13

Abstract: While patient-specific targeting of cellular growth and viability pathways dominates current approaches in anti-cancer therapeutics development, appreciation for the strategy of targeting transformation-dependent alterations in cellular organelle structure and function continues to grow. Here we discuss the lysosome as an anti-cancer target, highlighting its role as a key mediator of cell death. As the major degradative compartment of the cell, the lysosome houses dozens of destructive enzymes and is responsible for the breakdown of both internal and external molecules and particles; however, until relatively recently the contribution of the lysosome to cellular death mechanisms has been largely overlooked. Renewed interest in the therapeutic potential of lysosomal rupture to combat cancer has led to development of lysosome-disrupting agents that induce lysosomal membrane permeabilization (LMP), cathepsin protease release, and subsequent lysosome-dependent cell death (LDCD), now distinguished as a bona fide cell death process. Here, we present the basic biology, structure, and function of the lysosome, with particular emphasis on the transformation-associated alterations that sensitize cancer cell lysosomes to membrane rupture. We further describe the lysosome’s role in cell death and comprehensively outline emerging therapeutic strategies that exploit lysosomes for the treatment of a variety of malignancies.

Keywords: cancer therapeutic targeting; cationic amphiphilic drugs; lysosomal membrane permeabilization; lysosome; lysosome-dependent cell death

INTRODUCTION

Using subcellular fractionation based purely on biochemical criteria, de Duve and colleagues (1955) made the serendipitous discovery of the lysosome (1, 2), an achievement deemed worthy of the Nobel Prize in Physiology (1974). Upon observation that liver-derived acid phosphatase exhibited latent activity following homogenization, it was deduced that a membrane-bound structure must normally sequester it, and potentially other degradative hydrolases, from their substrates (2, 3). Indeed, the lysosome harbors some 60 lytic enzymes (4) capable of degrading proteins, nucleic acids, polysaccharides, and lipids. Subsequent investigations determined that lysosomes serve as the terminal compartment for the degradation of extracellular materials taken up by endocytosis and phagocytosis and the digestion of intracellular constituents isolated during autophagy (5, 6). We now appreciate that lysosomes are more than cellular refuse depots; they are fundamental components of dynamic physiologic processes such as plasma membrane repair, bone and tissue remodeling, matrix degradation, inflammatory responses, antigen presentation, cholesterol homeostasis, nutrient sensing and metabolism, cell signaling, growth factor recycling, and programmed cell death (7–23). These processes differentially rely on fusion with endocytic vesicles or the regulated release of lysosomal hydrolases into the cytosol via lysosomal membrane permeabilization (LMP) or into the extracellular space via lysosomal exocytosis. A summary of the lysosome’s various functions is illustrated in Figure 1.

Figure 1. Functions of the lysosome. Lysosomes regulate cell function by internalizing and degrading pathogens, receptors, cellular debris, etc. via endocytosis and phagocytosis (upper left) and autophagy (upper right). They also transmit materials to the cell surface and extracellular space by exocytosis (lower left). Lysosomal membrane permeabilization resulting from various stimuli (e.g., reactive oxygen species (ROS) and iron accumulation) promotes cathepsin release and subsequent lysosomal cell death (lower right). Figure created with BioRender.com (adapted with permission from ref 163).

Of note are observations that the quantity, composition, and complement of lysosomal hydrolases are often augmented with cancer pathologies. Along these lines, lysosomal heparanase and cathepsins promote cancer cell proliferation, angiogenesis, and metastasis, suggesting that these and other lysosomal enzymes are of potential clinical significance (24, 25). The therapeutic implications of lysosomal hydrolases were recognized decades ago, with seminal investigations describing enhanced activities of lysosomal enzymes in solid tumors as compared to their tissues of origin, with specific enzymes (i.e., beta-glucuronidase) favoring tumor cell invasiveness (26). Perhaps it is not surprising then that cancer cells often exhibit an expansion of the lysosomal compartment (24, 27, 28), a feature that would enhance tumor aggressiveness. However, such distinction may also provide a rational basis for therapeutic intervention. Cancer-associated lysosomes are more fragile than their normal counterparts due in some measure to increases in hydrolytic enzymes and fundamental changes in the composition of the lysosomal membrane (29–31). Based on the concept that the lysosome may represent a ‘suicide-bag’ as first proposed by de Duve (32), the instability of cancer-associated lysosomes may lend to enhanced cellular susceptibility to LMP, coincident release of destructive hydrolases into the cytosol and ultimate cell demise by either apoptotic or non-apoptotic cell death mechanisms.

Herein, we review the fundamentals of lysosome physiology, composition, and function in cell death, and connect cancer-associated changes in the expression and activity of lysosomal components with a particular focus on the therapeutic opportunities they may provide for breast and other tumor types.

LYSOSOMAL STRUCTURE, DISTRIBUTION, AND IDENTIFICATION

Lysosomes are typically less than 1 μm in diameter and contribute up to 0.5% of the total intracellular volume of many eukaryotic cells, although this may vary depending upon cell type (i.e., macrophages), energetic state, or degradative requirements (33–35). Unlike other organelles, lysosomes cannot be identified based on uniform morphologic criteria, as there is significant variation in their size, architecture, and morphology depending upon nutrient availability, for example, autophagy (36–39). Significant augmentation of lysosome volume, abundance, and structure also occurs during certain pathologic states, for example, lysosomal storage diseases (40) and cancer (24, 41), or following experimental manipulations that inhibit enzymatic digestion such as overloading with non-physiologic substrates such as sucrose (42), administration of cationic amphiphilic drugs (CADs (43–45)), or treatment with aminoglycoside antibiotics (46). Interestingly, lysosomes at peripheral locations can partially change their intracellular pH (47, 48), a feature that may be co-opted by some cancer cells to facilitate constitutive mTOR signaling (38) or modulate extracellular acidity to enhance invasion (49, 50).

By electron microscopy, lysosomes are identifiable as either tubular or spherical membrane-bound structures with cores of variable densities, amorphous granular material, or membrane whorls (34, 51). Biochemically, lysosomes are defined by the presence of multiple hydrolytic enzymes (32) which may vary between tissue type (51) and pathology (25, 40). Lysosomes may be distinguished from endosomes by their pH, calcium content, abundance of lysosomal-associated membrane proteins LAMP-1 and LAMP-2, and lack of mannose-6-phosphate receptors (34, 52–54). Fluorescent dyes that accumulate in acidic vesicles––such as LysoTracker Red and Acridine Orange––effectively label lysosomes, however other acidic vesicles such as endosomes and autophagosomes may be concomitantly labeled to varying degrees (55).

LYSOSOME COMPOSITION

Lysosomes can degrade a vast array of structurally diverse macromolecules into their constituent components. Following degradation, substances either diffuse or are transported out of the lysosome into the cytosol where they become fuel for metabolism or substrates for biosynthetic pathways (5). This dynamic recycling process requires the coordinated action of the lysosomal acid hydrolases with integral, peripheral, and transiently-associated proteins as discussed in the following sections.

Lysosomal hydrolases

To achieve efficient breakdown of complex substrates, lysosomes contain several acid hydrolases such as proteases, glycosidases, nucleases, sulfatases, and lipases. In addition to the degradation of material delivered via endosomes, phagocytic vesicles, and autophagosomes, lysosomal hydrolases are involved in diverse processes such as pro-protein and antigen processing, degradation of extracellular matrix, stimulation of angiogenesis, and the initiation of cell death (13, 56–61).

Chief among the acid hydrolases are the aspartic, serine, and cysteine proteases, with the most widely studied being the cathepsins (62). While cathepsins are most recognized for their activity within the lysosomal compartment, a number of studies have indicated their localization to any vesicle along the endocytic pathway (early and late endosomes, phagosomes), within the nucleus or cytosol, at the cell surface, or secreted into the extracellular matrix, depending on physiologic or pathologic state (61–64). Cathepsins are synthesized as inactive precursors that are then processed to their mature and active form by proteolytic removal of the N-terminal propeptide (65). Removal of the propeptide may occur by autolysis within acidic lysosomes or by activation of other proteases in a chain-like reaction (66–70) and may be enhanced in the presence of glycosaminoglycans or polysaccharides (71–74). While most cathepsins become destabilized at neutral pH, several interacting partners such as heparin and catalase may prolong cathepsin activity by promoting structural integrity and inhibiting peroxidation (75, 76).

Originally considered to function only within the lysosome in general protein turnover, it has become exceedingly clear from gene knockout models that cathepsins have non-redundant and diverse functions and may be expressed ubiquitously in a tissue-specific or even context-specific manner (77–82). The diversity of this class of proteases is beyond the scope of this review and has been extensively examined elsewhere (63, 83–85). Notably, cysteine proteases B, L, S, X, and K, as well as aspartic cathepsin D, have all been implicated to varying degrees in cancer progression. Along these lines, cathepsins in cancer cells are often translocated to the plasma membrane along with pH regulators such as v-ATPases and Na⁺/H⁺ exchangers (86), where they associate with microdomains or are secreted in an active form (87). In this respect, cancer cells effectively exploit cathepsins to remodel the extracellular environment to potentiate invasion and metastasis (61, 88–93). Alternatively, infiltrating macrophages may supply cathepsins to stimulate angiogenesis and promote the growth and invasion of associated tumor cells (94).

Cathepsins may be specifically regulated by interactions with endogenous inhibitors, including cytosolic stefins, extracellular cystatins, and kininogens (63, 93). As such, blunted cystatin often accompanies enhanced cathepsin levels during the acquisition of invasive capacity (95). More recently, cathepsins have been implicated in the development of intrinsic therapeutic resistance and adaptive responses to treatment (93, 96–98).

Membrane-associated proteins and lipids

The lysosomal membrane contains more than one hundred proteins, with LAMP-1 and -2 comprising nearly 50% of the total protein content (99). The oligosaccharide side chains on LAMPs and LIMPs (lysosomal integral membrane proteins) form a thick polysaccharide coat, or glycocalyx, that lines the inner surface of the lysosomal membrane to ensure protection of sensitive lysosomal and extralysosomal substrates from degradative hydrolases (100, 101). In addition to ensuring compartmentalization of acid hydrolases and maintaining structural integrity, peripheral and integral membrane-associated proteins are vital to lysosomal (LAMP-1) and the plasma membrane (RABs and SNAREs) trafficking, transport of ions and soluble substrates (cation channel mucolipin1, chloride channel CLCN7, protein transporter LAMP-2A, amino acid transporter LAAT1), and nutrient sensing (v-ATPase), as previously reviewed (52, 102–106). In addition, lysosomes may contain multiple internal vesicles that harbor their own unique complement of proteins and lipids (107, 108), imparting further functional diversity. Given this diversity and the propensity for alterations during cancer, it is worth detailing a few key membrane constituents. For a more in-depth discussion, the reader is referred to several excellent topical reviews (108–113).

Several observations suggest that LAMPs may contribute to the fragility of cancer-associated lysosomal membranes. Oncogenic transformation of fibroblasts is accompanied by a decrease in LAMP expression, redistribution of lysosomes to the cell periphery, and increased sensitivity to lysosomal cell death and to agents that induce LMP (114). Conversely, LAMP overexpression was found to be protective against LMP (114), and a role for LAMP in cytoprotective autophagy has been proposed (115). While overall LAMP expression is reportedly increased in a number of cancers (27, 116–118), it is likely these observations are indicative of an increase in total lysosome content and not changes in their activity per se. Given the role of LAMPs in the formation of the protective glycocalyx, it is conceivable that their overall loss augments internal hydrolase-mediated damage to other lysosomal membrane constituents and propensity toward LMP.

Lysosomes are bound by a single bilayer membrane, which is comprised of a primary lipid matrix of glycerophospholipids, sphingolipids, and cholesterol (110). In general, glycerophospholipids such as phosphatidylcholine, phosphatidylethanolamine, and phosphatidylinositol dictate the fluidity of biomembranes and participate in trafficking, fission, and fusion events (110, 119). Lysosomal and late endosomal membranes are uniquely enriched in the glycerophospholipid bis(monoacylglycero) phosphate (BMP), permitting a heightened capacity for cholesterol transport and sphingolipid degradation (108, 120, 121). Along these lines, depletion of cellular cholesterol results in an increase in lysosomal density and affects resistance to agents such as sucrose and lytic compounds known to perturb membrane structure (122). Moreover, cholesterol contributes to the formation of detergent-resistant lipid rafts within lysosomal membranes, which are focal centers for sorting and concentrating complexes of proteins vital for trafficking and signal transduction (123) as shown by proteomic (18) and biochemical (124) analyses. Cellular repressor of E1A-stimulated genes (CREG), a secreted glycoprotein that promotes the differentiation of pluripotent stem cells (125) and inhibits cell growth (126), concentrates specifically at lipid rafts (18). Lipid rafts have garnered particular interest in recent years because of their role in metastasis and various cell death pathways (127).

Although originally considered a source of structural support, mounting evidence implicates sphingolipids like sphingomyelin, ceramide, and glycosphingolipids as important agents of lipid raft cell signaling cascades (110, 128, 129). Different sphingolipid species have been implicated in regulating cell survival, angiogenesis, inflammation, proliferation, autophagy, and programmed cell death (130–133). For example, ceramide, which is hydrolyzed from sphingomyelin by lysosomal acid sphingomyelinase (aSMase) (134) or other mechanisms (135, 136), has been intensively studied following observations that aSMase-deficient mice were resistant to cell death induction (137, 138). The aSMase/ceramide pathway has since been identified as a central component of cellular response to various stressors and chemotherapeutics (111, 135, 139–141), potentiation of redox signaling (142, 143), autophagy (144–148) and regulation of proteins involved in programmed cell death (i.e., phospholipase A2 (149), cathepsin D (150, 151), Jun-N-terminal kinases (152).

LYSOSOMES IN CELL DEATH

Cell death is classically defined by morphological criteria: apoptotic cells display cellular shrinkage, nuclear fragmentation, and condensation into apoptotic bodies for clearance by phagocytosis; autophagy-dependent cell death involves cytoplasmic vacuolization and autophagosome formation, followed by lysosomal degradation; necrosis manifests as organelle swelling, plasma membrane breakdown, and disintegration of cellular structures (153). However, investigations into the precise biochemical and functional underpinnings of cell death processes have revealed distinct “regulated” cytotoxic programs and prompted a diversification of nomenclature. Lysosome-dependent cell death (LDCD), characterized by lysosomal destabilization and requiring LMP, is now distinguished as a subclass of programmed cell death (153). Though lysosomal rupture has been observed as an ultimate consequence of canonical cell death processes (154), primary LMP activates a death program in LDCD and can be differentiated by novel assay systems (155, 156). LMP does not generate defining morphological alterations (157) and is therefore classified at the molecular level by release of lysosomal luminal contents including proteolytic cathepsin enzymes to the cytosol, where cathepsins function in a variety of contexts as cell death executioners (158). However, the precise mechanisms leading to loss of lysosomal membrane integrity and protease translocation to the cytosol are not fully elucidated for the majority of LMP stimulants. Activities of pore-forming toxins such as venoms, bacterial toxins, and viral entry proteins are fairly straightforward; these compounds can disrupt membrane dynamics from within the lysosome following their uptake into the endolysosomal system and activation at low-pH, or alternatively induce pore formation from the cytosol (158, 159). Lysosomotropic detergents are well-studied LMP-promoting agents that function by directly disrupting membrane dynamics leading to organelle leakage or by impairing function of lysosomal lipases (158). Under physiological conditions, LDCD contributes to tissue remodeling during mammary gland involution (160) and regulates immune cell clearance following inflammation (161) or bacterial infection (162). Moreover, LDCD is associated with a variety of pathological states (153).

Consequences of LMP

LMP may either initiate or amplify a cell death cascade, and it can trigger distinct pathways depending on cellular context and the nature of lysosomal injury. The molecular players and morphological outcomes of a given lethal subroutine featuring LMP can be classified as apoptotic or necrotic, and it is widely accepted that the degree of lysosomal rupture––with respect to number of lysosomes impacted and extent of membrane damage––dictates the specificity of lysosomal component release and downstream cellular responses (158). Extensive LMP allows rapid release of lysosome contents to the cytosol and lethal cytoplasmic acidification, resulting in rampant hydrolysis of cytoplasmic contents and cell death by necrosis following plasma membrane breakdown. Conversely, the cytosolic translocation of select cathepsins with limited LMP initiates a regulated signaling cascade and death resembling apoptosis (163–165). Indeed, cathepsin inhibition can revert the effects of limited LMP. Cathepsins were identified as principal mediators of LMP-dependent cell death by studies demonstrating cell viability rescue with pharmacological or genetic manipulation of cathepsins and their endogenous inhibitors (158, 166). Moreover, partial LMP may trigger a cytoprotective lysophagy response and cell survival if the degree of damage is sufficiently limited (167, 168). Lysosomal stress sensors activate endolysosomal damage-response mechanisms (163) whereby injured lysosomes are eliminated and recycled before the cell is committed to die. A greater understanding of the precise lysosomal membrane alterations leading to LMP is required to elucidate consequent cell fate determinations.

