3.A.3 The P-type ATPase (P-ATPase) Superfamily

Nearly all of the members of this superfamily, found in bacteria, archaea and eukaryotes, catalyze cation uptake and/or efflux driven by ATP hydrolysis. Clustering on the phylogenetic tree is usually in accordance with specificity for the transported ion(s). Many of these protein complexes are multisubunit with a large subunit serving the primary ATPase and ion translocation functions. In eukaryotes, they are present in the plasma membranes or endoplasmic reticular membranes. In prokaryotes, they are localized to the cytoplasmic membranes. Gastric H+-translocating ATPases (ouabain-insensitive) comprise a subgroup of the larger and more diverse Na+/K+ ATPase family (ouabain-sensitive) (Family 1). The ouabain-binding pocket can be functionally reconstituted in the gastric H+,K+ ATPase by substitution of only seven residues (Qiu et al., 2005). Ca2+ ATPases of prokaryotes and eukaryotes comprise a very diverse family (Family 2) including in eukaryotes plasma membrane, golgi, and sarcoplasmic reticular types. Sarcoplasmic reticular Ca2+ ATPases (SERCA) in brown adipose tissue can uncouple ATP hydrolysis from Ca2+ transport and be thermogenic (de Meis, 2003). H+-translocating P-type ATPases of plants and fungi comprise their own family (Family 3). Plant P-ATPases have been reviewed by Wdowikowska and Klobus, (2011). Distinct bacterial enzymes specific for K+ (Family 7; only in prokaryotes) or Mg2+ (Family 4, mostly in prokaryotes; uptake), Cu2+, Ag+, Zn2+, Co2+, Pb2+, Ni2+, and/or Cd2+ (Family 6; efflux) and Cu2+ or Cu+ (Family 5; uptake or efflux, depending on the system) have been characterized, and each of these types of enzymes comprises a distinct family. Cu2+ or Cu+-translocating ATPases from bacteria, archaea and animals cluster together, and at least some of these also transport Ag+. The Cu+/Ag+ (Family 5 and heavy metal (Family 6)) ATPases have an 8 TMS topology (Mandal et al., 2002). A cys-pro-cys motif in CopA of E. coli (TC #3.A.3.5.5) is essential for Cu+/Ag+ efflux and phosphoenzyme formation (Fan and Rosen, 2002).

P-type ATPases play essential roles in numerous processes, which in humans include nerve impulse propagation, relaxation of muscle fibers, secretion and absorption in the kidney, acidification of the stomach and nutrient absorption in the intestine. Published evidence suggests that uncharacterized families of P-type ATPases with novel specificities exist. Thever & Saier (2009) analyzed the fully sequenced genomes of 26 eukaryotes including animals, plants, fungi and unicellular eukaryotes for P-type ATPases. They reported the organismal distributions, phylogenetic relationships, probable topologies and conserved motifs of nine functionally characterized families and 13 uncharacterized families of these enzyme transporters. A similar but more extensive study on Prokaryotic P-type ATPase was presented by Chan et al. 2010 (see below). Family 9 Na+- or K+-ATPases can be found in fungi, plants bryophytes and protozoa (Rodríguez-Navarro and Benito, 2010). The Na+/K+-ATPase acts as a receptor for cardiotonic steroids such as ouabain. Cardiotonic steroids inhibit this enzyme's activity, resulting in modulation of Ca2+ levels, which are essential for homeostasis in neurons. Two points regarding the interplay between the Na+/K+-ATPase and Ca2+ signalling in the brain are (1) Na+/K+-ATPase impairment causes illness and neuronal death due to deficient Ca2+ signalling and (2) benefits to the brain result from modulating Na+/K+-ATPase activity. These interactions play an essential role in neuronal cell fate determination (Kinoshita et al. 2022).

Chan et al. (2010) analyzed P-type ATPases in all major prokaryotic phyla for which complete genome sequence data were available and compared the results with those for eukaryotic P-type ATPases. Topological type I (heavy metal) P-type ATPases predominate in prokaryotes (approx. tenfold) while type II ATPases (specific for Na+,K+, H+ Ca2+, Mg2+ and phospholipids) predominate in eukaryotes (approx. twofold). Many P-type ATPase families are found exclusively in prokaryotes (e.g. Kdp-type K+ uptake ATPases (type III) and all ten prokaryotic functionally uncharacterized P-type ATPase (FUPA) familes), while others are restricted to eukaryotes (e.g. phospholipid flippases and all 13 eukaryotic FUPA families) (Thever and Saier, 2009). Horizontal gene transfer has occurred frequently among bacteria and archaea, which have similar distributions of these enzymes, but rarely between most eukaryotic kingdoms, and even more rarely between eukaryotes and prokaryotes. In some bacterial phyla (e.g. Bacteroidetes, Flavobacteria and Fusobacteria), ATPase gene gain and loss as well as horizontal transfer occurred seldom in contrast to most other bacterial phyla. Some families (i.e., Kdp-type ATPases) underwent far less horizontal gene transfer than other prokaryotic families, possibly due to their multisubunit characteristics. Functional motifs are better conserved across family lines than across organismal lines, and these motifs can be family specific, facilitating functional predictions. In some cases, gene fusion events created P-type ATPases covalently linked to regulatory catalytic enzymes. In one family (FUPA Family 24), a type I ATPase gene (N-terminal) is fused to a type II ATPase gene (C-terminal) with retention of function only for the latter. Several pseudogene-encoded nonfunctional ATPases were identified. Genome minimalization led to preferential loss of P-type ATPase genes. Chan et al. (2010) suggested that in prokaryotes and some unicellular eukaryotes, the primary function of P-type ATPases is protection from extreme environmental stress conditions. The classification of P-type ATPases of unknown function into phylogenetic families provides guides for future molecular biological studies (Chan et al., 2010).

