M.; Wang, Y.; Zhang, F.; Rothermund, K.; Zhang, Y.; Lu, J.; Sims-Lucas, S.; Beer-Stolz, D.; van Houten, B.E.; Vockley, J.; et al. Alterations in c-Myc phenotypes resulting from dynamin-related protein 1 (Drp1)-mediated mitochondrial fission. Cell Death Dis. 2013, 4, e670. [Google Scholar] [CrossRef] [Green Version]
  129. Zhang, H.; Gao, P.; Fukuda, R.; Kumar, G.; Krishnamachary, B.; Zeller, K.I.; Dang, C.V.; Semenza, G.L. HIF-1 inhibits mitochondrial biogenesis and cellular respiration in VHL-deficient renal cell carcinoma by repression of C-MYC activity. Cancer Cell 2007, 11, 407–420. [Google Scholar] [CrossRef] [Green Version]
  130. Daks, A.; Petukhov, A.; Fedorova, O.; Shuvalov, O.; Kizenko, A.; Tananykina, E.; Vasileva, E.; Semenov, O.; Bottrill, A.; Barlev, N. The RNA-binding protein HuR is a novel target of Pirh2 E3 ubiquitin ligase. Cell Death Dis. 2021, 12, 581. [Google Scholar] [CrossRef]
  131. Ho, J.J.D.; Balukoff, N.C.; Theodoridis, P.R.; Wang, M.; Krieger, J.R.; Schatz, J.H.; Lee, S. A network of RNA-binding proteins controls translation efficiency to activate anaerobic metabolism. Nat. Commun. 2020, 11, 2677. [Google Scholar] [CrossRef]
  132. Marchesi, N.; Thongon, N.; Pascale, A.; Provenzani, A.; Koskela, A.; Korhonen, E.; Smedowski, A.; Govoni, S.; Kauppinen, A.; Kaarniranta, K.; et al. Autophagy Stimulus Promotes Early HuR Protein Activation and p62/SQSTM1 Protein Synthesis in ARPE-19 Cells by Triggering Erk1/2, p38(MAPK), and JNK Kinase Pathways. Oxid. Med. Cell Longev. 2018, 2018, 4956080. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  133. Viiri, J.; Amadio, M.; Marchesi, N.; Hyttinen, J.M.; Kivinen, N.; Sironen, R.; Rilla, K.; Akhtar, S.; Provenzani, A.; D’Agostino, V.G.; et al. Autophagy activation clears ELAVL1/HuR-mediated accumulation of SQSTM1/p62 during proteasomal inhibition in human retinal pigment epithelial cells. PLoS ONE 2013, 8, e69563. [Google Scholar] [CrossRef] [Green Version]
  134. Liu, S.; Jiang, X.; Cui, X.; Wang, J.; Liu, S.; Li, H.; Yang, J.; Zhang, C.; Zhang, W. Smooth muscle-specific HuR knockout induces defective autophagy and atherosclerosis. Cell Death Dis. 2021, 12, 385. [Google Scholar] [CrossRef]
  135. Tian, M.; Wang, J.; Liu, S.; Li, X.; Li, J.; Yang, J.; Zhang, C.; Zhang, W. Hepatic HuR protects against the pathogenesis of non-alcoholic fatty liver disease by targeting PTEN. Cell Death Dis. 2021, 12, 236. [Google Scholar] [CrossRef] [PubMed]
  136. Chinopoulos, C. Succinate in ischemia: Where does it come from? Int. J. Biochem. Cell Biol. 2019, 115, 105580. [Google Scholar] [CrossRef] [PubMed]
  137. Ryan, D.G.; Murphy, M.P.; Frezza, C.; Prag, H.A.; Chouchani, E.T.; O’Neill, L.A.; Mills, E.L. Coupling Krebs cycle metabolites to signalling in immunity and cancer. Nat. Metab. 2019, 1, 16–33. [Google Scholar] [CrossRef]
  138. Sulkowski, P.L.; Oeck, S.; Dow, J.; Economos, N.G.; Mirfakhraie, L.; Liu, Y.; Noronha, K.; Bao, X.; Li, J.; Shuch, B.M.; et al. Oncometabolites suppress DNA repair by disrupting local chromatin signalling. Nature 2020, 582, 586–591. [Google Scholar] [CrossRef]
  139. King, A.; Selak, M.A.; Gottlieb, E. Succinate dehydrogenase and fumarate hydratase: Linking mitochondrial dysfunction and cancer. Oncogene 2006, 25, 4675–4682. [Google Scholar] [CrossRef] [Green Version]
  140. Matlac, D.M.; Hadrava Vanova, K.; Bechmann, N.; Richter, S.; Folberth, J.; Ghayee, H.K.; Ge, G.B.; Abunimer, L.; Wesley, R.; Aherrahrou, R.; et al. Succinate Mediates Tumorigenic Effects via Succinate Receptor 1: Potential for New Targeted Treatment Strategies in Succinate Dehydrogenase Deficient Paragangliomas. Front. Endocrinol. 2021, 12, 589451. [Google Scholar] [CrossRef]