LMP is most widely studied in relation to caspase-mediated apoptosis-like death. Select cathepsins that remain functional at neutral pH, including cathepsins B, D, and L, can activate apoptotic effectors following limited release from leaky lysosomes (166). Indeed, cathepsins are implicated in apoptotic cancer cell death in a variety of tumor models (169). Apoptotic pathways triggered by intrinsic factors such as DNA damage, endoplasmic reticulum stress, and LMP ultimately converge on mitochondrial membrane permeabilization (MOMP) and subsequent release of pro-apoptotic factors to the cytosol (166, 170–172). Mechanistic understanding of primary LMP in apoptosis was developed largely from studies using the lysosomotropic agent Leu-Leu-methyl ester (LLOMe) (173) and other apoptotic stimuli (153). Following their cytosolic release, cathepsins can cleave Bid to generate a pro-apoptotic t-Bid fragment, thereby initiating the common intrinsic apoptosis pathway that includes t-Bid activation of pore-forming Bax and Bak proteins, MOMP, mitochondrial cytochrome c release, and activation of executioner caspases (163, 173). In fact, cathepsin inhibition reduced Bid processing and alleviated LMP, rescuing cancer cell viability (174). Cathepsins can play a variety of other roles in LMP-mediated apoptotic death involving MOMP. Cathepsins amplify signaling upstream of MOMP via proteolytic Bax activation (165) or inactivation of anti-apoptotic Bcl-2 proteins (175), and they have been shown to cleave caspases directly (158) or degrade the caspase inhibitor XIAP (176). Of note, Bax may directly permeabilize the lysosomal membrane to initiate primary LMP (153, 177, 178). LMP has also been observed downstream of MOMP as a consequence of apoptotic signaling pathways (179, 180). Reactive oxygen species (ROS) production generated by MOMP induces lysosomal membrane lipid peroxidation and LMP to perpetuate apoptotic cell death (176), while various caspases themselves play causal roles in secondary LMP (158). Under certain conditions, cathepsins regulate apoptotic death independent of caspases, such as by direct cleavage of apoptosis inducing factor (AIF) (181) or in cells with defective apoptotic machinery (182).

LMP-mediated cell death can alternatively take the form of necrosis in the absence of caspase activation, whereby cathepsins serve as the principal cell death executioner proteases (63). The caspase dependence of LDCD pathways may shift depending on the cellular context or severity of cellular insult, as some lysosomotropic agents induce either apoptotic or necrotic cell death in a dose-dependent manner or in various cell types (173). Increased levels of oxidative stress or ATP depletion have been reported to promote a necrotic LDCD phenotype in several studies (166, 171). Necrosis, long considered an accidental and irreparable consequence of extreme chemical or physical cell stress, is now understood to be a highly regulated process with defined molecular drivers (183). Though cathepsin substrates in non-apoptotic death are not well characterized (63, 158), it is proposed that extensive LMP unleashes widespread cathepsin proteolysis and rapid breakdown of cellular structures, as cathepsin inhibition can mitigate necrotic LDCD (184–187). LMP may be an early and activating event in response to lysosome disruptors such as H₂O₂ (154), though it is also observed as a late-stage consequence of signaling in receptor interacting protein (RIP) kinase-dependent necroptosis (154). Furthermore, lysosomal ROS generation has been implicated in the execution of ferroptosis, a form of regulated cell death involving iron-dependent ROS accumulation which displays necrotic morphology (153, 188). Lysosomes degrade iron-containing proteins including ferritin during autophagy and serve as major storage sites of chelatable iron within the cell. Overloaded iron can catalyze Fenton reactions in redox cycling to produce ROS, which damage lysosomal membranes and increase the cell’s susceptibility to LMP (164).

Lysosomal disruption is critical to activation of the nod-like receptor (NLR)-dependent ‘inflammasome’ in pyroptosis, an inflammatory cell death pathway observed in macrophages that culminates in cellular swelling, plasma membrane rupture, and cytokine release. Pyroptosis is characterized by recruitment and activation of pro-inflammatory caspase-1 by the multimodular inflammasome platform and is mediated by gasdermins, which form plasma membrane pores to drive lytic death (189, 190). Various crystals and chemical compounds induce LMP-mediated pyroptosis, and LMP is reportedly critical to NLRP3 inflammasome function, though the precise molecular pathway linking LMP and lysosomal content release with inflammasome activation is debated (191, 192). Inflammasome activation was shown to potentiate tumor invasion and stimulate angiogenesis in cases where suppressive immune cells were favorably recruited to the tumor site, such as in the absence of IL-12 (193). The pro- and anti-tumor functions of inflammasomes may thus be context-dependent, reflected by responses of NLRP3 that differ significantly depending on cell lineage (e.g., hematopoietic vs. structural epithelium) or phenotype (193).

Lysosomes serve a principal function in autophagy, an adaptive cellular stress response that is normally cytoprotective but contributes to cell death in many pathophysiological conditions including cancer (153). During autophagy, the cell digests and recycles macromolecules and whole organelles by forming double membrane-bound autophagosomes that deliver engulfed material to lysosomes (194). Considering the requirement for functional lysosomes in autophagy execution, it is perhaps unsurprising that lysosomal damage can prevent autophagosome fusion and dysregulate autophagic flux, precipitating cell death (195, 196). Cell death is commonly the consequence of experimental or pharmacological autophagy blockade, and targeting autophagic processes has emerged as a promising therapeutic strategy for treatment of diseases including cancer (197, 198). For example, lysosome-disrupting chloroquine derivatives kill tumor cells by inhibiting autophagosome-lysosome fusion and chemotherapy-induced autophagy and demonstrate promise in clinical trials (199–201), while an aborted autophagic response following co-administration of lovastatin and farnesyl transferase inhibitor leads to non-apoptotic tumor cell death such that protein prenylation may be required for complete autophagy (202). Similarly, LMP may occur after inhibition of autophagic flux by a sophoridine analog, leading to apoptosis in pancreatic cancer cells (203). However, autophagy regulation is complex, and plays roles in cell survival, cell death, and other cytotoxic processes in a number of developmental and disease contexts (153, 204). In fact, LMP and cysteine cathepsin activity have been implicated in autophagy regulation and facilitate autophagy-mediated apoptosis in cancer cells (205–207). Indeed, numerous cell death mechanisms involve lysosome dysfunction, but the molecular interactions underlying phenotypic consequences of LMP remain largely uncharacterized.

LYSOSOMES IN CANCER

As cancer is broadly characterized by rapid cell proliferation and upregulated cell survival mechanisms to combat cellular damage, many transformation-associated changes at the level of the lysosome serve to protect the cell from LMP and LDCD. Paradoxically, the opposite effect is also observed, where neoplastic cells can sacrifice lysosomal stability in order to increase their tumorigenic potential or aggressiveness (24). Therefore, it is critical to understand the complex regulation of LMP in cancer cells in order to pharmacologically hedge the balance towards LDCD for therapeutic benefit.

Cancer cells rely on increased metabolism to sustain their rapid proliferation. The lysosome serves as a key regulator in this process and helps satisfy the catabolic need for building blocks for growth and neoplastic anabolic drive. The outer lysosomal membrane serves as a docking site for mTORC1, a signaling complex regulated by available nutrients in the cytoplasm. mTORC1 signals the upregulation of lipid and protein biosynthesis, as well as the transcription of pro-tumorigenic and anti-apoptotic regulators of the cell cycle (102, 208). Loss of several tumor suppressors including p53, PTEN, NF1 and TSC1/2 have been shown to activate mTORC1 (209). Furthermore, the lysosome is responsible for executing autophagy, one of the cell’s primary mechanisms of catabolism (210). Several cancers have engaged mechanisms to constitutively activate autophagy, such as in Ras-driven pancreatic cancers. This increased baseline level of autophagic flux allows for more rapid clearance of toxic metabolites that build up as a consequence of increased metabolism (211–213). Therefore, inhibition of autophagy could potentiate induction of LMP in such tumors.

The high metabolic activity of cancer cells presents another weakness in lysosomal regulation. Swift protein turnover demanded by rapidly dividing cancer cells leads to excessive intralysosomal accumulation of iron (214). Iron accretion alone is sufficient to sensitize the lysosome to LMP; however, Fenton-type reactions with H₂O₂ can generate additional ROS and further destabilize lysosomal membranes through lipid oxidation. Coupled with the well-documented increase in ROS production in cancer cells and augmented cytoplasmic levels of cathepsins, lysosomal iron accumulation sensitizes tumor cells to LMP and LDCD (215). Other transformation-associated alterations in cancer cell lysosomes include increases in expression of lysosomal enzymes, changes in lysosomal morphology and localization, and modifications in lysosome-associated proteins. Cathepsins are highly expressed in cancer cells and are localized to the cell periphery in secretory lysosomes, where they can be secreted into the surrounding extracellular environment through a mechanism similar to the lysosomal exocytosis that allows the formation of invasive protrusions in C. elegans development (216). Once in the extracellular space, they wreak havoc by cleaving a variety of adhesion proteins, degrading the basement membrane, and releasing sequestered growth factors such as platelet-derived growth factor (PDGF) and vascular endothelial growth factor (VEGF), leading to neoplastic progression through invasion, angiogenesis, and metastasis (41, 217, 218). Additionally, downregulation of a potent regulator of pericellular cathepsin accumulation, M6PR, in a rat cell model of hepatocellular carcinoma abrogates inhibition of pro-neoplastic cathepsin activity (219). High concentrations of cathepsins predict increased tumor aggressiveness and poor prognosis in many tumor types such as breast cancer, lung and colorectal carcinomas, and gliomas (220–223). However, increased levels of cytosolic cathepsins also sensitize the cell to LDCD (224). Increased lysosomal cathepsin activity leads to decreased LAMP-1 and -2 levels in ERK-, ErbB2-, and K-Ras-driven models of cancer, reducing lysosomal membrane stability and rendering the organelle susceptible to LMP (114). These cancer-associated vulnerabilities illuminate a potential therapeutic window to selectively target tumor cells.

As previously noted, lysosomes are alternatively localized within transformed cells; the switch from a perinuclear position to the plasma membrane facilitates secretion of toxic contents into the extracellular space, promoting extracellular acidification and activation of secreted lysosomal hydrolases (41). Increased expression of v-ATPase in metastatic tumor cells also contributes to extracellular acidification (225). Additionally, cancer cells exhibit increased lysosomal size, a phenomenon correlated with the metastatic potential of breast cancer cells (28). Once again, these tumorigenic changes compromise lysosomal stability, sensitizing tumor cells to LMP (28). Problematically, some forms of cancer have developed mechanisms to overcome this increased sensitivity to LMP. Breast cancer cells show elevated expression of Hsp70 (226), which has been shown to rescue lysosomal membrane integrity by stabilizing intralysosomal aSMase interaction with the critical lipase cofactor BMP (30, 215). Likewise, mammary-derived growth inhibitor (MDGI), which contributes to maintenance of lysosomal integrity, is a marker of invasiveness in human glioblastoma patient-derived cells that are resistant to chemoradiation. Silencing MDGI leads to alterations in lysosomal membrane lipid composition through reduced trafficking of polyunsaturated fatty acids into the lysosomes, leading to eventual LMP-dependent cell death (227). Along these lines, targeting Hsp70 or other molecules critical to lysosome stability and function could prove to be promising therapeutic strategies.

Localization of transport proteins to the lysosomal membrane in transformation confers therapeutic resistance via active sequestration and inactivation of antineoplastics within the lysosomal lumen. Canonically, members of the ATP-binding cassette (ABC) family of transporters reside in the plasma membrane where they expel cytoplasmic antineoplastics, but some cancers (e.g., leukemia, breast and cervical cancers) exhibit lysosomal expression of transporters like P-glycoprotein (P-gp or ABC1), leading to drug sequestration in the lysosome (228–230). Inhibition of P-gp in cancer cells restores sensitivity to the sequestered drugs and hyper-sensitizes cells to chemotherapeutic death (228, 230). Overexpression of ABCA3 correlates with poor prognosis in acute myeloid leukemia patients, and ABCA3 localizes to lysosomes in a chronic myeloid leukemia cell line (231, 232). The transport protein ATP7B, a copper transporter, is overexpressed in many cancers (233) and serves to sequester and exocytose platinum-based antineoplastics (234, 235). Overall, the dynamic changes in cancer-associated lysosomes reveals a wide range of possible therapeutic options for exploration (236).

MODULATION OF LMP AND ITS THERAPEUTIC POTENTIAL

Depletion of cancer cells via LMP is an attractive therapeutic strategy, holding particular promise for combating apoptosis-resistant cancer cell populations (237). Initial interest in this regard was sparked by realization of the degradative potential of lysosomes (1), catalyzing the search for pharmacologic agents that would destabilize the lysosomal membrane to kill cancer cells from the inside. Cholesterol and hydrocortisone were identified as stabilizing agents (32), while weakly basic amines with long hydrophobic tails disturbed membrane structure and induced LMP (238–240). However, enthusiasm quickly waned upon recognition that lysosomes were a ubiquitous feature of nearly all cells (excluding erythrocytes (34)), and would not permit the distinction between normal and cancer cells (32). Interest has since reignited following more recent studies that suggest cancer-associated lysosomes express unique features (i.e., size, hydrolase content, membrane fragility) that may make them suitable for targeting. Numerous lysosome-disrupting agents are currently under investigation or in clinical development for cancer and other indications (237, 241), though clinical data demonstrating efficacy of these approaches is limited. This section outlines select agents known to induce LMP, with a particular emphasis on their potential as anti-cancer therapeutics. General mechanisms of LMP induction are illustrated in Figure 2.

Figure 2. Mechanisms of LMP induction. (A) The lysosomal membrane is expanded to show detail. Carbohydrates on lysosomal associated membrane proteins (LAMPs) and lysosomal integral membrane proteins (LIMPs) form the protective glycocalyx. Also, resident within lysosomal membranes are ion channels and the vacuolar ATPase (v-ATPase) that maintains the acidic interior of the lysosome. A number of hydrolases are found within the lysosomal lumen. Dysregulation of these endogenous factors can contribute to LMP. (B) LMP-inducing agents including many lysosomotropic detergents and cationic amphiphilic drugs (CADs) contain hydrophilic side chains with charged cationic amine groups, allowing them to passively partition across cellular membranes in their non-ionized state. Within the acidic lysosomal lumen, these agents become protonated and sequestered in a process termed ‘lysosomal trapping.’ (C) The degree of LMP often dictates the ensuing course of cell death (apoptosis or necrosis). (D) Expansion of the lysosomal membrane to show details of lipid constituents: the lysosomal membrane is composed of phospholipids, glycerophospholipids such as bis(monoacylglycero)phosphate (BMP), cholesterol, and sphingolipids such as sphingomyelin and ceramide. Sphingomyelin is hydrolyzed to ceramide by acid sphingomyelinase (aSMase). Positively charged amino acids on aSMase allow it to interact with negatively charged head groups on lipids such as phosphatidylcholine or phosphatidylserine. aSMase is displaced by agents that interfere with this binding domain. (E) Mitochondrial reactive oxygen species (ROS) production stimulates lipid peroxidation and LMP. Lysosomal membrane peroxidation is augmented by production of reactive hydroxyl radicals generated through degradation of iron-containing molecules within the lysosomal lumen in the presence of reducing agents.

Lysosomotropic Agents

Weakly basic amine compounds rapidly accumulate within lysosomal lumens and are thus referred to as ‘lysosomotropic’ (242). Sequestration of amine-containing agents occurs by a non-enzymatic and non-transporter mediated cation-trapping mechanism, referred to as ‘lysosomal trapping’ (44, 243, 244), but may also occur by endocytosis or facilitated transport (245). In particular, CADs and lysosomotropic detergents feature a hydrophobic ring structure and a hydrophilic side chain with a charged cationic amine group that allow them to readily diffuse across cellular membranes in their non-ionized state. However, exposure to the acidic interior of the lysosome leads to protonation and entrapment (242). As these agents accumulate within the lysosomal lumen, they interact with negatively charged intra-lysosomal vesicles, displacing associated enzymes and lipid binding proteins and inducing swelling and vacuolization due to an influx of water into the lysosomal lumen (242). The appearance of lamellar bodies that occurs in response to some CADs (43) signals the accumulation of lipids within the lysosome that may occur following the interaction of drugs with phospholipids (246) or via inhibition of phospholipid metabolism (247, 248). The anti-tumor efficacy of CADs, including FDA-approved anti-histamines (227), anti-depressants (249), and anti-malarials (250), has been documented in both experimental and observational studies. The CAD 5-(N,N-hexamethylene) amiloride (HMA) showed robust induction of necrotic cell death in breast cancer cells regardless of molecular profile, with little toxicity against untransformed cells (184). Another CAD, the anti-histamine clemastine, killed patient-derived glioblastoma cells by LMP but was minimally toxic in normal human astrocytes and murine brain endothelial cells, pointing to a critical therapeutic window for the treatment of this highly aggressive and chemo-refractory disease (227). Lysosomotropic detergents, which combine weakly basic amines (e.g., imidazole, morpholine) with long (9–14 carbons) and straight hydrocarbon tails, are characterized by membrane disruptive surfactant properties that progress with continued lysosomal accumulation (238–240). Derivatives of imidazole and morpholine lysosomotropic detergents were originally developed as anticancer therapeutics, inducing apoptosis or necrosis with LMP in a dose-dependent fashion across a range of cancer cell types (239, 251–253).

Interestingly, basic amines are a ubiquitous feature of therapeutic agents, conferring varying degrees of lysosomotropic potential (254). As such, lysosomal drug sequestration may either be cytotoxic or cytoprotective depending upon whether it potentiates LMP or prohibits interactions with the intended target. Indeed, substantial evidence supports the notion that the lysosome contributes to chemotherapeutic resistance (255–258). In one such mechanism, cellular stress resulting from accumulation of lysosomotropic drugs was shown to trigger exocytosis, leading to lysosome-mediated multidrug resistance (259). Furthermore, the degree of resistance to the topoisomerase II inhibitor C-1330 and the receptor tyrosine kinase inhibitor sunitinib was directly associated with the total number of lysosomes (258) and, perhaps more importantly, the degree to which the normal cytosol-to-lysosome pH gradient is altered within a given cancer cell (260). Similar to other anti-cancer therapeutics (doxorubicin, mitoxantrone), C-1330 and sunitinib were shown to preferentially accumulate within lysosomes, triggering substantial, dose-dependent increases in lysosome number, size, and their ability to uptake the lysosomal marker LysoTracker Red (258). Poor lysosomal accumulation and retained drug sensitivity was associated with intrinsic disruption of the physiologic pH of some cancer cell lysosomes (i.e., MCF7 breast cancer cells) (260). Accordingly, diminished lysosomal entrapment of weakly basic amines could be replicated with pharmacological disruption of the pH gradient following v-ATPase inhibition by bafilomycin A (255, 260) or administration of the lysosomotropic agent chloroquine (255), suggesting that modulation of lysosomal pH may be an effective strategy to overcome chemoresistance.

