It is believed that I/R stimulates autophagic cascades which provide the elimination of intracellular proteinaceous aggregates and damaged organelles formed during I/R [34]. These processes play an important role after renal I/R since the knockout of *Atg5* and *Atg7* leads to a more severe AKI [48]. Relying on the LC3 II/LC3 I ratio, we may assume that, in old rats, autophagy is activated not as effectively as it takes place in young animals. Earlier, we demonstrated impaired autophagy in response to the damaging effect of I/R in premature senescent OXYS rats [25]; that could be another argument in favor of the dysfunction of the autophagic system caused by aging. In our experiments, even a preliminary exposure to DR, which is one of the most powerful autophagy-stimulating factors, did not restore the effective work of that system in old rats after I/R. On the contrary, in young animals, DR increased the LC3 II/LC3 I ratio and the amount of lysosomes that correlated with a decrease in the severity of AKI.

The deterioration of autophagy with aging also worsens the functioning of mitophagy, which is the only mechanism for mitochondrial degradation [49]. The elimination of mitochondria with low membrane potential is very important for the cells since dysfunctional mitochondria may be the source of pathological ROS [50]. For instance, in mice with a knockout of *Atg5*, an elevated accumulation

of damaged mitochondria after I/R was shown, which correlated with an increased AKI [48]. It is also suggested that the accumulation of damaged mitochondria as a result of the ineffective work of mitophagy leads to the progression of tissue aging [51]. In many organisms, aging is associated with the morphological alterations and the increased size of mitochondria [52], as well as with the accumulation of age-associated proteins [53].

In this study, we evaluated the intensity of mitophagy by the levels of PINK1 in isolated mitochondria. This protein is considered to be the marker of damaged mitochondria and the initiator of mitophagy. PINK1 carries a mitochondrial address and is delivered into mitochondria. When the mitochondrial membrane potential decreases, this protein is unable to penetrate into the mitochondria and cannot be cleaved by presenilin-associated rhomboid-like protein (PARL). Consequently, PINK1 remains located on the outer membrane of the mitochondria with a low potential, where it is marked by the ubiquitin ligase Parkin, which serves as a signal for the onset of mitophagy [54].

An increase in the levels of PINK1 is usually associated with the accumulation of damaged mitochondria in the cell. For instance, a high-calorie diet causes the rise in the levels of PINK1 whereas DR decreases the amount of this protein in mitochondria [55]. In our experiments, we observed a two-fold decrease in PINK1 levels in the mitochondrial fraction after 4 weeks of 35% DR in young rats, which shows the result of the effective elimination of low functional mitochondria. In old rats, the PINK1 levels were initially lower than in young animals, and DR did not affect the levels of PINK1. We speculate that the lower content of PINK1 in older rats may indicate a deficiency in this protein and the malfunctioning of the mechanism of mitophagy. A similar decrease of the PINK1 levels was observed in lung tissue with aging [56] and was accompanied by a pulmonary pathology. Moreover, the decrease in PINK1 activity was described in many pathological conditions, for example, in cells of patients with Parkinson's disease [57]. It was shown that decreased PINK1 synthesis led to the fragmentation of mitochondria, decrease of the mitochondrial membrane potential [58], the development of fibrosis [59], which indicates the importance of the PINK1 protein in the normal functioning of mitochondria.

Since mitophagy is the only way to eliminate poorly functioning mitochondria, deterioration in the work of this system must inevitably lead to the accumulation of mitochondria with low membrane potentials [60]. Indeed, a large number of studies have demonstrated that mitochondria isolated from the tissues of old organisms had lower membrane potentials than mitochondria from young [52,61]. Taken together with higher levels of ROS in the tissues of old animals, it is assumed that age impacts either on the functioning of mitochondria or on the process of their quality control [50]. Our findings fit this assumption as we showed a lower mitochondrial membrane potential in kidney cells isolated from old rats, as well as a higher heterogeneity in mitochondria of cells obtained from old rats' kidneys revealed by flow cytometry. A similar heterogeneity of mitochondrial membrane potential was observed in hepatocytes from old rats [62]. Earlier, the phenomenon of the augmentation of the mitochondrial membrane potential heterogeneity in the kidney tissue after I/R was reported [63]. We also revealed an increase in LysoTracker Green distribution heterogeneity in the tubules of old rats. Probably, the increase in heterogeneity is a process that accompanies the aging of tissues.

Another important factor reflecting the state of mitochondria is the activity of deacetylase SIRT-3. This NAD+-dependent deacetylase is localized predominantly in mitochondria and is considered to be a mitochondrial stress sensor that can modulate the activity of several mitochondrial proteins involved in metabolism and oxidative stress regulatory pathways [64]. SIRT-3 is suggested to be one of the mediators of the protective effect of DR [65]. In this study, DR increased the levels of SIRT-3 more than 2 times in the kidneys of young rats. In kidneys of old rats, the levels of SIRT-3 were significantly lower than in young, while DR significantly increased the content of this deacetylase. An age-associated drop of mitochondrial SIRT-3 was previously reported for some tissues of old people and old animals [66,67]. Contrarily, some variants of the *SIRT-3* gene were associated with longevity in humans [68], proving the key role of this deacetylase in the aging process.

Recent studies have demonstrated that increased levels of SIRT-3 protect the kidney from cisplatin nephrotoxicity [69], whereas the knockout of *SIRT-3* led to more severe damage after myocardial infarction [70]. The levels of SIRT-3 depended on physical activity and diet, e.g., the levels of the deacetylase rose in muscles after exercise [71] and in cells exposed to decreased glucose concentration in culture medium [72]. In addition, the knockout of *SIRT-3* in the kidneys of mice caused fibrosis, increased in the TGF-β1 level and the hyperacetylation of the kinase of glycogen synthase 3β (GSK3β) that adversely affected its phosphorylating activity [73]. Similarly, another deacetylase, SIRT-1, was detected to be lower in old animals, while 12-month-long DR increased the levels of this enzyme [44].

Our experiments with cell cultures showed that primary RTCs obtained from kidneys of old rats proliferated 4 times slower compared to cells from the kidneys of young rats. Such a dependence of proliferation rate on the age of the donor was demonstrated for various cells. For instance, hematopoietic stem cells derived from neonatal rats proliferated much better, and, unlike stem cells derived from adult rats, they were able to exhibit protective properties under myocardial I/R [74]. Although cardiomyocytes obtained from adult animals are thought to represent processes in the tissue better than neonatal cardiomyocytes, they proliferate much more slowly in culture [75].

Furthermore, primary kidney cells from old rats not only proliferated worse than those of young rats, but we also observed their less pronounced recovering ability after OGD. We believe that this impairment could be explained by deterioration in the functioning of the autophagy system, which could lead to the accumulation of lipofuscin, as well as to more pronounced oxidative stress.

#### **5. Conclusions**

In this study, we showed that a short-term 35% DR for 4 weeks has a significant nephroprotective effect against renal I/R in young rats, but DR does not provide such a beneficial effect in old rats. The lack of effectiveness in old animals can be explained by the deterioration in the work of both autophagy and mitophagy. Impairment in these recycling mechanisms results in accumulation of poorly functioning mitochondria, lipofuscin granules, and increased levels of oxidative stress. All these processes affect the normal functioning of the kidney and lead to a decrease in the ischemic tolerance of renal tissue. Molecular and structural alterations occurring in aging renal tissue make the manifestation of beneficial effects of DR in aged kidneys not as pronounced. Thus, an important problem is to develop approaches that would be helpful in the therapy of elderly patients.

**Author Contributions:** Conceptualization, S.S.J., E.Y.P. and N.V.A.; Methodology, S.S.J., I.B.P. and L.D.Z.; Software, V.A.P.; Validation, I.B.P. and L.D.Z.; Formal Analysis, N.V.A., L.D.Z. and I.B.P.; Investigation, N.V.A., L.D.Z. and E.Y.P.; Resources, D.N.S.; Data Curation, N.V.A. and S.S.J.; Writing-Original Draft Preparation, N.V.A.; Writing-Review & Editing, E.Y.P., D.N.S. and D.B.Z.; Visualization, A.N.V. and V.A.P.; Supervision, E.Y.P. and D.B.Z.; Project Administration, E.Y.P. and D.B.Z.

**Funding:** This research was funded by Russian science foundation grant number 18-15-00058.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

© 2018 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Review* **Autophagy in Chronic Kidney Diseases**

#### **Tien-An Lin 1, Victor Chien-Chia Wu 2 and Chao-Yung Wang 2,3,\***

Received: 8 December 2018; Accepted: 9 January 2019; Published: 16 January 2019

**Abstract:** Autophagy is a cellular recycling process involving self-degradation and reconstruction of damaged organelles and proteins. Current evidence suggests that autophagy is critical in kidney physiology and homeostasis. In clinical studies, autophagy activations and inhibitions are linked to acute kidney injuries, chronic kidney diseases, diabetic nephropathies, and polycystic kidney diseases. Oxidative stress, inflammation, and mitochondrial dysfunction, which are implicated as important mechanisms underlying many kidney diseases, modulate the autophagy activation and inhibition and lead to cellular recycling dysfunction. Abnormal autophagy function can induce loss of podocytes, damage proximal tubular cells, and glomerulosclerosis. After acute kidney injuries, activated autophagy protects tubular cells from apoptosis and enhances cellular regeneration. Patients with chronic kidney diseases have impaired autophagy that cannot be reversed by hemodialysis. Multiple nephrotoxic medications also alter the autophagy signaling, by which the mechanistic insights of the drugs are revealed, thus providing the unique opportunity to manage the nephrotoxicity of these drugs. In this review, we summarize the current concepts of autophagy and its molecular aspects in different kidney cells pathophysiology. We also discuss the current evidence of autophagy in acute kidney injury, chronic kidney disease, toxic effects of drugs, and aging kidneys. In addition, we examine therapeutic possibilities targeting the autophagy system in kidney diseases.

**Keywords:** autophagy; kidney diseases; oxidative stress; inflammation; mitochondria

#### **1. Introduction**

Autophagy is a dynamic cellular balancing mechanism for energy and resource. The word autophagy is derived from the Greek word, where *auto* means "self" and *phagy* means "eating". This "self-eating" process helps cells recycle their endogenous materials and build essential macromolecules to maintain cellular homeostasis and reutilize energy [1,2]. Autophagy was initially referred to the catabolic process that could provide nutrition and energy to cells during starvation. Recently, more evidence has shown that autophagy plays a critical role in synthesis and degradation and has complex cross-talks to apoptosis and cell cycle regulations [3]. Thus, autophagy acts as a protective mechanism in living organisms and can interfere in pathogenesis [4,5].

In 1963, Christian de Duve first described the autophagy as the degradation process that occurred after cytoplasmic materials were delivered to the lysosome. In the 1990 s the autophagy research bloomed after the team of Dr. Yoshinori Ohsumi identified the important genes related to the autophagy-defective mutants in yeasts called the autophagy-related gene (Atg) and its related protein [6]. Soon after, the research advances that focused on the genetic aspects and several Atg proteins were discovered [4,5], providing a better understanding of the function and mechanisms of autophagy. Recently, autophagy has been widely implicated not only in yeasts and animal models

but also in human pathophysiological processes [5,7]. In 2016, Yoshinori Ohsumi was awarded the Nobel Prize in Physiology or Medicine in recognition for his work on laying the foundation for a better understanding of the ability of cells to manage starvation, stress, and diseases [4].

Autophagy works through intracellular lysosomal degradation and recycling, and in turn, helps to maintain cellular integrity more efficiently by regenerating metabolic precursors and clearing subcellular debris [5]. Autophagy is a series of catabolic processes, starting with a small membrane phagophore in the cytoplasm and elongating to form a cup-shaped structure. It then matures and becomes a double-membrane structure called autophagosome that engulfs the damaged components. The autophagosome then fuses with lysosomes to form autophagolysosome [4,8] (Figure 1). After forming the autophagolysosome, the mTOR (mammalian target of rapamycin) signaling is activated to degenerate the intercellular components and transport back to the cytoplasm to reuse the macromolecule [9].

**Figure 1.** Schematic overview of the normal autophagy function in the kidney. Multiple steps in autophagy are modulated in kidney diseases, including autophagy initiation, elongation, maturation, fusion, and final degradation and recycling.

Defective autophagy signaling is now found in multiple diseases, such as autoimmune diseases, infectious diseases, metabolic diseases, muscular disorders, neurodegenerative diseases, cardiovascular and pulmonary diseases [5,7,10]. Many pathophysiologic mechanisms, including ischemic, toxic, infection, oxidative stress, circadian rhythm, and aging, are also confirmed to have close interactions with autophagy. In certain stressed situations, selective autophagy occurs in order to remove toxic materials within cells and organs [1,11]. Currently, only a few studies review the relationship between kidney diseases and autophagy. Therefore, our aim is to summarize the recent advances in understanding the role of autophagy in acute and chronic kidney disease patients [12].

#### **2. Autophagy in Normal Kidney**

Previous studies from human and animals provide the evidence that autophagy has a great impact on the maintenance of renal functions and homeostasis [13]. However, autophagy is nonessential for embryonic renal development. The embryonic *Atg5* knockout mice show no significant impairment of glomerular development, no change in podocyte maturation, normal tubular function, and normal nephrons development [3,14]. In terms of kidney physiology in adult animals, the autophagy affects different renal cell types and helps maintain kidney pathophysiology and homeostasis [13].

#### *2.1. Autophagy in Glomerular Mesangial Cells*

Glomerular mesangial cells are located in the mesangium of the glomerulus, in the centrilobular region of the kidneys. Mesangial cells are specialized pericytes with contractile activities. They regulate glomerular filtration and act as primary producers of the extracellular matrix that constitutes the mesangium, thus playing a vital role in maintaining mesangial matrix homeostasis [3]. Autophagy plays dual roles in modulating mesangial cell survival. After mesangial cells are exposed to stress, autophagy is activated and directed to type II programmed cell death. At the same time, autophagy can also serve as a protective role through transforming growth factor-β1 (TGF-β1) activation and inhibit mesangial cells from apoptosis and necrosis [3].