  141. Hackenbeck, T.; Knaup, K.X.; Schietke, R.; Schodel, J.; Willam, C.; Wu, X.; Warnecke, C.; Eckardt, K.U.; Wiesener, M.S. HIF-1 or HIF-2 induction is sufficient to achieve cell cycle arrest in NIH3T3 mouse fibroblasts independent from hypoxia. Cell Cycle 2009, 8, 1386–1395. [Google Scholar] [CrossRef] [Green Version]
  142. Ho, J.J.D.; Schatz, J.H.; Uniacke, J.; Lee, S. Jekyll and Hyde: Activating the Hypoxic Translational Machinery. Trends Biochem. Sci. 2021, 46, 171–174. [Google Scholar] [CrossRef]
  143. Ho, J.J.D.; Balukoff, N.C.; Cervantes, G.; Malcolm, P.D.; Krieger, J.R.; Lee, S. Oxygen-Sensitive Remodeling of Central Carbon Metabolism by Archaic eIF5B. Cell Rep. 2018, 22, 17–26. [Google Scholar] [CrossRef] [Green Version]
  144. Elorza, A.; Soro-Arnaiz, I.; Melendez-Rodriguez, F.; Rodriguez-Vaello, V.; Marsboom, G.; de Carcer, G.; Acosta-Iborra, B.; Albacete-Albacete, L.; Ordonez, A.; Serrano-Oviedo, L.; et al. HIF2alpha acts as an mTORC1 activator through the amino acid carrier SLC7A5. Mol. Cell 2012, 48, 681–691. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  145. Kuwano, A.; Tanaka, M.; Suzuki, H.; Kurokawa, M.; Imoto, K.; Tashiro, S.; Goya, T.; Kohjima, M.; Kato, M.; Ogawa, Y. Upregulated expression of hypoxia reactive genes in peripheral blood mononuclear cells from chronic liver disease patients. Biochem. Biophys. Rep. 2021, 27, 101068. [Google Scholar] [CrossRef] [PubMed]
  146. Kuwano, A.; Kurokawa, M.; Kohjima, M.; Imoto, K.; Tashiro, S.; Suzuki, H.; Tanaka, M.; Okada, S.; Kato, M.; Ogawa, Y. Microcirculatory disturbance in acute liver injury. Exp. Ther. Med. 2021, 21, 596. [Google Scholar] [CrossRef] [PubMed]
  147. Komatsu, M.; Waguri, S.; Koike, M.; Sou, Y.S.; Ueno, T.; Hara, T.; Mizushima, N.; Iwata, J.; Ezaki, J.; Murata, S.; et al. Homeostatic levels of p62 control cytoplasmic inclusion body formation in autophagy-deficient mice. Cell 2007, 131, 1149–1163. [Google Scholar] [CrossRef] [Green Version]
  148. Nan, L.; Wu, Y.; Bardag-Gorce, F.; Li, J.; French, B.A.; Fu, A.N.; Francis, T.; Vu, J.; French, S.W. p62 is involved in the mechanism of Mallory body formation. Exp. Mol. Pathol. 2004, 77, 168–175. [Google Scholar] [CrossRef]
  149. Lahiri, P.; Schmidt, V.; Smole, C.; Kufferath, I.; Denk, H.; Strnad, P.; Rulicke, T.; Frohlich, L.F.; Zatloukal, K. p62/Sequestosome-1 Is Indispensable for Maturation and Stabilization of Mallory-Denk Bodies. PLoS ONE 2016, 11, e0161083. [Google Scholar] [CrossRef]
  150. Manley, S.; Williams, J.A.; Ding, W.X. Role of p62/SQSTM1 in liver physiology and pathogenesis. Exp. Biol. Med. 2013, 238, 525–538. [Google Scholar] [CrossRef] [Green Version]
  151. Denk, H.; Stumptner, C.; Abuja, P.M.; Zatloukal, K. Sequestosome 1/p62-related pathways as therapeutic targets in hepatocellular carcinoma. Expert Opin. Ther. Targets 2019, 23, 393–406. [Google Scholar] [CrossRef]
  152. Jiang, T.; Harder, B.; Rojo de la Vega, M.; Wong, P.K.; Chapman, E.; Zhang, D.D. p62 links autophagy and Nrf2 signaling. Free Radic. Biol. Med. 2015, 88, 199–204. [Google Scholar] [CrossRef] [Green Version]
  153. Komatsu, M.; Kurokawa, H.; Waguri, S.; Taguchi, K.; Kobayashi, A.; Ichimura, Y.; Sou, Y.S.; Ueno, I.; Sakamoto, A.; Tong, K.I.; et al. The selective autophagy substrate p62 activates the stress responsive transcription factor Nrf2 through inactivation of Keap1. Nat. Cell Biol. 2010, 12, 213–223. [Google Scholar] [CrossRef] [PubMed]
  154. Yamada, T.; Dawson, T.M.; Yanagawa, T.; Iijima, M.; Sesaki, H. SQSTM1/p62 promotes mitochondrial ubiquitination independently of PINK1 and PRKN/parkin in mitophagy. Autophagy 2019, 15, 2012–2018. [Google Scholar] [CrossRef] [PubMed]