Indeed, the pH-disrupting agents chloroquine and hydroxychloroquine have been investigated as anti-cancer therapeutics, with dozens of clinical trials in progress (241, 261–265). Long used as an antimalarial, chloroquine sensitizes cancer cells to radiation and chemotherapy. Although thought to convey therapeutic sensitivity through the inhibition of protective autophagy, recent evidence suggests its role may be more complicated (266–268), and may at least partially involve the capacity to overcome drug sequestration within lysosomes (255, 269, 270). Nevertheless, as a nonspecific inhibitor of autophagy, potential side effects may arise from a loss of protective autophagy within normal tissues (i.e., brain, liver, heart, kidney) that occurs during therapeutic intervention (271). Along these lines, chloroquine-treated mice are more likely to suffer from kidney damage in a model of ischemic–reperfusion than untreated animals (272). Chloroquine uptake may also be reduced in the external acidic milieu of some tumors, reducing its efficacy under these conditions (273). However, particular derivatives of chloroquine (273) or other small molecule inhibitors of autophagy (264) may offer improved stability and potency and warrant further study.

Nanoparticles

While nanoparticles have been extensively investigated for efficient tumor-site delivery of anti-cancer drugs in recent years, issues of non-specific cell toxicity are often attributed to lysosome dysfunction, as nanoparticles can accumulate in lysosomes and induce LDCD themselves. However, the precise mechanisms underlying nanoparticle toxicity are debated, with the involvement of autophagy in question in various cancers (274, 275). Several studies reported that early induction of autophagy allowed rapid nanoparticle uptake and delivery to lysosomes—a requirement for zinc oxide nanoparticle (ZnONP)-mediated cytotoxicity, i.e., nanoparticle dissolution and content release within the lysosomal lumen (276)––but that subsequent lysosomal damage resulting from nanoparticle buildup impaired autophagic flux and ultimately resulted in cell death (276, 277). Moreover, ZnONP cytotoxicity was not attributed to nanoparticle dissolution and zinc ion release within lysosomes in cellular models of leukemia and normal red blood cells but rather to LMP triggered by intact nanoparticles (278). A variety of nanoparticle formulations are currently under investigation as novel lysosome-disrupting cancer therapeutics (279, 280). Interestingly, a recent report demonstrated that nanoparticles carrying small interfering RNA (siRNA) therapeutics can become sequestered in lysosomes and exhibit inefficient endolysosomal escape (281), while CAD administration promotes LMP and cytosolic siRNA delivery.

Sphingolipids

Mounting evidence suggests that LMP occurs following specific changes in the composition of membrane lipids and major lysosomal proteins (114). Particularly interesting data has recently come to light suggesting that cancer cells have perturbed lipid species as compared to their normal counterparts, a feature that may permit their selective depletion. Along these lines, the lysosomotropic detergent siramesine and similar compounds directly induce LMP and non-apoptotic cell death in transformed cells but not in oncogene-depleted (‘detransformed’) or non-transformed variants (29, 282). These effects were universal across all cancer types tested (breast, ovary, prostate, cervix and bone (29)). Intriguingly, similar cancer specific effects were not found for other compounds that also induce LMP (i.e., LLOMe, sphingosine) or neutralize pH (concanamycin A) (29), although the lack of specificity may be related to the dose used (283).

Ostensibly, siramesine cytotoxicity is mediated by the displacement of aSMase (29), normally responsible for the breakdown of sphingomyelin to ceramide at the inner lysosomal membrane (134). Augmentation of aSMase activity occurs in stressed normal cells following induction of heat shock protein 70 (Hsp70), which then binds the glycerophospholipid BMP to activate aSMase at the lysosomal membrane (30). The constitutive elevation of Hsp70 has been detailed for a number of cancer types, and is associated with resistance to caspase-dependent and -independent cell death and poor prognosis (284–287). Bolstered aSMase activity appears to contribute to lysosomal integrity (29), making it a particularly attractive therapeutic target. Accordingly, exposure to siramesine (29) or Hsp70 small molecule inhibitors (288) results in reductions in aSMase activity, induction of LMP, caspase-independent cell death and enhanced sensitivity to chemotherapeutics. In agreement with siramesine-mediated aSMase inhibition, depleted Hsp70 induces effects that are cancer cell specific (286, 289–292). It is likely that aSMase inhibition perturbs lipid ratios (293), leading to membrane fragility and a propensity for LMP (29). This postulate is further supported by studies suggesting that augmented sphingomyelin content alone destabilizes lysosomal membranes (30), is selectively toxic to transformed cells (294), inhibits autophagic flux (295), and impairs intracellular vesicle and plasma membrane fusion events (296). Destabilized lipid content following aSMase inhibition also affects signaling events at the plasma membrane, including the clustering and signaling of K-Ras (293), a protein with a dominant role in cell proliferation and survival. These data suggest that aSMase inhibition may not only be cancer cell specific (29), but may also be particularly well suited for K-Ras-driven cancers (i.e., pancreas, colon and lung), which currently lack targeted therapeutic options (297).

The clinical significance of sphingolipid species is further substantiated by observations that perturbed ceramide clearance was directly correlated with reduced chemotherapeutic sensitivity (298). As such, modification of ceramide levels by enhancing de novo biosynthesis or modulating aSMase activity have also been suggested as potential anti-cancer strategies to overcome imbalances in lysosomal ceramide (298–301). Although a clear link has yet to be established, a recent study demonstrated that markedly enhanced aSMase potentiates the accumulation of ceramide and triggers cathepsin B release via LMP upstream of apoptosis (176). Importantly, cathepsin B catalyzes the degradation of XIAP (X-linked inhibitor of apoptosis) (176), the upregulation of which has been associated with therapeutic resistance and poor survival (302–304).

It is worthwhile to note that experimental modulation of aSMase activity appears to produce seemingly contradictory results. On the one hand, the depletion of aSMase results in sphingolipid accumulation, ceramide depletion, LMP, and non-apoptotic cell death (29), whereas on the other hand its overexpression precipitates ceramide accumulation, LMP, and apoptosis (176). As such, it is likely that lysosome integrity depends upon the precise balance of sphingolipid species. Moreover, it is conceivable that there are factors modified under conditions of aSMase depletion or overexpression that await further investigation. For example, sphingosine accumulation inhibits cholesterol export (305), which may have implications for signaling events at lysosome-associated lipid raft domains or in membrane dynamics. Moreover, modifications to sphingosine or ceramide may generate either sphingosine-1-phosphate or sphingosine, respectively. Sphingosine-1-phosphate binds to G-coupled protein receptors to regulate growth, survival and migration of cells, and is associated with malignant transformation (306). Sphingosine was shown to induce LMP and programmed cell death (307–309), albeit nonspecifically (29). It was proposed that accumulated sphingosine permeabilizes the membrane in a detergent-like fashion, resulting in cell death (310). A potential mechanism is outlined in Figure 3 that discusses the intricate balance of sphingolipids in LMP induction.

Figure 3. Membrane stabilization and permeabilization is dependent on the balance of sphingolipids in lysosomes. Negatively charged bis(monoacylglycero)phosphate (BMP; shown in teal) is enriched in membranes of intraluminal lysosomal vesicles and provides docking points for positively charged acid sphingomyelinase, acid ceramidase, and other co-factors such as heat shock protein 70 (HSP70) and saposins (SAP). Proper recruitment of these enzymes facilitates lipid metabolism and membrane stabilization. Cationic amphiphilic drugs (CADs) disrupt BMP-enzyme interactions, inhibiting lipid metabolism and resulting in lysosomal membrane permeabilization. Figure created with BioRender.com.

Calcium

Interestingly, sphingosine was also shown to affect calcium release from the lysosomal membrane (311). Although not often considered a major calcium storage site, the reported calcium concentration of 500μM within the lysosomal lumen (312) is comparable with that of the endoplasmic reticulum (313). In studies addressing the pathology of Niemann-Pick disease type C (NPC), a rare progressive disorder characterized by the accumulation of sphingolipid species and cholesterol, sphingosine was shown to induce the specific release of calcium from lysosomes and late endosomes but not release from other intracellular storage sites (311). This rise in cytosolic calcium was dependent on interaction of sphingosine with lysosome and endosome calcium channels (two-pore channel 1) (311). Lysosome-released calcium is vital for vesicle fusion and secretion, autophagy, and lysosomal biogenesis (39, 314–316), as well as apoptosis-dependent phosphatidylserine externalization (317), an early recognition signal for engulfment by phagocytic cells (318, 319). Intriguingly, prolonged endoplasmic reticulum stress results in pancreatic cancer cell death mediated by LMP and accumulation of cytosolic calcium (320). General increases in intracellular calcium are also associated with the activation of cytosolic calpain proteases and the initiation of LMP. Along these lines, μ-calpain is capable of permeabilizing the membrane of isolated lysosomes (321). Further, activated calpain species were shown to localize to the lysosomal membrane prior to cathepsin release (186, 322, 323), whereas their pharmacological inhibition effectively abolishes LMP (324).

Reactive oxygen species

Increased production of ROS (i.e., singlet oxygen and hydrogen peroxide) stimulates lipid peroxidation and destabilization of the lysosomal membrane (325–327). Cell death depends on the degree of LMP, where minimal leakage of cathepsins is nonlethal, reversible (328), and primarily impacts cell proliferation, while moderate or high membrane destabilization respectively induce apoptosis and necrosis (170). Peroxidation of the lysosomal membrane is likely mediated by local production of highly reactive hydroxyl radicals. Since iron-containing macromolecules are degraded within the lysosomal lumen in the presence of reducing agents (i.e., glutathione, ascorbic acid and cysteine), there is a capacity to generate reactive radicals with exposure to H₂O₂ (329). Accordingly, iron chelation is protective against LMP whereas accumulation of iron-containing proteins or iron complexes sensitizes lysosomes to membrane damage (330–336). Cancer-associated lysosomes may have a heightened propensity to accumulate iron, given the enhanced turnover of iron-containing proteins that accompanies their rapid proliferation (170). Consequently, cancer cells may be more sensitive to ROS-induced LMP (214). Iron has also been implicated in maintenance of the cancer stem cell (CSC) state, and salinomycin selectively kills cancer stem cells by sequestering iron in lysosomes, leading to ROS-mediated LMP and ferroptotic cell death (337). Unsurprisingly, aSMase was implicated in ferroptosis initiation through a positive feedback loop of ROS production, further connecting key lysosomal enzymes to ferroptosis (338).

While iron-loading has not been vetted as an anti-cancer strategy, such an approach may offer particular benefit to hypoxic regions of tumors where limited ROS production might contribute to insensitivity to LMP (329). Iron loading of hypoxic cells may sensitize them to ionizing radiation or other ROS-generating agents. On the other hand, redox-active iron has been suggested as a mutagen, contributing to persistent oxidative stress of DNA damage (339–341). Iron-chelation also has important clinical applications, protecting normal tissues from radiation damage during cancer treatment (342). An exception is CAD iron-chelating compounds, as they remain trapped within lysosomes and lead to iron-starvation and death (343, 344).

CONCLUSION

The lysosome is clearly more than a cellular garbage disposal, contributing to dynamic processes that are essential to both normal physiologic function and disease pathology. Cancer-associated changes in lysosomal structure and function may bolster therapeutic resistance but may also be the gateway to cancer cells’ ultimate demise. Indeed, emerging data suggests the lysosome is potentially a powerful anti-cancer target, given specific alterations in cancer cells that are not seen in non-transformed counterparts. Moreover, cancer cells frequently evade therapy-induced apoptosis due to intrinsic or acquired mutations in caspase-dependent pathways; thus, utilization of LDCD and other caspase-independent cell death programs will be critical in the future development of cancer therapeutics. Nevertheless, a number of questions still remain with respect to cancer-associated lysosome signaling, membrane dynamics, ion regulation, and the precise role of LMP in cell death, which have limited the number of lysosome-targeted therapeutics transitioning from preclinical study to clinical development. The dual functionality of cathepsin activation in cancer cell invasion and LDCD is a critical area requiring further investigation, as cathepsins are important mediators of both pro-tumorigenic and pro-death processes, depending on context. Substantial progress on these and other fronts will likely be fueled by continued advances in methods for the detection and quantification of lysosome-associated events.

Acknowledgement: This work was supported by NIH grant R01 CA250211 (K.L.C.).

Conflict of Interest: The authors declare no potential conflict of interest with respect to research, authorship and/or publication of this chapter.

Copyright and Permission Statement: The authors confirm that the materials included in this chapter do not violate copyright laws. Where relevant, appropriate permissions have been obtained from the original copyright holders, and all original sources have been appropriately acknowledged or referenced.