The advanced glycation end-products (AGEs) induced mesangial cells injury is one of the leading causes of glomerular dysfunction in diabetic nephropathy [15]. Autophagy could serve as a protective manner through increased LC3 cleavage (LC3-II/LC3-I ratio), enhanced Atg5 protein expression, and decreased p62 level in mesangial cells in dose- and time-dependent manners after exposed to AGEs. Also, inhibition of Atg5 expression could aggravate AGEs related mesangial cells injury [16]. This indicates that autophagy may protect mesangial cells from apoptosis. Previous studies suggest that the AGEs could induce autophagy through a RAGE/PI3K/AKT/mTOR signaling pathway in cardiomyocytes [17].

After the environmental toxin cadmium exposure, the reactive oxygen species (ROS) increases and activates glycogen synthase kinase-3β (GSK-3β) to trigger autophagy that promotes mesangial cells death [3,18]. Cadmium exposure can also increase autophagy through Ca2+ signaling and mitochondrial depolarization pathway and serve as a housekeeping process to protect the kidney and an early biomarker for cadmium toxicity [18] (Figure 2A).

**Figure 2.** *Cont.*

(**C**)

**Figure 2.** (**A**). Autophagy in glomerular mesangial cells. AGEs: advanced glycation end-products; Akt: also stand for PKB (Protein kinase B); GSK-3β: glycogen synthase kinase-3β; PI3K: Phosphoinositide 3-kinase; ROS: reactive oxygen species; TGF-β1: transforming growth factor-β1; ER stress: endoplasmic reticulum stress; (**B**). Autophagy in podocytes. PI3K: Phosphoinositide 3-kinase; AMPK: AMP-activated protein kinase Akt: also stand for PKB (Protein kinase B); ROS: reactive oxygen species; ER stress: endoplasmic reticulum stress; (**C**). Autophagy in proximal tubular cells. IRI: ischemic reperfusion injury; ROS: reactive oxygen species; ER stress: endoplasmic reticulum stress; PI3K: Phosphoinositide 3-kinase; BUN, blood urea nitrogen; sCr, serum creatinine.

#### *2.2. Autophagy in Podocytes*

Autophagy is vital in post-mitotic cells, such as neuron cells and podocytes. [19]. Podocytes are highly differentiated epithelial cells located in the visceral site of the Bowman's capsule and surrounding capillaries of the glomerulus. Podocytes have characteristic foot processes called pedicles that wrap around the capillaries and extend into them to form a filtration slit diaphragm. The podocyte foot processes and slit diaphragm control the selective permeability of the glomerular filtration barrier that filters circulating blood to form urine [20]. Podocytes can also act as glomerular filtration rate regulators through contraction and filtration slit closure [21]. Podocyte injuries are typical hallmarks of acute kidney injury and can cause proteinuria and nutrient loss [19,20]. The degree of podocytes damages parallels the severity of glomerulosclerosis, proteinuria, and kidney injury [22].

Studies in mice suggested that podocytes exhibit a high basal level of autophagy with abundant autophagosomes [3]. Nine-week-old mice with podocyte-specific deletion of Class III PI3K vacuolar protein sorting 34 (Vps34), which helps to maintain autophagic regulation, develop early proteinuria, progressive glomerulosclerosis, and renal failure [3]. The podocyte-specific depletion of the Atg5 gene leads to glomerulopathy in aging mice, whose oxidative and ubiquitinated protein accumulation and podocyte endoplasmic reticulum stress eventually lead to loss of podocytes, increased proteinuria, and glomerulosclerosis [20,22]. Moreover, the inadvertently increased autophagosomes in podocytes can be found in different glomerular diseases such as IgA nephropathy, membranous glomerulonephritis, and focal segmental glomerulosclerosis [20,23].

The autophagy activated by mTOR pathway protects podocytes from apoptosis, foot process effacement, and chronic kidney diseases progression [24]. In podocyte-specific mTor knockout mice, the proteinuria and end-stage renal diseases (ESRD) occur in 3–5 weeks. Their podocytes accumulate with autophagosomes, microtubule-associated protein 1A/1B-light chain 3 (LC3), and damaged mitochondria [25]. The evidence suggest that autophagy could be regulated through the mTOR pathway in podocytes in both rat models and human. The physiologic level of mTOR activity inhibits autophagy and maintains autophagosomes at a basal level to remove damaged organelles, excessive lipids, and long-lived or misfolded proteins in podocytes. The decrease of growth factor signal or exogenous stimulation such as starvation or rapamycin could inhibit mTOR pathway and upregulate autophagy function (Figure 2B). The activation of autophagy as demonstrated by increase LC3 and Lysotracker markers enhanced autophagosome and autophagolysosomes formation [25]. The renal biopsy study demonstrates that podocyte markers such as synaptopodin, podocin, CD2AP, and nephrin were also decreased after being treated with sirolimus in podocytes [26]. Although the mechanism is not completely understood, the evidence indicates that autophagy plays a major role in maintaining podocyte homeostasis and renal function.

#### *2.3. Autophagy in Renal Tubular Cells*

Autophagy is important in proximal tubular cells. Proximal tubular cells consume much energy during electrolyte reabsorption, which requires high lysosomal activity and mitochondrial turnover [27]. The Atg5 deletion in both proximal and distal tubules results in severe tubular damage and renal dysfunction. However, distal tubules only-specific Atg5 knockout mice show no tubular damage and have normal renal functions [20,28,29]. This indicates that autophagy is critical and integral in proximal tubule function while distal tubular cells rely less on autophagy for their homeostasis due to its function being more passive and less energy dependent [30]. The tubular cells are vulnerable to renal toxic agents, such as cisplatin, cyclosporin, and cadmium [3,31]. Autophagy activation protects renal tubular cells from these renal toxic agents and eliminates the damaged proteins and DNA [3,11]. Selective tubular cell *Atg5* and *Atg7* knockout mice develop more severe tubular cells damages and acute kidney injury (AKI) after ischemia-reperfusion injury [30]. Another study that applied PI3K inhibitor 3-MA to reduced autophagosome sequestration, revealed that it can significantly reduce autophagy function and cause more severe elevation of BUN and serum Cr while rapamycin treatments showed the opposite effect after ischemia-reperfusion injury in vivo and

in vitro [8] (Figure 2C). Moreover, constant autophagy activations lead to tubular cells atrophy and promote kidney fibrosis. A delicate balance of the autophagy effects will be required to protect the renal proximal tubular cells from nephrotoxicity drugs and ischemic-reperfusion injury [13].

#### **3. Autophagy in Acute Kidney Injury**

AKI is a common clinical condition in critical care units. It presents as an abrupt decline of kidney function and imbalance of water, electrolytes, and protein homeostasis [9]. Patients with AKI are associated with higher morbidity and mortality. Major causes of AKI are infections, nephrotoxins, and ischemia-reperfusion injury all inducing inflammation. These diseases will result in direct tubular cell damage, accumulation of oxidative stress (ROS), and endothelium microvasculature dysfunction [9]. Among all, the ischemia-reperfusion injury is considered the most frequent and important etiology that usually causes severe injury to the renal tubular cells [32,33].

#### *3.1. The Kidney Ischemia-Reperfusion Injury and Autophagy*

The ischemia-reperfusion injury occurs when an organ is exposed to a prolonged duration of blood flow restriction with subsequent restoration of perfusion. The reoxygenation after ischemia will exacerbate the tissue injury and inflammation response [34]. This pathophysiological process is common in many diseases, such as myocardial infarction [35], ischemic stroke, acute kidney injury, trauma, sleep apnea, hypovolemic shock, surgery, and organ dysfunction after transplantation [34]. After ischemia-reperfusion occurs, reactive oxygen species (ROS) in mitochondria will increase and alter cell cycle, damage DNA, and lead to cell dysfunction and death [36]. These ROS and damaged mitochondria are the major upstream cellular signals for the autophagy during renal injury [37,38]. Previous data suggested that ischemia-reperfusion injury to the renal tubular cells results in upregulating the autophagic activity [20,28]. The accumulation of apoptotic cells also activates the autophagy. The activation of autophagy after ischemia is rapid and proceeds to tissue damages or tubular apoptosis [39]. However, newer evidence suggested that prolonged autophagy activations may have adverse effects after ischemic injury in mice. The persistent autophagy activation may trigger renal cell death pathways and exaggerate the kidney damage [9,20]. Although the exact mechanism responsible for the autophagy activation after AKI is still controversial, the autophagy activation after AKI is crucial for the renal protections after AKI.

#### *3.2. Autophagy Protects the Renal Cells from Acute Injury*

Many studies have proven that the autophagy has renoprotective effects on the proximal tubular cells during AKI [37,39]. Deletion of global Atg5 in mice results in a more vulnerable tubular cell phenotype after exposure to hypoxemia and ROS [27]. Mice with the proximal tubules-specific Atg5 knockout exhibits an accumulation of the damaged organelles and proteins in the proximal tubules and irreversible kidney injury [30,40]. The proximal tubule-specific Atg7 knockout mice also have increased renal injuries [31]. Renal protective role of autophagy has also been shown in cisplatin-induced AKI and sepsis-induced AKI. During the resolution phase of the AKI, modulation of autophagy can promote tubular cell regeneration and repair [8,27]. Moreover, the severity of AKI is associated with the possibility to progress to chronic kidney disease (CKD). Around 15% to 20% of patients with AKI advance to end-stage renal disease [41]. It is important to study the role of autophagy affecting the transition from AKI to CKD.

#### **4. Autophagy in Chronic Kidney Disease**

The incidence and prevalence of the CKD are increasing in the past two decades [32,42,43]. The CKD is a multifactorial disease with two major causes of CKD being diabetes and hypertension [43]. Other common causes of CKD include glomerulonephritis, polycystic kidney disease, kidney stones, urinary infections, drugs, and nephrotoxins [44]. The increasing prevalence of CKD has become a great burden to the healthcare system worldwide [32,44].

#### *4.1. Pathophysiology*

Currently, the exact mechanism of CKD is still unknown. The final common pathway of CKD involves glomerulosclerosis, vascular sclerosis, and tubulointerstitial fibrosis. The progression of the CKD involves complex mechanisms, including glomerular hypertension, renin-angiotensin-aldosterone signaling, podocyte homeostasis, dyslipidemia, tubulointerstitial fibrosis, and genetic factors. Recent advances also show that autophagy has an important role in CKD [45–47]. CKD patients have elevated oxidative stress and increased ROS production in mitochondria in addition to altered body homeostasis, protein aggregation, and inflammation. Autophagy is essential in keeping the balance of body homeostasis and protein recycling. Autophagy activation is critical in inflammatory responses. Moreover, oxidative stress and ROS are both important regulators of autophagy. Clinical data also show that patients with CKD have altered autophagy responses [48].

#### *4.2. Our Previous Research*

According to the 2016 guideline for autophagy monitoring, there is no absolute criteria that are applicable in every clinical or experimental context for determining autophagic status. Although some autophagy markers have been used to estimate the autophagic activity in patients, such as increase synthesis or lipidation of LC3 and increase autophagosomes formation [49]. However, it is difficult to measure the exact autophagy flux in clinical settings.

In 2013, we have designed a method to measure the autophagic function in leukocytes from patients. LC3 proteins are involved in phagophore formation and characterized as autophagosome markers. A cytosolic form of LC3 (LC3-I) is conjugated to phosphatidylethanolamine to form LC3-II, which usually reflects the formation of autophagosomes. Previous studies showed that the LC3-I level is very stable during starvation and that the LC3-II level is reflective of changes in the autophagic function and flow. We, therefore, postulated that LC3-I can serve as an ideal control in human leukocytes. The ratio of the 14 kDa LC3-II versus the 16 kDa LC3-I (LC3-II/LC3-I) in leukocytes can serve as an indicator of autophagy flux. The ratio of LC3-II/LC3-I after fasting for 12 h (LC3-II/LC3-I-AC) versus LC3-II/LC3-I 2 h after breakfast (LC3-II/LC3-I-PC) in the same subject can be calculated as γLC3 and regarded as an indicator of autophagy flux or activation. We enrolled 60 patients diagnosed with stages 4–5 CKD (30 with hemodialysis and 30 without hemodialysis), and 30 healthy volunteers as the control group who were sex- and age-matched. In the CKD with hemodialysis group, the blood sample was collected one day after hemodialysis. Using γLC3 as the marker, we have measured the autophagy flux in CKD patients. The isolated LC3-I or LC3-II after fasting or feeding showed no significant associations with healthy subjects and CKD patients with or without hemodialysis. Overnight fasting increased autophagy flux and γLC3 in healthy subjects, which were nearly absent in CKD patients. Moreover, hemodialysis could not correct the autophagy flux deficiency in CKD patients. The *Atg5* and *Beclin-1* transcript levels also increased after starvation in the healthy group, while there were no significant changes in the CKD group. Our data thus provided the direct evidence supporting that CKD patients have impaired autophagy activation and could not be reversed by hemodialysis. The γLC3 was a better autophagic activity indicator then the isolated LC3-I or LC3-II [48].

We also highlighted the relationship between cardiovascular diseases in CKD patients using echocardiography to measure the cardiac functions and structures. The γLC3 was negatively associated with left atrial sizes; changes of the *Atg5* transcript were negatively associated with LVEDD, and; changes of the *Beclin-1* transcript were negatively associated with diastolic mitral inflow E- and A-wave values. The different autophagy markers are associated with different echocardiographic parameters [48]. Because the increased LA size correlates with the increased incidence of atrial fibrillation, stroke, acute myocardial infarction, and congestive heart failure [50], our observation suggested a close relationship of autophagy and CKD-related cardiovascular diseases. The exact mechanism of how CKD has autophagy deficiency is still unclear and will require further studies.