  155. Williams, J.A.; Ding, W.X. Targeting Pink1-Parkin-mediated mitophagy for treating liver injury. Pharmacol. Res. 2015, 102, 264–269. [Google Scholar] [CrossRef] [Green Version]
  156. Wang, H.; Ni, H.M.; Chao, X.; Ma, X.; Rodriguez, Y.A.; Chavan, H.; Wang, S.; Krishnamurthy, P.; Dobrowsky, R.; Xu, D.X.; et al. Double deletion of PINK1 and Parkin impairs hepatic mitophagy and exacerbates acetaminophen-induced liver injury in mice. Redox Biol. 2019, 22, 101148. [Google Scholar] [CrossRef] [PubMed]
  157. Park, J.S.; Lee, D.H.; Lee, Y.S.; Oh, E.; Bae, K.H.; Oh, K.J.; Kim, H.; Bae, S.H. Dual roles of ULK1 (unc-51 like autophagy activating kinase 1) in cytoprotection against lipotoxicity. Autophagy 2020, 16, 86–105. [Google Scholar] [CrossRef] [PubMed]
  158. Duran, A.; Amanchy, R.; Linares, J.F.; Joshi, J.; Abu-Baker, S.; Porollo, A.; Hansen, M.; Moscat, J.; Diaz-Meco, M.T. p62 is a key regulator of nutrient sensing in the mTORC1 pathway. Mol. Cell 2011, 44, 134–146. [Google Scholar] [CrossRef] [Green Version]
  159. Linares, J.F.; Duran, A.; Reina-Campos, M.; Aza-Blanc, P.; Campos, A.; Moscat, J.; Diaz-Meco, M.T. Amino Acid Activation of mTORC1 by a PB1-Domain-Driven Kinase Complex Cascade. Cell Rep. 2015, 12, 1339–1352. [Google Scholar] [CrossRef] [Green Version]
  160. Linares, J.F.; Duran, A.; Yajima, T.; Pasparakis, M.; Moscat, J.; Diaz-Meco, M.T. K63 polyubiquitination and activation of mTOR by the p62-TRAF6 complex in nutrient-activated cells. Mol. Cell 2013, 51, 283–296. [Google Scholar] [CrossRef] [Green Version]
  161. Lau, A.; Wang, X.J.; Zhao, F.; Villeneuve, N.F.; Wu, T.; Jiang, T.; Sun, Z.; White, E.; Zhang, D.D. A noncanonical mechanism of Nrf2 activation by autophagy deficiency: Direct interaction between Keap1 and p62. Mol. Cell Biol. 2010, 30, 3275–3285. [Google Scholar] [CrossRef] [Green Version]
  162. Ichimura, Y.; Waguri, S.; Sou, Y.S.; Kageyama, S.; Hasegawa, J.; Ishimura, R.; Saito, T.; Yang, Y.; Kouno, T.; Fukutomi, T.; et al. Phosphorylation of p62 activates the Keap1-Nrf2 pathway during selective autophagy. Mol. Cell 2013, 51, 618–631. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  163. Fan, W.; Tang, Z.; Chen, D.; Moughon, D.; Ding, X.; Chen, S.; Zhu, M.; Zhong, Q. Keap1 facilitates p62-mediated ubiquitin aggregate clearance via autophagy. Autophagy 2010, 6, 614–621. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  164. Ishii, T.; Itoh, K.; Yamamoto, M. Roles of Nrf2 in activation of antioxidant enzyme genes via antioxidant responsive elements. Methods Enzymol. 2002, 348, 182–190. [Google Scholar]
  165. Itoh, K.; Tong, K.I.; Yamamoto, M. Molecular mechanism activating Nrf2-Keap1 pathway in regulation of adaptive response to electrophiles. Free Radic. Biol. Med. 2004, 36, 1208–1213. [Google Scholar] [CrossRef]
  166. Kageyama, S.; Saito, T.; Obata, M.; Koide, R.H.; Ichimura, Y.; Komatsu, M. Negative regulation of the Keap1-Nrf2 pathway by a p62/Sqstm1 splicing variant. Mol. Cell Biol. 2018, 38, 00642-17. [Google Scholar] [CrossRef] [Green Version]
  167. Zhu, C.; Tabas, I.; Schwabe, R.F.; Pajvani, U.B. Maladaptive regeneration—The reawakening of developmental pathways in NASH and fibrosis. Nat. Rev. Gastroenterol. Hepatol. 2021, 18, 131–142. [Google Scholar] [CrossRef]
  168. Zhu, C.; Kim, K.; Wang, X.; Bartolome, A.; Salomao, M.; Dongiovanni, P.; Meroni, M.; Graham, M.J.; Yates, K.P.; Diehl, A.M.; et al. Hepatocyte Notch activation induces liver fibrosis in nonalcoholic steatohepatitis. Sci. Transl. Med. 2018, 10, eaat0344. [Google Scholar] [CrossRef]