Many eukaryotic P-type ATPases are monomeric or homodimeric enzymes of the catalytic subunit that hydrolyzes ATP. They contain the aspartyl phosphorylation site and catalyzes ion transport. The Na+,K+-ATPases, the Ca2+-ATPases and the (fungal) H+-ATPases of higher organisms exhibit 10 transmembrane α helical spanners (TMSs), some of them highly tilted. Additional subunits that appear to lack catalytic activity may be present in the ATPase complex. For example, the 10 TMS catalytic α-subunit of the Na+,K+-ATPase of animals is tightly complexed to the 1 TMS β-subunit and the tissue-specific, regulatory, 1 TMS γ-subunit. The β-subunit, which may influence the activity of the α-subunit, probably functions to facilitate proper insertion of the α-subunit into the membrane, to allow proper targeting to a subcellular membrane site in post-translational processing, and to stabilize the catalytic subunit. The β-subunit can therefore be considered to be an auxiliary protein of the Na+,K+-ATPase catalytic subunit. The γ-subunit of the Na+,K+-ATPase has been reported to influence kinetic parameters and is homologous to a family of pore-forming peptides, the peptides of the phospholemman family (TC #1.A.27), and the C-subunits of V-type ATPases (TC #3.A.2). This γ-subunit is induced under stress conditions and modulates Na+,K+-ATPase activity and cell growth (Wetzel et al., 2004). The Na+, K+-ATPase can serve as a steroid hormone receptor (Schoner, 2002). Several other P-type ATPases also depend on small proteolipids, the functions of which are uncertain.

The annular lipid-protein stoichiometry in a native pig kidney Na+/K+ -ATPase preparation has been studied by [125I]TID-PC/16 labeling, giving results that indicated that the transmembrane domain of the Na+/K+ -ATPase in the E1 state is less exposed to the lipids than in the E2 state, i.e., the conformational transitions are accompanied by changes in the numbers of annular lipids but not in the affinity of these lipids for the protein (Mangialavori et al. 2011). The lipid-protein stoichiometry was 23 ± 2 (α subunit) and 5.0 ± 0.4 (β subunit) in the E1 conformation and 32 ± 2 (α subunit) and 7 ± 1 (β subunit) in the E2 conformation.

The stoichiometries of transport are sometimes known and complex. In the case of the Na+,K+-ATPases, 3 Na+ are exchanged for 2 K+ per ATP molecule hydrolyzed. The gastric H+-translocating ATPases replace H+ for K+ but with an H+/K+ stoichiometry of 2:2. Thus, although these two enzymes are ~65% identical, the Na+,K+-ATPases are electrogenic while the H+,K+-ATPases are electroneutral. Gastric H+, K+-ATPase transports 2 moles of H+ together with two H2O (two H3O+) per mole of ATP hydrolyzed in isolated hog gastric vesicles. Protons are charge-transferred from the cytosolic side to H2O in sites 2 and 1, the H2O coming from the cytosol, and H3O+ in these sites are transported into the lumen during the conformational transition from E1P to E2P (Morii et al., 2008). Ca2+ ATPases may catalyze Ca2+/K+ or Ca2+/H+ antiport. A single organism often possesses multiple isoforms of these enzymes. Caloxin-derived peptides in the micromolar range are inhibitors of plasma membrane calcium ATPases (Boutin et al. 2022).

Considerable evidence is available showing that animals have Cl- translocating, Cl- stimulating P-type ATPases. Although extensive biochemical data are available, the protein sequence of any one such Cl- ATPase has not yet been determined (Gerencser, 1993; Inagaki et al., 1996; Zeng et al., 1999). Evidence for mammalian iron-inducible, iron-transporting ATPases is also available (Baranano et al., 2000). Finally bacterial Na+-transporting P-type ATPases probably exist (Ueno et al., 2000). Evidence for a Na+-P-type ATPase has been obtained for the halotolerant cyanobacterium, Aphanothece halophytica (Wiangon et al., 2007). Thus the breadth of substrates transported by P-type ATPases is likely to be much greater than currently recognized.

The Na+,K+-ATPase acts as a signal transducer and transcription activator, modulating cell growth, apoptosis, and cell motility. A prominent binding motif linking the Na+,K+-ATPase to intracellular signaling effectors is the N-terminal tail of the Na+,K+-ATPase catalytic α-subunit which binds directly to the N-terminus of the inositol 1,4,5-trisphosphate receptor (Zhang et al., 2006). Three amino acyl residues, LKK, conserved in most species and most α-isoforms, are essential for binding. In wild-type cells, low concentrations of ouabain trigger low frequency calcium oscillations that activate NF-κB and protect from apoptosis. Thus, the LKK motif binds the inositol 1,4,5-trisphosphate receptor and triggers an anti-apoptotic calcium signal. However, the N-terminal hydrophilic region in front of the first TMS does not interact with the transported cation (Na+, K+, or Ca2+) although this first TMS does (Einholm et al., 2007). Of the P-type ATPases, only Na+, K+-ATPases are receptors that respond to endogenous cardiotonic steroids such as ouabain and marinobufagenin (steroid 'hormones') which regulate Na+ excretion and blood pressure (Liu and Xie, 2010).