#### *4.3. Diabetic Nephropathy*

Diabetic nephropathy is a major cause of CKD and end-stage renal disease worldwide [51,52]. Olivia Lenoir and colleagues have reported that high glucose concentration environs activated autophagy in podocytes and protected the podocytes from hyperglycemia-related apoptosis [47]. Deficiency of autophagy activation by knockout Atg5 in diabetic mice resulted in more severe proteinuria and impaired renal function [52,53]. Impaired autophagy in the kidney also resulted in podocyte loss and massive proteinuria in diabetic nephropathy [54]. Decreased mTORC1 activation in diabetic mice could stimulate autophagy and decrease glomerulosclerosis, proteinuria, and podocyte loss to slow down progression to diabetic nephropathy [55].

However, there were some studies suggesting the opposite effect of high glucose on podocyte. Through CASP3 activation, high glucose lead to podocyte apoptosis [56]. With the high glucose stimulation, human podocytes exhibited a dramatically reduced LC3-II and Beclin-1 and decreased autophagy activation [57]. The controversies are still unsolved, and more evidence is needed for making the conclusion.

Currently, only a few studies have described the relationship between autophagic markers and podocyte-specific proteins. The podocin protein is a key protein of the slit diaphragm of podocytes. In patients with diabetic nephropathy and severe proteinuria, their kidney biopsy samples express podocin with a granular and irregularly scattered pattern under immunofluorescent study, whereas intense accumulation of p62 proteins is presented in glomeruli. In the 50-week-old diabetic nephropathy rats model with massive proteinuria, the podocytes showed a reduction in podocin-positive areas, p62 accumulation, a decrease of LC3-II, and alteration of foot processes. These suggest that the insufficient autophagic function could cause podocytes injury in diabetic nephropathy with severe proteinuria [58].

Autophagy also protects mesangial cells from undergoing apoptosis in diabetic nephropathy after induced by TGF-β1 via TAK1 and PI3K–AKT-dependent pathways [55]. In proximal tubular cells, autophagy activation is reduced by hyperglycemia while p62 is increased in both type 1 and type 2 diabetes animal models [59,60]. Studies indicated that diabetic nephropathy is associated with decreased autophagy activity and increased apoptosis [54,55,61]. These evidence support that autophagy can be a therapeutic target for diabetic nephropathy [60].

#### *4.4. Autoimmune Kidney Disease*

Autophagy may regulate autoimmune responses by modulating innate immunity and lymphocyte homeostasis [62]. Several studies have manifested the relationship between autoimmune diseases and autophagy in both animal models and human studies. Autophagy helps to understand the mechanism of autoimmune diseases and open the possible therapeutic strategies in systemic lupus erythematosus (SLE), Sjögren's syndrome, Crohn's disease, rheumatoid arthritis, multiple sclerosis, and type 1 diabetes mellitus. [63,64]. The Lupus nephritis is the most common of all and manifested as severe complications of the SLE. Severe lupus nephritis can lead to end-stage renal disease and is an important predictor of mortality in SLE patients [65]. The lupus nephritis results from complement activation, autoantibody formation, immune complexes formation, and dysfunctional adaptive immune responses. The autophagy interacts with these processes and preserves renal function [66]. Some SLE patients have activated autophagic genes, such as *Atg5* and *Atg7*. They also have increased autophagic vacuoles in B cells, T cells, and macrophage in peripheral blood mononuclear cells [64,67]. Current evidence suggest that lupus nephritis may be associated with renal Epstein–Barr virus infection, which can induce autophagy in B cells in a dose-dependent manner [68,69]. In the mice lupus nephritis model, podocytes exhibit autophagy activation that protects renal function from deterioration. The mice podocytes from lupus nephritis have increased autophagosomes, increased LC3-II/LC3-I ratios, and decreased p62 [70]. The aggregated lupus autoantibodies can assist injured podocytes being degraded by autophagy activation [71]. Several drugs given to SLE patients are mTOR inhibitors which can

induce the autophagy activity, suggesting autoimmune diseases are related to autophagy dysfunction and therefore give rise to future possible therapeutic intervention options [64].

#### *4.5. Infection*

Severe systemic infection and sepsis induce a cytokine storm and influence multiple tissues and organs including the kidney. Autophagy is up-regulated early after sepsis and protects organs from pathogen by modulating the immune systems and regulating macrophage, dendritic cells, B cells, CD4+, and CD8+ T cells functions. [63,72]. Autophagy has been suggested to protect kidneys against septic kidney injury [41,66,73]. In the cecal ligation and puncture mice model, the sepsis induced by the peritoneal infections activates the autophagy. The activated autophagy protects the kidney function by decreasing circulating cytokines and endothelial activation [41]. In a sepsis rat model, the decline of autophagy response is associated with the development of kidney injury. Knockdown of *Atg7* decreases tumor necrosis factor α-related proximal tubular cell death, which can be reversed with rapamycin [73]. Autophagy harbors the capacity to both pro- and anti-inflammatory responses to suppress sepsis-induced kidney injury through regulation of infection and through targeting inflammasome and type I interferon responses [66]. These observations suggest that autophagy may be a therapeutic target to protect kidney injury from sepsis.

#### *4.6. Renal Tubulointerstitial Diseases and Ureter Obstruction*

On 2010, Li and colleagues demonstrated that autophagy was significantly activated in a unilateral ureteral obstruction mice model. The conversion of LC3-I to LC3-II, activation of Beclin-1, and accumulation of autophagosomes with massive autophagic vesicles were observed in atrophic tubules with tubulointerstitial injury [74]. Mice with LC3B knockout exhibited a deficit in autophagy activation and severe tubulointerstitial fibrosis after ureter obstruction [75]. The development of tubular atrophy and nephron loss correlated with autophagy in a time-dependent manner [13]. However, persistent activation of autophagy in kidney tubular cells would promote renal interstitial fibrosis through fibroblast growth factor 2 [76]. These studies suggested that the balance of autophagy activation is important in regulating the tubulointerstitial function and renal fibrosis in obstructive kidney disease [27].

#### *4.7. Toxic Effects of Drugs*

Nephrotoxicity of many therapeutic medications can cause acute kidney injuries and worsening renal function in patients with CKD. The non-steroidal anti-inflammatory drugs, iodinated contrast medium, and cisplatin are important drugs causing nephrotoxicity related acute kidney injuries. Recently, in vitro studies have shown that iodinated contrast leads to enhancement of mitophagy, and may protect kidneys from iodinated contrast related renal tubular epithelial injury. The mitophagy is one type of the autophagy that can selectively remove the damaged mitochondria [77]. Cisplatin-induced AKI involves multiple mechanisms, including proximal tubular injury, oxidative stress, inflammation, and vascular injury. The injury is predominantly acute tubular necrosis in the proximal tubules [78–80]. Previous studies suggest that the autophagic responses to cisplatin treatment may protect many types of cancer cells and result in cisplatin resistance [78]. Cisplatin also activated autophagy in the proximal tubules with massive autophagosome formation and LC3-II accumulation for protection purpose [79,81]. Also, in proximal tubule-specific Atg5-knockout mice, cisplatin prompted more severe DNA damage and p53 activation, as well as accumulated more protein aggregates in proximal tubules [79,82]. Furthermore, rapamycin, an mTOR inhibitor, could treat the cisplatin-induced AKI in mice to improve renal function [78,79]. Oral anti-diabetic agents, metformin, also activated the autophagy and protected against cisplatin-induced tubular injury by activating autophagy cascades and slowing down the apoptosis of tubular cells [83]. These data indicated that autophagy can protect renal tubule injury against cisplatin [29].

The Adriamycin- and Puromycin aminonucleoside-induced podocyte apoptosis is widely used for studying the pathophysiology of glomerular diseases in vitro and in vivo. The activation of autophagy with rapamycin could suppress the Adriamycin-induced apoptosis while inhibiting autophagy with chloroquine enhanced apoptosis. The podocyte-specific Atg7 knockout mouse model described the aggravated podocyte injury, glomerulopathy, and proteinuria after adriamycin treatment [84]. The upregulation of LC3-positive autophagosomes protects puromycin aminonucleoside-induced nephrosis in rats in vivo and immortalized mouse podocytes in vitro [85]. Puromycin aminonucleoside reduces the autophagy in human podocytes with the activation of mTORC1. When inhibiting autophagy with the 3-methyladenine or chloroquine, the podocyte apoptosis increased significantly along with the elevation of active caspase-3 [86]. The rapamycin activated autophagy led to decreased proteinuria and less severe foot-process effacement [87] (Figure 3). Collectively, this evidence supports autophagy may be an early adaptive cytoprotective mechanism for podocytes under Adriamycin and puromycin aminonucleoside-induced apoptosis intervention.

Although evidence has explicated the relationship between toxin/drug and the autophagy in the kidney, future studies are still necessary to investigate the mechanism of autophagy in many other nephrotoxic agents.

**Figure 3.** Overview of the different medications that regulate autophagy in different steps, including Rapamycin, Adriamycin, Puromycin, 3-methyladenine and Chloroquine.

#### *4.8. Cystic Disease (Polycystic Kidney Disease)*

Autosomal dominant polycystic kidney disease (ADPKD) is one of the most prevalent inherited renal cyst diseases and frequently leads to end-stage renal disease. The main cause of this disease is gene mutation. Approximately 85% of the mutations in ADPKD occur in the *pkd1* gene encoding for polycystin-1, and 10–15% in its interaction partner *pkd2*, encoding for polycystin-2. These ciliary proteins are involved in cellular repair and growth mechanisms [88].

Both in vitro and in vivo studies have shown that multiple autophagic molecular parameters and signaling pathways are involved in ADPKD, including mTOR, cyclic adenosine monophosphate, and several growth factors [89]. The current evidence suggests that autophagy processes may relate to the cystic formation and size growth by activating the mTOR signaling pathway [89,90]. ADPKD is a cilia-related disease and autophagy activation is essential in ciliogenesis [12]. Studies revealed that autophagy regulates cilia length by modulating protein synthesis and degradation. Through modulating autophagy activation, the primary cilium controls epithelial cell volume in a fluid flow dependent manner [91]. The ADPKD exhibiting autophagosome increases with LC3-II and Beclin-1 overexpression in tubular cyst-lining cells, suggesting autophagic flux dysregulation [1]. In a mice model with mutated PKD1 and patients with ADPKD, autophagy activation is impaired [92]. Polycystin-1 negatively regulates polycystin-2 via the autophagosomes-dependent pathway. Failure of pathogenic polycystin-1 mutants to induce this function may lead to ADPKD [93]. All this evidence suggests that the ADPKD may be present with dysfunctional autophagy [1,94]. Currently, the regulation of autophagy in ADPKD is incompletely understood. Autophagy may be one of the therapeutic targets in the future.

#### *4.9. Autophagy in Aging-Associated CKD*

Aging is associated with a deterioration and imbalance of the general homeostasis ability, which results in loss of the compensation mechanism in the kidneys. In the elderly population, the prevalence of non-dialysis CKD is markedly higher. The normal aging kidneys exhibit decreases in renal mass, increases in parenchymal tissue fibrosis and fat deposition, and accumulation of glomerular sclerosis [95,96]. Development of CKD is a major risk factor of ESRD in the elderly population. The mortality among the age of 75 with CKD is double compared to the healthy population [97,98].

The podocyte-specific *Atg5* knockout mice develop glomerulopathy gradually during aging. The podocytes without *Atg5* have decreased organelles turnover with the accumulation of ubiquitinated and oxidative protein. The deficiency of autophagy and proteasome pathways leads to proteinuria, loss of podocytes, and development of glomerulosclerosis in aging mice [22]. In aging mice, podocyte-specific autophagy-deficient mice have mild forms of glomerulosclerosis compared with tubular cell-specific autophagy-deficient mice [66]. Yamamoto and colleagues demonstrated that proximal tubule-specific deletion of Atg5 in mice resulted in significant deteriorations of kidney function and fibrosis at 24 months of age [99]. These results indicate that tubular cells play an important role in aging-related autophagy regulation of kidney function. The inability of older mice to recover from AKI has been attributed to an age-dependent loss of autophagy resulting in CKD [41]. These results suggest that autophagy is vital for maintaining kidney tissue homeostasis and aging-associated injury [3].

Caloric restriction has been shown to strongly induce autophagy in kidneys and slow down the process of interstitial fibrosis and tubular atrophy [100]. Moreover, caloric restriction prolongs lifespan in animals and decreases the age-related mitochondrial oxidative damage and the kidney tissue injury by enhancing autophagy [101,102]. Caloric restriction in AKI with rats of different ages suggests that the caloric restriction can significantly increase autophagy activation by increasing LC3-I/LC3-II ratio and improving renal function. The nephroprotection effects gradually decline with the age due to deteriorations of the autophagic system [103]. The mTOR pathway acts as a two-way regulator during starvation and caloric restriction in rat. At first, the mTOR inhibition activates autophagy to form autophagolysosome and breakdown the cellular content for regeneration to further protect the kidney function. However, prolonged starvation will reactivate the mTOR pathway and inhibit autophagy by negative feedback and accumulation of autophagolysosome degradation content. [104]. Therefore, a balanced caloric restriction strategy taking into consideration of the autophagy activation is one of the therapeutic options to delay the progression of CKD among elderly patients [100].

#### **5. Autophagy in Dialysis and Renal Transplantation**

The end-stage renal disease is the most severe outcome of CKD and can only be treated with dialysis or renal transplantation. Patients with end-stage renal disease have increased risks of cardiovascular disease, cerebrovascular accident, infection, and cognitive impairment [44].

Until recently, very few studies have investigated the relationship between dialysis and autophagy. Our previous study indicated that autophagy flux could not be rescued by hemodialysis in patients with end-stage renal disease, suggesting that hemodialysis has no role of modulating autophagy [48]. When it comes to peritoneal dialysis (PD), one study suggests that autophagy stimulates the fibrosis and apoptosis in peritoneal mesothelium cells due to long-term exposure to the high-glucose peritoneal dialysis solution. The activation of Beclin 1-dependent autophagy results in decreases of the viability of peritoneal mesothelium cells and finally PD failure [105]. Another study, however, suggests that

the PD solution promotes autophagosome formation and decreases cells death to maintain the PD function [106]. Therefore, future studies are required to understand the exact relationship of dialysis and autophagy regulation.