REFERENCES

1. de Duve C, Pressman BC, Gianetto R, Wattiaux R, Appelmans F. Tissue fractionation studies. 6. Intracellular distribution patterns of enzymes in rat-liver tissue. Biochemical Journal. 1955;60(4):604–17. https://doi.org/10.1042/bj0600604
2. de Duve C. The lysosome turns fifty. Nat Cell Biol. 2005;7(9):847–9. https://doi.org/10.1038/ncb0905-847
3. Bowers WE. Christian de Duve and the discovery of lysosomes and peroxisomes. Trends Cell Biol. 1998;8(8):330–3. https://doi.org/10.1016/S0962-8924(98)01314-2
4. Ballabio A. The awesome lysosome. EMBO Mol Med. 2016;8(2):73–6. https://doi.org/10.15252/emmm.201505966
5. Schulze H, Kolter T, Sandhoff K. Principles of lysosomal membrane degradation: Cellular topology and biochemistry of lysosomal lipid degradation. Biochim Biophys Acta. 2009;1793(4):674–83. https://doi.org/10.1016/j.bbamcr.2008.09.020
6. Kolter T, Sandhoff K. Lysosomal degradation of membrane lipids. FEBS Lett. 2010;584(9):1700–12. https://doi.org/10.1016/j.febslet.2009.10.021
7. Reddy A, Caler EV, Andrews NW. Plasma Membrane Repair Is Mediated by Ca2+-Regulated Exocytosis of Lysosomes. Cell. 2001;106(2):157–69. https://doi.org/10.1016/S0092-8674(01)00421-4
8. Gerasimenko JV, Gerasimenko OV, Petersen OH. Membrane repair: Ca(2+)-elicited lysosomal exocytosis. Curr Biol. 2001;11(23):R971–4. https://doi.org/10.1016/S0960-9822(01)00577-2
9. Rath NC, Hand AR, Reddi AH. Activity and distribution of lysosomal enzymes during collagenous matrix-induced cartilage, bone, and bone marrow development. Dev Biol. 1981;85(1):89–98. https://doi.org/10.1016/0012-1606(81)90238-4
10. Miller NR, Wolfe HJ. The nature and localization of acid phosphatase during the early phases of urodele limb regeneration. Dev Biol. 1968;17(4):447–81. https://doi.org/10.1016/0012-1606(68)90074-2
11. Bowen ID, den Hollander JE, Lewis GHJ. Cell Death and Acid Phosphatase Activity in the Regenerating Planarian Polycelis tenuis Iijima. Differentiation. 1982;21(1–3):160–7. https://doi.org/10.1111/j.1432-0436.1982.tb01209.x
12. Bowen ID, Lewis GHJ. Acid phosphatase activity and cell death in mouse thymus. Histochemistry. 1980;65(2):173–9. https://doi.org/10.1007/BF00493166
13. Honey K, Rudensky AY. Lysosomal cysteine proteases regulate antigen presentation. Nat Rev Immunol. 2003;3(6):472–82. https://doi.org/10.1038/nri1110
14. Maeda FY, Van Haaren JJH, Langley DB, Christ D, Andrews NW, Song W. Surface-associated antigen induces B-cell permeabilization and lysosome exocytosis facilitating antigen uptake and presentation to T-cells. 2020. https://doi.org/10.1101/2020.07.24.220418
15. He Y, Xu Y, Zhang C, Gao X, Dykema KJ, Martin KR, et al. Identification of a lysosomal pathway that modulates glucocorticoid signaling and the inflammatory response. Science signaling. 2011;4(180):ra44. https://doi.org/10.1126/scisignal.2001450
16. Sancak Y, Bar-Peled L, Zoncu R, Markhard AL, Nada S, Sabatini DM. Ragulator-Rag complex targets mTORC1 to the lysosomal surface and is necessary for its activation by amino acids. Cell. 2010;141(2):290–303. https://doi.org/10.1016/j.cell.2010.02.024
17. Schahs P, Weidinger P, Probst OC, Svoboda B, Stadlmann J, Beug H, et al. Cellular repressor of E1A-stimulated genes is a bona fide lysosomal protein which undergoes proteolytic maturation during its biosynthesis. Exp Cell Res. 2008;314(16):3036–47. https://doi.org/10.1016/j.yexcr.2008.06.015
18. Bagshaw RD, Mahuran DJ, Callahan JW. A proteomic analysis of lysosomal integral membrane proteins reveals the diverse composition of the organelle. Mol Cell Proteomics. 2005;4(2):133–43. https://doi.org/10.1074/mcp.M400128-MCP200
19. Appelqvist H, Wäster P, Kågedal K, Öllinger K. The lysosome: from waste bag to potential therapeutic target. J Mol Cell Biol. 2013;5(4):214–26. https://doi.org/10.1093/jmcb/mjt022
20. Reinheckel T, Deussing J, Roth W, Peters C. Towards specific functions of lysosomal cysteine peptidases: phenotypes of mice deficient for cathepsin B or cathepsin L. Biol Chem. 2001;382(5):735–41. https://doi.org/10.1515/BC.2001.089
21. Lawrence RE, Zoncu R. The lysosome as a cellular centre for signalling, metabolism and quality control. Nature cell biology. 2019;21(2):133–42. https://doi.org/10.1038/s41556-018-0244-7
22. Hesketh GG, Wartosch L, Davis LJ, Bright NA, Luzio JP. The Lysosome and Intracellular Signalling. Prog Mol Subcell Biol. 2018;57:151–80. https://doi.org/10.1007/978-3-319-96704-2_6
23. Saftig P, Puertollano R. How Lysosomes Sense, Integrate, and Cope with Stress. Trends in Biochemical Sciences. 2021;46(2):97–112. https://doi.org/10.1016/j.tibs.2020.09.004
24. Kallunki T, Olsen OD, Jaattela M. Cancer-associated lysosomal changes: friends or foes[quest]. Oncogene. 2013;32(16):1995–2004. https://doi.org/10.1038/onc.2012.292
25. Gocheva V, Joyce JA. Cysteine Cathepsins and the Cutting Edge of Cancer Invasion. Cell Cycle. 2007;6(1):60–4. https://doi.org/10.4161/cc.6.1.3669
26. Allison AC. Lysosomes in cancer cells. Journal of Clinical Pathology Supplement (Royal College of Pathologists). 1974;7:43–50. https://doi.org/10.1136/jcp.27.Suppl_7.43
27. Jensen SS, Aaberg-Jessen C, Christensen KG, Kristensen B. Expression of the lysosomal-associated membrane protein-1 (LAMP-1) in astrocytomas. Int J Clin Exp Pathol. 2013;6(7):1294–305.
28. Glunde K, Guggino SE, Solaiyappan M, Pathak AP, Ichikawa Y, Bhujwalla ZM. Extracellular acidification alters lysosomal trafficking in human breast cancer cells. Neoplasia. 2003;5(6):533–45. https://doi.org/10.1016/S1476-5586(03)80037-4
29. Petersen N, Olsen O, Groth-Pedersen L, Ellegaard AM, Bilgin M, Redmer S, et al. Transformation-Associated Changes in Sphingolipid Metabolism Sensitize Cells to Lysosomal Cell Death Induced by Inhibitors of Acid Sphingomyelinase. Cancer Cell. 2013;24(3):379–93. https://doi.org/10.1016/j.ccr.2013.08.003
30. Kirkegaard T, Roth AG, Petersen NHT, Mahalka AK, Olsen OD, Moilanen I, et al. Hsp70 stabilizes lysosomes and reverts Niemann-Pick disease-associated lysosomal pathology. Nature. 2010;463(7280):549–53. https://doi.org/10.1038/nature08710
31. Ono K, Kim SO, Han J. Susceptibility of lysosomes to rupture is a determinant for plasma membrane disruption in tumor necrosis factor alpha-induced cell death. Mol Cell Biol. 2003;23(2):665–76. https://doi.org/10.1128/MCB.23.2.665-676.2003
32. de Duve C. Lysosomes revisited. Eur J Biochem. 1983;137(3):391–7. https://doi.org/10.1111/j.1432-1033.1983.tb07841.x
33. Puertollano R. mTOR and lysosome regulation. F1000Prime Reports. 2014;6. https://doi.org/10.12703/P6-52
34. Lüllmann-Rauch R. History and Morphology of the Lysosome. In: Saftig P, editor. Lysosomes. Medical Intelligence Unit. Boston, MA: Springer US; 2005. p. 1–16. https://doi.org/10.1007/0-387-28957-7_1
35. Holtzman E. Lysosomes. New York: Plenum Press; 1989. https://doi.org/10.1007/978-1-4899-2540-4
36. Bandyopadhyay D, Cyphersmith A, Zapata JA, Kim YJ, Payne CK. Lysosome Transport as a Function of Lysosome Diameter. PLoS One. 2014;9(1):e86847. https://doi.org/10.1371/journal.pone.0086847
37. Yu L, McPhee CK, Zheng L, Mardones GA, Rong Y, Peng J, et al. Termination of autophagy and reformation of lysosomes regulated by mTOR. Nature. 2010;465(7300):942–6. https://doi.org/10.1038/nature09076
38. Korolchuk VI, Saiki S, Lichtenberg M, Siddiqi FH, Roberts EA, Imarisio S, et al. Lysosomal positioning coordinates cellular nutrient responses. Nat Cell Biol. 2011;13(4):453–60. https://doi.org/10.1038/ncb2204
39. Settembre C, Di Malta C, Polito VA, Arencibia MG, Vetrini F, Erdin S, et al. TFEB Links Autophagy to Lysosomal Biogenesis. Science. 2011;332(6036):1429–33. https://doi.org/10.1126/science.1204592
40. Parkinson-Lawrence EJ, Shandala T, Prodoehl M, Plew R, Borlace GN, Brooks DA. Lysosomal Storage Disease: Revealing Lysosomal Function and Physiology. Physiology. 2010;25(2):102–15. https://doi.org/10.1152/physiol.00041.2009
41. Palermo C, Joyce JA. Cysteine cathepsin proteases as pharmacological targets in cancer. Trends Pharmacol Sci. 2008;29. https://doi.org/10.1016/j.tips.2007.10.011
42. Swanson J, Yirinec B, Burke E, Bushnell A, Silverstein SC. Effect of alterations in the size of the vacuolar compartment on pinocytosis in J774.2 macrophages. J Cell Physiol. 1986;128(2):195–201. https://doi.org/10.1002/jcp.1041280209
43. Lüllmann H, Lüllmann-Rauch R, Wassermann O. Lipidosis induced by amphiphilic cationic drugs. Biochem Pharmacol. 1978;27(8):1103–8. https://doi.org/10.1016/0006-2952(78)90435-5
44. Kazmi F, Hensley T, Pope C, Funk RS, Loewen GJ, Buckley DB, et al. Lysosomal Sequestration (Trapping) of Lipophilic Amine (Cationic Amphiphilic) Drugs in Immortalized Human Hepatocytes (Fa2N-4 Cells). Drug Metabolism and Disposition. 2013;41(4):897–905. https://doi.org/10.1124/dmd.112.050054
45. Funk RS, Krise JP. Cationic amphiphilic drugs cause a marked expansion of apparent lysosomal volume: Implications for an intracellular distribution-based drug interaction. Molecular Pharmaceutics. 2012;9(5):1384–95. https://doi.org/10.1021/mp200641e
46. Laurent G, Kishore BK, Tulkens PM. Aminoglycoside-induced renal phospholipidosis and nephrotoxicity. Biochem Pharmacol. 1990;40(11):2383–92. https://doi.org/10.1016/0006-2952(90)90078-Y
47. Korolchuk VI, Rubinsztein DC. Regulation of autophagy by lysosomal positioning. Autophagy. 2011;7(8):927–8. https://doi.org/10.4161/auto.7.8.15862
48. Heuser J. Changes in lysosome shape and distribution correlated with changes in cytoplasmic pH. J Cell Biol. 1989;108(3):855–64. https://doi.org/10.1083/jcb.108.3.855
49. Cardone RA, Casavola V, Reshkin SJ. The role of disturbed pH dynamics and the Na+/H+ exchanger in metastasis. Nature Reviews Cancer. 2005;5(10):786–95. https://doi.org/10.1038/nrc1713
50. Steffan JJ, Williams BC, Welbourne T, Cardelli JA. HGF-induced invasion by prostate tumor cells requires anterograde lysosome trafficking and activity of Na+-H+ exchangers. J Cell Sci. 2010;123(Pt 7):1151–9. https://doi.org/10.1242/jcs.063644
51. Ghadially FN. 7 - Lysosomes. Ultrastructural Pathology of the Cell and Matrix (Third Edition): Butterworth-Heinemann; 1988. p. 589–765. https://doi.org/10.1016/B978-0-407-01572-2.50008-3
52. Luzio JP, Pryor PR, Bright NA. Lysosomes: fusion and function. Nat Rev Mol Cell Biol. 2007;8(8):622–32. https://doi.org/10.1038/nrm2217
53. Saftig P, Klumperman J. Lysosome biogenesis and lysosomal membrane proteins: trafficking meets function. Nat Rev Mol Cell Biol. 2009;10(9):623–35. https://doi.org/10.1038/nrm2745
54. Morgan AJ, Platt FM, Lloyd-Evans E, Galione A. Molecular mechanisms of endolysosomal Ca2+ signalling in health and disease. Biochem J. 2011;439(3):349–74. https://doi.org/10.1042/BJ20110949
55. Bampton ET, Goemans CG, Niranjan D, Mizushima N, Tolkovsky AM. The dynamics of autophagy visualized in live cells: from autophagosome formation to fusion with endo/lysosomes. Autophagy. 2005;1(1):23–36. https://doi.org/10.4161/auto.1.1.1495
56. Conus S, Simon H-U. Cathepsins: key modulators of cell death and inflammatory responses. Biochem Pharmacol. 2008;76(11):1374–82. https://doi.org/10.1016/j.bcp.2008.07.041
57. Chapman HA, Riese RJ, Shi GP. Emerging roles for cysteine proteases in human biology. Annu Rev Physiol. 1997;59. https://doi.org/10.1146/annurev.physiol.59.1.63
58. Stahl S, Reinders Y, Asan E, Mothes W, Conzelmann E, Sickmann A, et al. Proteomic analysis of cathepsin B- and L-deficient mouse brain lysosomes. Biochim Biophys Acta. 2007;1774(10):1237–46. https://doi.org/10.1016/j.bbapap.2007.07.004
59. Gocheva V, Zeng W, Ke D, Klimstra D, Reinheckel T, Peters C, et al. Distinct roles for cysteine cathepsin genes in multistage tumorigenesis. Genes Dev. 2006;20(5):543–56. https://doi.org/10.1101/gad.1407406
60. Akkari L, Gocheva V, Quick ML, Kester JC, Spencer AK, Garfall AL, et al. Combined deletion of cathepsin protease family members reveals compensatory mechanisms in cancer. Genes Dev. 2016;30(2):220–32. https://doi.org/10.1101/gad.270439.115
61. Benes P, Vetvicka V, Fusek M. Cathepsin D-Many functions of one aspartic protease. Critical Reviews in Oncology/Hematology. 2008;68(1):12–28. https://doi.org/10.1016/j.critrevonc.2008.02.008
62. Brix K. Lysosomal Proteases. Lysosomes. Boston, MA: Springer US; 2005. p. 50–9. https://doi.org/10.1007/0-387-28957-7_5
63. Turk V, Stoka V, Vasiljeva O, Renko M, Sun T, Turk B, et al. Cysteine cathepsins: From structure, function and regulation to new frontiers. Biochimica et Biophysica Acta (BBA) - Proteins and Proteomics. 2012;1824(1):68–88. https://doi.org/10.1016/j.bbapap.2011.10.002
64. Joyce JA, Hanahan D. Multiple roles for cysteine cathepsins in cancer. Cell Cycle. 2004;3(12):1516–619. https://doi.org/10.4161/cc.3.12.1289
65. Taylor MA, Pratt KA, Revell DF, Baker KC, Sumner IG, Goodenough PW. Active papain renatured and processed from insoluble recombinant propapain expressed in Escherichia coli. Protein Eng. 1992;5(5):455–9. https://doi.org/10.1093/protein/5.5.455
66. Turk B, Turk D, Turk V. Lysosomal cysteine proteases: more than scavengers. Biochim Biophys Acta. 2000;1477(1–2):98–111. https://doi.org/10.1016/S0167-4838(99)00263-0
67. Turk V, Turk B, Turk D. Lysosomal cysteine proteases: facts and opportunities. EMBO J. 2001;20(17):4629–33. https://doi.org/10.1093/emboj/20.17.4629
68. Kominami E, Tsukahara T, Hara K, Katunuma N. Biosyntheses and processing of lysosomal cysteine proteinases in rat macrophages. FEBS Lett. 1988;231(1):225–8. https://doi.org/10.1016/0014-5793(88)80736-1
69. Nishimura Y, Kawabata T, Kato K. Identification of latent procathepsins B and L in microsomal lumen: Characterization of enzymatic activation and proteolytic processing in vitro. Archives of Biochemistry and Biophysics. 1988;261(1):64–71. https://doi.org/10.1016/0003-9861(88)90104-X
70. Rowan AD, Mason P, Mach L, Mort JS. Rat procathepsin B: Proteolytic processing to the mature form in vitro. J Biol Chem. 1992;267(22):15993–9. https://doi.org/10.1016/S0021-9258(19)49632-4
71. Mason RW, Massey SD. Surface activation of pro-cathepsin L. Biochemical and Biophysical Research Communications. 1992;189(3):1659–66. https://doi.org/10.1016/0006-291X(92)90268-P
72. Ishidoh K, Kominami E. Procathepsin L Degrades Extracellular Matrix Proteins in the Presence of Glycosaminoglycans in Vitro. Biochemical and Biophysical Research Communications. 1995;217(2):624–31. https://doi.org/10.1006/bbrc.1995.2820
73. Caglic D, Pungercar JR, Pejler G, Turk V, Turk B. Glycosaminoglycans facilitate procathepsin B activation through disruption of propeptide-mature enzyme interactions. J Biol Chem. 2007;282(45):33076–85. https://doi.org/10.1074/jbc.M705761200
74. Vasiljeva O, Dolinar M, Pungercar JR, Turk V, Turk B. Recombinant human procathepsin S is capable of autocatalytic processing at neutral pH in the presence of glycosaminoglycans. FEBS Lett. 2005;579(5):1285–90. https://doi.org/10.1016/j.febslet.2004.12.093
75. Costa MG, Batista PR, Shida CS, Robert CH, Bisch PM, Pascutti PG. How does heparin prevent the pH inactivation of cathepsin B? Allosteric mechanism elucidated by docking and molecular dynamics. BMC Genomics. 2010;11(5):1–15. https://doi.org/10.1186/1471-2164-11-S5-S5