P-type ATPases provide a polar transmembrane pathway, to which access is strictly controlled by coupled gates that are constrained to open alternately, thereby enabling thermodynamically uphill ion transport. Reyes and Gadsby (2006) have examined the ion pathway through the N+, K+-ATPase, a representative P-type pump, after uncoupling its extra- and intracellular gates with the marine toxin palytoxin. They found a wide outer vestibule penetrating deep into the Na+, K+-ATPase, where the pathway narrows and leads to a charge-selectivity filter. Acidic residues in this region, which are conserved to coordinate pumped ions, allow the approach of cations but exclude anions. Reversing the charge at just one of those positions converts the pathway from cation selective to anion selective. Cysteine scans from TM1 to TM6 in the Na+, K+-ATPase revealed a single unbroken cation pathway that traverses palytoxin-bound Na+,K+-pump-channels from one side of the membrane to the other (Takeuchi et al. 2008). This pathway comprises residues from TM1, TM2, TM4 and TM6, passes through ion-binding site II, and is probably conserved in structurally and evolutionarily related P-type pumps. Close structural homology among the catalytic subunits of Ca2+-, Na+, K+- and H+, K+-ATPases argues that their extracytosolic cation exchange pathways all share these physical characteristics (Reyes and Gadsby, 2006). The mechanistic details of type II ATPases, notably those for which 3-d structures are available (Na, K+-, gastri H+, K+-, Ca2+ and H+, K+-ATPase); TC Families 1,2 and 3, respectively, as well as prepared cation translocation pathways, have been discussed by Bublitz et al. (2010). P-type ATPase in several Rosaceae species including the pear have been identified (Zhang et al. 2020).

The X-ray crystal structure at 3.5 Å resolution of the pig renal Na+,K+-ATPase has been determined with two rubidium ions bound in an occluded state in the transmembrane part of the α-subunit (Morth et al., 2007). Several of the residues forming the cavity for rubidium/potassium occlusion in the Na+,K+-ATPase are homologous to those binding calcium in the Ca2+-ATPase of the sarco(endo)plasmic reticulum. The β- and γ-subunits specific to the Na+,K+-ATPase are associated with transmembrane helices αM7/αM10 and αM9, respectively. The γ-subunit corresponds to a fragment of the V-type ATPase c subunit. The carboxy terminus of the α-subunit is contained within a pocket between transmembrane helices and seems to be a novel regulatory element controlling sodium affinity, possibly influenced by the membrane potential.

The Na+,K+-ATPase can be transformed into an ion channel using pharmacological agents. Palytoxin (PTX), produced by soft coral of the genus Palythoa, binds to the ATPase with a Kd of 20 pM and creates a monovalent cation-selective channel with a single channel conductance of 10 pS (Rossini and Bigiani, 2011). The presence of external Na+ seems to be essential for channel activation (Wu et al., 2003). When the N-terminal 35 residues are removed from the ATPase, the toxin-activated channel does not exhibit a time-dependent inactivation gating at positive potentials as is characteristic of the wild-type protein. The truncated pump exhibits no electrogenic current, and the ion stoichiometry for active transport is altered. Addition of the synthetic peptide restores activity towards wild type. The N-terminal peptide therefore appears to act as an inactivation gate (similar to Shaker B channels of the VIC family (TC #1.A.1)). It may also play a critical role in determining the ion stoichiometry (Wu et al., 2003). Fluorometric studies indicate that under normal conditions, α- and β-subunits move towards each other during the E2 to E1 transition (Dempski et al., 2006).

The structures of the sarcoplasmic reticular Ca2+-ATPase have been solved at 2.6 Å resolution for the complex to which 2 Ca2+ are bound, and at 3.1 Å resolution for the complex lacking Ca2+ (Toyoshima et al., 2000; Toyoshima and Nomura, 2002). A total of eight different states of the Ca2+ -ATPase, representing the many steps in the reaction cycle, have been visualized by high resolution x-ray crystallography (Toyoshima et al., 2007; Toyoshima, 2008). The two Ca2+ are located side by side, surrounded by 4 transmembrane helices, two of which are unwound for efficient coordination geometry. There are 3 cytoplasmic domains, one, the central catalytic domain, bearing the phosphorylation site, a second bearing the adenosine nucleotide binding site, and a third of unknown function. The central domain has the same fold as haloacid dehydrogenases (Aravind et al., 1998; Stokes and Green, 2000). The Ca2+-free form shows large conformational differences from the Ca2+-bound form with the three cytoplasmic domains tightly associated to form a single headpiece and six of the ten TMSs largely rearranged. These latter rearrangements guarantee the release of external Ca2+ and create a pathway for entry of Ca2+ from the cytoplasm. ATPase activity and Ca2+ binding are cooperatively interdependent, but the two processes can be separated by mutations (Zhang et al., 2002).