Renal transplantation is considered as the definitive treatment to ESRD. Currently, some evidence supports the connections between autophagy and renal transplantation. The mTOR inhibitor is frequently used in renal transplantation as an immunosuppressant, including rapamycin, sirolimus, and everolimus. Rapamycin inhibits T cell proliferation and activates autophagy to maintain the renal homeostasis [107,108]. Cyclosporine is another immunosuppressive drug commonly used in renal transplantation patients [109]. Cyclosporine induces autophagy in primary cultured renal tubular cells. After treatment of cyclosporine in mice, the kidney proximal tubule cells exhibit an increase in numbers and sizes of autophagosomes and autophagy flow [110]. Deficiency in autophagy function will lead to an exacerbation of cyclosporine-related endoplasmic reticulum stress and renal injury [110,111]. Another indirect clue is that a kidney from elderly donors is now considered to have a higher risk of post-transplant ischemic injury, which may be due to the decline of autophagy activation with aging [107,112].

#### **6. Therapeutic Consideration and Conclusion**

Autophagy defects can occur at different stages of the pathway in CKD, and this may influence treatment strategies. Early studies examine the rapamycin through upregulating autophagy to enable the clearance of intracytoplasmic aggregation-prone proteins. Rapamycin is currently the only drug that could be used to target autophagy in CKD treatments. However, the long-term side effects of rapamycin should be taken into consideration [7]. Some studies have used the rapamycin under intermittent dosing protocols in mice that could create pulsatile upregulation of autophagosome formations. This gives us a possible therapeutic regimen to reduce rapamycin side effects in patients.

#### *6.1. Future Therapeutic Considerations*

Several candidate drugs targeting autophagy have been shown to inhibit or activate the autophagy function. However, many of them are still under animal trials or early phase clinical trials and are not ready to be used clinically. Some drugs that are used clinically for other diseases also are found to have the potential to regulate autophagy through different autophagic pathways. The psychotropic drug lithium, carbamazepine, and valproic acid can activate autophagy through the phosphatidylinositol signaling pathway. The clonidine and rilmenidine act as the imidazoline receptor agonists to enhance autophagy. Verapamil targets L-type Ca2+ channels to modulate autophagic flux. Metformin upregulates AMPK signaling and induces autophagy function. In contrary, chloroquine is a lysosomotropic agent that inhibits autophagosome-lysosome fusion and lead to autophagy downregulation [113]. Ultimately, whether autophagy represents a useful target in CKD prevention or treatment will need to be addressed by conducting clinical trials in patients in the future.

#### *6.2. Conclusions*

We summarize the autophagy pathophysiology in chronic kidney disease (Figure 4). The exposure of normal kidneys to selective stresses, such as ischemic-reperfusion injury, toxin, and sepsis, will result in acute kidney injury, ROS accumulation, and autophagy activation. The protective mechanisms of autophagy take place in podocytes, mesangial cells, and tubular cells, help repair and regenerate the damaged kidneys. After several episodes of stresses, the balancing by the autophagic repair mechanism cannot keep up and CKD ensues. Different CKDs also modulate autophagy in diverse pathways and help to slow down the progression to ESRD as in Table 1.

**Figure 4.** Diagram depicting the roles of autophagy in various chronic kidney disease. DN: diabetic nephropathy; LN: lupus nephropathy; APCKD: adult polycystic kidney disease.

**Table 1.** Overview of the autophagy in chronic kidney diseases (CKD).

**Table 1.** *Cont.*

At the turning point of autophagy research in the early 1990s with the identification of the autophagy-related gene, many researchers have devoted their efforts to finding the potential therapeutic use of autophagy in human diseases. The relationships between autophagy and kidney diseases are well-founded, but there is still no clinically useable agent targeting the autophagic pathway for patients with CKD or AKI. The complex nature of CKD or AKI further complicates the development of therapeutic advancement. The evidence from mice or cells have provided us with comprehensive mechanistic insights into the etiology and pathophysiology of the disease. Since autophagic machinery has a vital role in controlling immunity, it will be crucial for clinicians to monitor the occurrence of infection and autoimmune diseases with autophagy inducers or inhibitors [104]. Besides, inhibition or activation of the autophagy will impact multiple aspects of organism physiology with various effects [114–116]. Therefore, it is necessary to take these diverse interactions into consideration when developing autophagy-targeting drugs. In the end, medications developed by tackling these

mechanistic pathways still require prospective and randomized trials to verify their therapeutic potentials and adverse events when administered in humans.

**Author Contributions:** T.-A.L. and C.-Y.W. construct the manuscript structure and wrote the manuscript. V.C.-C.W. proofread and polished the manuscript.

**Funding:** C.Y.W. received support from the National Health Research Institute (NHRI-EX106-10617SI), National Science Council (105-2628-B-182-009-MY4), and Chang Gung Memorial Hospital (CMRPG3H0131, CMRPG3H0271, CMRPG3I0321, and CMRPG3H0841).

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

## *Review* **Autophagy in Neurotrauma: Good, Bad, or Dysregulated**

**Junfang Wu 1,2,3,\* and Marta M. Lipinski 1,2**

Received: 31 May 2019; Accepted: 9 July 2019; Published: 10 July 2019

**Abstract:** Autophagy is a physiological process that helps maintain a balance between the manufacture of cellular components and breakdown of damaged organelles and other toxic cellular constituents. Changes in autophagic markers are readily detectable in the spinal cord and brain following neurotrauma, including traumatic spinal cord and brain injury (SCI/TBI). However, the role of autophagy in neurotrauma remains less clear. Whether autophagy is good or bad is under debate, with strong support for both a beneficial and detrimental role for autophagy in experimental models of neurotrauma. Emerging data suggest that autophagic flux, a measure of autophagic degradation activity, is impaired in injured central nervous systems (CNS), and interventions that stimulate autophagic flux may provide neuroprotection in SCI/TBI models. Recent data demonstrating that neurotrauma can cause lysosomal membrane damage resulting in pathological autophagosome accumulation in the spinal cord and brain further supports the idea that the impairment of the autophagy–lysosome pathway may be a part of secondary injury processes of SCI/TBI. Here, we review experimental work on the complex and varied responses of autophagy in terms of both the beneficial and detrimental effects in SCI and TBI models. We also discuss the existing and developing therapeutic options aimed at reducing the disruption of autophagy to protect the CNS after injuries.

**Keywords:** spinal cord injury; traumatic brain injury; autophagy; autophagic flux; neuronal cell death; lysosomal damage

#### **1. Introduction**

The role of autophagy after the central nervous system (CNS) insults is under perusal, as investigation has begun to determine how autophagy and related pathways contribute to secondary injury and functional recovery following neurotrauma [1–5], including spinal cord injury (SCI) and traumatic brain injury (TBI). Whether detected autophagic responses to injury are protective or detrimental remains controversial, with experimental reports revealing protective effects of both enhancing and hindering autophagy [1,6].

Autophagy is a vital intracellular degradation pathway that delivers cytoplasmic constituents to the lysosomes for degradation [7]. There are three different types of autophagy that have been descripted so far, including macroautophagy, microautophagy, and chaperone-mediated autophagy [8–10]. In addition, several types of targeted autophagy have been described, including mitophagy, which is a selective form of macroautophagy that specifically targets and degrades damaged mitochondria. Macroautophagy involves the formation of a cytoplasmic membrane that engulfs cargo and eventually elongates to form double-membrane vesicles termed autophagosomes. It culminates in the fusion of cargo-containing autophagosomes to the lysosomes and the degradation of the cargo by lysosomal

hydrolases [11]. As macroautophagy is the best characterized form of autophagy, hereafter, the term autophagy refers to macroautophagy unless otherwise specified. Under normal conditions, autophagy is an essential physiological process that maintains a balance between the manufacture of cellular components and breakdown of damaged organelles and other toxic cellular constituents. Its processing is strictly regulated under physiological conditions [12]. Recent considerable progress has demonstrated that autophagy is dysregulated in injured CNS following trauma, and may play either beneficial or detrimental functions after injury, depending on the context and the mechanisms leading to its perturbation [1,6,13,14]. Emerging data also suggest that the restoration and/or augmentation of proper autophagy function, for example by inducing lysosomal biogenesis, may be a potential therapeutic target for TBI and SCI.

This review explores the current research on the function and mechanisms of autophagy in two models of CNS injury: SCI and TBI. Cell-type specific responses of autophagy are discussed according to different locations, severity, and time windows of traumatic injury. We discuss recent findings suggesting that impairment of the autophagy–lysosomal pathway may be part of the secondary injury processes of SCI/TBI. We also review recent studies and novel mechanistic discoveries on cytosolic phospholipase A2 (cPLA2) participation in lysosomal damage, and provide therapeutic options, with an emphasis on the pharmacological modulation of autophagy and lysosomal biogenesis for neuroprotection and the prevention of neuroinflammation after CNS trauma.

#### **2. Traumatic Spinal Cord and Brain Injury and Their Injury Mechanisms**

According to the National Spinal Cord Injury Statistical Center [NSCISC, 2018], in the United States (U.S.), there are approximately 288,000 people living with SCI, and 17,000 new SCI cases occur each year. The impact of these injuries is devastating for individuals, and the health care costs associated with the injury are some of the highest in the U.S. [15,16]. Experimental and clinical studies [17–21] have indicated that acute SCI is a two-step process involving primary and secondary mechanisms. Primary injury of the spinal cord refers to the initial mechanical tissue damage. It also includes ruptured microvessels in the spinal cord, and subsequent hemorrhage and edema that are associated with secondary injury. SCI-mediated secondary injury processes include a cascade of biochemical and cellular events [22], such as free-radical formation, excitatory amino acid release, axonal damage, neuronal and oligodendroglial cell death, the infiltration of macrophages and peripheral immune cells, microglia, and astrocyte activation/glial scar formation. These delayed secondary injury processes can occur over hours, days, and months after initial impact [23,24], contributing to progressive neuronal degeneration and neurological dysfunction. Despite considerable research over the past 30 years, there is still no established effective treatment to improve recovery following SCI. In part, this reflects an incomplete understanding of the complex secondary pathobiological mechanisms involved.

TBI represents a major public health problem, with more than 1.7 million new cases annually in the United States [25] and accounting for 60% of all trauma deaths in the U.S. [26]. Similar to SCI, TBI causes cell loss and neurological functional deficits through both the direct physical damage of tissue (primary injury), and through subsequent biochemical changes (secondary injury). The latter reflects delayed and potentially reversible molecular and cellular pathophysiological mechanisms [27,28], which begin within seconds to minutes after the primary insult, and may continue for months to years [29,30]. Such secondary injury processes, similar in SCI, lead to cellular changes that include neuronal and oligodendroglial cell death, as well as microglia and astrocyte activation and glial scar formation. Although most of the research to date has been directed at early cellular and molecular events, both experimental and clinical evidence suggests that CNS trauma-mediated pathophysiological changes may continue for years [30–33], leading to chronic post-mitotic cell loss and microglial/astrocytes activation, and contributing to chronic functional deficits.

Cell death after neurotrauma is a major cause of neurological deficits and mortality. Although CNS injury induces changes in multiple cell types such as neurons and oligodendrocytes, the mechanisms of neuronal cell death have been the predominant focus. There are multiple cell death mechanisms in injured CNS following trauma [34,35]. Three main morphological types of cell death include apoptosis, autophagic cell death, and necrosis. Molecular and biochemical pathways that are involved in cell death after both TBI and SCI include, among others, caspases-dependent apoptosis (the Bcl-2 family), cell cycle activation-dependent pathways, the autophagy-related (ATG) protein family-mediated cell death (autophagy), caspase-independent programmed cell death [including extracellular-signal-regulated kinase 2 (ERK2), poly[Adenosine diphosphate (ADP)-ribose] polymerase (PARP)-1, apoptosis-inducing factor (AIF)], calpains and cathepsis-mediated calcium-dependent cell death, and the c-Jun N-terminal kinase-mediated non-apoptotic cell death (oncosis) [34,36,37]. Interestingly, apoptosis and autophagy share many regulatory factors, including the Bcl-2 family proteins [38]. The pro-apoptotic B-cell lymphoma 2 (Bcl-2) proteins Bim, Bid, and Bax, are known mediators of lysosomal membrane permeabilization (LMP) [39–42], while the anti-apoptotic Bcl-1 and Mcl-1 negatively regulate autophagy through interaction with Beclin 1. Additionally, necroptosis, the regulated receptor interacting protein kinase (RIPK)-dependent necrosis pathway, may also contribute to secondary neuronal cell death after CNS insults [3,43,44]. Obstacles to successful therapy against neurotrauma-induced neuronal cell death include the diversity of cell death pathways, which have both overlapping and distinct molecular mechanisms, and the narrow therapeutic windows for some types of neuronal cell death [34,45]. Thus, we emphasize that the effective neuroprotective strategies will need to concurrently modulate multiple signaling pathways to reflect the spatial and temporal changes underlying the diversity of neuronal cell death.

Stimulation by pro-inflammatory cytokines, chemokines, or alterations in the CNS environment induces microglial activation and the attraction of macrophages to the injury site [46,47]. Activated microglia undergo a transition from a resting, ramified phenotype to a phagocytosis-capable, 'macrophage-like' phenotype that is virtually indistinguishable from blood-borne macrophages [48–50]. Microglia and macrophages are found in the injured CNS within 12 to 24 h post-injury, with maximal concentration at four to eight days post-injury [49,51]. These cells produce free radicals, nitric oxide, and arachidonic acid derivatives, as well as a number of cytokines and chemokines in an effort to remove debris and dysfunctional cells. However, excessive inflammation may also result in the death of neighboring undamaged cells [52–56]. It is reported [57] that the activation of microglia-associated inflammatory factors may continue indefinitely after CNS trauma: certainly, it lasts for many months in rodents.