  169. Syn, W.K.; Choi, S.S.; Liaskou, E.; Karaca, G.F.; Agboola, K.M.; Oo, Y.H.; Mi, Z.; Pereira, T.A.; Zdanowicz, M.; Malladi, P.; et al. Osteopontin is induced by hedgehog pathway activation and promotes fibrosis progression in nonalcoholic steatohepatitis. Hepatology 2011, 53, 106–115. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  170. Verdelho Machado, M.; Diehl, A.M. Role of Hedgehog Signaling Pathway in NASH. Int. J. Mol. Sci. 2016, 17, 857. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  171. Canto, C.; Menzies, K.J.; Auwerx, J. NAD(+) Metabolism and the Control of Energy Homeostasis: A Balancing Act between Mitochondria and the Nucleus. Cell Metab. 2015, 22, 31–53. [Google Scholar] [CrossRef] [Green Version]
  172. Zhu, Y.; Yan, Y.; Gius, D.R.; Vassilopoulos, A. Metabolic regulation of Sirtuins upon fasting and the implication for cancer. Curr. Opin. Oncol. 2013, 25, 630–636. [Google Scholar] [CrossRef] [Green Version]
  173. He, J.; Hu, B.; Shi, X.; Weidert, E.R.; Lu, P.; Xu, M.; Huang, M.; Kelley, E.E.; Xie, W. Activation of the aryl hydrocarbon receptor sensitizes mice to nonalcoholic steatohepatitis by deactivating mitochondrial sirtuin deacetylase Sirt3. Mol. Cell Biol. 2013, 33, 2047–2055. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  174. Pantazi, E.; Zaouali, M.A.; Bejaoui, M.; Folch-Puy, E.; Ben Abdennebi, H.; Rosello-Catafau, J. Role of sirtuins in ischemia-reperfusion injury. World J. Gastroenterol. 2013, 19, 7594–7602. [Google Scholar] [CrossRef] [Green Version]
  175. Hubbi, M.E.; Hu, H.; Kshitiz; Gilkes, D.M.; Semenza, G.L. Sirtuin-7 inhibits the activity of hypoxia-inducible factors. J. Biol. Chem. 2013, 288, 20768–20775. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  176. Xu, C.; Wang, L.; Fozouni, P.; Evjen, G.; Chandra, V.; Jiang, J.; Lu, C.; Nicastri, M.; Bretz, C.; Winkler, J.D.; et al. SIRT1 is downregulated by autophagy in senescence and ageing. Nat. Cell Biol. 2020, 22, 1170–1179. [Google Scholar] [CrossRef] [PubMed]
  177. Aravinthan, A.; Pietrosi, G.; Hoare, M.; Jupp, J.; Marshall, A.; Verrill, C.; Davies, S.; Bateman, A.; Sheron, N.; Allison, M.; et al. Hepatocyte expression of the senescence marker p21 is linked to fibrosis and an adverse liver-related outcome in alcohol-related liver disease. PLoS ONE 2013, 8, e72904. [Google Scholar] [CrossRef] [Green Version]
  178. Aravinthan, A.; Scarpini, C.; Tachtatzis, P.; Verma, S.; Penrhyn-Lowe, S.; Harvey, R.; Davies, S.E.; Allison, M.; Coleman, N.; Alexander, G. Hepatocyte senescence predicts progression in non-alcohol-related fatty liver disease. J. Hepatol. 2013, 58, 549–556. [Google Scholar] [CrossRef]
  179. Aravinthan, A.D.; Alexander, G.J.M. Senescence in chronic liver disease: Is the future in aging? J. Hepatol. 2016, 65, 825–834. [Google Scholar] [CrossRef] [Green Version]
  180. Ogrodnik, M.; Jurk, D. Senescence explains age- and obesity-related liver steatosis. Cell Stress 2017, 1, 70–72. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  181. Ogrodnik, M.; Miwa, S.; Tchkonia, T.; Tiniakos, D.; Wilson, C.L.; Lahat, A.; Day, C.P.; Burt, A.; Palmer, A.; Anstee, Q.M.; et al. Cellular senescence drives age-dependent hepatic steatosis. Nat. Commun. 2017, 8, 15691. [Google Scholar] [CrossRef]
  182. Cazzagon, N.; Sarcognato, S.; Floreani, A.; Corra, G.; de Martin, S.; Guzzardo, V.; Russo, F.P.; Guido, M. Cholangiocyte senescence in primary sclerosing cholangitis is associated with disease severity and prognosis. JHEP Rep. 2021, 3, 100286. [Google Scholar] [CrossRef] [PubMed]
  183. Ferreira-Gonzalez, S.; Lu, W.Y.; Raven, A.; Dwyer, B.; Man, T.Y.; O’Duibhir, E.; Lewis, P.J.S.; Campana, L.; Kendall, T.J.; Bird, T.G.; et al. Paracrine cellular senescence exacerbates biliary injury and impairs regeneration. Nat. Commun. 2018, 9, 1020. [Google Scholar] [CrossRef]
