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    Topic review

    The V-ATPase a3 Subunit

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    Submitted by: Anh Chu

    Definition

    This entry focuses on one of the 16 proteins composing the V-ATPase complex responsible for resorbing bone: the a3 subunit. The rationale for focusing on this biomolecule is that mutations in this one protein account for over 50% of osteopetrosis cases, highlighting its critical role in bone physiology. Despite its essential role in bone remodeling and its involvement in bone diseases, little is known about the way in which this subunit is targeted and regulated within osteoclasts. To this end, this review is broadened to include the three other mammalian paralogues (a1, a2 and a4) and the two yeast orthologs (Vph1p and Stv1p). By examining the literature on all of the paralogues/orthologs of the V-ATPase a subunit, we hope to provide insight into the molecular mechanisms and future research directions specific to a3. This review starts with an overview on bone, highlighting the role of V-ATPases in osteoclastic bone resorption. We then cover V-ATPases in other location/functions, highlighting the roles which the four mammalian a subunit paralogues might play in differential targeting and/or regulation. Author review the ways in which the energy of ATP hydrolysis is converted into proton translocation, and go in depth into the diverse role of the a subunit, not only in proton translocation but also in lipid binding, cell signaling and human diseases. Finally, the therapeutic implication of targeting a3 specifically for bone diseases and cancer is discussed, with concluding remarks on future directions.

    1. Bone

    Bone is a remarkable dynamic tissue which is involved in a variety of roles besides providing structural support. Bone exhibits endocrine, immune, mineral storage, growth factor, organ protection and repair functions [1][2][3]. Most of these functions can be attributed to the presence of three distinct major cell types, the osteoblast (OB), the osteoclast (OC) and the osteocytes. Osteoblasts are derived from mesenchymal stem cells during embryogenesis, and are responsible for the secretion of a proteinaceous matrix, including growth factors, which becomes mineralized [1][4]. OBs are found lining the bone surface and also become encased in the mineralized matrix, where they differentiate into osteocytes [5][6]. Osteocytes communicate with each other and other cell types via canaliculi found in bone. Osteocytes are capable of detecting stresses on the skeleton, and are able to activate OBs lining the bone surface, as well as OCs to start the repair process [7]. Osteoclasts have been thought to arise from hematopoietic cells exclusively; however, recent lineage tracing studies using mice have shown that there is also an extraembryonic component to this [8][9]. Cells derived from erythromyeloid-progenitors (EMP) in the embryonic yolk sac are the first wave of OC to differentiate, followed later by a distinct second wave derived from hematopoietic stem cells (HSCs). These two stem cell populations occupy two different niches in the adult, with the EMP homing to the spleen while the HSCs seed the bone marrow [8][9]. OCs are capable of resorbing bone via their ability to secrete acid to dissolve the mineral component and proteinases in order to digest the now exposed proteinaceous matrix [10][11]. This is a highly organized process that involves pre-OC cells fusing with each other, the formation of a sealed bone compartment underneath the now multinucleated OC sequestered by the sealing zone, and the formation of a ruffled membrane contained within the sealing zone [12][13][14]. The ruffled border acts as the gateway for the secretion of the acid and proteinases, and allows for the uptake of the dissolved mineral and digested proteins, which are mostly transcytosed by vesicles to the apical cell membrane for eventual disposal via the circulation [15]. Experiments performed in RAW 264 cells showed that the formation of an actin ring redirects intracellular vesicles, mostly secretory lysosomes, to transport large quantities of proteinases (e.g., Cathepsin K, alkaline phosphatase) and the acid generating machinery (made up from chloride channel 7 and the V-ATPase) to the OC plasma membrane adjacent to the bone surface [16][17]. Bone remodeling is a highly coordinated process that involves constant communication between OBs and OCs, and any interference with this can lead to disease [10][18][19]. V-ATPases are involved in pre-pro-protein processing (including glycosylation) [20], secretion [21], the internalization and degradation of molecules [22], vesicle transport and fusion [23][24], modulate signaling complexes, participate in distinct signalosomes [25], and promote cell migration in cancer [26]. To this end, mutations that interfere with V-ATPase function underlie diseases affecting a number of organ systems.