#### **3. Autophagy, Autophagy Flux, and the Lysosomal Functions in Neurotrauma**

#### *3.1. Autophagy and Autophagy Flux*

The process of autophagy includes several essential steps, including the induction, sequestration of cargo within the autophagosomal membrane, maturation of the autophagosomes, and degradation [12,58]. While baseline autophagy proceeds at all times, in adverse conditions such as environmental stress, nutrient starvation, or acute cellular injury, it is further activated. Cytoplasmic components including damaged organelles and toxic protein aggregates are sequestered by a unique membrane called the phagophore or isolation membrane. Complete sequestration by the elongating phagophore results in the formation of the autophagosome, which is typically a double-membraned vesicle. Then, autophagosomes are transported within the cell and fuse with lysosomes. Then, the inner membrane of the autophagosome and the cytoplasm-derived materials contained in the autophagosome are degraded by lysosomal hydrolases. The whole process of autophagy is referred to autophagic flux [7], which represents the dynamic process of autophagy from cargo sequestration to its degradation.

Completion of the autophagic flux requires the coordinated activity of various members of the autophagy-related (ATG) protein family [59,60]. For example, ATG13, ATG101, and FIP200 (homolog of yeast ATG17) are part of the ULK1 kinase complex that initiates autophagy. ATG9 also mediates an essential function in early autophagosome formation. Expansion of the autophagosomal membrane depends on two ubiquitin-like conjugating systems: the ATG12–ATG5–ATG16L system and the phosphatidylethanolamine (PE)–LC3 system. LC3 is the commonly used name for microtubule-associated protein 1 light chain 3β (MAP1LC3B). Upon activation of the autophagic pathway, the cytosolic, proteolytically processed form of LC3 (termed LC3-I) is lipidated to form LC3–PE (termed LC3-II), which is specifically recruited to the phagophore membrane. LC3-II accumulates on autophagosomal membranes and—through interaction with adaptor proteins—helps recruit substrates that are tagged for autophagic degradation. Thus, LC-II levels correlate well with autophagosome numbers. In addition, rapamycin complex 1 (MTORC1) operates as a central suppressor of autophagy. Conversely, beclin 1 (BECN1), a regulatory subunit of the type III phosphatidylinositol (PI) 3 kinase, promotes autophagy by promoting the formation of phosphatidylinositol 3 phosphate, which is necessary for autophagosome formation [60]. Moreover, ubiquitylated structures, including protein aggregated and in some cases mitochondria, are bound by the autophagic adaptor sequestosome 1 (SQSTM1; also known as p62), which enables their uptake by autophagosomes through LC3-II [61]. p62/SQSTM1 is one of several factors that target specific cargoes for autophagy, also including NBR1 [neighbor of breast cancer type 1 (BRCA1) gene], NDP52 (nuclear domain 10 protein 52), and OPTN (optineurin).

A number of methods are currently utilized to assess autophagic flux [62]. Transmission electron microscopy can be used for the partial analysis of the autophagosome pool size in vitro and in vivo without a time dimension. One of the most common ways to monitor autophagy is by measuring LC3-II protein turnover, which is incorporated into autophagosomes and then degraded in the lysosome. There are many different ways to measure LC3-II protein levels, including Western blot and immunofluorescent microscopy. However, these methods are complicated by the LC3-II levels increasing with autophagy induction due to increased autophagosomal formation, but also decreasing as these autophagosomes are turned over. Thus, a more reliable way to accurately monitor LC3-II levels is with a flux assay that uncouples autophagosome formation from its degradation by using lysosomal inhibitors such as bafilomycin-A1 (V-ATPase inhibitor) or chloroquine (a lysosomotropic compound that neutralizes lysosomal pH). As the flux is a rate, an analysis of both LC3-II and p62 levels over time in the presence and absence of lysosomal inhibitor is required. Another possibility in vitro is monitoring the half-life of labeled long-lived proteins that are known to be degraded via autophagy [63]. Fluorescence microscopy is an extremely valuable tool for the assessment of autophagic flux and the autophagosome pool size per cell. Use of the green fluorescent protein (GFP)–LC3 reporter allowing direct visualization of autophagosomes has been especially valuable. It is widely used for the evaluation of autophagy flux in vitro via image-based flux assay and live cell microscopy, and of the autophagosome pool in vivo using transgenic *GFP-LC3* reporter mice. Recent developments of the dual RFP-GFP-LC3 reporter further improved the evaluation of autophagy flux. It takes advantage of differential pH sensitivity of GFP (acid labile) and RFP (acid stable) to directly assess the fusion of autophagosomes (red and green) with lysosomes to form autolysosomes (red only) [62]. However, it remains challenging to employ these techniques in vivo in a manner that uniformly expresses autophagic flux quantitatively and assesses the magnitude of change in flux.

#### *3.2. Impairment of Autophagy Flux in SCI and TBI*

Increased markers of autophagy have been observed in different experimental SCI models [64–68]. Several reports indicate that the accumulation of autophagosomes is initiated very early during secondary injury, in some cases within hours after the initial impact. In SCI, depending on the injury models and severities as well as different species and sex, some reported significant increases in expression levels of the autophagosome marker LC3-II during the first 24 h after SCI [3,4,67,69,70], whereas others have only detected a delayed increase of this marker three to seven days after injury [64,65,68,71–73]. In some cases, the expression of Beclin 1, a key protein in the initiation of autophagy [74], has been reported to increase after SCI, supporting the hypothesis that SCI may lead to an increase of autophagy initiation [64,65,73,75,76]. However, other works have reported unaltered levels of Beclin 1 and other upstream mediators and regulators of autophagy [4,68], suggesting that

increased autophagy initiation is unlikely to account for an increase in the number of autophagosomes. The p62/SQSTM1 protein, similar to all autophagy cargo adaptors, is degraded by autophagy along with its cargo. Therefore, provided no changes in transcription/synthesis are observed, p62 levels can be often used to approximate the autophagy-dependent degradation rate, with low levels of p62 protein associated with high autophagy flux (high degradation) and high p62 levels associated with low autophagy flux (inhibited degradation). Most reports describe the accumulation of both LC3 and p62 in the damaged spinal cord tissues [3,4,68]. Since p62/*Sqstm1* mRNA expression is not changed after SCI, this suggests that an accumulation of autophagosomes after SCI is likely a result of inhibited autophagy flux rather than its increased initiation [1]. This includes diverse injury models such as acute contusion SCI in male rat [4] and male mouse [3] and chronic spinal cord compression in both sexes [66,77]. The inhibition of autophagy flux was also recently reported following moderate contusion SCI in *GFP–LC3* male mice [3], and demonstrated the accumulation of both GFP–LC3 positive autophagosomes and p62 in the same cells near the injury site. Although autophagic changes in response to SCI show some discrepancies observed among studies, species, injury models, and injury severities, the vast majority of data support the inhibition of autophagic flux in the damaged spinal cord.

Increased markers of autophagy, including LC3-II, beclin1, p62, and autophagosomes, have been reported in human TBI autopsy samples [78,79], as well as in cerebrospinal fluid (CSF) [80]. TBI-mediated accumulation of p62 in both brain autopsy samples [78] and CSF [80] is associated with injury severity and worse recovery [80], and likely reflects an impairment of autophagic flux. Similarly, in experimental models of TBI, LC3-II expression levels as well as the accumulation of autophagosomes observed by electron microscope [81] has been demonstrated in the injured brain [79,82–84]. However, autophagy flux has not been directly assessed until more recently [62]. Interestingly, these data indicate that unlike in the spinal cord, in the brain, the autophagic flux capacity may be dependent on injury severity [14]. Supporting this possibility, the accumulation of p62 appears most pronounced at or near the lesion area after moderate to severe injury in both rodent TBI models [5,85] and human TBI brain samples [78]. The initiation of autophagy is not changed after moderate/severe injury in a controlled cortical impact (CCI) model [85]. Recently, an inhibition of autophagy flux was also reported in *GFP–LC3* mice after moderate CCI [5], and further confirmed by ex vivo flux assay in organotypic brain slices in the presence or absence of a lysosomal inhibitor (chloroquine). Conversely, in a mild CCI model [85], p62 was reduced in the ipsilateral cortex accompanied by increased beclin 1, ATG5, and ATG12, suggesting the enhancement of autophagy initiation. Increased autophagy has also been reported in rat models of TBI following a fluid percussion injury [83] and CCI [86,87]. Bayir et al. [88] reported that increased LC3-II after moderate CCI injury on postnatal day 17 rats is more prominent in male versus female rats, suggesting that trauma-induced autophagy is not limited to the mature mammalian brain, and that similar to nutrient deprivation studies in vitro [89], there are sex-dependent differences in the autophagic response. The divergent findings in autophagy flux following TBI may be explained by the different lesion paradigms and tissue sampling or time windows after injury. Nonetheless, autophagy flux is impaired in most models of moderate to severe brain injury.

#### *3.3. Lysosomal Functions In Neurotrauma*

Lysosomes are membrane-enclosed organelles that contain an array of enzymes that are capable of breaking down all types of biological polymers: proteins, nucleic acids, carbohydrates, and lipids [90–93]. Lysosomes function as the digestive system of the cell, serving both to degrade material taken up from outside the cell and the obsolete or superfluous components of the cell itself. Autophagic degradation is dependent on lysosomal proteases. Defects in the lysosomal function have been demonstrated in various neurodegenerative diseases and aging. In many of these disease paradigms, lysosomal dysfunction can be caused by the increase in lysosomal membrane permeability (LMP) [94–99]. We and others [3–5,100] demonstrated that decreased protein levels and activity of lysosomal enzymes accompanied the inhibition of autophagy flux in both TBI and SCI. The altered

intracellular localization of soluble lysosomal enzymes including cathepsin D (CTSD) diffuse rather than discrete punctate, which further suggests the possibility that LMP allows the leakage of CTSD into cytosol, resulting in decreased lysosomal activity and the inhibition of autophagy flux after neurotrauma. In addition to its detrimental effect on autophagy, the leakage of the highly degradative lysosomal enzymes into the cytosol could lead to significant damage to other cellular components.

Similar to most cellular organelles, lysosomes are surrounded by a single-layer phospholipid membrane. Therefore, they are vulnerable to the activation of phospholipases. There are three major phospholipases present in the CNS: namely, calcium-dependent secretory phospholipase A2 (sPLA2), calcium-dependent cytosolic phospholipase A2 (cPLA2), and calcium-independent phospholipase A2 (iPLA2) [101,102]. Among these, cPLA2 is considered to be the most important PLA2 isoform, because it exhibits a strong preference for the deacylation of arachidonic acid (AA) over other fatty acids, and has been implicated as an effector in the receptor-mediated release of AA [103,104]. Both the expression levels and activity of cPLA2 are increased in several neurodegenerative diseases [101,105–108] as well as following SCI [109,110] and TBI [111]. cPLA2 cleaves the fatty acyl linkage of glycerophospholipids at the sn-2 position, releasing AA and leaving lysophospholipids remaining in the membrane [101,102,112–115]. Then, AA is oxygenated and further transformed into a variety of products such as prostaglandins, leukotrienes, and thromboxanes, which mediate or modulate inflammatory reactions. The accumulation of lysophospholids can affect the membrane properties, including its fluidity and permeability [112,113,115]. Although the signaling function of cPLA2 metabolites has been widely studied, the consequences of its action on the properties of the affected membranes and organelles are less well understood.

We recently used mass spectrometry (MS)-based in vivo lysosomal lipidomics to demonstrate an increase in several classes of lysosophospholipids, which are the products of phospholipases A (PLAs), as well as the accumulation of PLA activators and ceramides in lysosomes purified from the injured brain [111] and spinal cord [110]. Further in vitro and in vivo data indicated that cPLA2-mediated LMP leads to release of lysosomal enzymes into the cytosol, the inhibition of autophagy flux, and neuronal cell death. Taken together, our data implicate cPLA2 in the mediation of the lysosomal defects observed in the pathophysiology of neurotrauma. cPLA2-mediated lysosomal damage in turn causes the inhibition of autophagy flux and autophagosome accumulation, which are associated with neuronal cell death (Figure 1). Interestingly, our in vitro results indicate that the inhibition of cPLA2 can also limit amyloid-β-induced LMP and the inhibition of autophagy [111], suggesting that similar mechanisms may also contribute to Alzheimer's and potentially other neurodegenerative diseases.

**Figure 1.** Spinal cord injury (SCI) and traumatic brain injury (TBI) activate cytosolic phospholipase A2 (cPLA2), mediating increased lysosomal membrane permeability (LMP), and leading to lysosomal damage and the inhibition of autophagy after SCI and TBI.

#### **4. Beneficial or Detrimental E**ff**ects of Autophagy in CNS Cells After Trauma**

#### *4.1. Function of Autophagy in Neurons after Neurotrauma*

Neurons, the fundamental units of the CNS, are post-mitotic cells, and most neurons have a cell body, an axon, and dendrites. CNS trauma causes progressive secondary injury processes including neuronal cell death and axonal damage, contributing to neurological dysfunction. Neurons are also the cell type that are most commonly associated with dysregulated autophagy after both TBI and SCI [1]. Afterwards, TBI markers of autophagy are predominantly reported in the neurons of the ipsilateral cortex and hippocampus, and appear within 24 h after injury [5,79,83]. In a mouse closed head injury model, Diskin et al. [82] reported that 17–37% of Beclin-1+ neurons were also terminal deoxynucleotidyl transferase dUTP (2'-deoxyuridine 5'-triphosphate) nick end labeling (TUNEL) positive, suggesting that cell death was associated with autophagy. In agreement with this observation, we observed significant numbers of GFP-LC3+ and p62+ neurons expressing cleaved caspase-3 in the injured cortex at day (d) 1 and 3 after injury in a mouse controlled cortical impact CCI model [5]. In addition, neuronal GFP-LC3 and p62 also co-localized with markers of caspase-independent cell death (such as AIF), indicating that impaired autophagy flux may contribute to both apoptotic and non-apoptotic neuronal cell death.