  184. Chapman, J.; Fielder, E.; Passos, J.F. Mitochondrial dysfunction and cell senescence: Deciphering a complex relationship. FEBS Lett. 2019, 593, 1566–1579. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  185. Correia-Melo, C.; Marques, F.D.; Anderson, R.; Hewitt, G.; Hewitt, R.; Cole, J.; Carroll, B.M.; Miwa, S.; Birch, J.; Merz, A.; et al. Mitochondria are required for pro-ageing features of the senescent phenotype. EMBO J. 2016, 35, 724–742. [Google Scholar] [CrossRef] [PubMed]
  186. Korolchuk, V.I.; Miwa, S.; Carroll, B.; von Zglinicki, T. Mitochondria in Cell Senescence: Is Mitophagy the Weakest Link? EBioMedicine 2017, 21, 7–13. [Google Scholar] [CrossRef] [Green Version]
  187. Wiley, C.D.; Velarde, M.C.; Lecot, P.; Liu, S.; Sarnoski, E.A.; Freund, A.; Shirakawa, K.; Lim, H.W.; Davis, S.S.; Ramanathan, A.; et al. Mitochondrial Dysfunction Induces Senescence with a Distinct Secretory Phenotype. Cell Metab. 2016, 23, 303–314. [Google Scholar] [CrossRef] [Green Version]
  188. Ristow, M.; Zarse, K. How increased oxidative stress promotes longevity and metabolic health: The concept of mitochondrial hormesis (mitohormesis). Exp. Gerontol. 2010, 45, 410–418. [Google Scholar] [CrossRef]
  189. Yi, W.; Lan, H.; Wen, Y.; Wang, Y.; He, D.; Bai, Z.; Zhang, Y.; Jiang, W.; Liu, B.; Shen, J.; et al. HO-1 overexpression alleviates senescence by inducing autophagy via the mitochondrial route in human nucleus pulposus cells. J. Cell Physiol. 2020, 235, 8402–8415. [Google Scholar] [CrossRef]
  190. Dou, F.; Wu, B.; Chen, J.; Liu, T.; Yu, Z.; Chen, C. PPARalpha Targeting GDF11 Inhibits Vascular Endothelial Cell Senescence in an Atherosclerosis Model. Oxid. Med. Cell Longev. 2021, 2021, 2045259. [Google Scholar] [CrossRef]
  191. Hanada, S.; Harada, M.; Abe, M.; Akiba, J.; Sakata, M.; Kwan, R.; Taniguchi, E.; Kawaguchi, T.; Koga, H.; Nagata, E.; et al. Aging modulates susceptibility to mouse liver Mallory-Denk body formation. J. Histochem. Cytochem. 2012, 60, 475–483. [Google Scholar] [CrossRef] [Green Version]
  192. Ding, J.; Li, H.Y.; Zhang, L.; Zhou, Y.; Wu, J. Hedgehog Signaling, a Critical Pathway Governing the Development and Progression of Hepatocellular Carcinoma. Cells 2021, 10, 123. [Google Scholar] [CrossRef] [PubMed]
  193. Chan, I.S.; Guy, C.D.; Chen, Y.; Lu, J.; Swiderska-Syn, M.; Michelotti, G.A.; Karaca, G.; Xie, G.; Kruger, L.; Syn, W.K.; et al. Paracrine Hedgehog signaling drives metabolic changes in hepatocellular carcinoma. Cancer Res. 2012, 72, 6344–6350. [Google Scholar] [CrossRef] [Green Version]
  194. Bruschi, F.V.; Tardelli, M.; Herac, M.; Claudel, T.; Trauner, M. Metabolic regulation of hepatic PNPLA3 expression and severity of liver fibrosis in patients with NASH. Liver Int. 2020, 40, 1098–1110. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  195. Bruschi, F.V.; Tardelli, M.; Einwallner, E.; Claudel, T.; Trauner, M. PNPLA3 I148M Up-Regulates Hedgehog and Yap Signaling in Human Hepatic Stellate Cells. Int. J. Mol. Sci. 2020, 21, 8711. [Google Scholar] [CrossRef] [PubMed]
  196. Rao, M.S.; Reddy, J.K. PPARalpha in the pathogenesis of fatty liver disease. Hepatology 2004, 40, 783–786. [Google Scholar] [CrossRef]
  197. Souza-Mello, V. Peroxisome proliferator-activated receptors as targets to treat non-alcoholic fatty liver disease. World J. Hepatol. 2015, 7, 1012–1019. [Google Scholar] [CrossRef]
  198. Pawlak, M.; Lefebvre, P.; Staels, B. Molecular mechanism of PPARalpha action and its impact on lipid metabolism, inflammation and fibrosis in non-alcoholic fatty liver disease. J. Hepatol. 2015, 62, 720–733. [Google Scholar] [CrossRef] [Green Version]