76. Almeida PC, Nantes IL, Chagas JR, Rizzi CCA, Faljoni-Alario A, Carmona E, et al. Cathepsin B Activity Regulation: heparin-like glycosaminoglycans protect human cathepsin b from alkaline ph-induced inactivation. J Biol Chem. 2001;276(2):944–51. https://doi.org/10.1074/jbc.M003820200
77. Guicciardi ME, Miyoshi H, Bronk SF, Gores GJ. Cathepsin B knockout mice are resistant to tumor necrosis factor-alpha-mediated hepatocyte apoptosis and liver injury: implications for therapeutic applications. Am J Pathol. 2001;159(6):2045–54. https://doi.org/10.1016/S0002-9440(10)63056-8
78. Gowen M, Lazner F, Dodds R, Kapadia R, Feild J, Tavaria M, et al. Cathepsin K Knockout Mice Develop Osteopetrosis Due to a Deficit in Matrix Degradation but Not Demineralization. Journal of Bone and Mineral Research. 1999;14(10):1654–63. https://doi.org/10.1359/jbmr.1999.14.10.1654
79. Petermann I, Mayer C, Stypmann J, Biniossek ML, Tobin DJ, Engelen MA, et al. Lysosomal, cytoskeletal, and metabolic alterations in cardiomyopathy of cathepsin L knockout mice. The FASEB Journal. 2006;20(8):1266–8. https://doi.org/10.1096/fj.05-5517fje
80. Leissring MA, Reinstatler L, Sahara T, Sevlever D, Roman R, Ji Z, et al. Cathepsin D knockout mice harbor large and highly selective increases in cerebral Aß42 and tau: Implications for Alzheimer’s disease pathogenesis. Alzheimer’s & Dementia. 2009;5(4):P155-P6. https://doi.org/10.1016/j.jalz.2009.05.537
81. Kitamoto S, Sukhova GK, Sun J, Yang M, Libby P, Love V, et al. Cathepsin L Deficiency Reduces Diet-Induced Atherosclerosis in Low-Density Lipoprotein Receptor-Knockout Mice. Circulation. 2007;115(15):2065–75. https://doi.org/10.1161/CIRCULATIONAHA.107.688523
82. Hua Y, Robinson TJ, Cao Y, Shi GP, Ren J, Nair S. Cathepsin K knockout alleviates aging-induced cardiac dysfunction. Aging Cell. 2015;14(3):345–51. https://doi.org/10.1111/acel.12276
83. Tan G-J, Peng Z-K, Lu J-P, Tang F-Q. Cathepsins mediate tumor metastasis. World J Biol Chem. 2013;4(4):91–101. https://doi.org/10.4331/wjbc.v4.i4.91
84. Pišlar A, Perišić Nanut M, Kos J. Lysosomal cysteine peptidases - Molecules signaling tumor cell death and survival. Semin Cancer Biol. 2015;35:168–79. https://doi.org/10.1016/j.semcancer.2015.08.001
85. Bose SJ, Ayagama T, Burton RAB. Chapter 3 - Lysosomal proteases and their role in signaling pathways. In: Zelanis A, editor. Proteolytic Signaling in Health and Disease: Academic Press; 2022. p. 41–61. https://doi.org/10.1016/B978-0-323-85696-6.00007-X
86. Brisson L, Reshkin SJ, Goré J, Roger S. PH regulators in invadosomal functioning: Proton delivery for matrix tasting. Eur J Cell Biol. 2012;91(11–12):847–60. https://doi.org/10.1016/j.ejcb.2012.04.004
87. Mohamed MM, Sloane BF. Cysteine cathepsins: multifunctional enzymes in cancer. Nature reviews. 2006;6. https://doi.org/10.1038/nrc1949
88. Krueger S, Kellner U, Buehling F, Roessner A. Cathepsin L antisense oligonucleotides in a human osteosarcoma cell line: Effects on the invasive phenotype. Cancer Gene Ther. 2001;8(7):522–8. https://doi.org/10.1038/sj.cgt.7700341
89. Bervar A, Zajc I, Sever N, Katunuma N, Sloane BF, Lah TT. Invasiveness of transformed human breast epithelial cell lines is related to cathepsin B and inhibited by cysteine proteinase inhibitors. Biol Chem. 2003;384(3):447–55. https://doi.org/10.1515/BC.2003.050
90. Gole B, Durán Alonso MB, Dolenc V, Lah T. Post-translational regulation of cathepsin B, but not of other cysteine cathepsins, contributes to increased glioblastoma cell invasiveness in vitro. Pathology and Oncology Research. 2009;15(4):711–23. https://doi.org/10.1007/s12253-009-9175-8
91. Kenig S, Alonso MBD, Mueller MM, Lah TT. Glioblastoma and endothelial cells cross-talk, mediated by SDF-1, enhances tumour invasion and endothelial proliferation by increasing expression of cathepsins B, S, and MMP-9. Cancer Lett. 2010;289(1):53–61. https://doi.org/10.1016/j.canlet.2009.07.014
92. Withana NP, Blum G, Sameni M, Slaney C, Anbalagan A, Olive MB, et al. Cathepsin B inhibition limits bone metastasis in breast cancer. Cancer Res. 2012;72(5):1199–209. https://doi.org/10.1158/0008-5472.CAN-11-2759
93. Olson OC, Joyce JA. Cysteine cathepsin proteases: regulators of cancer progression and therapeutic response. Nat Rev Cancer. 2015;15(12):712–29. https://doi.org/10.1038/nrc4027
94. Gocheva V, Wang HW, Gadea BB, Shree T, Hunter KE, Garfall AL, et al. IL-4 induces cathepsin protease activity in tumor-associated macrophages to promote cancer growth and invasion. Genes Dev. 2010;24(3):241–55. https://doi.org/10.1101/gad.1874010
95. Cox JL. Cystatins and cancer. Frontiers in bioscience (Landmark edition). 2009;14:463–74. https://doi.org/10.2741/3255
96. Bruchard M, Mignot G, Derangere V, Chalmin F, Chevriaux A, Vegran F, et al. Chemotherapy-triggered cathepsin B release in myeloid-derived suppressor cells activates the Nlrp3 inflammasome and promotes tumor growth. Nat Med. 2013;19(1):57–64. https://doi.org/10.1038/nm.2999
97. Shree T, Olson OC, Elie BT, Kester JC, Garfall AL, Simpson K, et al. Macrophages and cathepsin proteases blunt chemotherapeutic response in breast cancer. Genes Dev. 2011;25(23):2465–79. https://doi.org/10.1101/gad.180331.111
98. Seo HR, Bae S, Lee YS. Radiation-induced cathepsin S is involved in radioresistance. Int J Cancer. 2009;124(8):1794–801. https://doi.org/10.1002/ijc.24095
99. Wartosch L, Bright NA, Luzio JP. Lysosomes. Curr Biol. 2015;25(8):R315-R6. https://doi.org/10.1016/j.cub.2015.02.027
100. Peters C, von Figura K. Biogenesis of lysosomal membranes. FEBS Lett. 1994;346(1):108–14. https://doi.org/10.1016/0014-5793(94)00499-4
101. Carlsson SR, Roth J, Piller F, Fukuda M. Isolation and characterization of human lysosomal membrane glycoproteins, h-lamp-1 and h-lamp-2. Major sialoglycoproteins carrying polylactosaminoglycan. J Biol Chem. 1988;263(35):18911–9. https://doi.org/10.1016/S0021-9258(18)37369-1
102. Settembre C, Fraldi A, Medina DL, Ballabio A. Signals from the lysosome: a control centre for cellular clearance and energy metabolism. Nat Rev Mol Cell Biol. 2013;14(5):283–96. https://doi.org/10.1038/nrm3565
103. Nishi T, Forgac M. The vacuolar (H+)-ATPases--nature’s most versatile proton pumps. Nat Rev Mol Cell Biol. 2002;3(2):94–103. https://doi.org/10.1038/nrm729
104. Marshansky V, Futai M. The V-type H+-ATPase in vesicular trafficking: targeting, regulation and function. Curr Opin Cell Biol. 2008;20(4):415–26. https://doi.org/10.1016/j.ceb.2008.03.015
105. Ohkuma S, Poole B. Fluorescence probe measurement of the intralysosomal pH in living cells and the perturbation of pH by various agents. Proc Natl Acad Sci U S A. 1978;75(7):3327–31. https://doi.org/10.1073/pnas.75.7.3327
106. Luzio JP, Pryor PR, Bright NA. Lysosomes: fusion and function. Nature Reviews Molecular Cell Biology. 2007;8(8):622–32. https://doi.org/10.1038/nrm2217
107. Kobayashi T, Beuchat MH, Chevallier J, Makino A, Mayran N, Escola JM, et al. Separation and characterization of late endosomal membrane domains. J Biol Chem. 2002;277(35):32157–64. https://doi.org/10.1074/jbc.M202838200
108. Hamer I, Van Beersel G, Arnould T, Jadot M. Lipids and lysosomes. Curr Drug Metab. 2012;13(10):1371–87. https://doi.org/10.2174/138920012803762684
109. Garcia-Ruiz C, Morales A, Fernandez-Checa JC. Glycosphingolipids and cell death: one aim, many ways. Apoptosis. 2015;20(5):607–20. https://doi.org/10.1007/s10495-015-1092-6
110. van Meer G, Voelker DR, Feigenson GW. Membrane lipids: where they are and how they behave. Nat Rev Mol Cell Biol. 2008;9(2):112–24. https://doi.org/10.1038/nrm2330
111. Young MM, Kester M, Wang H-G. Sphingolipids: regulators of crosstalk between apoptosis and autophagy. J Lipid Res. 2013;54(1):5–19. https://doi.org/10.1194/jlr.R031278
112. Jenkins RW, Canals D, Hannun YA. Roles and Regulation of Secretory and Lysosomal Acid Sphingomyelinase. Cell Signal. 2009;21(6):836–46. https://doi.org/10.1016/j.cellsig.2009.01.026
113. Rudnik S, Damme M. The lysosomal membrane-export of metabolites and beyond. The FEBS Journal. 2021;288(14):4168–82. https://doi.org/10.1111/febs.15602
114. Fehrenbacher N, Bastholm L, Kirkegaard-Sørensen T, Rafn B, Bøttzauw T, Nielsen C, et al. Sensitization to the Lysosomal Cell Death Pathway by Oncogene-Induced Down-regulation of Lysosome-Associated Membrane Proteins 1 and 2. Cancer Res. 2008;68(16):6623–33. https://doi.org/10.1158/0008-5472.CAN-08-0463
115. Kon M, Kiffin R, Koga H, Chapochnick J, Macian F, Varticovski L, et al. Chaperone-mediated autophagy is required for tumor growth. Sci Transl Med. 2011;3(109):109ra17. https://doi.org/10.1126/scitranslmed.3003182
116. Furuta K, Ikeda M, Nakayama Y, Nakamura K, Tanaka M, Hamasaki N, et al. Expression of lysosome-associated membrane proteins in human colorectal neoplasms and inflammatory diseases. Am J Pathol. 2001;159(2):449–55. https://doi.org/10.1016/S0002-9440(10)61716-6
117. Ozaki K, Nagata M, Suzuki M, Fujiwara T, Ueda K, Miyoshi Y, et al. Isolation and characterization of a novel human lung-specific gene homologous to lysosomal membrane glycoproteins 1 and 2: significantly increased expression in cancers of various tissues. Cancer Res. 1998;58(16):3499–503.
118. Kunzli BM, Berberat PO, Zhu ZW, Martignoni M, Kleeff J, Tempia-Caliera AA, et al. Influences of the lysosomal associated membrane proteins (Lamp-1, Lamp-2) and Mac-2 binding protein (Mac-2-BP) on the prognosis of pancreatic carcinoma. Cancer. 2002;94(1):228–39. https://doi.org/10.1002/cncr.10162
119. Lippincott-Schwartz J, Phair RD. Lipids and cholesterol as regulators of traffic in the endomembrane system. Annual review of biophysics. 2010;39:559–78. https://doi.org/10.1146/annurev.biophys.093008.131357
120. Chevallier J, Chamoun Z, Jiang G, Prestwich G, Sakai N, Matile S, et al. Lysobisphosphatidic Acid Controls Endosomal Cholesterol Levels. J Biol Chem. 2008;283(41):27871–80. https://doi.org/10.1074/jbc.M801463200
121. Kobayashi T, Stang E, Fang KS, de Moerloose P, Parton RG, Gruenberg J. A lipid associated with the antiphospholipid syndrome regulates endosome structure and function. Nature. 1998;392(6672):193–7. https://doi.org/10.1038/32440
122. Jadot M, Andrianaivo F, Dubois F, Wattiaux R. Effects of methylcyclodextrin on lysosomes. Eur J Biochem. 2001;268(5):1392–9. https://doi.org/10.1046/j.1432-1327.2001.02006.x
123. Brown DA, London E. Functions of lipid rafts in bioogical membranes. Annual Review of Cell and Developmental Biology. 1998;14(1):111–36. https://doi.org/10.1146/annurev.cellbio.14.1.111
124. Taute A, Wätzig K, Simons B, Lohaus C, Meyer HE, Hasilik A. Presence of detergent-resistant microdomains in lysosomal membranes. Biochemical and Biophysical Research Communications. 2002;298(1):5–9. https://doi.org/10.1016/S0006-291X(02)02387-2
125. Veal E, Groisman R, Eisenstein M, Gill G. The secreted glycoprotein CREG enhances differentiation of NTERA-2 human embryonal carcinoma cells. Oncogene. 2000;19(17):2120–8. https://doi.org/10.1038/sj.onc.1203529
126. Di Bacco A, Gill G. The secreted glycoprotein CREG inhibits cell growth dependent on the mannose-6-phosphate//insulin-like growth factor II receptor. Oncogene. 2003;22(35):5436–45. https://doi.org/10.1038/sj.onc.1206670
127. Greenlee JD, Subramanian T, Liu K, King MR. Rafting Down the Metastatic Cascade: The Role of Lipid Rafts in Cancer Metastasis, Cell Death, and Clinical Outcomes. Cancer Res. 2020:canres.2199.202. https://doi.org/10.1158/0008-5472.CAN-20-2199
128. Hannun YA. Functions of ceramide in coordinating cellular responses to stress. Science. 1996;274(5294):1855–9. https://doi.org/10.1126/science.274.5294.1855
129. Simons K, Ikonen E. Functional rafts in cell membranes. Nature. 1997;387(6633):569–72. https://doi.org/10.1038/42408
130. Hannun YA, Obeid LM. The Ceramide-centric Universe of Lipid-mediated Cell Regulation: Stress Encounters of the Lipid Kind. J Biol Chem. 2002;277(29):25847–50. https://doi.org/10.1074/jbc.R200008200
131. Kota V, Hama H. 2’-Hydroxy ceramide in membrane homeostasis and cell signaling. Advances in biological regulation. 2014;54:223–30. https://doi.org/10.1016/j.jbior.2013.09.012
132. Spiegel S, Milstien S. Functions of the Multifaceted Family of Sphingosine Kinases and Some Close Relatives. J Biol Chem. 2007;282(4):2125–9. https://doi.org/10.1074/jbc.R600028200
133. Hannun YA, Obeid LM. Principles of bioactive lipid signalling: lessons from sphingolipids. Nat Rev Mol Cell Biol. 2008;9(2):139–50. https://doi.org/10.1038/nrm2329
134. Kornhuber J, Tripal P, Reichel M, Muhle C, Rhein C, Muehlbacher M, et al. Functional Inhibitors of Acid Sphingomyelinase (FIASMAs): a novel pharmacological group of drugs with broad clinical applications. Cellular physiology and biochemistry : international journal of experimental cellular physiology, biochemistry, and pharmacology. 2010;26(1):9–20. https://doi.org/10.1159/000315101
135. Smith EL, Schuchman EH. The unexpected role of acid sphingomyelinase in cell death and the pathophysiology of common diseases. The FASEB Journal. 2008;22(10):3419–31. https://doi.org/10.1096/fj.08-108043
136. Taniguchi M, Okazaki T. Role of ceramide/sphingomyelin (SM) balance regulated through “SM cycle” in cancer. Cell Signal. 2021;87:110119. https://doi.org/10.1016/j.cellsig.2021.110119
137. Haimovitz-Friedman A, Cordon-Cardo C, Bayoumy S, Garzotto M, McLoughlin M, Gallily R, et al. Lipopolysaccharide Induces Disseminated Endothelial Apoptosis Requiring Ceramide Generation. The Journal of Experimental Medicine. 1997;186(11):1831–41. https://doi.org/10.1084/jem.186.11.1831
138. Santana P, Pena LA, Haimovitz-Friedman A, Martin S, Green D, McLoughlin M, et al. Acid sphingomyelinase-deficient human lymphoblasts and mice are defective in radiation-induced apoptosis. Cell. 1996;86(2):189–99. https://doi.org/10.1016/S0092-8674(00)80091-4
139. Gulbins E, Li PL. Physiological and pathophysiological aspects of ceramide. American journal of physiology Regulatory, integrative and comparative physiology. 2006;290(1):R11–26. https://doi.org/10.1152/ajpregu.00416.2005
140. Palma CD, Perrotta C. Ceramide as a target of chemotherapy: its role in apoptosis and autophagy. Clin Lipidol. 2012;7(1):111–9. https://doi.org/10.2217/clp.11.71
141. Gomez-Larrauri A, Das Adhikari U, Aramburu-Nuñez M, Custodia A, Ouro A. Ceramide Metabolism Enzymes-Therapeutic Targets against Cancer. Medicina. 2021;57(7):729. https://doi.org/10.3390/medicina57070729
142. Zhang AY, Yi F, Jin S, Xia M, Chen QZ, Gulbins E, et al. Acid sphingomyelinase and its redox amplification in formation of lipid raft redox signaling platforms in endothelial cells. Antioxid Redox Signal. 2007;9(7):817–28. https://doi.org/10.1089/ars.2007.1509
143. Li X, Han WQ, Boini KM, Xia M, Zhang Y, Li PL. TRAIL death receptor 4 signaling via lysosome fusion and membrane raft clustering in coronary arterial endothelial cells: evidence from ASM knockout mice. J Mol Med (Berl). 2013;91(1):25–36. https://doi.org/10.1007/s00109-012-0968-y
144. Daido S, Kanzawa T, Yamamoto A, Takeuchi H, Kondo Y, Kondo S. Pivotal role of the cell death factor BNIP3 in ceramide-induced autophagic cell death in malignant glioma cells. Cancer Res. 2004;64(12):4286–93. https://doi.org/10.1158/0008-5472.CAN-03-3084