    2. V-ATPase Functions

    V-ATPases are ATP-driven proton pumps found in the endomembrane of the intracellular compartments in all of the eukaryotic cells and the plasma membrane of several specialized cells [27]. V-ATPases are responsible for acidifying and maintaining the pH of intracellular organelles, including the Golgi apparatus, endosome, lysosome and secretory vesicles [28][29]. V-ATPases pump protons into the Golgi apparatus, which become more acidic from the cis-Golgi to the trans-Golgi [20]. As newly synthesized proteins traverse the Golgi apparatus, they undergo post-translational modification including glycosylation, sulfation and phosphorylation. The maintenance of the pH gradient in the Golgi apparatus by V-ATPases is crucial for the function and localization of the glycosyltransferases required for the modification processes [30]. V-ATPase activity in the intracellular membrane is important for membrane trafficking processes such as receptor-mediated endocytosis [21][22]. The V-ATPase-dependent acidification of the endocytic compartments is required for the dissociation of ligand–receptor complexes, allowing the receptors to recycle to the cell surface. The released ligands are subsequently targeted to the lysosomes, where the low pH maintained by V-ATPases facilitates their degradation [31][32]. This process is important for the continued uptake of ligands such as low-density lipoprotein (LDL), a main carrier of plasma membrane cholesterol [33]. Many pathogens employ the V-ATPase-mediated acidification of the endocytic compartments to gain entry into cells, including diphtheria and anthrax toxins, as well as viruses such as influenza and Ebola [34][35]. After entering the host cells, viruses also require a low pH to trigger fusion and to deliver their viral genome into the host. V-ATPases are also involved in the intracellular trafficking of lysosomal enzymes by establishing a luminal pH gradient between compartments [21][32]. Lysosomes are more acidic than late endosomes, which in turn are more acidic than the trans-Golgi network (TGN). This gradient allows the binding of lysosomal proteases to the mannose-6-phosphate receptor at the TGN, facilitating the enzyme delivery to the lysosomes, and the dissociation of enzyme-receptor complexes in late endosomes, allowing the receptors to recycle to the TGN [36]. V-ATPases play a key role in cellular nutrient homeostasis by providing the acidic environment within lysosomes which is necessary for proteolysis, which is a major way in which cells generate free amino acids [37][38][39]. In addition to maintaining the lysosomal pH, V-ATPases also associate with the nutrient-sensing machinery in the lysosomal membrane, and are involved in the recruitment of the metabolic regulators mTORC1 and AMPK [38][39][40]. Within secretory vesicles, V-ATPases generate a proton gradient driving the uptake of small molecules such as the neurotransmitter glutamate [41][42][43], and they facilitate the processing of prohormones like proinsulin [33][44][45].
    V-ATPases are targeted to the plasma membrane of specialized cells such as kidney intercalated cells [46][47], epididymis clear cells [48][49] and osteoclasts [50][51], where they function to transport protons from the cytoplasm to the extracellular space [30][52][53][54]. In the kidneys, V-ATPases are localized to the apical membrane of the alpha-intercalated cells to facilitate the secretion of protons into the urine in order to maintain pH homeostasis [54]. Osteoclasts rely on V-ATPases at the ruffle border for the demineralization of bone and the activation of the proteolytic enzymes required for bone resorption [55][56]. V-ATPases targeting the plasma membrane of epididymis clear cells are involved in the establishment of the acidic luminal pH necessary for sperm maturation and storage [49]. Recently, plasma membrane V-ATPases have been shown to be overexpressed in breast cancer cells, and to facilitate invasion by promoting the activity of acid-dependent proteases that degrade the extracellular matrix [57]. The inhibition of V-ATPases by concanamycin in prostate cancer cells results in a decreased level of mRNA for prostate-specific antigens [58]. Increasing evidence implicates the important role of V-ATPases in cancer cells’ growth and metastasis, and suggests a potential therapeutic treatment of metastatic cancer by the inhibition of V-ATPases activity.
    In addition to the conventional functions of V-ATPases in intracellular signalling and membrane trafficking by generating pH gradients, recent findings suggest novel emerging roles of V-ATPases in the modulation of the function of receptors and their regulatory complexes through direct protein–protein interactions. For example, it was recently uncovered that Wnt/β-catenin signal transmission requires the interaction of co-receptor LRP6 with V-ATPase lysosomal accessory protein-2 (ATP6AP2) in late endosomes [59]. In Drosophila, V-ATPases have been suggested to be involved in the membrane fusion of synaptic vesicles via direct interaction with calmodulin [60]. Emerging studies propose the importance of V-ATPases in modulating various signalling pathways, including Notch, mTOR and AMPK via unconventional mechanisms [39][61].
    In summary, V-ATPase-dependent acidification is essential for cellular metabolism, membrane trafficking and intracellular signalling. Moreover, the importance and novel emerging roles of V-ATPases in many signalling pathways and diseases, including cancers, makes them promising targets for drug development.

    3. V-ATPase Structure

    V-ATPases share their structure with mitochondrial and chloroplast F-type ATPases [27]. Both enzymes are composed of a peripheral catalytic sector (V1 or F1) and a membrane-bound proton channel sector (VO or FO). They are evolutionarily related, and are functionally conserved as rotary proton pumps [44]. The eukaryotic V-ATPase is a 900 kDa complex consisting of sixteen subunits: A3B3CDE3FG3H comprising the V1 sector, and ac9cdefAP1AP2 forming the membrane-bound VO (the subscript numbers represent the subunits’ stoichiometry in the complex) (Figure 1) [62][63]. Subunits A and B are arranged in a hexameric configuration and contain the nucleotide binding sites responsible for ATP hydrolysis [64]. ATP hydrolysis creates a driving force to induce the rotation of the central stalk composed of subunits D, F and d, and the membrane-bound proteolipid c-ring c9c[65].
    Figure 1. Mammalian V-ATPase complex. Cytosolic sector V1, comprised of A3B3CDE3FG3H, is responsible for the ATP hydrolysis, which generates the force required to drive the rotation of the proteolipid c-ring (c9c) of the membrane-bound VO consisting of ac9cdefAP1AP2. The aCT forms two half-channels that create a pathway for protons to cross the lipid bilayer as the c-ring rotates. Both a2 and a3 orthologs are glycosylated twice on the first luminal loop within the C-terminus (depicted here), whereas a1 and a4 are only glycosylated once [66].
    Each proteolipid subunit c, c has a conserved glutamate residue which is essential for proton translocation (E139 in c, and E98 in c) [64][67][68]. The glutamate residues are protonated when the subunit rotates past the membrane-embedded C-terminal domain of the a subunit (aCT). The aCT forms two half-channels that create a pathway for protons to cross from cytoplasm to the organellar lumen or the extracellular space [64]. Protons access the glutamate residue of subunit c upon entering the cytosolic half-channel, and the protonated glutamate residue carries the proton through the lipid bilayer as the c-ring rotates. The proton is released through the luminal half-channel following the deprotonation of the glutamate residue and stabilization by the critical arginine residue R740 in the a subunit (R735 in Vph1p, an S. cerevisiae ortholog of the a subunit) [69]. The AB hexamer is held stationary relative to the a subunit by three peripheral stalk EG heterodimers which connect the V1 sector to subunits C and H, and the N-terminal domain of the a subunit (aNT) [70].
    V-ATPase activity is tightly controlled both spatially and temporally. One example of the temporal modes of V-ATPase regulation is the reversible assembly/disassembly upon environmental cues, which was first described in yeast [71]. The dissociation of V1-VO sectors is regulated by nutrient availability, as the dissociated complex is inactive in both ATP hydrolysis and proton translocation, reflecting the cells’ attempt to conserve cellular ATP. In yeast, the dissociation occurs in response to glucose starvation, involves an intact microtubular network, and is reversible without new protein synthesis [72]. Upon V1–VO disassembly, the C subunit dissociates from the V-ATPase complex, and the H subunit undergoes a conformational change resulting in the loss of the interaction with aNT [73]. The reassembly of the complex requires the RAVE complex (Rav1, Rav2 and Skp1). The RAVE complex binds to subunits E and G, the dissociated C subunit of V1, and to the VO subunit a, thereby positioning them to promote assembly [71]. The glucose-induced reassembly of V-ATPases requires the interaction of the protein complexes with regulatory proteins, such as the RAVE complex. Moreover, studies in yeast and RAW 264 osteoclast-like cells suggest a direct interaction between the glycolytic enzyme aldolase and V-ATPase subunits in a glucose-dependent manner [74]. The deletion of the aldolase gene in yeast resulted in V-ATPase disassembly and a reduction in V-ATPase activity [75]. In the presence of glucose, aldolase and V-ATPase interactions increase, inducing the reassembly of V1 and VO; hence, aldolase can act as a glucose sensor mediating V-ATPase assembly [76]. Several other determinants of V-ATPase assembly have been identified, including the membrane environment [77] and the interaction with regulatory factors such as HRG-1 [78] and viral infection [34]. The spatial regulation of V-ATPases is observed in the luminal pH gradients between compartments [79][80]. This mechanism of controlling V-ATPase activity is through the regulation of the trafficking of the complex, which is facilitated by different isoforms of the a subunit [81][82].