Structures are available for both the E1 and E2 states of the Ca2+ ATPase showing that Ca2+ binding induces major changes in all three cytoplasmic domains relative to each other (Xu et al., 2002). Xu et al. proposed how Ca2+ binding induces conformational changes in TMS4 and 5 in the membrane domain (M) that in turn induce rotation of the phosphorylation domain (P). The nucleotide binding (N) and β-sheet (β) domains are highly mobile, with N flexibly linked to P, and β flexibly linked to M. Modeling of the fungal H+ ATPase, based on the structures of the Ca2+ pump, suggested a comparable 70º rotation of N relative to P to deliver ATP to the phosphorylation site (Kühlbrandt et al., 2002). One report suggests that this S.R. Ca2+ ATPase is homodimeric (Ushimaru and Fukushima, 2008).

Crystal structures (Gadsby, 2007) have shown that the conserved TGES loop of the Ca2+-ATPase is isolated in the Ca2E1 state but becomes inserted in the catalytic site in E2 states. Anthonisen et al. (2006) characterized the kinetics of the partial reaction steps of the transport cycle and the binding of the phosphoryl analogs BeF, AlF, MgF, and vanadate in mutants with alterations to the TGES residues. The data provide functional evidence supporting a role of Glu183 in activating the water molecule involved in the E2P → E2 dephosphorylation and suggest a direct participation of the side chains of the TGES loop in the control and facilitation of the insertion of the loop in the catalytic site. The interactions of the TGES loop furthermore seem to facilitate its disengagement from the catalytic site during the E2 → Ca2E1 transition.

Olesen et al. (2007) have described functional studies and three new crystal structures of the rabbit skeletal muscle Ca2+-ATPase. These represent the phosphoenzyme intermediates associated with (1) Ca2+ binding, (2) Ca2+ translocation and (3) dephosphorylation. They are based on complexes with a functional ATP analogue, beryllium fluoride or aluminium fluoride. The structures complete the cycle of nucleotide binding and cation transport of Ca2+-ATPase. Phosphorylation of the enzyme triggers a conformational change that leads to opening of a luminal exit pathway defined by the transmembrane segments M1 through M6. M1-M6 represent the canonical membrane domain of P-type pumps. Ca2+ release is promoted by translocation of the M4 helix, exposing Glu 309, Glu 771 and Asn 796 to the lumen. The mechanism explains how P-type ATPases are able to form the steep electrochemical gradients required for key functions in eukaryotic cells (Olesen et al., 2007). Moller et al. (2010) reveiwed structural studies of various conformers of the Ca2+ ATPase, (SERCA1a), present in skeletal muscle. The structures corresponding to the various intermediary states. They have been obtained after stabilization with structural analogues of ATP or metal fluorides as mimicks of inorganic phosphate. It is possible to provide a detailed structural description of both ATP hydrolysis and Ca2+ transport through the membrane.

The structure of a P-type proton pump was determined by X-ray crystallography by Pederson et al., (2007). Ten transmembrane helices and three cytoplasmic domains define the functional unit of ATP-coupled proton transport across the plasma membrane. The structure is locked in a functional state not previously observed in P-type ATPases. The transmembrane domain reveals a large cavity, which is likely to be filled with water, located near the middle of the membrane plane where it is lined by conserved hydrophilic and charged residues. Proton transport against a high membrane potential is readily explained by this structural arrangement. The plasma membrane H+-ATPase, SmPHA4, negatively regulates the biosynthesis of tanshinones in Salvia miltiorrhiza.(Li et al. 2021).

Detailed high resolution x-ray structures of heavy metal P1B-type ATPases were not available prior to 2011 when Gourdon et al. (2011) reported the structure of CopA, a Cu+-ATPase from Legionella pneumophile at 3.2 Å resolution. The results provided the first in depth description of a heavy metal-translocatin P1B-type ATPase. A three-stage copper transport pathway involves several well conserved residues. A P1B-specific transmembrane helix kinks at a double-glycine motif displaying an amphipathic helix that lines a putative copper entry point at the intracellular interface. An ATPase-coupled copper release mechanism from the binding sites in the membrane via an extracellular exit site is probable (Gourdon et al. 2011).

As in other P-type ATPases, metal binding to transmembrane metal-binding sites (TM-MBS) in Cu+-ATPases is required for enzyme phosphorylation and subsequent transport. However, Cu+ does not access Cu+-ATPases in a free (hydrated) form but is bound to a chaperone protein. The delivery of Cu+ by Archaeoglobus fulgidus Cu+ chaperone CopZ to the corresponding Cu+-ATPase, CopA, has been studied (González-Guerrero and Argüello, 2008). CopZ interacted with and delivered the metal to the N-terminal metal binding domain(s) of CopA (MBDs). Cu+-loaded MBDs, acting as metal donors, were unable to activate CopA or a truncated CopA lacking MBDs. Conversely, Cu+-loaded CopZ activated the CopA ATPase and CopA constructs in which MBDs were rendered unable to bind Cu+. Furthermore, under nonturnover conditions, CopZ transferred Cu+ to the TM-MBS of a CopA lacking MBDs altogether. Thus, MBDs may serve a regulatory function without participating directly in metal transport, and the chaperone delivers Cu+ directly to transmembrane transport sites of Cu+-ATPases (González-Guerrero and Argüello, 2008). Wu et al (2008) have determined structures of two constructs of the Cu (CopA) pump from Archaeoglobus fulgidus by cryoelectron microscopy of tubular crystals, which revealed the overall architecture and domain organization of the molecule. They localized its N-terminal MBD within the cytoplasmic domains that use ATP hydrolysis to drive the transport cycle and built a pseudoatomic model by fitting existing crystallographic structures into the cryoelectron microscopy maps for CopA. The results also similiarly suggested a Cu-dependent regulatory role for the MBD.