145. Scarlatti F, Bauvy C, Ventruti A, Sala G, Cluzeaud F, Vandewalle A, et al. Ceramide-mediated macroautophagy involves inhibition of protein kinase B and up-regulation of beclin 1. J Biol Chem. 2004;279(18):18384–91. https://doi.org/10.1074/jbc.M313561200
146. Guenther GG, Peralta ER, Rosales KR, Wong SY, Siskind LJ, Edinger AL. Ceramide starves cells to death by downregulating nutrient transporter proteins. Proc Natl Acad Sci U S A. 2008;105(45):17402–7. https://doi.org/10.1073/pnas.0802781105
147. Sims K, Haynes CA, Kelly S, Allegood JC, Wang E, Momin A, et al. Kdo(2)-Lipid A, a TLR4-specific Agonist, Induces de Novo Sphingolipid Biosynthesis in RAW264.7 Macrophages, Which Is Essential for Induction of Autophagy. The Journal of Biological Chemistry. 2010;285(49):38568–79. https://doi.org/10.1074/jbc.M110.170621
148. Pattingre S, Bauvy C, Carpentier S, Levade T, Levine B, Codogno P. Role of JNK1-dependent Bcl-2 Phosphorylation in Ceramide-induced Macroautophagy. The Journal of Biological Chemistry. 2009;284(5):2719–28. https://doi.org/10.1074/jbc.M805920200
149. Huwiler A, Johansen B, Skarstad A, Pfeilschifter J. Ceramide binds to the CaLB domain of cytosolic phospholipase A2 and facilitates its membrane docking and arachidonic acid release. FASEB J. 2001;15(1):7–9. https://doi.org/10.1096/fj.00-0370fje
150. Heinrich M, Wickel M, Schneider-Brachert W, Sandberg C, Gahr J, Schwandner R, et al. Cathepsin D targeted by acid sphingomyelinase-derived ceramide. EMBO J. 1999;18(19):5252–63. https://doi.org/10.1093/emboj/18.19.5252
151. Edelmann B, Bertsch U, Tchikov V, Winoto-Morbach S, Perrotta C, Jakob M, et al. Caspase-8 and caspase-7 sequentially mediate proteolytic activation of acid sphingomyelinase in TNF-R1 receptosomes. The EMBO Journal. 2011;30(2):379–94. https://doi.org/10.1038/emboj.2010.326
152. Westwick JK, Bielawska AE, Dbaibo G, Hannun YA, Brenner DA. Ceramide activates the stress-activated protein kinases. J Biol Chem. 1995;270(39):22689–92. https://doi.org/10.1074/jbc.270.39.22689
153. Galluzzi L, Vitale I, Aaronson SA, Abrams JM, Adam D, Agostinis P, et al. Molecular mechanisms of cell death: recommendations of the Nomenclature Committee on Cell Death 2018. Cell Death Differ. 2018;25(3):486–541. https://doi.org/10.1038/s41418-017-0012-4
154. Vanden Berghe T, Vanlangenakker N, Parthoens E, Deckers W, Devos M, Festjens N, et al. Necroptosis, necrosis and secondary necrosis converge on similar cellular disintegration features. Cell Death Differ. 2010;17(6):922–30. https://doi.org/10.1038/cdd.2009.184
155. Aits S, Jaattela M, Nylandsted J. Methods for the quantification of lysosomal membrane permeabilization: a hallmark of lysosomal cell death. Methods Cell Biol. 2015;126:261–85. https://doi.org/10.1016/bs.mcb.2014.10.032
156. Hu M, Carraway KL, III. Repurposing Cationic Amphiphilic Drugs and Derivatives to Engage Lysosomal Cell Death in Cancer Treatment. Front Oncol. 2020;10. https://doi.org/10.3389/fonc.2020.605361
157. Brunk UT, Ericsson JL. Cytochemical evidence for the leakage of acid phosphatase through ultrastructurally intact lysosomal membranes. Histochem J. 1972;4(6):479–91. https://doi.org/10.1007/BF01011128
158. Aits S, Jaattela M. Lysosomal cell death at a glance. J Cell Sci. 2013;126(Pt 9):1905–12. https://doi.org/10.1242/jcs.091181
159. Matsuda S, Okada N, Kodama T, Honda T, Iida T. A cytotoxic type III secretion effector of Vibrio parahaemolyticus targets vacuolar H+-ATPase subunit c and ruptures host cell lysosomes. PLoS Pathog. 2012;8(7):e1002803. https://doi.org/10.1371/journal.ppat.1002803
160. Arnandis T, Ferrer-Vicens I, Garcia-Trevijano ER, Miralles VJ, Garcia C, Torres L, et al. Calpains mediate epithelial-cell death during mammary gland involution: mitochondria and lysosomal destabilization. Cell Death Differ. 2012;19(9):1536–48. https://doi.org/10.1038/cdd.2012.46
161. Loison F, Zhu H, Karatepe K, Kasorn A, Liu P, Ye K, et al. Proteinase 3-dependent caspase-3 cleavage modulates neutrophil death and inflammation. J Clin Invest. 2014;124(10):4445–58. https://doi.org/10.1172/JCI76246
162. Zhu W, Tao L, Quick ML, Joyce JA, Qu JM, Luo ZQ. Sensing cytosolic RpsL by macrophages induces lysosomal cell death and termination of bacterial infection. PLoS Pathog. 2015;11(3):e1004704. https://doi.org/10.1371/journal.ppat.1004704
163. Wang F, Gomez-Sintes R, Boya P. Lysosomal membrane permeabilization and cell death. Traffic. 2018;19(12):918–31. https://doi.org/10.1111/tra.12613
164. Boya P, Kroemer G. Lysosomal membrane permeabilization in cell death. Oncogene. 2008;27(50):6434–51. https://doi.org/10.1038/onc.2008.310
165. Bidere N, Lorenzo HK, Carmona S, Laforge M, Harper F, Dumont C, et al. Cathepsin D triggers Bax activation, resulting in selective apoptosis-inducing factor (AIF) relocation in T lymphocytes entering the early commitment phase to apoptosis. J Biol Chem. 2003;278(33):31401–11. https://doi.org/10.1074/jbc.M301911200
166. Serrano-Puebla A, Boya P. Lysosomal membrane permeabilization in cell death: new evidence and implications for health and disease. Ann N Y Acad Sci. 2016;1371(1):30–44. https://doi.org/10.1111/nyas.12966
167. Papadopoulos C, Meyer H. Detection and Clearance of Damaged Lysosomes by the Endo-Lysosomal Damage Response and Lysophagy. Curr Biol. 2017;27(24):R1330-R41. https://doi.org/10.1016/j.cub.2017.11.012
168. Skowyra ML, Schlesinger PH, Naismith TV, Hanson PI. Triggered recruitment of ESCRT machinery promotes endolysosomal repair. Science. 2018;360(6384). https://doi.org/10.1126/science.aar5078
169. Vasiljeva O, Turk B. Dual contrasting roles of cysteine cathepsins in cancer progression: apoptosis versus tumour invasion. Biochimie. 2008;90(2):380–6. https://doi.org/10.1016/j.biochi.2007.10.004
170. Brunk UT, Neuzil J, Eaton JW. Lysosomal involvement in apoptosis. Redox report : communications in free radical research. 2001;6(2):91–7. https://doi.org/10.1179/135100001101536094
171. Boya P, Andreau K, Poncet D, Zamzami N, Perfettini JL, Metivier D, et al. Lysosomal membrane permeabilization induces cell death in a mitochondrion-dependent fashion. J Exp Med. 2003;197(10):1323–34. https://doi.org/10.1084/jem.20021952
172. Boya P, Gonzalez-Polo RA, Poncet D, Andreau K, Vieira HL, Roumier T, et al. Mitochondrial membrane permeabilization is a critical step of lysosome-initiated apoptosis induced by hydroxychloroquine. Oncogene. 2003;22(25):3927–36. https://doi.org/10.1038/sj.onc.1206622
173. Repnik U, Hafner Cesen M, Turk B. Lysosomal membrane permeabilization in cell death: concepts and challenges. Mitochondrion. 2014;19 Pt A:49–57. https://doi.org/10.1016/j.mito.2014.06.006
174. Cirman T, Oresic K, Mazovec GD, Turk V, Reed JC, Myers RM, et al. Selective disruption of lysosomes in HeLa cells triggers apoptosis mediated by cleavage of Bid by multiple papain-like lysosomal cathepsins. J Biol Chem. 2004;279(5):3578–87. https://doi.org/10.1074/jbc.M308347200
175. Droga-Mazovec G, Bojic L, Petelin A, Ivanova S, Romih R, Repnik U, et al. Cysteine cathepsins trigger caspase-dependent cell death through cleavage of bid and antiapoptotic Bcl-2 homologues. J Biol Chem. 2008;283(27):19140–50. https://doi.org/10.1074/jbc.M802513200
176. Taniguchi M, Ogiso H, Takeuchi T, Kitatani K, Umehara H, Okazaki T. Lysosomal ceramide generated by acid sphingomyelinase triggers cytosolic cathepsin B-mediated degradation of X-linked inhibitor of apoptosis protein in natural killer/T lymphoma cell apoptosis. Cell Death Dis. 2015;6:e1717. https://doi.org/10.1038/cddis.2015.82
177. Bove J, Martinez-Vicente M, Dehay B, Perier C, Recasens A, Bombrun A, et al. BAX channel activity mediates lysosomal disruption linked to Parkinson disease. Autophagy. 2014;10(5):889–900. https://doi.org/10.4161/auto.28286
178. Guan JJ, Zhang XD, Sun W, Qi L, Wu JC, Qin ZH. DRAM1 regulates apoptosis through increasing protein levels and lysosomal localization of BAX. Cell Death Dis. 2015;6:e1624. https://doi.org/10.1038/cddis.2014.546
179. Huai J, Vogtle FN, Jockel L, Li Y, Kiefer T, Ricci JE, et al. TNFalpha-induced lysosomal membrane permeability is downstream of MOMP and triggered by caspase-mediated NDUFS1 cleavage and ROS formation. J Cell Sci. 2013;126(Pt 17):4015–25. https://doi.org/10.1242/jcs.129999
180. Oberle C, Huai J, Reinheckel T, Tacke M, Rassner M, Ekert PG, et al. Lysosomal membrane permeabilization and cathepsin release is a Bax/Bak-dependent, amplifying event of apoptosis in fibroblasts and monocytes. Cell Death Differ. 2010;17(7):1167–78. https://doi.org/10.1038/cdd.2009.214
181. Yuste VJ, Moubarak RS, Delettre C, Bras M, Sancho P, Robert N, et al. Cysteine protease inhibition prevents mitochondrial apoptosis-inducing factor (AIF) release. Cell Death Differ. 2005;12(11):1445–8. https://doi.org/10.1038/sj.cdd.4401687
182. Kirkegaard T, Jaattela M. Lysosomal involvement in cell death and cancer. Biochim Biophys Acta. 2009;1793(4):746–54. https://doi.org/10.1016/j.bbamcr.2008.09.008
183. Vanden Berghe T, Linkermann A, Jouan-Lanhouet S, Walczak H, Vandenabeele P. Regulated necrosis: the expanding network of non-apoptotic cell death pathways. Nat Rev Mol Cell Biol. 2014;15(2):135–47. https://doi.org/10.1038/nrm3737
184. Rowson-Hodel AR, Berg AL, Wald JH, Hatakeyama J, VanderVorst K, Curiel DA, et al. Hexamethylene amiloride engages a novel reactive oxygen species- and lysosome-dependent programmed necrotic mechanism to selectively target breast cancer cells. Cancer Lett. 2016;375(1):62–72. https://doi.org/10.1016/j.canlet.2016.02.042
185. Lipton P. Lysosomal membrane permeabilization as a key player in brain ischemic cell death: a “lysosomocentric” hypothesis for ischemic brain damage. Transl Stroke Res. 2013;4(6):672–84. https://doi.org/10.1007/s12975-013-0301-2
186. Yamashima T, Kohda Y, Tsuchiya K, Ueno T, Yamashita J, Yoshioka T, et al. Inhibition of ischaemic hippocampal neuronal death in primates with cathepsin B inhibitor CA-074: a novel strategy for neuroprotection based on ‘calpain-cathepsin hypothesis’. Eur J Neurosci. 1998;10(5):1723–33. https://doi.org/10.1046/j.1460-9568.1998.00184.x
187. Broker LE, Huisman C, Span SW, Rodriguez JA, Kruyt FA, Giaccone G. Cathepsin B mediates caspase-independent cell death induced by microtubule stabilizing agents in non-small cell lung cancer cells. Cancer Res. 2004;64(1):27–30. https://doi.org/10.1158/0008-5472.CAN-03-3060
188. Torii S, Shintoku R, Kubota C, Yaegashi M, Torii R, Sasaki M, et al. An essential role for functional lysosomes in ferroptosis of cancer cells. Biochem J. 2016;473(6):769–77. https://doi.org/10.1042/BJ20150658
189. Frank D, Vince JE. Pyroptosis versus necroptosis: similarities, differences, and crosstalk. Cell Death Differ. 2019;26(1):99–114. https://doi.org/10.1038/s41418-018-0212-6
190. Shi J, Gao W, Shao F. Pyroptosis: Gasdermin-Mediated Programmed Necrotic Cell Death. Trends Biochem Sci. 2017;42(4):245–54. https://doi.org/10.1016/j.tibs.2016.10.004
191. Hornung V, Bauernfeind F, Halle A, Samstad EO, Kono H, Rock KL, et al. Silica crystals and aluminum salts activate the NALP3 inflammasome through phagosomal destabilization. Nat Immunol. 2008;9(8):847–56. https://doi.org/10.1038/ni.1631
192. Duewell P, Kono H, Rayner KJ, Sirois CM, Vladimer G, Bauernfeind FG, et al. NLRP3 inflammasomes are required for atherogenesis and activated by cholesterol crystals. Nature. 2010;464(7293):1357–61. https://doi.org/10.1038/nature08938
193. Terlizzi M, Casolaro V, Pinto A, Sorrentino R. Inflammasome: Cancer’s friend or foe? Pharmacol Ther. 2014;143(1):24–33. https://doi.org/10.1016/j.pharmthera.2014.02.002
194. Zhao YG, Zhang H. Autophagosome maturation: An epic journey from the ER to lysosomes. J Cell Biol. 2019;218(3):757–70. https://doi.org/10.1083/jcb.201810099
195. Wang Y, Singh R, Massey AC, Kane SS, Kaushik S, Grant T, et al. Loss of macroautophagy promotes or prevents fibroblast apoptosis depending on the death stimulus. J Biol Chem. 2008;283(8):4766–77. https://doi.org/10.1074/jbc.M706666200
196. Mahapatra KK, Mishra SR, Behera BP, Patil S, Gewirtz DA, Bhutia SK. The lysosome as an imperative regulator of autophagy and cell death. Cellular and Molecular Life Sciences. 2021. https://doi.org/10.1007/s00018-021-03988-3
197. Mulcahy Levy JM, Thorburn A. Autophagy in cancer: moving from understanding mechanism to improving therapy responses in patients. Cell Death Differ. 2020;27(3):843–57. https://doi.org/10.1038/s41418-019-0474-7
198. Levy JMM, Towers CG, Thorburn A. Targeting autophagy in cancer. Nat Rev Cancer. 2017;17(9):528–42. https://doi.org/10.1038/nrc.2017.53
199. Barnard RA, Wittenburg LA, Amaravadi RK, Gustafson DL, Thorburn A, Thamm DH. Phase I clinical trial and pharmacodynamic evaluation of combination hydroxychloroquine and doxorubicin treatment in pet dogs treated for spontaneously occurring lymphoma. Autophagy. 2014;10(8):1415–25. https://doi.org/10.4161/auto.29165
200. Mauthe M, Orhon I, Rocchi C, Zhou X, Luhr M, Hijlkema KJ, et al. Chloroquine inhibits autophagic flux by decreasing autophagosome-lysosome fusion. Autophagy. 2018;14(8):1435–55. https://doi.org/10.1080/15548627.2018.1474314
201. Kondratskyi A, Kondratska K, Vanden Abeele F, Gordienko D, Dubois C, Toillon RA, et al. Ferroquine, the next generation antimalarial drug, has antitumor activity. Sci Rep. 2017;7(1):15896. https://doi.org/10.1038/s41598-017-16154-2
202. Wojtkowiak JW, Sane KM, Kleinman M, Sloane BF, Reiners JJ, Jr., Mattingly RR. Aborted autophagy and nonapoptotic death induced by farnesyl transferase inhibitor and lovastatin. J Pharmacol Exp Ther. 2011;337(1):65–74. https://doi.org/10.1124/jpet.110.174573
203. Liu L, Zhang N, Dou Y, Mao G, Bi C, Pang W, et al. Lysosomal dysfunction and autophagy blockade contribute to IMB-6G-induced apoptosis in pancreatic cancer cells. Sci Rep. 2017;7:41862. https://doi.org/10.1038/srep41862
204. Karch J, Schips TG, Maliken BD, Brody MJ, Sargent MA, Kanisicak O, et al. Autophagic cell death is dependent on lysosomal membrane permeability through Bax and Bak. Elife. 2017;6. https://doi.org/10.7554/eLife.30543
205. Bhoopathi P, Chetty C, Gujrati M, Dinh DH, Rao JS, Lakka S. Cathepsin B facilitates autophagy-mediated apoptosis in SPARC overexpressed primitive neuroectodermal tumor cells. Cell Death Differ. 2010;17(10):1529–39. https://doi.org/10.1038/cdd.2010.28
206. Hsu KF, Wu CL, Huang SC, Wu CM, Hsiao JR, Yo YT, et al. Cathepsin L mediates resveratrol-induced autophagy and apoptotic cell death in cervical cancer cells. Autophagy. 2009;5(4):451–60. https://doi.org/10.4161/auto.5.4.7666
207. Seo SU, Woo SM, Lee HS, Kim SH, Min KJ, Kwon TK. mTORC1/2 inhibitor and curcumin induce apoptosis through lysosomal membrane permeabilization-mediated autophagy. Oncogene. 2018;37(38):5205–20. https://doi.org/10.1038/s41388-018-0345-6
208. Hsieh AC, Costa M, Zollo O, Davis C, Feldman ME, Testa JR, et al. Genetic dissection of the oncogenic mTOR pathway reveals druggable addiction to translational control via 4EBP-eIF4E. Cancer Cell. 2010;17(3):249–61. https://doi.org/10.1016/j.ccr.2010.01.021
209. Leone RD, Amaravadi RK. Autophagy: a targetable linchpin of cancer cell metabolism. Trends Endocrinol Metab. 2013;24(4):209–17. https://doi.org/10.1016/j.tem.2013.01.008