    4. The V-ATPase a Subunit

    Each V-ATPase complex contains one copy of the ~100 kDa a subunit, which exists as two isoforms in yeast (Vph1p and Stv1p) and four isoforms (a1, a2, a3 and a4) in mammals [27][81][83]. The a subunit has a bipartite structure, with a cytoplasmic N-terminal half (aNT) and a membrane-integrated C-terminal half (aCT) which consists of eight transmembrane helices (Figure 2) [62][84]. As described above, two of the eight helices in aCT are tilted and interact with the proteolipid c-ring to form the two hemichannels for proton translocation [62][85]. Even though ATP hydrolysis-coupled proton translocation can tolerate numerous a subunit mutations, the arginine residue in aCT (R735 in Vph1p, and R740 in TCIRG1 encoding the mammalian a3 isoform) is absolutely essential [69]. The dominant R740S missense mutation of this critical arginine in mice uncouples the proton pumping activity from ATP hydrolysis, resulting in mice with a high bone mineral density [86]. The aNT, oriented parallel to the membrane, is essential for V-ATPase function as it couples V1 ATP hydrolysis to VO proton translocation [44].
    Figure 2. Mammalian V-ATPase a3 subunit. (a) Homology model of the a3 isoform generated using the Phyre2.0 server with constraint coordinates from the mammalian brain a1 isoform (PDB: 6vqc_3); (b) topology of the a3 isoform. The a subunit contains a cytoplasmic N-terminal half (aNT), which can be divided into three sub-domains—a distal domain (DD), connecting stalk (CS) and a proximal domain (PD)—and a membrane-bound C-terminal half (aCT) consisting of eight transmembrane helices (TM1-8), two of which are tilted and form the two hemichannels with the proteolipid c-ring. Cytosolic loops (CL1-3) connect TM2 and 3, TM4 and 5, and TM6 and 7, respectively; luminal loops 1 and 2 (LL1 and LL2) connect TM3 and 4, and TM5 and 6, respectively. Within luminal loop 1, a2 and a3 orthologs are glycosylated twice, whereas a1 and a4 are glycosylated once [66].
    Studies with chimeric forms of Vph1p and Stv1p suggest that organelle targeting information is located in aNT [87]. In yeast, V-ATPases are targeted to the vacuole and Golgi by Vph1p and Stv1p, respectively; when chimeric a subunits were made, the targeted organelles were determined by the aNT. Furthermore, mutagenesis studies revealed that the signal sequence W83KY within the aNT of Stv1p is necessary for V-ATPase Golgi localization [88].
    Similarly, in mammalian cells, different isoforms of the a subunit are enriched in specific organelles or cell types. However, the specific targeting signal of mammalian a isoforms has not been determined. V-ATPases containing the a1 isoform are found in the synaptic vesicles of neurons, and are relocated to the presynaptic plasma membrane at the nerve terminals [60][89]. The a2 isoform targets Golgi [90], and the a3 isoform is expressed in late endosomes and lysosome [16][91]. The a3 and a4 isoforms are also found on the plasma membrane of specialized cells, with a3 targeting the ruffle border of osteoclasts [50][92]; a4 is found in the apical membrane of kidney alpha intercalated cells and epididymal cells [49][93]. The upregulation of both a3 and a4 have been linked to the invasiveness of metastatic breast cancer cells [26]. Recently, the a4 isoform was shown to localize to the membrane of the invapodia of mouse breast cancer cells, where it plays a crucial role in the invasion and migration of the cancer cells [94]. While it is ubiquitously expressed in different organelles and cell types, the expression of a3 is approximately 100-fold greater in osteoclasts than in other cell types [95]. V-ATPases containing a3 are enriched in the membrane of the ruffled border, where they actively pump acid to dissolve bone and provide an acidic environment to activate the secreted proteases required for bone resorption. Furthermore, mutations in the a3 isoform in mammals—for example, the R740S in mice, mentioned above [86]—are associated with V-ATPase-related autosomal recessive osteopetrosis [53][96][97]. To this end, it is clear that the a3 isoform plays a crucial role in bone resorption by osteoclasts; therefore, the a3 isoform is a potential drug target for osteoporosis treatment, in which the excessive bone loss associated with this disease could be controlled by inhibiting a3-containing V-ATPases [50][98][99].