In the Archaeoglobus fulgidus CopA (TC# 3.A.3.5.7), invariant residues in helixes 6, 7 and 8 form two transmembrane metal binding sites (TM-MBSs). These bind Cu+ with high affinity in a trigonal planar geometry. The cytoplasmic Cu+ chaperone CopZ transfers the metal directly to the TM-MBSs; however, loading both of the TM-MBSs requires binding of nucleotides to the enzyme. In agreement with the classical transport mechanism of P-type ATPases, occupancy of both transmembrane sites by cytoplasmic Cu+ is a requirement for enzyme phosphorylation and subsequent transport into the periplasmic or extracellular milieu. Transport studies have shown that most Cu+-ATPases drive cytoplasmic Cu+ efflux, albeit with quite different transport rates in tune with their various physiological roles. Archetypical Cu+-efflux pumps responsible for Cu+ tolerance, like the Escherichia coli CopA, have turnover rates ten times higher than those involved in cuproprotein assembly (or alternative functions). This explains the incapability of the latter group to significantly contribute to the metal efflux required for survival in high copper environments.  Structural and mechanistic details of copper-transporting P-type ATPase functionhave been described (Meng et al. 2015).

Chintalapati et al. (2008) have characterized two copper-transporting ATPases, CtrA2 and CtrA3 from Aquifex aeolicus. CtrA2 has a CPC metal-binding sequence in TM6 and a CxxC metal-binding N-terminal domain, while CtrA3 has a CPH metal-binding motif in TM6 and a histidine-rich N-terminal metal-binding domain. CtrA2 is activated by Ag+ and Cu+ and presumably transports reduced Cu+, while CtrA3 is activated by, and presumably transports, the oxidized copper ion. Both CtrA2 and CtrA3 are thermophilic proteins with an activity maximum at 75 degrees C. Electron cryomicroscopy of two-dimensional crystals of CtrA3 yielded a projection map at approximately 7 A resolution with density peaks indicating eight membrane-spanning alpha-helices per monomer. A fit of the Ca-ATPase structure to the projection map indicates that the arrangement of the six central helices surrounding the ion-binding site in the membrane is conserved, and suggests the position of the two additional N-terminal transmembrane helices that are characteristic of the heavy metal, eight-helix P(1B)-type ATPases (Chintalapati et al., 2008). 

Transmembrane helices contain a cation-binding cysteine-proline-cysteine/histidine/serine (CPx) motif for catalytic activation and cation translocation. In addition, most Cu-ATPases possess the N-terminal Cu-binding CxxC motif required for regulation of enzyme activity. In cells, the Cu- ATPases receive copper from soluble chaperones and maintain intracellular copper homeostasis by efflux of copper from the cell or transport of the metal into the intracellular compartments (Migocka 2015).

The 8TMS CadA of Listeria monocytogenes (Family 6) confirs resistance to cadmium. Residues in TMS6 (Cys354 and Cys356), TMS8 (Asp692) and TMS3 (Met149) may bind Cd2+ (Wu et al., 2006). However, the two cysteine residues in the CPC motif act at different steps: Cys354 is involved in Cd2+ binding while Cys356 is involved in Cd2+ occlusion. The two equivalent cysteines in the yeast Cu2+ ATPase may also act at different steps. The conserved Glu164 may be required for Cd2+ release. Possibly two Cd2+ are involved in the reaction cycle of CadA (Wu et al., 2006).  A hemerythrin-like two iron-binding domain in a P1B-type transport ATPase from Acidothermus cellulolyticus has been identified (Traverso et al., 2010).

The phospholipid translocating P-type ATPases (Family 8; TC #3.A.3.8) are found only in eukaryotes. They appear to function with β-subunits of about 400 aas with 2 TMSs. These have been functionally characterized from yeast and protozoans (TC #8.A.27; Perez-Victoria et al., 2006). These enzymes catalyze the ATP-dependent flipping of phospholipids and lysophospholipids from the outer leaflet of the cytoplasmic membrane to the inner leaflet. Residues defining phospholipid flippase substrate specificity have been identified (Baldridge and Graham, 2012). In the yeast Saccharomyces cerevisiae, Family 3.A.3.8 lipid flipping ATPases play a pivotal role in the biogenesis of intracellular transport vesicles, polarized protein transport and protein maturation.  However, in mammals, these ATPases act in concert with members of the CDC50 protein family, putative beta-subunits for these ATPases, and many function as part of the vesicle-generating machinery (Paulusma and Oude Elferink, 2010). Family 8 ATPases may exert their cellular functions by combining enzymatic phospholipid translocation activity with an enzyme-independent action. The latter can involve the timely recruitment of proteins involved in cellular signalling, vesicle coat assembly and cytoskeleton regulation (van der Velden et al., 2010). The beta-subunit, CDC50A, allows the stable expression, assembly, subcellular localization, and lipid transport activity of the P4-ATPase ATP8A2 (Coleman and Molday, 2011).  Residues in phospholipid specificity have been identified (Baldridge and Graham 2012). 