210. Sabatini DM. mTOR and cancer: insights into a complex relationship. Nat Rev Cancer. 2006;6(9):729–34. https://doi.org/10.1038/nrc1974
211. Degenhardt K, Mathew R, Beaudoin B, Bray K, Anderson D, Chen G, et al. Autophagy promotes tumor cell survival and restricts necrosis, inflammation, and tumorigenesis. Cancer Cell. 2006;10(1):51–64. https://doi.org/10.1016/j.ccr.2006.06.001
212. White E, Mehnert JM, Chan CS. Autophagy, Metabolism, and Cancer. Clin Cancer Res. 2015;21(22):5037–46. https://doi.org/10.1158/1078-0432.CCR-15-0490
213. Perera RM, Stoykova S, Nicolay BN, Ross KN, Fitamant J, Boukhali M, et al. Transcriptional control of autophagy-lysosome function drives pancreatic cancer metabolism. Nature. 2015;524(7565):361–5. https://doi.org/10.1038/nature14587
214. Eaton JW, Qian M. Molecular bases of cellular iron toxicity. Free Radic Biol Med. 2002;32(9):833–40. https://doi.org/10.1016/S0891-5849(02)00772-4
215. Gyrd-Hansen M, Nylandsted J, Jaattela M. Heat shock protein 70 promotes cancer cell viability by safeguarding lysosomal integrity. Cell Cycle. 2004;3(12):1484–5. https://doi.org/10.4161/cc.3.12.1287
216. Naegeli KM, Hastie E, Garde A, Wang Z, Keeley DP, Gordon KL, et al. Cell Invasion In Vivo via Rapid Exocytosis of a Transient Lysosome-Derived Membrane Domain. Dev Cell. 2017;43(4):403–17 e10. https://doi.org/10.1016/j.devcel.2017.10.024
217. Tardy C, Codogno P, Autefage H, Levade T, Andrieu-Abadie N. Lysosomes and lysosomal proteins in cancer cell death (new players of an old struggle). Biochim Biophys Acta. 2006;1765(2):101–25. https://doi.org/10.1016/j.bbcan.2005.11.003
218. Bian B, Mongrain S, Cagnol S, Langlois MJ, Boulanger J, Bernatchez G, et al. Cathepsin B promotes colorectal tumorigenesis, cell invasion, and metastasis. Mol Carcinog. 2016;55(5):671–87. https://doi.org/10.1002/mc.22312
219. Puxbaum V, Nimmerfall E, Bauerl C, Taub N, Blaas PM, Wieser J, et al. M6P/IGF2R modulates the invasiveness of liver cells via its capacity to bind mannose 6-phosphate residues. J Hepatol. 2012;57(2):337–43. https://doi.org/10.1016/j.jhep.2012.03.026
220. Kang J, Yu Y, Jeong S, Lee H, Heo HJ, Park JJ, et al. Prognostic role of high cathepsin D expression in breast cancer: a systematic review and meta-analysis. Ther Adv Med Oncol. 2020;12:175883592092783. https://doi.org/10.1177/1758835920927838
221. Vetvicka V, Vetvickova J, Benes P. Role of enzymatically inactive procathepsin D in lung cancer. Anticancer Res. 2004;24(5A):2739–43.
222. Abdulla MH, Valli-Mohammed MA, Al-Khayal K, Al Shkieh A, Zubaidi A, Ahmad R, et al. Cathepsin B expression in colorectal cancer in a Middle East population: Potential value as a tumor biomarker for late disease stages. Oncol Rep. 2017;37(6):3175–80. https://doi.org/10.3892/or.2017.5576
223. Fukuda ME, Iwadate Y, Machida T, Hiwasa T, Nimura Y, Nagai Y, et al. Cathepsin D is a potential serum marker for poor prognosis in glioma patients. Cancer Res. 2005;65(12):5190–4. https://doi.org/10.1158/0008-5472.CAN-04-4134
224. Fehrenbacher N, Gyrd-Hansen M, Poulsen B, Felbor U, Kallunki T, Boes M, et al. Sensitization to the lysosomal cell death pathway upon immortalization and transformation. Cancer Res. 2004;64(15):5301–10. https://doi.org/10.1158/0008-5472.CAN-04-1427
225. Sennoune SR, Martinez-Zaguilan R. Plasmalemmal vacuolar H+-ATPases in angiogenesis, diabetes and cancer. Journal of Bioenergetics and Biomembranes. 2007;39(5):427–33. https://doi.org/10.1007/s10863-007-9108-8
226. Jagadish N, Agarwal S, Gupta N, Fatima R, Devi S, Kumar V, et al. Heat shock protein 70–2 (HSP70-2) overexpression in breast cancer. J Exp Clin Cancer Res. 2016;35(1). https://doi.org/10.1186/s13046-016-0425-9
227. Le Joncour V, Filppu P, Hyvonen M, Holopainen M, Turunen SP, Sihto H, et al. Vulnerability of invasive glioblastoma cells to lysosomal membrane destabilization. EMBO Mol Med. 2019;11(6). https://doi.org/10.15252/emmm.201809034
228. Ferrao P, Sincock P, Cole S, Ashman L. Intracellular P-gp contributes to functional drug efflux and resistance in acute myeloid leukaemia. Leuk Res. 2001;25(5):395–405. https://doi.org/10.1016/S0145-2126(00)00156-9
229. Fu D, Roufogalis BD. Actin disruption inhibits endosomal traffic of P-glycoprotein-EGFP and resistance to daunorubicin accumulation. American Journal of Physiology-Cell Physiology. 2007;292(4):C1543–C52. https://doi.org/10.1152/ajpcell.00068.2006
230. Yamagishi T, Sahni S, Sharp DM, Arvind A, Jansson PJ, Richardson DR. P-glycoprotein mediates drug resistance via a novel mechanism involving lysosomal sequestration. J Biol Chem. 2013;288(44):31761–71. https://doi.org/10.1074/jbc.M113.514091
231. Chapuy B, Panse M, Radunski U, Koch R, Wenzel D, Inagaki N, et al. ABC transporter A3 facilitates lysosomal sequestration of imatinib and modulates susceptibility of chronic myeloid leukemia cell lines to this drug. Haematologica. 2009;94(11):1528–36. https://doi.org/10.3324/haematol.2009.008631
232. Chapuy B, Koch R, Radunski U, Corsham S, Cheong N, Inagaki N, et al. Intracellular ABC transporter A3 confers multidrug resistance in leukemia cells by lysosomal drug sequestration. Leukemia. 2008;22(8):1576–86. https://doi.org/10.1038/leu.2008.103
233. Petruzzelli R, Polishchuk RS. Activity and Trafficking of Copper-Transporting ATPases in Tumor Development and Defense against Platinum-Based Drugs. Cells. 2019;8(9):1080. https://doi.org/10.3390/cells8091080
234. Pena K, Coblenz J, Kiselyov K. Brief exposure to copper activates lysosomal exocytosis. Cell Calcium. 2015;57(4):257–62. https://doi.org/10.1016/j.ceca.2015.01.005
235. Komatsu M, Sumizawa T, Mutoh M, Chen ZS, Terada K, Furukawa T, et al. Copper-transporting P-type adenosine triphosphatase (ATP7B) is associated with cisplatin resistance. Cancer Res. 2000;60(5):1312–6.
236. Zhitomirsky B, Assaraf YG. Lysosomes as mediators of drug resistance in cancer. Drug Resist Updat. 2016;24:23–33. https://doi.org/10.1016/j.drup.2015.11.004
237. Davidson SM, Vander Heiden MG. Critical Functions of the Lysosome in Cancer Biology. Annu Rev Pharmacol Toxicol. 2017;57:481–507. https://doi.org/10.1146/annurev-pharmtox-010715-103101
238. Firestone RA, Pisano JM, Bailey PJ, Sturm A, Bonney RJ, Wightman P, et al. Lysosomotropic agents. 4. Carbobenzoxyglycylphenylalanyl, a new protease-sensitive masking group for introduction into cells. J Med Chem. 1982;25(5):539–44. https://doi.org/10.1021/jm00347a012
239. Firestone RA, Pisano JM, Bonney RJ. Lysosomotropic agents. 1. Synthesis and cytotoxic action of lysosomotropic detergents. J Med Chem. 1979;22(9):1130–3. https://doi.org/10.1021/jm00195a026
240. Miller DK, Griffiths E, Lenard J, Firestone RA. Cell killing by lysosomotropic detergents. J Cell Biol. 1983;97(6):1841–51. https://doi.org/10.1083/jcb.97.6.1841
241. Bonam SR, Wang F, Muller S. Lysosomes as a therapeutic target. Nat Rev Drug Discov. 2019;18(12):923–48. https://doi.org/10.1038/s41573-019-0036-1
242. de Duve C, de Barsy T, Poole B, Trouet A, Tulkens P, Van Hoof F. Commentary. Lysosomotropic agents. Biochem Pharmacol. 1974;23(18):2495–531. https://doi.org/10.1016/0006-2952(74)90174-9
243. MacIntyre AC, Cutler DJ. The potential role of lysosomes in tissue distribution of weak bases. Biopharm Drug Dispos. 1988;9(6):513–26. https://doi.org/10.1002/bod.2510090602
244. Duvvuri M, Krise JP. Intracellular drug sequestration events associated with the emergence of multidrug resistance: a mechanistic review. Front Biosci. 2005;10:1499–509. https://doi.org/10.2741/1634
245. Kaufmann AM, Krise JP. Lysosomal Sequestration of Amine-Containing Drugs: Analysis and Therapeutic Implications. J Pharm Sci. 2007;96(4):729–46. https://doi.org/10.1002/jps.20792
246. Ceccarelli M, Germani R, Massari S, Petit C, Nurisso A, Wolfender JL, et al. Phospholipidosis effect of drugs by adsorption into lipid monolayers. Colloids Surf B Biointerfaces. 2015;136:175–84. https://doi.org/10.1016/j.colsurfb.2015.09.003
247. Handler JA, Badger A, Genell CA, Klinkner AM, Kassis S, Waites CR, et al. Selective inhibition of phospholipases by atiprimod, a macrophage targeting antiarthritic compound. Toxicology and applied pharmacology. 1999;159(1):9–17. https://doi.org/10.1006/taap.1999.8732
248. Anderson N, Borlak J. Drug-induced phospholipidosis. FEBS Lett. 2006;580(23):5533–40. https://doi.org/10.1016/j.febslet.2006.08.061
249. Chung C, Mader CC, Schmitz JC, Atladottir J, Fitchev P, Cornwell ML, et al. The vacuolar-ATPase modulates matrix metalloproteinase isoforms in human pancreatic cancer. Lab Invest. 2011;91(5):732–43. https://doi.org/10.1038/labinvest.2011.8
250. Das S, Dielschneider R, Chanas-LaRue A, Johnston JB, Gibson SB. Antimalarial drugs trigger lysosome-mediated cell death in chronic lymphocytic leukemia (CLL) cells. Leuk Res. 2018;70:79–86. https://doi.org/10.1016/j.leukres.2018.06.005
251. Dubowchik GM, Gawlak SL, Firestone RA. The in vitro effects of three lysosomotropic detergents against three human tumor cell lines. Bioorg Med Chem Lett. 1995;5(8):893–8. https://doi.org/10.1016/0960-894X(95)00136-H
252. Li W, Yuan X, Nordgren G, Dalen H, Dubowchik GM, Firestone RA, et al. Induction of cell death by the lysosomotropic detergent MSDH. FEBS letters. 2000;470(1):35–9. https://doi.org/10.1016/S0014-5793(00)01286-2
253. Brunk UT. Lysosomotropic detergents induce time- and dose-dependent apoptosis/necrosis in cultured cells. Redox Report. 2000;5(2-3):87–8. https://doi.org/10.1179/135100000101535609
254. Goldman SD, Funk RS, Rajewski RA, Krise JP. Mechanisms of amine accumulation in, and egress from, lysosomes. Bioanalysis. 2009;1(8):1445–59. https://doi.org/10.4155/bio.09.128
255. Hrabeta J, Groh T, Khalil MA, Poljakova J, Adam V, Kizek R, et al. Vacuolar-ATPase-mediated intracellular sequestration of ellipticine contributes to drug resistance in neuroblastoma cells. Int J Oncol. 2015;47(3):971–80. https://doi.org/10.3892/ijo.2015.3066
256. Adar Y, Stark M, Bram EE, Nowak-Sliwinska P, van den Bergh H, Szewczyk G, et al. Imidazoacridinone-dependent lysosomal photodestruction: a pharmacological Trojan horse approach to eradicate multidrug-resistant cancers. Cell Death Dis. 2012;3:e293. https://doi.org/10.1038/cddis.2012.30
257. Gotink KJ, Broxterman HJ, Labots M, de Haas RR, Dekker H, Honeywell RJ, et al. Lysosomal sequestration of sunitinib: a novel mechanism of drug resistance. Clin Cancer Res. 2011;17(23):7337–46. https://doi.org/10.1158/1078-0432.CCR-11-1667
258. Zhitomirsky B, Assaraf YG. Lysosomal sequestration of hydrophobic weak base chemotherapeutics triggers lysosomal biogenesis and lysosome-dependent cancer multidrug resistance. Oncotarget. 2015;6(2):1143–56. https://doi.org/10.18632/oncotarget.2732
259. Zhitomirsky B, Assaraf YG. Lysosomal accumulation of anticancer drugs triggers lysosomal exocytosis. Oncotarget. 2017;8(28):45117–32. https://doi.org/10.18632/oncotarget.15155
260. Zhitomirsky B, Assaraf YG. The role of cytoplasmic-to-lysosomal pH gradient in hydrophobic weak base drug sequestration in lysosomes. Cancer Cell & Microenvironment. 2015;2(3).
261. Amaravadi RK, Lippincott-Schwartz J, Yin X-M, Weiss WA, Takebe N, Timmer W, et al. Principles and Current Strategies for Targeting Autophagy for Cancer Treatment. Clinical Cancer Research. 2011;17(4):654–66. https://doi.org/10.1158/1078-0432.CCR-10-2634
262. Rosenfeldt MT, Ryan KM. The multiple roles of autophagy in cancer. Carcinogenesis. 2011;32(7):955–63. https://doi.org/10.1093/carcin/bgr031
263. Janku F, McConkey DJ, Hong DS, Kurzrock R. Autophagy as a target for anticancer therapy. Nat Rev Clin Oncol. 2011;8(9):528–39. https://doi.org/10.1038/nrclinonc.2011.71
264. Goodall ML, Wang T, Martin KR, Kortus MG, Kauffman AL, Trent JM, et al. Development of potent autophagy inhibitors that sensitize oncogenic BRAF V600E mutant melanoma tumor cells to vemurafenib. Autophagy. 2014;10(6):1120–36. https://doi.org/10.4161/auto.28594
265. Xiao M, Benoit A, Hasmim M, Duhem C, Vogin G, Berchem G, et al. Targeting Cytoprotective Autophagy to Enhance Anticancer Therapies. Front Oncol. 2021;11:180. https://doi.org/10.3389/fonc.2021.626309
266. Maycotte P, Aryal S, Cummings CT, Thorburn J, Morgan MJ, Thorburn A. Chloroquine sensitizes breast cancer cells to chemotherapy independent of autophagy. Autophagy. 2012;8(2):200–12. https://doi.org/10.4161/auto.8.2.18554
267. Sironi J, Aranda E, Nordstrom LU, Schwartz EL. Lysosome Membrane Permeabilization and Disruption of the Molecular Target of Rapamycin (mTOR)-Lysosome Interaction Are Associated with the Inhibition of Lung Cancer Cell Proliferation by a Chloroquinoline Analog. Mol Pharmacol. 2019;95(1):127–38. https://doi.org/10.1124/mol.118.113118
268. Zhou W, Guo Y, Zhang X, Jiang Z. Lys05 induces lysosomal membrane permeabilization and increases radiosensitivity in glioblastoma. J Cell Biochem. 2020;121(2):2027–37. https://doi.org/10.1002/jcb.29437
269. Circu M, Cardelli J, Barr MP, O’Byrne K, Mills G, El-Osta H. Modulating lysosomal function through lysosome membrane permeabilization or autophagy suppression restores sensitivity to cisplatin in refractory non-small-cell lung cancer cells. PLoS One. 2017;12(9):e0184922. https://doi.org/10.1371/journal.pone.0184922
270. Carew JS, Nawrocki ST, Kahue CN, Zhang H, Yang C, Chung L, et al. Targeting autophagy augments the anticancer activity of the histone deacetylase inhibitor SAHA to overcome Bcr-Abl-mediated drug resistance. Blood. 2007;110(1):313–22. https://doi.org/10.1182/blood-2006-10-050260
271. Kimura T, Takabatake Y, Takahashi A, Isaka Y. Chloroquine in Cancer Therapy: A Double-Edged Sword of Autophagy. Cancer Res. 2013;73(1):3–7. https://doi.org/10.1158/0008-5472.CAN-12-2464
272. Periyasamy-Thandavan S, Jiang M, Wei Q, Smith R, Yin XM, Dong Z. Autophagy is cytoprotective during cisplatin injury of renal proximal tubular cells. Kidney Int. 2008;74(5):631–40. https://doi.org/10.1038/ki.2008.214
273. Pellegrini P, Strambi A, Zipoli C, Hägg-Olofsson M, Buoncervello M, Linder S, et al. Acidic extracellular pH neutralizes the autophagy-inhibiting activity of chloroquine. Autophagy. 2014;10(4):562–71. https://doi.org/10.4161/auto.27901
274. Stern ST, Adiseshaiah PP, Crist RM. Autophagy and lysosomal dysfunction as emerging mechanisms of nanomaterial toxicity. Part Fibre Toxicol. 2012;9:20. https://doi.org/10.1186/1743-8977-9-20
275. Peynshaert K, Manshian BB, Joris F, Braeckmans K, De Smedt SC, Demeester J, et al. Exploiting intrinsic nanoparticle toxicity: the pros and cons of nanoparticle-induced autophagy in biomedical research. Chem Rev. 2014;114(15):7581–609. https://doi.org/10.1021/cr400372p
276. Zhang J, Qin X, Wang B, Xu G, Qin Z, Wang J, et al. Zinc oxide nanoparticles harness autophagy to induce cell death in lung epithelial cells. Cell Death Dis. 2017;8(7):e2954. https://doi.org/10.1038/cddis.2017.337
277. Wang F, Salvati A, Boya P. Lysosome-dependent cell death and deregulated autophagy induced by amine-modified polystyrene nanoparticles. Open Biol. 2018;8(4). https://doi.org/10.1098/rsob.170271
278. Ziglari T, Anderson DS, Holian A. Determination of the relative contribution of the non-dissolved fraction of ZnO NP on membrane permeability and cytotoxicity. Inhal Toxicol. 2020;32(2):86–95. https://doi.org/10.1080/08958378.2020.1743394