    The entry is from 10.3390/ijms22136934

    References

    1. Lin, X.; Patil, S.; Gao, Y.-G.; Qian, A. The Bone Extracellular Matrix in Bone Formation and Regeneration. Front. Pharmacol. 2020, 11, 757.
    2. Lee, D.; Kim, D.W.; Cho, J.-Y. Correction to: Role of growth factors in hematopoietic stem cell niche. Cell Biol. Toxicol. 2020, 36, 1.
    3. Kitaura, H.; Marahleh, A.; Ohori, F.; Noguchi, T.; Shen, W.-R.; Qi, J.; Nara, Y.; Pramusita, A.; Kinjo, R.; Mizoguchi, I. Osteocyte-Related Cytokines Regulate Osteoclast Formation and Bone Resorption. Int. J. Mol. Sci. 2020, 21, 5169.
    4. Roeder, E.; Matthews, B.G.; Kalajzic, I. Visual reporters for study of the osteoblast lineage. Bone 2016, 92, 189–195.
    5. Prideaux, M.; Findlay, D.M.; Atkins, G.J. Osteocytes: The master cells in bone remodelling. Curr. Opin. Pharmacol. 2016, 28, 24–30.
    6. Creecy, A.; Damrath, J.G.; Wallace, J.M. Control of Bone Matrix Properties by Osteocytes. Front. Endocrinol. 2021, 11, 578477.
    7. Chen, X.; Wang, Z.; Duan, N.; Zhu, G.; Schwarz, E.M.; Xie, C. Osteoblast-osteoclast interactions. Connect. Tissue Res. 2018, 59, 99–107.
    8. Jacome-Galarza, C.E.; Percin, G.I.; Muller, J.T.; Mass, E.; Lazarov, T.; Eitler, J.; Rauner, M.; Yadav, V.K.; Crozet, L.; Bohm, M.; et al. Developmental origin, functional maintenance and genetic rescue of osteoclasts. Nat. Cell Biol. 2019, 568, 541–545.
    9. Yahara, Y.; Barrientos, T.; Tang, Y.J.; Puviindran, V.; Nadesan, P.; Zhang, H.; Gibson, J.R.; Gregory, S.G.; Diao, Y.; Xiang, Y.; et al. Erythromyeloid progenitors give rise to a population of osteoclasts that contribute to bone homeostasis and repair. Nat. Cell Biol. 2020, 22, 49–59.
    10. Søe, K.; Delaisse, J.-M.; Borggaard, X.G. Osteoclast formation at the bone marrow/bone surface interface: Importance of structural elements, matrix, and intercellular communication. Semin. Cell Dev. Biol. 2020, 112, 8–15.
    11. Sims, N.A.; Martin, T.J. Osteoclasts Provide Coupling Signals to Osteoblast Lineage Cells Through Multiple Mechanisms. Annu. Rev. Physiol. 2020, 82, 507–529.
    12. Takito, J.; Nakamura, M. Heterogeneity and Actin Cytoskeleton in Osteoclast and Macrophage Multinucleation. Int. J. Mol. Sci. 2020, 21, 6629.
    13. Gambari, L.; Grassi, F.; Roseti, L.; Grigolo, B.; Desando, G. Learning from Monocyte-Macrophage Fusion and Multinucleation: Potential Therapeutic Targets for Osteoporosis and Rheumatoid Arthritis. Int. J. Mol. Sci. 2020, 21, 6001.
    14. Takito, J.; Inoue, S.; Nakamura, M. The Sealing Zone in Osteoclasts: A Self-Organized Structure on the Bone. Int. J. Mol. Sci. 2018, 19, 984.
    15. Mulari, M.T.K.; Zhao, H.; Lakkakorpi, P.T.; Väänänen, H.K. Osteoclast Ruffled Border Has Distinct Subdomains for Secretion and Degraded Matrix Uptake. Traffic 2003, 4, 113–125.
    16. Toyomura, T.; Murata, Y.; Yamamoto, A.; Oka, T.; Sun-Wada, G.H.; Wada, Y.; Futai, M. From lysosomes to the plasma membrane: Localization of vacuolar-type H+ -ATPase with the a3 isoform during osteoclast differentiation. J. Biol. Chem. 2003, 278, 22023–22030.
    17. Ng, P.Y.; Ribet, A.B.P.; Pavlos, N.J. Membrane trafficking in osteoclasts and implications for osteoporosis. Biochem. Soc. Trans. 2019, 47, 639–650.
    18. Kim, J.H.; Kim, N. Bone Cell Communication Factors Provide a New Therapeutic Strategy for Osteoporosis. Chonnam. Med. J. 2020, 56, 94–98.
    19. Wang, H.; Yang, G.; Xiao, Y.; Luo, G.; Li, G.; Li, Z. Friend or Foe? Essential Roles of Osteoclast in Maintaining Skeletal Health. BioMed Res. Int. 2020, 2020, 1–10.
    20. Corbacho, I.; Teixidó, F.; Olivero, I.; Hernández, L.M. Dependence of Saccharomyces cerevisiae Golgi functions on V-ATPase activity. FEMS Yeast Res. 2012, 12, 341–350.
    21. Nelson, N. Structure and function of V-ATPases in endocytic and secretory organelles. J. Exp. Biol. 1992, 172, 149–153.
    22. Perzov, N.; Padler-Karavani, V.; Nelson, H.; Nelson, N. Characterization of yeast V-ATPase mutants lacking Vph1p or Stv1p and the effect on endocytosis. J. Exp. Biol. 2002, 205, 1209–1219.