Asymmetric phosopholipid distribution in the plasma membranes of animals is disrupted during apoptosis, exposing phosphatidylserine (PtdSer) on the cell surface.  ATP11C (adenosine triphosphatase type 11C) and CDC50A (cell division cycle protein 50A) are required for aminophospholipid translocation from the outer to the inner plasma membrane leaflet due to their  flippase activity. ATP11C contain caspase recognition sites, and mutations at these sites generate caspase-resistant ATP11C without affecting its flippase activity. Cells expressing caspase-resistant ATP11C do not expose PtdSer during apoptosis and are not engulfed by macrophages, suggesting that inactivation of the flippase activity is required for apoptotic PtdSer exposure. CDC50A-deficient cells displayed PtdSer on their surface and were engulfed by macrophages, indicating that PtdSer is sufficient as an 'eat me' signal.  CDC50A serves as a chaparone protein for most phospholipid-flipping ATPases, targetting them (including ATP11C) to the plasma membrane (Segawa et al. 2014).

P4-ATPases mediate the translocation of phospholipids from the outer to the inner leaflet and maintain lipid asymmetry, which is critical for membrane trafficking and signaling pathways. Hiraizumi et al. 2019 reported cryo-EM structures of six distinct intermediates of the human ATP8A1-CDC50a heterocomplex at resolutions of 2.6 to 3.3 angstroms, elucidating the lipid translocation cycle of this P4-ATPase. ATP-dependent phosphorylation induces a large rotational movement of the actuator domain around the phosphorylation site in the phosphorylation domain, accompanied by lateral shifts of the first and second transmembrane helices, thereby allowing phosphatidylserine binding. The phospholipid head group passes through the hydrophilic cleft, while the acyl chain is exposed toward the lipid environment (Hiraizumi et al. 2019).

 

The generalized reactions for P-type ATPases are:

nMe1 (out) + mMe2 (in) + ATP → nMe1 (in) + mMe2 (out) + ADP + Pi.

Phospholipid (outer leaflet of the membrane) + ATP → Phospholipid (inner leaflet) + ADP + Pi



This family belongs to the P-type ATPase (P-ATPase) Superfamily.

 

References:

and Migocka M. (2015). Copper-transporting ATPases: The evolutionarily conserved machineries for balancing copper in living systems. IUBMB Life. 67(10):737-45.

Abe, K., J. Shimokawa, M. Naito, K. Munson, O. Vagin, G. Sachs, H. Suzuki, K. Tani, and Y. Fujiyoshi. (2017). The cryo-EM structure of gastric H,K-ATPase with bound BYK99, a high-affinity member of K-competitive, imidazo[1,2-a]pyridine inhibitors. Sci Rep 7: 6632.

Abe, K., K. Tani, T. Friedrich, and Y. Fujiyoshi. (2012). Cryo-EM structure of gastric H+,K+-ATPase with a single occupied cation-binding site. Proc. Natl. Acad. Sci. USA 109: 18401-18406.

Abeyrathna, N., , S. Abeyrathna, , M.T. Morgan, , C.J. Fahrni, , and G. Meloni,. (2020). Transmembrane Cu(I) P-type ATPase pumps are electrogenic uniporters. Dalton Trans 49: 16082-16094.

Adle, D.J., D. Sinani, H. Kim, and J. Lee. (2007). A cadmium-transporting P1B-type ATPase in yeast Saccharomyces cerevisiae. J. Biol. Chem. 282: 947-955.

Ahn, W., M.G. Lee, K.H. Kim, and S. Muallem. (2003). Multiple effects of SERCA2b mutations associated with Darier's disease. J. Biol. Chem. 278: 20795-20801.

Alevizopoulos, K., T. Calogeropoulou, F. Lang, and C. Stournaras. (2014). Na+/K+ ATPase inhibitors in cancer. Curr Drug Targets 15: 988-1000.

Anthonisen, A.N., J.D. Clausen, and J.P. Andersen. (2006). Mutational analysis of the conserved TGES loop of sarcoplasmic reticulum Ca2+-ATPase. J. Biol. Chem. 281: 31572-31582.

Antonaci, F., S. Ravaglia, G.S. Grieco, S. Gagliardi, C. Cereda, and A. Costa. (2021). Familial hemiplegic migraine type 2 due to a novel missense mutation in ATP1A2. J Headache Pain 22: 12.

Arashiki, N., Y. Takakuwa, N. Mohandas, J. Hale, K. Yoshida, H. Ogura, T. Utsugisawa, S. Ohga, S. Miyano, S. Ogawa, S. Kojima, and H. Kanno. (2016). ATP11C is a major flippase in human erythrocytes and its defect causes congenital hemolytic anemia. Haematologica 101: 559-565.

Aravind, L., M.Y. Galperin, and E.V. Koonin. (1998). The catalytic domain of the P-type ATPase has the haloacid dehalogenase fold. Trends Biochem. Sci. 23: 127-129.

Auer, M., G.A. Scarborough, and W. Kühlbrandt. (1999). Surface crystallisation of the plasma membrane H+-ATPase on a carbon support film for electron crystallography. J. Mol. Biol. 287: 961-968.

Autry, J.M., J.E. Rubin, S.D. Pietrini, D.L. Winters, S.L. Robia, and D.D. Thomas. (2011). Oligomeric interactions of sarcolipin and the Ca-ATPase. J. Biol. Chem. 286: 31697-31706.

Axelsen, K.B. and M.G. Palmgren. (1998). Evolution of substrate specificities in the P-type ATPase superfamily. J. Mol. Evol. 46: 84-101.

Baekgaard, L., L. Luoni, M.I. De Michelis, and M.G. Palmgren. (2006). The plant plasma membrane Ca2+ pump ACA8 contains overlapping as well as physically separated autoinhibitory and calmodulin-binding domains. J. Biol. Chem. 281: 1058-1065.