In SCI, changes in neuronal autophagy appear to be dependent on both the injury model and neuronal subtype. In rat and mouse contusion SCI models, the accumulation of both LC3 and p62 was greater in the ventral horn motor neurons as compared to the dorsal horn sensory neurons, despite the latter being located closer to the impact site [3,4,68]. This suggests that motor neurons may be relatively more vulnerable to the disruption of autophagy flux. Moreover, in these models, motor neurons with impaired autophagy also expressed higher levels of caspase12 and cleaved caspase 3 [4], supporting a role for impaired autophagy in mediating neuronal apoptosis. However, the inhibition of autophagy flux in the dorsal horn neurons was observed in spinal nerve ligation, which is a model of neuropathic pain [116,117], suggesting that neuronal subclasses may respond differentially to specific types of injury. That autophagy flux was inhibited even in this mild injury model also suggests that overall, spinal cord neurons may be more prone to the inhibition of autophagy flux as compared to brain neurons.

Endoplasmic reticulum (ER) stress induces a variety of neuronal cell death pathways, and has been implicated in the secondary injury processes after CNS trauma [118,119], but its mechanisms remain unclear. The induction of ER stress and activation of caspase 12 following TBI and SCI have been previously reported [4,5,118]. ER stress is a potent inducer of autophagy [120,121], and autophagy can protect cells from ER stress-mediated cell death [122]. Consistently, several studies indicate that markers of inhibited autophagy flux after injury correlate with the exacerbation of ER stress, suggesting a connection between the inhibition of autophagy and induction of ER stress-mediated neuronal cell death after neurotrauma [4,5] (Figure 2).

Necroptosis is a programmed form of necrosis, or inflammatory cell death, mediated by the RIPK1/RIPK3 complex [123]. The inhibition of necroptosis can improve functional recovery after both SCI and TBI, suggesting that it is involved in the mediation of the secondary injury [43,44]. A recent study [3] demonstrated the accumulation of markers of necroptosis specifically in neurons displaying signs of autophagy flux inhibition and lysosomal damage, pointing to a previously unexplored link between the inhibition of the autophagy-lysosomal pathway and the induction of neuronal necroptosis after SCI. Additionally, the inhibition of lysosomal function in vitro resulted in the accumulation of necroptosis mediators RIPK1 and RIPK3. Therefore, the inhibition of lysosomal function and autophagy in CNS trauma could also contribute to neuronal necroptosis (Figure 2).

**Figure 2.** Lysosomal damage after spinal cord injury (SCI) and traumatic brain injury (TBI) cause endoplasmic reticulum (ER) stress and the accumulation of receptor interacting protein kinases (RIPK1/RIPK3), leading to neuronal apoptosis and necroptosis.

Recent findings have identified another type of cell death called 'autosis', which is an autophagy-dependent non-apoptotic form of cell death with unique features. It is characterized by enhanced cell-substrate adhesion, focal ballooning of the perinuclear space, and the dilation and fragmentation of ER. Autosis is mediated by the Na+, K+-ATPase pump in the mitochondria, and it is triggered by autophagy-inducing peptides, starvation, and neonatal cerebral hypoxia–ischemia. Whether or not autophagy-dependent autosis participates in neuronal cell death in injured CNS is unknown. While it's known that autophagy can mediate the execution of cell death in the absence of other death pathways, it is difficult to determine in vivo whether cell death is caused by high autophagy or if it is the by-product of other processes that happen alongside autophagy.

In addition to neuronal cell bodies, autophagosome accumulation has been reported to occur in damaged axons after both TBI and SCI [68,79,124]. While autophagy has been proposed to participate in the process of axonal degeneration in vitro, it is currently not known whether it is involved in axonal damage after neurotrauma. Another possibility is that axonal accumulation of autophagosomes could reflect the inhibition of autophagy flux due to lysosomal defects and cytoskeletal collapse, similar to what is observed in neurodegenerative diseases.

#### *4.2. Role of Autophagy in Oligodendrocyte Survival after Neurotrauma*

Oligodendrocytes (OLs), also called oligodendroglia, are a type of neuroglia whose main functions are to provide support and insulation to axons, and are the myelinating cells of the CNS [125]. Mature OLs are the end product of oligodendrocyte precursor cells (OPCs), which have to undergo a complex and precisely timed program of proliferation, migration, differentiation, and myelination to finally produce the insulating sheath of axons. Due to this complex differentiation program, and due to their unique metabolism and physiology, OLs count among the most vulnerable cells of the CNS. It is known that OLs undergo apoptosis following CNS trauma, thus the loss and demyelination of OLs contribute as major pathological processes to secondary damages after injury [126]. Increased numbers of autophagosomes have been reported in OLs after both TBI and SCI [4,5,65,66,68,127]. The accumulation of phagophores and autophagosomes in OLs was observed at three to seven days post-injury [4,5,68], which is later than in neurons in most reports. In a mouse contusion SCI model [127], myelin fractions purified from injured spinal cord tissues at eight days post-injury showed enrichment in autophagic proteins, including LC3-II, Atg5, and beclin 1. This study [127] has also

demonstrated the increased co-localization of p62 with Olig2 at 3 days post-SCI, indicating impaired autophagic flux in OLs. Additionally, after TBI, increased LC3+ cells have been observed not only in mature OLs, but also in NG2+ OPCs [5].

While the current data strongly support changes in autophagy in OLs during functional recovery after CNS trauma, much less is known about either the mechanisms or the function of autophagy in OL and OPC cells. It has been previously demonstrated in a rat model of demyelination that autophagy is necessary to support oligodendrocyte precursor survival and myelin development [128]. Thus, it is possible that autophagy in OPCs/OLs may play a similar function after CNS trauma. Using transgenic mice with OL-specific loss of the essential autophagy gene *Atg5*, Saraswat et al. reported [127] that the loss of autophagy in OLs exacerbates the inhibition of the autophagic flux in OLs and correlates with worse functional recovery after SCI and greater myelin loss.

ER stress is also a potent inducer of autophagy in OLs [127]. In vitro, a pharmacological blockade of ER stress-induced autophagy in OPCs increases survival. *Atg5* deletion specifically in OLs increased ER stress and reduced cell viability in vitro and in vivo after SCI, indicating a critical regulation of ER stress by autophagy in OLs. This is consistent with the beneficial contribution of *Atg5* and autophagy to OPC/OL health after SCI. Whether or not *Atg5* and ER stress in OLs/OPCs play a similar function following TBI needs to be determined.

#### *4.3. Function of Autophagy in Microglia and Astrocytes after CNS Trauma*

Microglia are a type of glial cell that is located throughout the CNS. They account for 10–15% of all cells found within the brain. As the resident macrophage cells, microglia act as the first and main form of active immune defense in the CNS. Microglia are activated in response to injury and are one of the main drivers of inflammatory responses after both TBI and SCI [129]. An increase in markers of autophagy has been reported in microglia after both TBI and SCI [4,5,66]. Interestingly, only the most activated CD68-expressing microglia with amoeboid morphology were reported to accumulate autophagosomes and p62 after TBI in a mouse CCI model [5]. This is consistent with the recently discovered function for autophagy in the regulation of inflammatory responses in macrophages and other immune cells [130,131], including microglia [132]. In general, high levels of autophagy flux are associated with anti-inflammatory properties and the inhibition of flux, with pro-inflammatory phenotypes [133]. Several autophagy genes are also linked to inflammatory and autoimmune diseases [133,134]. On the mechanistic level, autophagy is thought to control the activity of the inflammasomes, both directly by degrading components of activated NLRP3 (nucleotide-binding oligomerization domain, leucine rich repeat and pyrin domain containing 3) and AIM2 (absent in melanoma 2) inflammasomes, and indirectly by limiting mitochondrial damage and reactive oxygen species (ROS) production [135–137]. Additionally, autophagy can modulate inflammatory polarization through its influence on NFκB (nuclear factor kappa B subunit) activity by directly targeting the RELA/p65 protein and by affecting the availability of the p62 protein, which is needed for NFκB activation [138–140]. Conversely, a pro-inflammatory environment may also regulate levels of autophagy, as in primary cultured mouse microglia, where interleukin 1 Beta (IL-1β) induced the accumulation of many acidic vesicles loaded with autophagic markers (p62 and LC3) [141]. It is possible that autophagy may similarly be regulated by and contribute to the regulation of inflammatory responses in microglia and/or infiltrating macrophages after CNS injury.

Astrocytes are the largest and most numerous types of glial cells in the CNS, and are known to have a wide variety of physiological functions, including the maintenance of neurons, formation of the blood–brain barrier, and regulation of synaptic function. Similarly, to microglia, in response to injury, astrocytes are activated and become hypertrophic. Several groups reported that glial fibrillary acid protein (GFAP)-positive astrocytes accumulate autophagosomes after neurotrauma [65,66,82,84]. In mouse contusion SCI, reactive astrocytes accumulated phagophores and autophagosomes at seven d post-injury. Unlike other cell types, this was associated with increased autophagy initiation rather than its inhibition [68]. Much less is known about either the mechanisms or the function of autophagy

in astrocytes. Autophagy has been implicated in protective cellular responses in astrocytes [142–144]; however, further work will be necessary to determine how it may affect astrocyte survival and function after CNS trauma.

#### *4.4. Beneficial or Detrimental E*ff*ects of Autophagy Activation in Neurotrauma*

Defects in autophagy are thought to contribute to Parkinson's disease [145], Alzheimer's disease [146], and other age-related dementias. The accumulated evidence suggests that autophagy deficiency contributes to neurodegenerative diseases by perturbing protein homeostasis, as well as by directly contributing to neuronal cell death, axonal degeneration, and synaptic dysfunction. Conversely, the upregulation of autophagy has been proposed as a potential therapeutic strategy and shows considerable promise in both pre-clinical and clinical studies. However, the function of autophagy in neurodegeneration after traumatic CNS injury has been more controversial [1], with both beneficial and detrimental roles proposed. This may reflect that, as discussed above, either the induction or inhibition of autophagy flux may occur after acute CNS injury, especially in the brain. Since the function of autophagy is dependent on its flux, in cases where flux is inhibited, such as moderate to severe TBI or most cases of SCI, it would be expected to contribute to neuronal cell death and exacerbate other pathological phenotypes. Conversely, in cases where autophagy flux is increased, such as mild TBI, it would be expected to promote cell survival and improve outcomes. Thus, in each neurotrauma model, it is pertinent to ascertain whether observed changes in autophagy are the result of its induction or the inhibition of flux.

#### **5. Therapeutic Potential of Autophagy–Lysosomal Pathway Modulation in Neurotrauma**

#### *5.1. Neuroprotection*

The most common drugs used to manipulate autophagy in vivo are the mTOR (mechanistic target of Rapamycin kinase) inhibitor rapamycin, which is used to stimulate autophagy, and the type III PI3 kinase inhibitor 3-methyladenine (3-MA), which inhibits autophagosome formation. In general, pharmacological modulations that promote autophagy flux have been shown to provide neuroprotection in both TBI and SCI. However, in addition to autophagy, rapamycin is known to affect other cellular functions such protein synthesis, cell proliferation, and immune responses. Therefore, it is important to confirm that the neuroprotective effects of rapamycin are mediated via the restoration of the autophagy–lysosomal pathway. In a neonatal hypoxia–ischemia injury model, the protective effects of rapamycin were attenuated in animals treated with 3-MA [147]. Rapamycin-induced neuroprotection was also attenuated by AKT1 inhibition, suggesting that both autophagy and AKT1 signaling may be involved downstream of mTOR. Involvement of this pathway in TBI and SCI remains to be confirmed. However, as an improvement in functional recovery was reported after both pharmacologically enhancing [64,148] or blocking autophagy [149,150] after a thoracic SCI, further studies are warranted.

Another complicated factor is that activity of the target of rapamycin, mTOR, plays a vital role in oligodendrocyte differentiation [151] and myelination, and is also involved in axonal sprouting [152], which are both necessary for functional recovery after CNS injury, especially in the spinal cord. Therefore, the inhibition of mTOR may not be optimal as a treatment for CNS injury in general, and SCI in particular. In agreement with this, some [127,149] reported no effects of rapamycin on functional recovery after SCI. This suggests that drugs that are able to promote autophagy flux without inhibiting mTOR function may offer the best therapeutic benefits after neurotrauma. Trehalose is a naturally occurring sugar containing two glucose molecules, and has been reported as a novel mTOR-independent autophagy enhancer [153,154]. Trehalose has been shown to improve outcomes in rodent models of neurodegenerative diseases [155,156] and in a rabbit model of spinal cord ischemia [157]. However, trehalose is known to also act as a chemical chaperone [153], which could attenuate injury and improve recovery independently of autophagy. A specific mTOR inhibitor, Torin 1 [158,159], has been reported to increase autophagy flux. Both trehalose and Torin 1 induce lysosomal biogenesis via activation of

the transcriptional factor E-box (TFEB) [160] and increase autophagy flux but are known to act through different molecular pathways (mTOR-independent and mTOR-dependent, respectively). However, whether or not a combination of trehalose and Torin 1 is able to overcome the block of autophagy flux after neurotrauma and lead to improved functional outcomes needs to be investigated.

As lysosomal function is necessary to support autophagy flux, increasing lysosomal biogenesis has been shown to augment autophagy flux and improve outcomes in animal models of neurodegenerative diseases [161]. Enhancing lysosomal function by reducing lysosomal damage or promoting lysosomal biogenesis may also provide an attractive approach for therapy after neurotrauma. In both contusion TBI and SCI mice models, we recently demonstrated that the early administration of cPLA2 inhibitor arachidonyl trifluoromethyl ketone (AACOCF3) reduced lysosomal damage, improved autophagy flux, limited neuronal cell death, and improved functional outcomes [110,111]. Another possibility is the enhancing activity of the transcription factor EB (TFEB). TFEB is the master regulator of lysosomal biogenesis and is negatively regulated by mTOR [162]. The enhancement of autophagic flux by activating TFEB is protective in experimental brain injuries produced by cadmium [163]. This supports the notion that enhancing autophagy flux by stimulating lysosomal biogenesis may represent a potential treatment strategy after neurotrauma.