    23. Márquez-Sterling, N.; Herman, I.M.; Pesacreta, T.; Arai, H.; Terres, G.; Forgac, M. Immunolocalization of the vacuolar-type (H+)-ATPase from clathrin-coated vesicles. Eur. J. Cell Biol. 1991, 56, 19–33.
    24. Kissing, S.; Hermsen, C.; Repnik, U.; Nesset, C.K.; von Bargen, K.; Griffiths, G.; Ichihara, A.; Lee, B.S.; Schwake, M.; De Brabander, J.; et al. Vacuolar ATPase in Phagosome-Lysosome Fusion. J. Biol. Chem. 2015, 290, 14166–14180.
    25. Sun-Wada, G.-H.; Wada, Y. Role of vacuolar-type proton ATPase in signal transduction. Biochim. et Biophys. Acta (BBA) Bioenerg. 2015, 1847, 1166–1172.
    26. Stransky, L.; Cotter, K.; Forgac, M. The Function of V-ATPases in Cancer. Physiol. Rev. 2016, 96, 1071–1091.
    27. Kibak, H.; Taiz, L.; Starke, T.; Bernasconi, P.; Gogarten, J.P. Evolution of structure and function of V-ATPases. J. Bioenerg. Biomembr. 1992, 24, 415–424.
    28. Sun-Wada, G.-H.; Wada, Y.; Futai, M. Vacuolar H+ pumping ATPases in luminal acidic organelles and extracellular compartments: Common rotational mechanism and diverse physiological roles. J. Bioenerg. Biomembr. 2003, 35, 347–358.
    29. Futai, M.; Oka, T.; Sun-Wada, G.; Moriyama, Y.; Kanazawa, H.; Wada, Y. Luminal acidification of diverse organelles by V-ATPase in animal cells. J. Exp. Biol. 2000, 203, 107–116.
    30. Esmail, S.; Kartner, N.; Yao, Y.; Kim, J.W.; Reithmeier, R.A.; Manolson, M.F. Molecular mechanisms of cutis laxa– and distal renal tubular acidosis–causing mutations in V-ATPase a subunits, ATP6V0A2 and ATP6V0A4. J. Biol. Chem. 2018, 293, 2787–2800.
    31. Mindell, J.A. Lysosomal Acidification Mechanisms. Annu. Rev. Physiol. 2012, 74, 69–86.
    32. Futai, M.; Sun-Wada, G.H.; Wada, Y.; Matsumoto, N.; Nakanishi-Matsui, M. Vacuolar-type ATPase: A proton pump to lysosomal trafficking. Proc. Jpn. Acad. Ser. B Phys. Biol. Sci. 2019, 95, 17.
    33. Lu, X.; Meima, M.E.; Nelson, J.K.; Sorrentino, V.; Loregger, A.; Scheij, S.; Dekkers, D.H.; Mulder, M.T.; Demmers, J.A.; M-Dallinga-Thie, G.; et al. Identification of the (Pro)renin Receptor as a Novel Regulator of Low-Density Lipoprotein MetabolismNovelty and Significance. Circ. Res. 2016, 118, 222–229.
    34. Yeganeh, B.; Ghavami, S.; Kroeker, A.L.; Mahood, T.H.; Stelmack, G.L.; Klonisch, T.; Coombs, K.; Halayko, A.J. Suppression of influenza A virus replication in human lung epithelial cells by noncytotoxic concentrations bafilomycin A1. Am. J. Physiol. Cell. Mol. Physiol. 2015, 308, L270–L286.
    35. Müller, K.H.; E Kainov, D.; El Bakkouri, K.; Saelens, X.; De Brabander, J.K.; Kittel, C.; Samm, E.; Muller, C.P. The proton translocation domain of cellular vacuolar ATPase provides a target for the treatment of influenza A virus infections. Br. J. Pharmacol. 2011, 164, 344–357.
    36. Gary-Bobo, M.; Nirdé, P.; Jeanjean, A.; Morère, A.; Garcia, M. Mannose 6-Phosphate Receptor Targeting and its Applications in Human Diseases. Curr. Med. Chem. 2007, 14, 2945–2953.
    37. Abu-Remaileh, M.; Wyant, G.A.; Kim, C.; Laqtom, N.N.; Abbasi, M.; Chan, S.H.; Freinkman, E.; Sabatini, D.M. Lysosomal metabolomics reveals V-ATPase- and mTOR-dependent regulation of amino acid efflux from lysosomes. Science 2017, 358, 807–813.
    38. Efeyan, A.; Zoncu, R.; Sabatini, D.M. Amino acids and mTORC1: From lysosomes to disease. Trends Mol. Med. 2012, 18, 524–533.
    39. Zoncu, R.; Bar-Peled, L.; Efeyan, A.; Wang, S.; Sancak, Y.; Sabatini, D.M. mTORC1 Senses Lysosomal Amino Acids Through an Inside-Out Mechanism That Requires the Vacuolar H+-ATPase. Science 2011, 334, 678–683.
    40. Stransky, L.A.; Forgac, M. Amino Acid Availability Modulates Vacuolar H+-ATPase Assembly. J. Biol. Chem. 2015, 290, 27360–27369.
    41. Rama, S.; Boumedine-Guignon, N.; Sangiardi, M.; Youssouf, F.; Maulet, Y.; Lévêque, C.; Belghazi, M.; Seagar, M.; Debanne, D.; El Far, O. Chromophore-Assisted Light Inactivation of the V-ATPase V0c Subunit Inhibits Neurotransmitter Release Downstream of Synaptic Vesicle Acidification. Mol. Neurobiol. 2018, 56, 3591–3602.
    42. Morel, N.; Poëa-Guyon, S. The membrane domain of vacuolar H+ATPase: A crucial player in neurotransmitter exocytotic release. Cell. Mol. Life Sci. 2015, 72, 2561–2573.