279. Wang J, Yu Y, Lu K, Yang M, Li Y, Zhou X, et al. Silica nanoparticles induce autophagy dysfunction via lysosomal impairment and inhibition of autophagosome degradation in hepatocytes. Int J Nanomedicine. 2017;12:809–25. https://doi.org/10.2147/IJN.S123596
280. Ding L, Zhu X, Wang Y, Shi B, Ling X, Chen H, et al. Intracellular Fate of Nanoparticles with Polydopamine Surface Engineering and a Novel Strategy for Exocytosis-Inhibiting, Lysosome Impairment-Based Cancer Therapy. Nano Lett. 2017;17(11):6790–801. https://doi.org/10.1021/acs.nanolett.7b03021
281. Du Rietz H, Hedlund H, Wilhelmson S, Nordenfelt P, Wittrup A. Imaging small molecule-induced endosomal escape of siRNA. Nat Commun. 2020;11(1):1809. https://doi.org/10.1038/s41467-020-15300-1
282. Ostenfeld MS, Fehrenbacher N, Hoyer-Hansen M, Thomsen C, Farkas T, Jaattela M. Effective tumor cell death by sigma-2 receptor ligand siramesine involves lysosomal leakage and oxidative stress. Cancer Res. 2005;65(19):8975–83. https://doi.org/10.1158/0008-5472.CAN-05-0269
283. Yuan N, Song L, Zhang S, Lin W, Cao Y, Xu F, et al. Bafilomycin A1 targets both autophagy and apoptosis pathways in pediatric B-cell acute lymphoblastic leukemia. Haematologica. 2015;100(3):345–56. https://doi.org/10.3324/haematol.2014.113324
284. Brodsky JL, Chiosis G. Hsp70 molecular chaperones: emerging roles in human disease and identification of small molecule modulators. Curr Top Med Chem. 2006;6(11):1215–25. https://doi.org/10.2174/156802606777811997
285. Garrido C, Brunet M, Didelot C, Zermati Y, Schmitt E, Kroemer G. Heat shock proteins 27 and 70: anti-apoptotic proteins with tumorigenic properties. Cell Cycle. 2006;5(22):2592–601. https://doi.org/10.4161/cc.5.22.3448
286. Schmitt E, Maingret L, Puig PE, Rerole AL, Ghiringhelli F, Hammann A, et al. Heat shock protein 70 neutralization exerts potent antitumor effects in animal models of colon cancer and melanoma. Cancer Res. 2006;66(8):4191–7. https://doi.org/10.1158/0008-5472.CAN-05-3778
287. Guzhova I, Margulis B. Hsp70 chaperone as a survival factor in cell pathology. Int Rev Cytol. 2006;254:101–49. https://doi.org/10.1016/S0074-7696(06)54003-3
288. Granato M, Lacconi V, Peddis M, Lotti LV, Di Renzo L, Gonnella R, et al. HSP70 inhibition by 2-phenylethynesulfonamide induces lysosomal cathepsin D release and immunogenic cell death in primary effusion lymphoma. Cell Death Dis. 2013;4:e730. https://doi.org/10.1038/cddis.2013.263
289. Nylandsted J, Brand K, Jaattela M. Heat shock protein 70 is required for the survival of cancer cells. Ann N Y Acad Sci. 2000;926:122–5. https://doi.org/10.1111/j.1749-6632.2000.tb05605.x
290. Rohde M, Daugaard M, Jensen MH, Helin K, Nylandsted J, Jaattela M. Members of the heat-shock protein 70 family promote cancer cell growth by distinct mechanisms. Genes Dev. 2005;19(5):570–82. https://doi.org/10.1101/gad.305405
291. Aghdassi A, Phillips P, Dudeja V, Dhaulakhandi D, Sharif R, Dawra R, et al. Heat shock protein 70 increases tumorigenicity and inhibits apoptosis in pancreatic adenocarcinoma. Cancer Res. 2007;67(2):616–25. https://doi.org/10.1158/0008-5472.CAN-06-1567
292. Powers MV, Clarke PA, Workman P. Dual targeting of HSC70 and HSP72 inhibits HSP90 function and induces tumor-specific apoptosis. Cancer Cell. 2008;14(3):250–62. https://doi.org/10.1016/j.ccr.2008.08.002
293. Cho K-j, van der Hoeven D, Zhou Y, Maekawa M, Ma X, Chen W, et al. Inhibition of Acid Sphingomyelinase Depletes Cellular Phosphatidylserine and Mislocalizes K-Ras from the Plasma Membrane. Molecular and Cellular Biology. 2016;36(2):363–74. https://doi.org/10.1128/MCB.00719-15
294. Barceló-Coblijn G, Martin ML, de Almeida RFM, Noguera-Salvà MA, Marcilla-Etxenike A, Guardiola-Serrano F, et al. Sphingomyelin and sphingomyelin synthase (SMS) in the malignant transformation of glioma cells and in 2-hydroxyoleic acid therapy. Proceedings of the National Academy of Sciences. 2011;108(49):19569–74. https://doi.org/10.1073/pnas.1115484108
295. Ostenfeld MS, Hoyer-Hansen M, Bastholm L, Fehrenbacher N, Olsen OD, Groth-Pedersen L, et al. Anti-cancer agent siramesine is a lysosomotropic detergent that induces cytoprotective autophagosome accumulation. Autophagy. 2008;4(4):487–99. https://doi.org/10.4161/auto.5774
296. Utermohlen O, Herz J, Schramm M, Kronke M. Fusogenicity of membranes: the impact of acid sphingomyelinase on innate immune responses. Immunobiology. 2008;213(3–4):307–14. https://doi.org/10.1016/j.imbio.2007.10.016
297. Prior IA, Lewis PD, Mattos C. A comprehensive survey of Ras mutations in cancer. Cancer Res. 2012;72(10):2457–67. https://doi.org/10.1158/0008-5472.CAN-11-2612
298. Litvak DA, Bilchik AJ, Cabot MC. Modulators of ceramide metabolism sensitize colorectal cancer cells to chemotherapy: a novel treatment strategy. J Gastrointest Surg. 2003;7(1):140–8; discussion 8. https://doi.org/10.1016/S1091-255X(02)00126-9
299. Lucci A, Giuliano AE, Han TY, Dinur T, Liu YY, Senchenkov A, et al. Ceramide toxicity and metabolism differ in wild-type and multidrug-resistant cancer cells. Int J Oncol. 1999;15(3):535–40. https://doi.org/10.3892/ijo.15.3.535
300. Lucci A, Han TY, Liu YY, Giuliano AE, Cabot MC. Modification of ceramide metabolism increases cancer cell sensitivity to cytotoxics. Int J Oncol. 1999;15(3):541–6. https://doi.org/10.3892/ijo.15.3.541
301. Gouaze-Andersson V, Yu JY, Kreitenberg AJ, Bielawska A, Giuliano AE, Cabot MC. Ceramide and glucosylceramide upregulate expression of the multidrug resistance gene MDR1 in cancer cells. Biochim Biophys Acta. 2007;1771(12):1407–17. https://doi.org/10.1016/j.bbalip.2007.09.005
302. Lin H, Chen C, Li X, Chen BD. Activation of the MEK/MAPK Pathway Is Involved in Bryostatin1-Induced Monocytic Differenciation and Up-regulation of X-Linked Inhibitor of Apoptosis Protein. Exp Cell Res. 2002;272(2):192–8. https://doi.org/10.1006/excr.2001.5417
303. Ibrahim AM, Mansour IM, Wilson MM, Mokhtar DA, Helal AM, Al Wakeel HM. Study of survivin and X-linked inhibitor of apoptosis protein (XIAP) genes in acute myeloid leukemia (AML). Lab Hematol. 2012;18(1):1–10. https://doi.org/10.1532/LH96.11005
304. Augello C, Caruso L, Maggioni M, Donadon M, Montorsi M, Santambrogio R, et al. Inhibitors of apoptosis proteins (IAPs) expression and their prognostic significance in hepatocellular carcinoma. BMC Cancer. 2009;9:125. https://doi.org/10.1186/1471-2407-9-125
305. Abdul-Hammed M, Breiden B, Adebayo MA, Babalola JO, Schwarzmann G, Sandhoff K. Role of endosomal membrane lipids and NPC2 in cholesterol transfer and membrane fusion. J Lipid Res. 2010;51(7):1747–60. https://doi.org/10.1194/jlr.M003822
306. Pyne NJ, Pyne S. Sphingosine 1-phosphate and cancer. Nat Rev Cancer. 2010;10(7):489–503. https://doi.org/10.1038/nrc2875
307. Ullio C, Casas J, Brunk UT, Sala G, Fabriàs G, Ghidoni R, et al. Sphingosine mediates TNFα-induced lysosomal membrane permeabilization and ensuing programmed cell death in hepatoma cells. J Lipid Res. 2012;53(6):1134–43. https://doi.org/10.1194/jlr.M022384
308. Suzuki E, Handa K, Toledo MS, Hakomori S. Sphingosine-dependent apoptosis: A unified concept based on multiple mechanisms operating in concert. Proc Natl Acad Sci U S A. 2004;101(41):14788–93. https://doi.org/10.1073/pnas.0406536101
309. Cuvillier O. Sphingosine in apoptosis signaling. Biochim Biophys Acta. 2002;1585(2–3):153–62. https://doi.org/10.1016/S1388-1981(02)00336-0
310. Kagedal K, Zhao M, Svensson I, Brunk UT. Sphingosine-induced apoptosis is dependent on lysosomal proteases. Biochem J. 2001;359(Pt 2):335–43. https://doi.org/10.1042/bj3590335
311. Höglinger D, Haberkant P, Aguilera-Romero A, Riezman H, Porter FD, Platt FM, et al. Intracellular sphingosine releases calcium from lysosomes. eLife. 2015;4:e10616. https://doi.org/10.7554/eLife.10616
312. Christensen KA, Myers JT, Swanson JA. pH-dependent regulation of lysosomal calcium in macrophages. J Cell Sci. 2002;115(3):599–607. https://doi.org/10.1242/jcs.115.3.599
313. Miyakawa T, Maeda A, Yamazawa T, Hirose K, Kurosaki T, Iino M. Encoding of Ca2+ signals by differential expression of IP3 receptor subtypes. EMBO J. 1999;18(5):1303–8. https://doi.org/10.1093/emboj/18.5.1303
314. Medina DL, Di Paola S, Peluso I, Armani A, De Stefani D, Venditti R, et al. Lysosomal calcium signalling regulates autophagy through calcineurin and TFEB. Nat Cell Biol. 2015;17(3):288–99. https://doi.org/10.1038/ncb3114
315. Patel S, Docampo R. Acidic calcium stores open for business: expanding the potential for intracellular Ca2+ signaling. Trends Cell Biol. 2010;20(5):277–86. https://doi.org/10.1016/j.tcb.2010.02.003
316. Li P, Gu M, Xu H. Lysosomal Ion Channels as Decoders of Cellular Signals. Trends Biochem Sci. 2019;44(2):110–24. https://doi.org/10.1016/j.tibs.2018.10.006
317. Mirnikjoo B, Balasubramanian K, Schroit AJ. Mobilization of Lysosomal Calcium Regulates the Externalization of Phosphatidylserine during Apoptosis. The Journal of Biological Chemistry. 2009;284(11):6918–23. https://doi.org/10.1074/jbc.M805288200
318. Ravichandran KS. “Recruitment signals” from apoptotic cells: invitation to a quiet meal. Cell. 2003;113(7):817–20. https://doi.org/10.1016/S0092-8674(03)00471-9
319. Fadok VA, Voelker DR, Campbell PA, Cohen JJ, Bratton DL, Henson PM. Exposure of phosphatidylserine on the surface of apoptotic lymphocytes triggers specific recognition and removal by macrophages. J Immunol. 1992;148(7):2207–16.
320. Dauer P, Gupta VK, McGinn O, Nomura A, Sharma NS, Arora N, et al. Inhibition of Sp1 prevents ER homeostasis and causes cell death by lysosomal membrane permeabilization in pancreatic cancer. Sci Rep. 2017;7(1):1564. https://doi.org/10.1038/s41598-017-01696-2
321. Guicciardi ME, Deussing J, Miyoshi H, Bronk SF, Svingen PA, Peters C, et al. Cathepsin B contributes to TNF-alpha-mediated hepatocyte apoptosis by promoting mitochondrial release of cytochrome c. J Clin Invest. 2000;106(9):1127–37. https://doi.org/10.1172/JCI9914
322. Yamashima T, Saido TC, Takita M, Miyazawa A, Yamano J, Miyakawa A, et al. Transient brain ischaemia provokes Ca2+, PIP2 and calpain responses prior to delayed neuronal death in monkeys. Eur J Neurosci. 1996;8(9):1932–44. https://doi.org/10.1111/j.1460-9568.1996.tb01337.x
323. Yamashima T, Tonchev AB, Tsukada T, Saido TC, Imajoh-Ohmi S, Momoi T, et al. Sustained calpain activation associated with lysosomal rupture executes necrosis of the postischemic CA1 neurons in primates. Hippocampus. 2003;13(7):791–800. https://doi.org/10.1002/hipo.10127
324. Yap YW, Whiteman M, Bay BH, Li Y, Sheu FS, Qi RZ, et al. Hypochlorous acid induces apoptosis of cultured cortical neurons through activation of calpains and rupture of lysosomes. J Neurochem. 2006;98(5):1597–609. https://doi.org/10.1111/j.1471-4159.2006.03996.x
325. Antunes F, Cadenas E, Brunk UT. Apoptosis induced by exposure to a low steady-state concentration of H2O2 is a consequence of lysosomal rupture. Biochem J. 2001;356(Pt 2):549–55. https://doi.org/10.1042/bj3560549
326. Waster PK, Ollinger KM. Redox-dependent translocation of p53 to mitochondria or nucleus in human melanocytes after UVA- and UVB-induced apoptosis. J Invest Dermatol. 2009;129(7):1769–81. https://doi.org/10.1038/jid.2008.421
327. Ollinger K, Brunk UT. Cellular injury induced by oxidative stress is mediated through lysosomal damage. Free Radic Biol Med. 1995;19(5):565–74. https://doi.org/10.1016/0891-5849(95)00062-3
328. Brunk UT, Zhang H, Dalen H, Ollinger K. Exposure of cells to nonlethal concentrations of hydrogen peroxide induces degeneration-repair mechanisms involving lysosomal destabilization. Free Radic Biol Med. 1995;19(6):813–22. https://doi.org/10.1016/0891-5849(95)02001-Q
329. Kurz T, Eaton JW, Brunk UT. The role of lysosomes in iron metabolism and recycling. The International Journal of Biochemistry & Cell Biology. 2011;43(12):1686–97. https://doi.org/10.1016/j.biocel.2011.08.016
330. Garner B, Li W, Roberg K, Brunk UT. On the cytoprotective role of ferritin in macrophages and its ability to enhance lysosomal stability. Free Radic Res. 1997;27(5):487–500. https://doi.org/10.3109/10715769709065788
331. Persson HL, Nilsson KJ, Brunk UT. Novel cellular defenses against iron and oxidation: ferritin and autophagocytosis preserve lysosomal stability in airway epithelium. Redox report : communications in free radical research. 2001;6(1):57–63. https://doi.org/10.1179/135100001101536049
332. Persson HL, Yu Z, Tirosh O, Eaton JW, Brunk UT. Prevention of oxidant-induced cell death by lysosomotropic iron chelators. Free Radic Biol Med. 2003;34(10):1295–305. https://doi.org/10.1016/S0891-5849(03)00106-0
333. Persson HL, Kurz T, Eaton JW, Brunk UT. Radiation-induced cell death: importance of lysosomal destabilization. Biochem J. 2005;389(Pt 3):877–84. https://doi.org/10.1042/BJ20050271
334. Yu Z, Persson HL, Eaton JW, Brunk UT. Intralysosomal iron: a major determinant of oxidant-induced cell death. Free Radic Biol Med. 2003;34(10):1243–52. https://doi.org/10.1016/S0891-5849(03)00109-6
335. Persson HL. Iron-dependent lysosomal destabilization initiates silica-induced apoptosis in murine macrophages. Toxicol Lett. 2005;159(2):124–33. https://doi.org/10.1016/j.toxlet.2005.05.002
336. Rizzollo F, More S, Vangheluwe P, Agostinis P. The lysosome as a master regulator of iron metabolism. Trends in Biochemical Sciences. 2021. https://doi.org/10.1016/j.tibs.2021.07.003
337. Mai TT, Hamai A, Hienzsch A, Caneque T, Muller S, Wicinski J, et al. Salinomycin kills cancer stem cells by sequestering iron in lysosomes. Nat Chem. 2017;9(10):1025–33. https://doi.org/10.1038/nchem.2778
338. Thayyullathil F, Cheratta AR, Alakkal A, Subburayan K, Pallichankandy S, Hannun YA, et al. Acid sphingomyelinase-dependent autophagic degradation of GPX4 is critical for the execution of ferroptosis. Cell Death Dis. 2021;12(1). https://doi.org/10.1038/s41419-020-03297-w
339. Yamaguchi K, Mandai M, Oura T, Matsumura N, Hamanishi J, Baba T, et al. Identification of an ovarian clear cell carcinoma gene signature that reflects inherent disease biology and the carcinogenic processes. Oncogene. 2010;29(12):1741–52. https://doi.org/10.1038/onc.2009.470
340. Kajihara H, Yamada Y, Kanayama S, Furukawa N, Noguchi T, Haruta S, et al. Clear cell carcinoma of the ovary: potential pathogenic mechanisms (Review). Oncol Rep. 2010;23(5):1193–203. https://doi.org/10.3892/or_00000750
341. Yamada Y, Shigetomi H, Onogi A, Haruta S, Kawaguchi R, Yoshida S, et al. Redox-Active Iron-Induced Oxidative Stress in the Pathogenesis of Clear Cell Carcinoma of the Ovary. Int J Gynecol Cancer. 2011;21(7):1200–7. https://doi.org/10.1097/IGC.0b013e318222cfdd
342. Yu Z, Eaton JW, Persson HL. The radioprotective agent, amifostine, suppresses the reactivity of intralysosomal iron. Redox Report. 2003;8(6):347–55. https://doi.org/10.1179/135100003225003384
343. Kurz T, Terman A, Gustafsson B, Brunk UT. Lysosomes in iron metabolism, ageing and apoptosis. Histochem Cell Biol. 2008;129(4):389–406. https://doi.org/10.1007/s00418-008-0394-y
344. Kurz T, Terman A, Gustafsson B, Brunk UT. Lysosomes and oxidative stress in aging and apoptosis. Biochim Biophys Acta. 2008;1780(11):1291–303. https://doi.org/10.1016/j.bbagen.2008.01.009