There are also other autophagy inducers reported in the context of neurotruama. Baicalin (7-D-glucuronic acid-5,6-dihydroxyflavone) is a major flavonoid in traditional Chinese medicinal herb isolated from the radix of Scutellaria baicalensis that was found to increase expression levels of LC3, Beclin 1, and p62 at 24 h following the weight-drop TBI model in mice [164]. Salubrinal is the selective phosphatase inhibitor of p-eIF2α. Wang et al. reported [165] that salubrinal reduced the expression of the ER stress marker as well as the number of CHOP+/TUNEL+ and CHOP+/LC3+ cells at 48 h after mouse TBI, which was associated with neuroprotection and improved recovery. Omega-3 polyunsaturated fatty acids (ω-3 PUFA) are known to have anti-oxidative and anti-inflammatory effects. ω-3 PUFA supplementation increased Beclin-1 deacetylation and its nuclear export, and induced direct interactions between cytoplasmic Beclin-1 and Bcl-2 by increasing the sirtuin family of proteins (SIRT1) activity following a weight-drop TBI model in rat [166]. Valproic acid (VPA), a class I/II histone deacetylase inhibitor, is able to significantly increase expression levels of the autophagic markers (LC3-II, Beclin, ATG-3, and ATG-7) at one day after TBI in rat [167]. These data are consistent with the notion that enhancing autophagy may be beneficial after neurotrauma. However, while the treatments affected the expression of autophagy markers after neurotrauma, their influence on autophagy flux remains to be assessed. Additionally, none of the compounds used are specific for autophagy, and have many additional cellular and organismal effects. Therefore, it remains to be determined if their beneficial effects are in fact dependent on autophagy.

#### *5.2. Neuroinflammation*

The crosstalk between autophagy and inflammation has been reported [168] in diseases associated with inflammation, such as inflammatory bowel diseases [169], type 2 diabetes [170], cardiac disorders [171], and cystic fibrosis [172]. Autophagy is known to be involved in the development, homeostasis, and survival of all inflammatory cells, including macrophages, neutrophils, and lymphocytes, thus playing critical roles in the development and pathogenesis of inflammation. In a mouse Alzheimer's disease model, a recent study [173] demonstrated positive correlation between Beclin-1, IL-1β, and TNF-α (tumor necrosis factor alpha) in the cortex and/or hippocampus, suggesting a relationship between inflammatory responses and autophagy. It remains to be determined whether a similar relationship exists after neurotrauma, and whether enhancing autophagy can be used to manipulate neuroinflammatory responses in this context. Conversely, as inflammation can affect cellular autophagy [1], the anti-inflammatory treatments could also improve autophagy flux.

Neurotrauma-induced neuroinflammation can cause the excessive generation of reactive oxygen species (ROS), which non-selectively damages neurons and glia, and thus participates in pathophysiology [174]. ROS can also regulate autophagy in both a positive and negative manner, depending on the levels and context [175,176]. Excessive ROS as well as reactive nitrogen species (RNS) can inhibit autophagy through the S-nitrosylation of proteins in the JNK (c-Jun N-terminal kinase) and mTOR pathways, which are important for the regulation of autophagy [177]. Lysosomal membranes can also be directly subjected to oxidative damage, leading to LMP. This suggests that ROS could be an important factor contributing to alterations in autophagy levels after neurotrauma. Conversely, autophagy can also be stimulated by ROS through its influence on ATG4 activity. Therefore, depending on the injury severity and mechanism, CNS injury-mediated ROS and RNS could either stimulate or inhibit autophagy flux.

#### **6. Conclusions and Perspectives**

When the CNS is injured, many cellular processes are activated that attempt to heal the damage. The activation of other cellular responses can increase damage over time. In case of autophagy, both protective and pathological functions are possible. Although we need to more thoroughly understand the complex and heterogeneous autophagic response triggered by the CNS trauma, in general, the enhancement of autophagy is considered protective in many injury paradigms, and helps restore homeostasis. Autophagy-enhancing drugs could positively affect multiple cell types, promoting neuron and oligodendrocyte survival, oligodendrocyte differentiation, and decreasing neuroinflammation. So, while we need to understand the mechanisms in more detail, the enhancement of autophagy after TBI and SCI carries a promise of a potential pleiotropic treatment that could target multiple cell types and pathways.

**Author Contributions:** Writing—original draft preparation, J.W.; figure drawing, J.W.; writing—review and editing, M.M.L. and J.W.

**Funding:** This research was funded by the National Institutes of Health (NIH), R01NS094527, R01NR013601, R01 NS110635, and R01 NS110567 to J.W., and R01NS091218 to M.M.L. and the APC was funded by R01NS094527.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

#### *Cells* **2019**, *8*, 693

© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

## *Review* **Recent Data on Cellular Component Turnover: Focus on Adaptations to Physical Exercise**

**Anthony MJ Sanchez 1,\*, Robin Candau 2 and Henri Bernardi 3**

Received: 15 May 2019; Accepted: 2 June 2019; Published: 5 June 2019

**Abstract:** Significant progress has expanded our knowledge of the signaling pathways coordinating muscle protein turnover during various conditions including exercise. In this manuscript, the multiple mechanisms that govern the turnover of cellular components are reviewed, and their overall roles in adaptations to exercise training are discussed. Recent studies have highlighted the central role of the energy sensor (AMP)-activated protein kinase (AMPK), forkhead box class O subfamily protein (FOXO) transcription factors and the kinase mechanistic (or mammalian) target of rapamycin complex (MTOR) in the regulation of autophagy for organelle maintenance during exercise. A new cellular trafficking involving the lysosome was also revealed for full activation of MTOR and protein synthesis during recovery. Other emerging candidates have been found to be relevant in organelle turnover, especially Parkin and the mitochondrial E3 ubiquitin protein ligase (Mul1) pathways for mitochondrial turnover, and the glycerolipids diacylglycerol (DAG) for protein translation and FOXO regulation. Recent experiments with autophagy and mitophagy flux assessment have also provided important insights concerning mitochondrial turnover during ageing and chronic exercise. However, data in humans are often controversial and further investigations are needed to clarify the involvement of autophagy in exercise performed with additional stresses, such as hypoxia, and to understand the influence of exercise modality. Improving our knowledge of these pathways should help develop therapeutic ways to counteract muscle disorders in pathological conditions.

**Keywords:** autophagy; mitophagy; mitochondria; exercise; AMPK; FOXO; MTOR; parkin

#### **1. Introduction**

Skeletal muscles are fundamental to the body's maintenance, and disorders in their function or metabolism are related to numerous diseases. Improved skeletal muscle activity has a significant effect on major processes in the body, such as the regulation of glucose homeostasis, contributing to enhanced health. Importantly, our capacity to recover from illness also depends on skeletal muscle oxidative capacity. Hence, skeletal muscle displays noteworthy adaptive responses from several stimuli, such as contractile activity, nutritional interventions, and environmental factors like hypoxia. These conditions may induce a transitory cellular stress leading to numerous adaptations, such as modifications in fiber composition, improvements of cell ability to renew cellular proteins and organelles, and modifications of muscle size [1–3].

Among the molecular sensors involved in adaptations to training, the adenosine monophosphate (AMP)-activated protein kinase (AMPK) is an enzyme composed of two regulatory domains (i.e., AMPK-ß, AMPK-γ) and a catalytic domain (i.e., AMPK-α). AMPK is a critical enzyme for preserving cellular homeostasis under conditions of low energy [4,5]. AMPK activity is increased by several energy stresses, including hypoxia/ischemia [6,7], electrical-stimulated muscle contraction [8,9], starvation [10], and physical exercise [11–13]. When cellular ATP is depleted, AMP modulates AMPK activity in an allosteric way, thereby promoting the phosphorylation of a threonine residue (Thr-172) within the α subunit by other enzymes called the "AMPK kinases" (AMPKK) [14]. There are three AMPKK proposed to date, the Ca2+/calmodulin- dependent protein kinase ß (CaMKKß) [15,16], the liver kinase B1 (LKB1) [17,18], and the transforming growth factor ß-activated kinase 1 (TAK-1) [19]. Of note, the binding of ADP, like AMP, prevents AMPK Thr-172 dephosphorylation [20]. On the contrary, AMPK is inhibited by ATP and glycogen [21,22]. AMPK is involved in cell metabolism and several data have highlighted the physiological relevance of its activation in skeletal muscle [4,23]. Thus, AMPK promotes energy production through the anaerobic and aerobic systems (i.e., glycolysis and oxidation of fatty acids) and, conversely, inhibits glycogenesis and cholesterol synthesis [5,24–28]. AMPK enhances mitochondrial biogenesis by stimulating PGC-1α (peroxisome proliferator-activated receptor gamma coactivator 1 alpha) expression [29]. A study by Jager et al. also showed that AMPK phosphorylates PGC-1α on two residues (Thr-117 and Ser-538) in vitro and in cells [30]. PGC-1α consecutively regulates the activity of PPARs (peroxisome proliferator-activated receptors) and NRFs (nuclear respiratory factors), leading to mitochondrial adaptations [30–32].

AMPK's biological functions are not limited to energy metabolism. In the last decade, AMPK was found to coordinate cell component turnover. AMPK decreases protein translation by reducing the activity of the mechanistic (or mammalian) target of rapamycin complex 1 (MTORC1) signaling, and promotes protein breakdown by regulating several component of the ubiquitin-proteasome and autophagosome-lysosome systems [5]. Major targets of AMPK are the forkhead box class O subfamily proteins 1 and 3 (FOXO1 and FOXO3, respectively). FOXO proteins are important transcription factors highly conserved through evolution and their various functions in skeletal muscle (i.e., cell cycle, DNA damage repair, apoptosis, energy metabolism, and oxidative stress resistance) have been recently reviewed [33]. In recent years, the AMPK-FOXO3 axis has been extensively studied with an important focus on processes regulating organelle turnover, especially mitophagy.

In this review, recent discoveries on AMPK-MTORC1 and AMPK-FOXO axes in the coordination of muscle organelle renewal and the importance of physical exercise on both acute and chronic adaptations are discussed. The multiple modes of regulation of these sensors are detailed, as their implication in the regulation of skeletal muscle protein and organelle turnover, especially mitophagy. Apparent discrepancies between the data are discussed in regard to the methodology used to access autophagy or mitophagy activity. The functions of newly identified actors in protein and organelle quality control, specifically the diacylglycerol kinase ζ (DGKζ), Parkin (RING-between-RING E3 ligase), and Mul1 (mitochondrial E3 ubiquitin protein ligase), are also presented. We finally discuss the impact of exercise modality, hypoxia, and examine the current limitations in the literature to suggest other perspectives.

#### **2. AMPK and MTORC1 Pathways**

#### *2.1. AMPK*/*MTORC1 Axis in Organelle Quality Control during Exercise*

Protein synthesis machinery is globally decreased during exercise. The MTORC1 pathway is a central regulator of protein turnover under conditions of increased external loading by the regulation of ribosomal translation. Thus, the kinase MTOR modulates mRNA translation and protein synthesis by regulating major regulators of ribosomal activity, 4E-BP1 (eukaryotic translation initiation factor 4E-binding protein 1) and S6K1 (ribosomal protein S6 kinase 1). MTOR phosphorylates S6K1 at Thr-389 (its hydrophobic motif), which, in turn, phosphorylates translational effectors such as rpS6 (ribosomal protein S6) and eIF4B (eukaryotic translation initiation factor 4B) [34,35]. In addition, PDK1 (phosphoinositide-dependent kinase-1) phosphorylates S6K1 at Thr-229 provides its full activation [36,37]. 4E-BP1 phosphorylation by MTOR on Thr-37/46 provides its disconnection from the preinitiation complex (PIC) promoting the transcription of protein-coding genes [38,39]. In a study conducted in mice, moderate

endurance exercise decreased the phosphorylation state of MTORC1 signaling (i.e., MTOR, S6K1, rpS6, and 4E-BP1) from 90 min [40]. These modulations were concomitant with a raise of the phosphorylation state of eiF2α (Ser-51), an indicator of endoplasmic reticulum stress and AMPK activation. The AMPK mediates inhibition of MTORC1 through phosphorylation of MTOR at Thr-2446 [41], of the tuberous sclerosis complex 2 (TSC2) at Thr-1227 and Ser-1345 [42] and the associated regulatory protein of MTOR complex-1 (RPTOR) on two well-conserved serine residues (Ser-722/792) [43,44] leads to the sequestration of the raptor by 14-3-3 proteins [44]. These events promote the inhibition of Rheb (Ras homolog enriched in brain) and overall MTORC1 inhibition (Figure 1). However, data on the involvement of AMPK/RPTOR axis for MTORC1 inhibition during endurance exercise are lacking.

**Figure 1.** (AMP)-activated protein kinase (AMPK), forkhead box class O subfamily protein (FOXO), and mechanistic (or mammalian) target of rapamycin complex 1 (MTORC1) in the regulation of protein and organelle turnover. FOXO proteins increase the transcription of the E3 ubiquitin protein ligases muscle atrophy F-box (MAFbx)/atrogin-1, muscle RING finger 1 (MuRF1), mitochondrial E3 ubiquitin protein ligase (Mul1), several autophagic genes (Atgs), and BCL2/adenovirus E1B 19 kDa protein-interacting protein 3 (BNIP3) in muscle cells. MTOR and AMPK differentially modulate autophagy initiation by phosphorylation of the Unc-51-like kinase (ULK1). AMPK also activates FOXO1 and FOXO3 and tuberous sclerosis complex 2 (TSC)1/TSC2 complex, and inhibits MTORC1 complex through phosphorylation of the associated regulatory protein of MTOR complex-1 (RPTOR). MAFbx/atrogin-1 and MuRF1 target sarcomeric proteins. MAFbx/atrogin-1 also targets factors involved in cell growth including the transcription factors Myogenin and MyoD, and the eukaryotic initiation factor 3f (eIF3f). PTEN-induced putative kinase protein 1 (PINK1)/Parkin and Mul1 axes enhance mitophagy through ubiquitination of mitochondrial proteins.