    43. Morel, N. Neurotransmitter release: The dark side of the vacuolar-H+ATPase. Biol. Cell 2003, 95, 453–457.
    44. Marshansky, V.R.; Grüber, G. Eukaryotic V-ATPase: Novel structural findings and functional insights. Biochim. Biophys. Acta 2014, 1837, 23.
    45. Sun-Wada, G.H.; Toyomura, T.; Murata, Y.; Yamamoto, A.; Futai, M.; Wada, Y. The a3 isoform of V-ATPase regulates insulin secretion from pancreatic beta-cells. J. Cell. Sci. 2006, 119, 4531–4540.
    46. Sun-Wada, G.-H.; Tabata, H.; Kawamura, N. Selective Assembly of V-ATPase Subunit Isoforms in Mouse Kidney. J. Bioenerg. Biomembr. 2005, 37, 415–418.
    47. Brown, D.; Sabolic, I.; Gluck, S. Polarized targeting of V-ATPase in kidney epithelial cells. J. Exp. Biol. 1992, 172, 231–243.
    48. Hermo, L.; I Adamali, H.; Andonian, S. Immunolocalization of CA II and H+ V-ATPase in epithelial cells of the mouse and rat epididymis. J. Androl. 2000, 21, 376–391.
    49. Pietrement, C.; Sun-Wada, G.-H.; Da Silva, N.; McKee, M.; Marshansky, V.; Brown, D.; Futai, M.; Breton, S. Distinct Expression Patterns of Different Subunit Isoforms of the V-ATPase in the Rat Epididymis1. Biol. Reprod. 2006, 74, 185–194.
    50. Qin, A.; Cheng, T.; Pavlos, N.; Lin, Z.; Dai, K.; Zheng, M. V-ATPases in osteoclasts: Structure, function and potential inhibitors of bone resorption. Int. J. Biochem. Cell Biol. 2012, 44, 1422–1435.
    51. Kartner, N.; Manolson, M. V-ATPase subunit interactions: The long road to therapeutic targeting. Curr. Protein Pept. Sci. 2012, 13, 164–179.
    52. Hinton, A.; Bond, S.; Forgac, M. V-ATPase functions in normal and disease processes. Pflügers Arch. Eur. J. Physiol. 2009, 457, 589–598.
    53. Frattini, A.; Orchard, P.J.; Sobacchi, C.; Giliani, S.; Abinun, M.; Mattsson, J.P.; Keeling, D.J.; Andersson, A.-K.; Wallbrandt, P.; Zecca, L.; et al. Defects in TCIRG1 subunit of the vacuolar proton pump are responsible for a subset of human autosomal recessive osteopetrosis. Nat. Genet. 2000, 25, 343–346.
    54. Nakhoul, N.L.; Hamm, L.L. Vacuolar H(+)-ATPase in the kidney. J. Nephrol. 2002, 15, 22–31.
    55. Kartner, N.; Yao, Y.; Li, K.; Crasto, G.J.; Datti, A.; Manolson, M.F. Inhibition of Osteoclast Bone Resorption by Disrupting Vacuolar H+-ATPase a3-B2 Subunit Interaction. J. Biol. Chem. 2010, 285, 37476–37490.
    56. Blair, H.C.; Teitelbaum, S.L.; Ghiselli, R.; Gluck, S. Osteoclastic bone resorption by a polarized vacuolar proton pump. Science 1989, 245, 855–857.
    57. Collins, M.P.; Forgac, M. Regulation of V-ATPase Assembly in Nutrient Sensing and Function of V-ATPases in Breast Cancer Metastasis. Front. Physiol. 2018, 9, 902.
    58. Michel, V.; Munoz, Y.L.; Trujillo, K.; Bisoffi, M.; Parra, K.J. Inhibitors of vacuolar ATPase proton pumps inhibit human prostate cancer cell invasion and prostate-specific antigen expression and secretion. Int. J. Cancer 2013, 132, E1–E10.
    59. Jung, Y.S.; Jun, S.; Kim, M.J.; Lee, S.H.; Suh, H.N.; Lien, E.M.; Jung, H.Y.; Lee, S.; Zhang, J.; Yang, J.I.; et al. TMEM9 promotes intestinal tumorigenesis through vacuolar-ATPase-activated Wnt/beta-catenin signalling. Nat. Cell Biol. 2018, 20, 1421–1433.
    60. Wang, D.; Epstein, D.; Khalaf, O.; Srinivasan, S.; Williamson, W.R.; Fayyazuddin, A.; Quiocho, F.A.; Hiesinger, P.R. Ca2+-Calmodulin regulates SNARE assembly and spontaneous neurotransmitter release via v-ATPase subunit V0a1. J. Cell Biol. 2014, 205, 21–31.
    61. Portela, M.; Yang, L.; Paul, S.; Li, X.; Veraksa, A.; Parsons, L.M.; Richardson, H.E. Lgl reduces endosomal vesicle acidification and Notch signaling by promoting the interaction between Vap33 and the V-ATPase complex. Sci. Signal. 2018, 11, eaar1976.
    62. Abbas, Y.M.; Wu, D.; Bueler, S.A.; Robinson, C.V.; Rubinstein, J.L. Structure of V-ATPase from the mammalian brain. Science 2020, 367, 1240–1246.
    63. Jansen, E.J.; Scheenen, W.J.; Hafmans, T.G.; Martens, G.J. Accessory subunit Ac45 controls the V-ATPase in the regulated secretory pathway. Biochim. Biophys. Acta (BBA) Bioenerg. 2008, 1783, 2301–2310.