  199. Huang, H.; McIntosh, A.L.; Martin, G.G.; Petrescu, A.D.; Landrock, K.K.; Landrock, D.; Kier, A.B.; Schroeder, F. Inhibitors of Fatty Acid Synthesis Induce PPAR alpha-Regulated Fatty Acid beta*Oxidative Genes: Synergistic Roles of L-FABP and Glucose. PPAR Res. 2013, 2013, 865604. [Google Scholar] [CrossRef] [Green Version]
  200. Harmon, G.S.; Lam, M.T.; Glass, C.K. PPARs and lipid ligands in inflammation and metabolism. Chem. Rev. 2011, 111, 6321–6340. [Google Scholar] [CrossRef] [Green Version]
  201. Mossa, A.H.; Velasquez Flores, M.; Cammisotto, P.G.; Campeau, L. Succinate, increased in metabolic syndrome, activates GPR91 receptor signaling in urothelial cells. Cell Signal. 2017, 37, 31–39. [Google Scholar] [CrossRef]
  202. Sadagopan, N.; Li, W.; Roberds, S.L.; Major, T.; Preston, G.M.; Yu, Y.; Tones, M.A. Circulating succinate is elevated in rodent models of hypertension and metabolic disease. Am. J. Hypertens 2007, 20, 1209–1215. [Google Scholar] [CrossRef]
  203. Gonzalez, F.J.; Xie, C.; Jiang, C. The role of hypoxia-inducible factors in metabolic diseases. Nat. Rev. Endocrinol. 2018, 15, 21–32. [Google Scholar] [CrossRef]
  204. Sundaram, S.S.; Halbower, A.; Pan, Z.; Robbins, K.; Capocelli, K.E.; Klawitter, J.; Shearn, C.T.; Sokol, R.J. Nocturnal hypoxia-induced oxidative stress promotes progression of pediatric non-alcoholic fatty liver disease. J. Hepatol. 2016, 65, 560–569. [Google Scholar] [CrossRef] [Green Version]
  205. Chen, J.; Chen, J.; Fu, H.; Li, Y.; Wang, L.; Luo, S.; Lu, H. Hypoxia exacerbates nonalcoholic fatty liver disease via the HIF-2alpha/PPARalpha pathway. Am. J. Physiol. Endocrinol. Metab. 2019, 317, E710–E722. [Google Scholar] [CrossRef] [PubMed]
  206. Chen, J.; Chen, J.; Huang, J.; Li, Z.; Gong, Y.; Zou, B.; Liu, X.; Ding, L.; Li, P.; Zhu, Z.; et al. HIF-2alpha upregulation mediated by hypoxia promotes NAFLD-HCC progression by activating lipid synthesis via the PI3K-AKT-mTOR pathway. Aging 2019, 11, 10839–10860. [Google Scholar] [CrossRef] [PubMed]
  207. Qu, A.; Taylor, M.; Xue, X.; Matsubara, T.; Metzger, D.; Chambon, P.; Gonzalez, F.J.; Shah, Y.M. Hypoxia-inducible transcription factor 2alpha promotes steatohepatitis through augmenting lipid accumulation, inflammation, and fibrosis. Hepatology 2011, 54, 472–483. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  208. Rankin, E.B.; Rha, J.; Selak, M.A.; Unger, T.L.; Keith, B.; Liu, Q.; Haase, V.H. Hypoxia-inducible factor 2 regulates hepatic lipid metabolism. Mol. Cell Biol. 2009, 29, 4527–4538. [Google Scholar] [CrossRef] [Green Version]
  209. Ham, P.B., 3rd; Raju, R. Mitochondrial function in hypoxic ischemic injury and influence of aging. Prog. Neurobiol. 2017, 157, 92–116. [Google Scholar] [CrossRef] [PubMed]
  210. Fuhrmann, D.C.; Wittig, I.; Heide, H.; Dehne, N.; Brune, B. Chronic hypoxia alters mitochondrial composition in human macrophages. Biochim. Biophys. Acta 2013, 1834, 2750–2760. [Google Scholar] [CrossRef]
  211. Keiran, N.; Ceperuelo-Mallafre, V.; Calvo, E.; Hernandez-Alvarez, M.I.; Ejarque, M.; Nunez-Roa, C.; Horrillo, D.; Maymo-Masip, E.; Rodriguez, M.M.; Fradera, R.; et al. SUCNR1 controls an anti-inflammatory program in macrophages to regulate the metabolic response to obesity. Nat. Immunol. 2019, 20, 581–592. [Google Scholar] [CrossRef]
  212. Ferreira-Gonzalez, S.; Rodrigo-Torres, D.; Gadd, V.L.; Forbes, S.J. Cellular Senescence in Liver Disease and Regeneration. Semin. Liver Dis. 2021, 41, 50–66. [Google Scholar] [CrossRef]
  213. Meijnikman, A.S.; Herrema, H.; Scheithauer, T.P.M.; Kroon, J.; Nieuwdorp, M.; Groen, A.K. Evaluating causality of cellular senescence in non-alcoholic fatty liver disease. JHEP Rep. 2021, 3, 100301. [Google Scholar] [CrossRef]