Baekgaard, L., M.D. Mikkelsen, D.M. Sørensen, J.N. Hegelund, D.P. Persson, R.F. Mills, Z. Yang, S. Husted, J.P. Andersen, M.J. Buch-Pedersen, J.K. Schjoerring, L.E. Williams, and M.G. Palmgren. (2010). A combined zinc/cadmium sensor and zinc/cadmium export regulator in a heavy metal pump. J. Biol. Chem. 285: 31243-31252.

Baldridge RD., Xu P. and Graham TR. (2013). Type IV P-type ATPases distinguish mono- versus diacyl phosphatidylserine using a cytofacial exit gate in the membrane domain. J Biol Chem. 288(27):19516-27.

Baldridge, R.D. and T.R. Graham. (2012). Identification of residues defining phospholipid flippase substrate specificity of type IV P-type ATPases. Proc. Natl. Acad. Sci. USA 109: E290-298.

Banuelos, M.A. and A. Rodríguez-Navarro. (1998). P-type ATPases mediate sodium and potassium effluxes in Schwanniomyces occidentalis. J. Biol. Chem. 273: 1640-1646.

Baranano, D.E., H. Wolosker, B. Bae, R. K. Barrow, S.H. Snyder, and C.D. Ferris. (2000). A mammalian iron ATPase induced by iron. J. Biol. Chem. 275: 15166-15173.

Barcelos, R.C.S., H.Z. Rosa, K. Roversi, C.D.S. Tibúrcio-Machado, P.T. Inchaki, M.E. Burger, and C.A.S. Bier. (2020). Apical periodontitis induces changes on oxidative stress parameters and increases Na/K-ATPase activity in adult rats. Arch Oral Biol 118: 104849.

Barnes, N., R. Tsivkovskii, N. Tsivkovskaia, and S. Lutsenko. (2005). The copper-transporting ATPases, Menkes and Wilson disease proteins, have distinct roles in adult and developing cerebellum. J. Biol. Chem. 280: 9640-9645.

Barry, A.N., A. Otoikhian, S. Bhatt, U. Shinde, R. Tsivkovskii, N.J. Blackburn, and S. Lutsenko. (2011). The lumenal loop Met672-Pro707 of copper-transporting ATPase ATP7A binds metals and facilitates copper release from the intramembrane sites. J. Biol. Chem. 286: 26585-26594.

Beard, S.J., R. Hashim, J. Membrillo-Hernández, M.N. Hughes, and R.K. Poole. (1997). Zinc(II) tolerance in Escherichia coli K-12: evidence that the zntA gene (o732) encodes a cation transport ATPase. Mol. Microbiol. 25: 883-891.

Benito B., Garciadeblas B., Perez-Martin J. and Rodriguez-Navarro A. (2009). Growth at high pH and sodium and potassium tolerance in media above the cytoplasmic pH depend on ENA ATPases in Ustilago maydis. Eukaryot Cell. 8(6):821-9.

Benito, B., B. Garciadeblás, and A. Rodríguez-Navarro. (2002). Potassium- or sodium-efflux ATPase, a key enzyme in the evolution of fungi. Microbiology 148: 933-941.

Benito, B., B. Garciadeblás, and A. Rodríguez-Navarro. (2000). Molecular cloning of the calcium and sodium ATPases in Neurospora crassa. Mol. Microbiol. 35: 1079-1088.

Berrocal, M., I. Corbacho, C. Gutierrez-Merino, and A.M. Mata. (2018). Methylene blue activates the PMCA activity and cross-interacts with amyloid β-peptide, blocking Aβ-mediated PMCA inhibition. Neuropharmacology 139: 163-172.

Bertini, I. and A. Rosato. (2008). Menkes disease. Cell Mol Life Sci 65(1): 89-91.

Bitter, R.M., S. Oh, Z. Deng, S. Rahman, R.K. Hite, and P. Yuan. (2022). Structure of the Wilson disease copper transporter ATP7B. Sci Adv 8: eabl5508.

Bock, K.W., D. Honys, J.M. Ward, S. Padmanaban, E.P. Nawrocki, K.D. Hirschi, D. Twell, and H. Sze. (2006). Integrating membrane transport with male gametophyte development and function through transcriptomics. Plant Physiol. 140: 1151-1168.

Bonza, M.C., H. Martin, M. Kang, G. Lewis, T. Greiner, S. Giacometti, J.L. Van Etten, M.I. De Michelis, G. Thiel, and A. Moroni. (2010). A functional calcium-transporting ATPase encoded by chlorella viruses. J Gen Virol 91: 2620-2629.

Boutin, J.A., S. Bedut, M. Jullian, M. Galibert, L. Frankiewicz, P. Gloanec, G. Ferry, K. Puget, and J. Leprince. (2022). Caloxin-derived peptides for the inhibition of plasma membrane calcium ATPases. Peptides 154: 170813.

Bowman, B.J., S. Abreu, E. Margolles-Clark, M. Draskovic, and E.J. Bowman. (2011). Role of four calcium transport proteins, encoded by nca-1, nca-2, nca-3, and cax, in maintaining intracellular calcium levels in Neurospora crassa. Eukaryot. Cell. 10: 654-661.