    64. Mazhab-Jafari, M.T.; Rohou, A.; Schmidt, C.; Bueler, S.A.; Benlekbir, S.; Robinson, C.; Rubinstein, J.L. Atomic model for the membrane-embedded VO motor of a eukaryotic V-ATPase. Nat. Cell Biol. 2016, 539, 118–122.
    65. Kawasaki-Nishi, S.; Nishi, T.; Forgac, M. Proton translocation driven by ATP hydrolysis in V-ATPases. FEBS Lett. 2003, 545, 76–85.
    66. Esmail, S.; Kartner, N.; Yao, Y.; Kim, J.W.; Reithmeier, R.A.F.; Manolson, M.F. N-linked glycosylation of a subunit isoforms is critical for vertebrate vacuolar H(+) -ATPase (V-ATPase) biosynthesis. J. Cell Biochem. 2018, 119, 861–875.
    67. Flannery, A.R.; Graham, L.A.; Stevens, T.H. Topological Characterization of the c, c′, and c″ Subunits of the Vacuolar ATPase from the Yeast Saccharomyces cerevisiae. J. Biol. Chem. 2004, 279, 39856–39862.
    68. Wang, Y.; Cipriano, D.J.; Forgac, M. Arrangement of Subunits in the Proteolipid Ring of the V-ATPase. J. Biol. Chem. 2007, 282, 34058–34065.
    69. Kawasaki-Nishi, S.; Nishi, T.; Forgac, M. Arg-735 of the 100-kDa subunit a of the yeast V-ATPase is essential for proton translocation. Proc. Natl. Acad. Sci. USA 2001, 98, 12397–12402.
    70. Oot, R.A.; Kane, P.M.; Berry, E.A.; Wilkens, S. Crystal structure of yeast V1-ATPase in the autoinhibited state. EMBO J. 2016, 35, 1694–1706.
    71. Smardon, A.M.; Tarsio, M.; Kane, P.M. The RAVE Complex Is Essential for Stable Assembly of the Yeast V-ATPase. J. Biol. Chem. 2002, 277, 13831–13839.
    72. Kane, P.M. Targeting Reversible Disassembly as a Mechanism of Controlling V-ATPase Activity. Curr. Protein Pept. Sci. 2012, 13, 117–123.
    73. Sharma, S.; Oot, R.A.; Wilkens, S. MgATP hydrolysis destabilizes the interaction between subunit H and yeast V1-ATPase, highlighting H’s role in V-ATPase regulation by reversible disassembly. J. Biol. Chem. 2018, 293, 10718–10730.
    74. Lu, M.; Holliday, L.S.; Zhang, L.; Dunn, W.A., Jr.; Gluck, S.L. Interaction between aldolase and vacuolar H+-ATPase: Evidence for direct coupling of glycolysis to the ATP-hydrolyzing proton pump. J. Biol. Chem. 2001, 276, 30407–30413.
    75. Lu, M.; Sautin, Y.; Holliday, L.S.; Gluck, S.L. The Glycolytic Enzyme Aldolase Mediates Assembly, Expression, and Activity of Vacuolar H+-ATPase. J. Biol. Chem. 2004, 279, 8732–8739.
    76. Li, M.; Zhang, C.-S.; Zong, Y.; Feng, J.-W.; Ma, T.; Hu, M.; Lin, Z.; Li, X.; Xie, C.; Wu, Y.; et al. Transient Receptor Potential V Channels Are Essential for Glucose Sensing by Aldolase and AMPK. Cell Metab. 2019, 30, 508–524.
    77. Banerjee, S.; Kane, P.M. Regulation of V-ATPase Activity and Organelle pH by Phosphatidylinositol Phosphate Lipids. Front. Cell Dev. Biol. 2020, 8, 510.
    78. Fogarty, F.M.; O’Keeffe, J.; Zhadanov, A.; Papkovsky, D.; Ayllón, V.; O’Connor, R. HRG-1 enhances cancer cell invasive potential and couples glucose metabolism to cytosolic/extracellular pH gradient regulation by the vacuolar-H+ ATPase. Oncogene 2013, 33, 4653–4663.
    79. Marshansky, V.; Futai, M. The V-type H+-ATPase in vesicular trafficking: Targeting, regulation and function. Curr. Opin. Cell Biol. 2008, 20, 415–426.
    80. Qi, J.; Wang, Y.; Forgac, M. The vacuolar (H+)-ATPase: subunit arrangement and in vivo regulation. J. Bioenerg. Biomembr. 2007, 39, 423–426.
    81. Rahman, S.; Yamato, I.; Murata, T. Function and Regulation of Mammalian V-ATPase Isoforms. In Regulation of Ca2+-ATPases, V-ATPases and F-ATPases; Springer Science and Business Media LLC: Cham, Switzerland, 2016; Volume 14, pp. 283–299.
    82. Lafourcade, C.; Sobo, K.; Kieffer-Jaquinod, S.; Garin, J.; Van Der Goot, F.G. Regulation of the V-ATPase along the Endocytic Pathway Occurs through Reversible Subunit Association and Membrane Localization. PLoS ONE 2008, 3, e2758.
    83. Manolson, M.F.; Wu, B.; Proteau, D.; Taillon, B.E.; Roberts, B.T.; Hoyt, M.A.; Jones, E.W. STV1 gene encodes functional homologue of 95-kDa yeast vacuolar H(+)-ATPase subunit Vph1p. J. Biol. Chem. 1994, 269, 14064–14074.
    84. Srinivasan, S.; Vyas, N.K.; Baker, M.L.; Quiocho, F.A. Erratum to “Crystal Structure of the Cytoplasmic N-Terminal Domain of Subunit I, a Homolog of Subunit a, of V-ATPase” [J. Mol. Biol. 412/1 (2011) 14–21]. J. Mol. Biol. 2011, 413, 523.