  214. Moustakas, I.I.; Katsarou, A.; Legaki, A.I.; Pyrina, I.; Ntostoglou, K.; Papatheodoridi, A.M.; Gercken, B.; Pateras, I.S.; Gorgoulis, V.G.; Koutsilieris, M.; et al. Hepatic Senescence Accompanies the Development of NAFLD in Non-Aged Mice Independently of Obesity. Int. J. Mol. Sci. 2021, 22, 3446. [Google Scholar] [CrossRef] [PubMed]
  215. Palmer, A.K.; Xu, M.; Zhu, Y.; Pirtskhalava, T.; Weivoda, M.M.; Hachfeld, C.M.; Prata, L.G.; van Dijk, T.H.; Verkade, E.; Casaclang-Verzosa, G.; et al. Targeting senescent cells alleviates obesity-induced metabolic dysfunction. Aging Cell 2019, 18, e12950. [Google Scholar] [CrossRef]
  216. Sanguino, E.; Roglans, N.; Alegret, M.; Sanchez, R.M.; Vazquez-Carrera, M.; Laguna, J.C. Atorvastatin reverses age-related reduction in rat hepatic PPARalpha and HNF-4. Br. J. Pharmacol. 2005, 145, 853–861. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  217. Yang, S.; Koteish, A.; Lin, H.; Huang, J.; Roskams, T.; Dawson, V.; Diehl, A.M. Oval cells compensate for damage and replicative senescence of mature hepatocytes in mice with fatty liver disease. Hepatology 2004, 39, 403–411. [Google Scholar] [CrossRef]
  218. Zimmermann, H.W.; Koch, A.; Seidler, S.; Trautwein, C.; Tacke, F. Circulating soluble urokinase plasminogen activator is elevated in patients with chronic liver disease, discriminates stage and aetiology of cirrhosis and predicts prognosis. Liver Int. 2012, 32, 500–509. [Google Scholar] [CrossRef]
  219. Wyss, R.K.; Mendez Carmona, N.; Arnold, M.; Segiser, A.; Mueller, M.; Dutkowski, P.; Carrel, T.P.; Longnus, S.L. Hypothermic, oxygenated perfusion (HOPE) provides cardioprotection via succinate oxidation prior to normothermic perfusion in a rat model of donation after circulatory death (DCD). Am. J. Transplant. 2021, 21, 1003–1011. [Google Scholar] [CrossRef]
  220. Fafian-Labora, J.A.; Rodriguez-Navarro, J.A.; O’Loghlen, A. Small Extracellular Vesicles Have GST Activity and Ameliorate Senescence-Related Tissue Damage. Cell Metab. 2020, 32, 71–86. [Google Scholar] [CrossRef] [PubMed]
  221. Fafian-Labora, J.A.; O’Loghlen, A. Classical and Nonclassical Intercellular Communication in Senescence and Ageing. Trends Cell Biol. 2020, 30, 628–639. [Google Scholar] [CrossRef] [PubMed]
  222. James, A.M.; Cocheme, H.M.; Smith, R.A.; Murphy, M.P. Interactions of mitochondria-targeted and untargeted ubiquinones with the mitochondrial respiratory chain and reactive oxygen species. Implications for the use of exogenous ubiquinones as therapies and experimental tools. J. Biol. Chem. 2005, 280, 21295–21312. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  223. James, A.M.; Sharpley, M.S.; Manas, A.R.; Frerman, F.E.; Hirst, J.; Smith, R.A.; Murphy, M.P. Interaction of the mitochondria-targeted antioxidant MitoQ with phospholipid bilayers and ubiquinone oxidoreductases. J. Biol. Chem. 2007, 282, 14708–14718. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  224. Fink, B.D.; Herlein, J.A.; Guo, D.F.; Kulkarni, C.; Weidemann, B.J.; Yu, L.; Grobe, J.L.; Rahmouni, K.; Kerns, R.J.; Sivitz, W.I. A mitochondrial-targeted coenzyme q analog prevents weight gain and ameliorates hepatic dysfunction in high-fat-fed mice. J. Pharmacol. Exp. Ther. 2014, 351, 699–708. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  225. Fink, B.D.; Herlein, J.A.; Yorek, M.A.; Fenner, A.M.; Kerns, R.J.; Sivitz, W.I. Bioenergetic effects of mitochondrial-targeted coenzyme Q analogs in endothelial cells. J. Pharmacol. Exp. Ther. 2012, 342, 709–719. [Google Scholar] [CrossRef] [PubMed]
  226. Suliman, H.B.; Piantadosi, C.A. Mitochondrial Quality Control as a Therapeutic Target. Pharmacol. Rev. 2015, 68, 20–48. [Google Scholar] [CrossRef] [PubMed]