A single bout of exercise may lead to a strong stimulation of muscle protein synthesis during the recovery period. Synthesis of myofibrillar proteins is elevated after resistance exercise [45–47] and seems to become more pronounced in trained athletes compared to untrained individuals [48]. Endurance training also increases protein synthesis during the recovery since the MTORC1 pathway is altered in response to this type of exercise. Indeed, increased phosphorylation of several MTOR targets occur in both mice and humans after moderate and exhaustive endurance exercises [40,49,50]. However, selective activation of MTORC1 by specific intracellular signaling pathways is involved according to the exercise modality. Acute sprint exercise or electrical stimulation at high frequency increases PKB (protein kinase B or Akt) and MTOR phosphorylation during recovery [51,52]. Importantly, a recent study conducted in rats compared different resistance exercise models and suggested that hypertrophic response is correlated with the phosphorylation level of MTOR and its regulators or targets (i.e., Akt, the extracellular signal-regulated kinases ERKs, p38, the mitogen-activated protein kinases MAPKs, 4E-BP1) [53]. Finally, the study by Ogasawara highlighted that both rapamycin-sensitive and rapamycin-insensitive MTOR signaling regulate MTOR-dependent muscle protein synthesis during resistance exercise [54]. Concerning endurance exercise, several studies failed to observe modulation of Akt during recovery, whatever exercise intensity [52,55]. Thus, it has been proposed that an AMPK-Akt switch may be involved in the occurrence of specific adaptations to resistance and endurance training [55], with a more specific contribution of Akt/TSC1/TSC2 cascade for resistance, strength, and sprint exercises. This also means that Akt is not essential for MTORC1 activation and protein synthesis during endurance exercise. AMPK activation also contributes to decrease the rate of protein production during resistance exercise. However, protein translation augments from 1 h post-exercise even if AMPK activity becomes less pronounced from 2h[56]. Finally, in individuals accustomed to training, MTORC1 signaling appears preferably involved for hypertrophy-inducing exercises and the AMPK axis seems to be more specific to endurance training adaptations [57]. It is interesting to note that a combination of endurance and resistance training may affect protein synthesis differentially. Indeed, AMPK activation via endurance exercise may negatively affect MTORC1 activation induced by resistance training. This effect is increased when the endurance exercise is carried-out after a bout of resistance exercise [58].

#### *2.2. MTORC1 Regulators and Exercise: Recent Data on DGKs, FOXO, eIF3f and Cellular Tra*ffi*cking*

Among MTORC1 regulators, the PI3K (phosphoinositide 3-kinase)/Akt axis is notably modulated by exogenous nutrients and the release of growth factors [59–62]. MTOR phosphorylation during mechanical overload, and at the early phase of recovery, is related to MEK/ERK signaling through phosphorylation of TSC2 at Ser-664 but not to PI3K/Akt signaling [63]. ERK1/2 also regulates nuclear transcriptional factors, such as Elk-1 (E-26-like protein 1), c-Myc, c-Jun, and c-Fos that play a role in muscle growth [64]. Activation of ERK1/2 and MTORC1 seems to be required for full stimulation of protein synthesis in humans [65]. However, none of these MTORC1 regulators are recognized as critical actors for fiber hypertrophy during resistance exercise [59–61].

It is also recognized that mechanical stimulus enhances the activation of PLD (phospholipase D) and production of PA (phosphatidic acid), promoting activation of MTORC1. PA is a lipid messenger that binds to MTOR's FKBP12- rapamycin binding domain, favoring its activation [61,66–68]. Nonetheless, PLD activity modulation does not seem only implicated in PA production or MTORC1 activation [69]. Thus, studies from Hornberger's lab investigated the role of diacylglycerol (DAG) and DAG kinases (DGKs) in PA production during mechanical stimulation. DGKζ (DAG zeta) is known to be involved in PA accumulation via DAG phosphorylation and was strongly suggested to be essential for PA production and enhanced MTORC1 activity during mechanical stimulation. More recently, the same group explored the importance of DGKζ in muscle adaptations in vivo with a hypertrophic model of mechanical loading [70]. Thus, the authors reported that DGKζ isoform is the most highly raised and is of importance for muscle growth and hypertrophy. Studies on the mechanisms underlying these adaptations highlighted that DGKζ also suppresses FOXO3 activity, leading to a decline of MAFbx (muscle atrophy F-box)/atrogin-1 and MuRF1 (muscle RING finger 1) induction. Conversely, the expression of these E3 ligases during exercise was increased in DGKζ knock-out muscles, confirming the role of DGKζ in limiting protein breakdown during mechanical overload. Even if the precise mechanism needs to be identified, DGKζ relocation in the nucleus seems to be essential for inhibiting FOXO3 transcriptional activity during mechanical overload. Interestingly, the authors also found that mechanical overload increases expression of the eukaryotic initiation factor 3 subunit f (eIF3f), and this increase is totally blunted in DGKζ knock-out muscles. Given the importance of eIF3f

in protein translation process, it can be hypothesized that DGKζ-dependent alteration of FOXO pathway during mechanical overload may also have effects on the translational machinery. Indeed, the FOXO-dependent E3 ligase MAFbx/atrogin-1 is well known to target eIF3f leading to its proteasomal degradation [71].

The role of eIF3f and MTORC1 intracellular trafficking in adaptation to exercise has been also recently investigated. eIF3f belongs to the translation initiation factor complex eIF3f among its 13 subunits involved in mRNA translation initiation. In the last two decades, studies from Leibovitch's lab highlighted eIF3f's involvement in skeletal muscle protein synthesis and hypertrophy [72]. It was demonstrated that a TOS (TOR signaling) motif in eIF3f operates as a scaffold to connect MTORC1 with its translational substrates and to support the initiation of cap-dependent translation [73]. Starvation muscle atrophy is suppressed when an eIF3f mutant insensitive to polyubiquitination by MAFbx is overexpressed, showing its critical role in muscle homeostasis during such a stress [74]. More recently, eIF3f was found to be essential for mouse embryonic development and its partial depletion reduces adult skeletal mass and amplifies muscle loss during disuse by mainly modulating protein synthesis [75]. Concerning exercise, an increase of eIF3f expression was evidenced during overload [70], suggesting a role in adaptations to exercise. Studies in human tissue model that used immunofluorescence approaches identified a type of cellular trafficking involving eIF3f that occurs during a single bout of resistance exercise [76]. Thus, the complex composed of MTORC1/eIF3f was found to co-localize with the lysosome, where the GTPase Rheb is known to trigger the kinase activity of MTOR enhancing MTORC1 substrates phosphorylation [77]. In accordance, studies in vitro and in animals highlighted that MTORC1 recruitment to the lysosome surface is critical to raise MTOR kinase activity [78,79]. After resistance exercise, the MTORC1/LAMP2 complex was found to rapidly translocate at the cell membrane in close proximity to capillaries [76]. Importantly, TSC2 abundance at the cell membrane was also reduced with a dissociation from Rheb, suggesting a decrease of MTOR inhibition favorable to its full activation. These innovative results suggest that, at least in humans, MTORC1 is recruited and activated at the cell periphery following resistance exercise. Interestingly, a protein–carbohydrate beverage post-exercise does not alter MTORC1/eIF3f translocation but increases the interaction between MTOR and eIF3f [76], an association well recognized to drive MTORC1 activation and enhance MTOR target activity [72]. Thus, a bout of resistance exercise may enhance mRNA translational capacity through the association and the translocation to the cell periphery of MTOR and its positive regulators eIF3f and Rheb.

Interestingly, nonprotein dietary factors also influence post-exercise myofibrillar protein synthesis. In humans, it was recently reported that the ingestion of egg whites alone results in lower myofibrillar protein synthesis activation than the ingestion of whole eggs during the recovery from resistance exercise [80]. In accordance, whole egg ingestion increases the phosphorylation level of MTOR, 4E-BP1, rpS6 to a greater extent than egg white ingestion. Whole egg ingestion was also found to increase MTORC1 co-localization with the lysosome after resistance exercise, and this result was correlated with higher rates of myofibrillar protein synthesis [81]. This observation suggests a better mRNA translational capacity after whole egg consumption than after egg white consumption. The underlying mechanisms have been partially studied and PA was proposed to have a role among the potential factors involved in MTORC1 recruitment to the lysosome. Egg yolk contains phosphatidylcholine and oleic acid that can be converted to PA via de novo synthesis, and DAG [82–84]. In addition, the egg yolk is enriched in low-density lipoprotein (LDL)-derived cholesterol that was shown to play a role in MTORC1 recruitment to the lysosome in a SLC38A9-Niemann-Pick C1 (a sterol transport system) signaling complex fashion [85]. In summary, these results highlighted the importance of a new intracellular trafficking mechanism and nonprotein dietary factors that drive optimal myofibrillar protein synthesis after resistance exercise (Figure 2).

**Figure 2.** Cellular trafficking of mechanistic (or mammalian) target of rapamycin complex 1 (MTORC1) and lysosome after resistance exercise. In response to resistance exercise, MTORC1 co-localizes with the lysosome and translocates at the cell membrane. MTORC1 recruitment to the lysosome surface is critical to increase MTOR kinase activity. Contractions lead to a decrease of tuberous sclerosis complex 2 (TSC2) abundance and, conversely, to the activation of MTOR by the GTPase Ras homolog enriched in brain (Rheb). A protein-carbohydrate beverage post-exercise increases the interaction between MTOR and eukaryotic initiation factor 3f (eIF3f) without altering MTORC1/eIF3f translocation. Whole eggs ingestion enhances MTORC1 co-localization to the lysosome after resistance exercise.

#### **3. AMPK and FOXO Transcription Factors**

#### *3.1. FOXO Homologues in Energy Metabolism and Post-Translational Modifications*

Four FOXO members (FOXO1/3/4/6) are expressed in muscle. FOXO6 represses PGC-1α expression and low intensity exercise reduces FOXO6 induction [86], suggesting that exercise-induced PGC-1α may be partially dependent on FOXO6. Nonetheless, data concerning this factor remain limited in this tissue. FOXO1, 3, and 4 have been more extensively investigated, notably due to their important roles in cell cycle, apoptosis, muscle growth, and muscle regeneration [33]. FOXO3 regulates MyoD (an essential myogenic differentiation factor) transcription [87]. FOXO1 and FOXO3 play critical roles in energy homeostasis by favoring mitochondrial metabolism, inhibiting glycolysis, and enhancing lipolytic flux [88–91]. FOXO3 plays a role in the regulation of mitochondrial genome through AMPK and the mitochondrial Sirtuin 3 (SIRT3) [92,93]. FOXO1 and FOXO3 also regulate exercise-induced angiogenesis. FOXOs contribute to the repression of muscle angiogenic response through the induction of thrombospondin 1 (THBS1) during the first days of chronic exercise. Importantly, FOXO repression is critical for long term adaptations to endurance training, especially for angiogenesis [94,95].

FOXO proteins are regulated by several post-translational mechanisms, and numerous enzymes have been identified as kinases of FOXO, including AMPK, Akt, and, more recently, DGKζ. In muscle cells, AMPK activates FOXO3 by phosphorylation at Ser413/588 [23,96]. We recently found that AMPK activation by 5-aminoimidazole-4-carboxamide-1-ß-d-ribofuranoside (AICAR) extends the FOXO3 protein half-life in skeletal muscle primary cells [97]. AMPK also increases the cellular NAD+ level, thus enhancing the activity of histone deacetylase Sirtuin 1 (SIRT1) [98]. This process induces the deacetylation of FOXO1 and 3 leading to their activation [98]. Conversely, Akt, when inhibited by the phosphatidylinositol 3-kinase (PI3K), is responsible for FOXO1 protein translocation from the nucleus to the cytoplasm, leading to its inactivation [99]. Akt phosphorylates FOXO3 on several residues (Thr-32, Ser-253/315), and the phosphorylation of residues Thr-32 and Ser-253 promotes its cytoplasmic retention by a mechanism involving the 14-3-3 chaperone protein [100]. FOXO regulation by Akt induces a decrease in the binding between FOXO and its DNA sequences targets and, consequently, decreases

FOXO transcriptional activity [101,102]. In addition, FOXO3 is targeted by the histone acetyl-transferase p300 for its ubiquitination by Mdm2 (E3 ligase murine double minute 2) and subsequent degradation by the proteasome [103]. In muscle, p300 alters differentially the expression of FOXO 1 and 4 without affecting the expression of FOXO3 [104]. Furthermore, p300 alters FOXO3 and FOXO4 activity but increases the nuclear localization of FOXO1 and the transcription of FOXO1-dependant genes underlining the differential regulation of the FOXO homologues [104]. Finally, DGKζ has been recently found to suppress FOXO3 activity in skeletal muscle [70], extending the role of the DGK to the regulation of catabolism (Figure 3).

**Figure 3.** (AMP)-activated protein kinase AMPK and forkhead box class O subfamily protein FOXO regulation in skeletal muscle. FOXO proteins are phosphorylated and inhibited by protein kinase B (PKB or Akt) in response to insulin or growth factor. Under the condition of energy stress, AMPK, which is activated by the AMPK kinases (AMPKK) Ca2+/calmodulin- dependent protein kinase ß (CaMKKß), liver kinase B1 (LKB1), and transforming growth factor ß-activated kinase 1 (TAK-1), phosphorylates and increases FOXO3 activity. AMPK is also involved in FOXO deacetylation through Sirtuin 1 (SIRT1) and Sirtuin 3 (SIRT3). AMPK activation by 5-aminoimidazole-4-carboxamide-1-ß-D-ribofuranoside (AICAR) stabilizes FOXO3 by increasing its protein half-life. Moreover, AMPK favors the association between FOXO3 and mitochondrial DNA through SIRT3, mediating the transcription of mitochondrial genes. During mechanical overload, FOXO3 transcriptional activity is inhibited by diacylglycerol kinase ζ (DGKζ) by a mechanism independent of DGKζ kinase activity within the nucleus. Cytosolic FOXO3 is targeted by E3 ligase murine double minute 2 (Mdm2) for degradation via the ubiquitin-proteasome system. FOXO3 and FOXO4 are acetylated by the histone acetyltransferase p300. Abbreviations undefined in the main text: IRS1, insulin receptor substrate 1; PIP2, phosphatidylinositol 4,5-bisphosphate; PIP3, phosphatidylinositol 3,4,5-triphosphate.

#### *3.2. AMPK*/*FOXO Axis in Organelle Quality Control during Exercise*