    85. Kawasaki-Nishi, S.; Nishi, T.; Forgac, M.; Garofano, A.; Zwicker, K.; Kerscher, S.; Okun, P.; Brandt, U. Interacting Helical Surfaces of the Transmembrane Segments of Subunits a and c′ of the Yeast V-ATPase Defined by Disulfide-mediated Cross-linking. J. Biol. Chem. 2003, 278, 41908–41913.
    86. Ochotny, N.; Flenniken, A.M.; Owen, C.; Voronov, I.; A Zirngibl, R.; Osborne, L.R.; E Henderson, J.; Adamson, S.L.; Rossant, J.; Manolson, M.; et al. The V-ATPase a3 subunit mutation R740S is dominant negative and results in osteopetrosis in mice. J. Bone Miner. Res. 2011, 26, 1484–1493.
    87. Kawasaki-Nishi, S.; Bowers, K.; Nishi, T.; Forgac, M.; Stevens, T.H. The Amino-terminal Domain of the Vacuolar Proton-translocating ATPase a Subunit Controls Targeting and in Vivo Dissociation, and the Carboxyl-terminal Domain Affects Coupling of Proton Transport and ATP Hydrolysis. J. Biol. Chem. 2001, 276, 47411–47420.
    88. Finnigan, G.C.; Cronan, G.E.; Park, H.J.; Srinivasan, S.; Quiocho, F.A.; Stevens, T.H. Sorting of the yeast vacuolar-type, proton-translocating ATPase enzyme complex (V-ATPase): Identification of a necessary and sufficient Golgi/endosomal retention signal in Stv1p. J. Biol. Chem. 2012, 287, 19487–194500.
    89. Zhang, W.; Wang, D.; Volk, E.; Bellen, H.J.; Hiesinger, P.R.; Quiocho, F.A. V-ATPase V0 sector subunit a1 in neurons is a target of calmodulin. J. Biol. Chem. 2008, 283, 294–300.
    90. Saw, N.M.N.; Kang, S.-Y.A.; Parsaud, L.; Han, G.A.; Jiang, T.; Grzegorczyk, K.; Surkont, M.; Sun-Wada, G.-H.; Wada, Y.; Li, L.; et al. Vacuolar H+-ATPase subunits Voa1 and Voa2 cooperatively regulate secretory vesicle acidification, transmitter uptake, and storage. Mol. Biol. Cell 2011, 22, 3394–3409.
    91. Matsumoto, N.; Sekiya, M.; Tohyama, K.; Ishiyama-Matsuura, E.; Sun-Wada, G.-H.; Wada, Y.; Futai, M.; Nakanishi-Matsui, M. Essential Role of the a3 Isoform of V-ATPase in Secretory Lysosome Trafficking via Rab7 Recruitment. Sci. Rep. 2018, 8, 1–18.
    92. Matsumoto, N.; Daido, S.; Sun-Wada, G.-H.; Wada, Y.; Futai, M.; Nakanishi-Matsui, M. Diversity of proton pumps in osteoclasts: V-ATPase with a3 and d2 isoforms is a major form in osteoclasts. Biochim. et Biophys. Acta (BBA) Bioenerg. 2014, 1837, 744–749.
    93. Oka, T.; Murata, Y.; Namba, M.; Yoshimizu, T.; Toyomura, T.; Yamamoto, A.; Sun-Wada, G.-H.; Hamasaki, N.; Wada, Y.; Futai, M. a4, a Unique Kidney-specific Isoform of Mouse Vacuolar H+-ATPase Subunit a. J. Biol. Chem. 2001, 276, 40050–40054.
    94. McGuire, C.M.; Collins, M.P.; Sun-Wada, G.; Wada, Y.; Forgac, M. Isoform-specific gene disruptions reveal a role for the V-ATPase subunit a4 isoform in the invasiveness of 4T1-12B breast cancer cells. J. Biol. Chem. 2019, 294, 11248–11258.
    95. Toyomura, T.; Oka, T.; Yamaguchi, C.; Wada, Y.; Futai, M. Three subunit a isoforms of mouse vacuolar H(+)-ATPase. Preferential expression of the a3 isoform during osteoclast differentiation. J. Biol. Chem. 2000, 275, 8760–8765.
    96. Kornak, U.; Schulz, A.; Friedrich, W.; Uhlhaas, S.; Kremens, B.; Voit, T.; Hasan, C.; Bode, U.; Jentsch, T.J.; Kubisch, C. Mutations in the a3 subunit of the vacuolar H(+)-ATPase cause infantile malignant osteopetrosis. Hum. Mol. Genet. 2000, 9, 2059–2063.
    97. Zirngibl, R.A.; Wang, A.; Yao, Y.; Manolson, M.; Krueger, J.; Dupuis, L.; Mendoza-Londono, R.; Voronov, I. Novel c.G630A TCIRG1 mutation causes aberrant splicing resulting in an unusually mild form of autosomal recessive osteopetrosis. J. Cell. Biochem. 2019, 120, 17180–17193.
    98. Duan, X.; Yang, S.; Zhang, L.; Yang, T. V-ATPases and osteoclasts: Ambiguous future of V-ATPases inhibitors in osteoporosis. Theranostics 2018, 8, 5379–5399.
    99. Kartner, N.; Manolson, M.F. The Vacuolar Proton ATPase (V-ATPase): Regulation and Therapeutic Targeting. In Regulation of Ca2+-ATPases, V-ATPases and F-ATPases, 1st ed.; Chakraborti, S., Dhalla, N.S., Eds.; Springer International Publishing: Cham, Switzerland, 2016; Volume 14, pp. 407–437.
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