  227. Hickson, L.J.; Langhi Prata, L.G.P.; Bobart, S.A.; Evans, T.K.; Giorgadze, N.; Hashmi, S.K.; Herrmann, S.M.; Jensen, M.D.; Jia, Q.; Jordan, K.L.; et al. Senolytics decrease senescent cells in humans: Preliminary report from a clinical trial of Dasatinib plus Quercetin in individuals with diabetic kidney disease. EBioMedicine 2019, 47, 446–456. [Google Scholar] [CrossRef] [Green Version]
  228. Sheard, M.A.; Vojtesek, B.; Simickova, M.; Valik, D. Release of cytokeratin-18 and -19 fragments (TPS and CYFRA 21-1) into the extracellular space during apoptosis. J. Cell Biochem. 2002, 85, 670–677. [Google Scholar] [CrossRef]
  229. Tarantino, G.; Conca, P.; Coppola, A.; Vecchione, R.; di Minno, G. Serum concentrations of the tissue polypeptide specific antigen in patients suffering from non-alcoholic steatohepatitis. Eur. J. Clin. Investig. 2007, 37, 48–53. [Google Scholar] [CrossRef]
Figure 1. Overview on the PTAS model for steatohepatitis.
Figure 1. Overview on the PTAS model for steatohepatitis.
Ijms 22 12545 g001
Table 2. Factors that trigger acute mitochondrial damage in the liver.
Table 2. Factors that trigger acute mitochondrial damage in the liver.
FactorMechanismModel
Porphyrinogens (DDC, griseofulvin); liver toxinsInhibition of ferrochelatase (haem deficiency)Intoxication mouse models [28,50,51,52,53,100]; DILI [12,13,14]
Excess keratin 8 1Impaired mitochondrial QC via Pirh2Keratin 18−/− mouse model, HFD mouse model, human NASH [65,74,78,79,80,81]
HFD 2Increased ROS production by β-oxidation of fatty acidsHFD model, human NASH [18,22,27,31,93,117]
1 Keratin 8 excess might prime due to impaired NADPH production in the TCA cycle, reducing mitochondrial GSH, and trigger through increased ROS production due to accumulating mitochondrial damage. 2 HFD might need a separate trigger since HFD mouse models do not produce the full SH phenotype [16].
Table 3. Outcome of priming and triggering, leading to the CDM.
Table 3. Outcome of priming and triggering, leading to the CDM.
StageOutcomeEffect
primingPersistently reduced PPARαpseudohypoxia
triggeringDamage to mitochondrial ETC
Reduced mitochondrial QC
succinate ↑
DAMP, ROS ↑, succinate ↑, pseudohypoxia, cell cycle arrest
DAMP, stellate cell activation, immune response, inflammation
Keratin 8 excessReduced mitochondrial QC, cell cycle arrest 1
1 The implication of Pirh2 and HuR in SH is presently not clear but represents a testable hypothesis.
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Abuja, P.M.; Zatloukal, K.; Denk, H. Priming, Triggering, Adaptation and Senescence (PTAS): A Hypothesis for a Common Damage Mechanism of Steatohepatitis. Int. J. Mol. Sci. 2021, 22, 12545. https://doi.org/10.3390/ijms222212545

AMA Style

Abuja PM, Zatloukal K, Denk H. Priming, Triggering, Adaptation and Senescence (PTAS): A Hypothesis for a Common Damage Mechanism of Steatohepatitis. International Journal of Molecular Sciences. 2021; 22(22):12545. https://doi.org/10.3390/ijms222212545

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Abuja, Peter M., Kurt Zatloukal, and Helmut Denk. 2021. "Priming, Triggering, Adaptation and Senescence (PTAS): A Hypothesis for a Common Damage Mechanism of Steatohepatitis" International Journal of Molecular Sciences 22, no. 22: 12545. https://doi.org/10.3390/ijms222212545

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Abuja, P. M., Zatloukal, K., & Denk, H. (2021). Priming, Triggering, Adaptation and Senescence (PTAS): A Hypothesis for a Common Damage Mechanism of Steatohepatitis. International Journal of Molecular Sciences, 22(22), 12545. https://doi.org/10.3390/ijms222212545

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