Braiterman, L., L. Nyasae, F. Leves, and A.L. Hubbard. (2011). Critical roles for the COOH terminus of the Cu-ATPase ATP7B in protein stability, trans-Golgi network retention, copper sensing, and retrograde trafficking. Am. J. Physiol. Gastrointest Liver Physiol 301: G69-81.

Braiterman, L.T., A. Murthy, S. Jayakanthan, L. Nyasae, E. Tzeng, G. Gromadzka, T.B. Woolf, S. Lutsenko, and A.L. Hubbard. (2014). Distinct phenotype of a Wilson disease mutation reveals a novel trafficking determinant in the copper transporter ATP7B. Proc. Natl. Acad. Sci. USA 111: E1364-1373.

Bramkamp, M., K. Altendorf, and J.C. Greie. (2007) Common patterns and unique features of P-type ATPases: a comparative view on the KdpFABC complex from Escherichia coli (Review). Mol. Membr. Biol. 24: 375-386.

Bryde, S., H. Hennrich, P.M. Verhulst, P.F. Devaux, G. Lenoir, and J.C. Holthuis. (2010). CDC50 proteins are critical components of the human class-1 P4-ATPase transport machinery. J. Biol. Chem. 285: 40562-40572.

Bublitz, M., H. Poulsen, J.P. Morth, and P. Nissen. (2010). In and out of the cation pumps: P-type ATPase structure revisited. Curr. Opin. Struct. Biol. 20: 431-439.

Burke, R., E. Commons, and J. Camakaris. (2008). Expression and localisation of the essential copper transporter DmATP7 in Drosophila neuronal and intestinal tissues. Int J Biochem. Cell Biol. 40: 1850-1860.

Cai, K., H. Gao, X. Wu, S. Zhang, Z. Han, X. Chen, G. Zhang, and F. Zeng. (2019). The Ability to Regulate Transmembrane Potassium Transport in Root Is Critical for Drought Tolerance in Barley. Int J Mol Sci 20:.

Cairo, E.R., T. Friedrich, H.G. Swarts, N.V. Knoers, R.J. Bindels, L.A. Monnens, P.H. Willems, J.J. De Pont, and J.B. Koenderink. (2008). Impaired routing of wild type FXYD2 after oligomerisation with FXYD2-G41R might explain the dominant nature of renal hypomagnesemia. Biochim. Biophys. Acta. 1778: 398-404.

Calì, T., M. Brini, and E. Carafoli. (2017). Regulation of Cell Calcium and Role of Plasma Membrane Calcium ATPases. Int Rev Cell Mol Biol 332: 259-296.

Calì, T., R. Lopreiato, J. Shimony, M. Vineyard, M. Frizzarin, G. Zanni, G. Zanotti, M. Brini, M. Shinawi, and E. Carafoli. (2015). A Novel Mutation in Isoform 3 of the Plasma Membrane Ca2+ Pump Impairs Cellular Ca2+ Homeostasis in a Patient with Cerebellar Ataxia and Laminin Subunit 1α Mutations. J. Biol. Chem. 290: 16132-16141.

Cao, Z.Z., X.Y. Lin, Y.J. Yang, M.Y. Guan, P. Xu, and M.X. Chen. (2019). Gene identification and transcriptome analysis of low cadmium accumulation rice mutant (lcd1) in response to cadmium stress using MutMap and RNA-seq. BMC Plant Biol 19: 250.

Carpinelli, M.R., M.G. Manning, B.T. Kile, and A.B. Rachel. (2013). Two ENU-induced alleles of Atp2b2 cause deafness in mice. PLoS One 8: e67479.

Catty, P., A.D. d’Exaerde, and A. Goffeau. (1997). The complete inventory of the yeast Saccharomyces cerevisiae P-type transport ATPases. FEBS Lett. 409: 325-332.

Čechová, P., K. Berka, and M. Kubala. (2016). Ion Pathways in the Na+/K+-ATPase. J Chem Inf Model 56: 2434-2444.

Chan, H., V. Babayan, E. Blyumin, C. Gandhi, K. Hak, D. Harake, K. Kumar, P. Lee, T.T. Li, H.Y. Liu, T.C. Lo, C.J. Meyer, S. Stanford, K.S. Zamora, and M.H. Saier, Jr. (2010). The p-type ATPase superfamily. J. Mol. Microbiol. Biotechnol. 19: 5-104.

Chantalat, S., S.K. Park, Z. Hua, K. Liu, R. Gobin, A. Peyroche, A. Rambourg, T.R. Graham, and C.L. Jackson. (2004). The Arf activator Gea2p and the P-type ATPase Drs2p interact at the Golgi in Saccharomyces cerevisiae. J Cell Sci 117: 711-722.

Chaoprasid, P., S. Nookabkaew, R. Sukchawalit, and S. Mongkolsuk. (2015). Roles of Agrobacterium tumefaciens C58 ZntA and ZntB and the transcriptional regulator ZntR in controlling Cd2+/Zn2+/Co2+ resistance and the peroxide stress response. Microbiology 161: 1730-1740.

Chaubey, P.M., L. Hofstetter, B. Roschitzki, and B. Stieger. (2016). Proteomic Analysis of the Rat Canalicular Membrane Reveals Expression of a Complex System of P4-ATPases in Liver. PLoS One 11: e0158033.

Chaumette, B., V. Ferrafiat, A. Ambalavanan, A. Goldenberg, A. Dionne-Laporte, D. Spiegelman, P.A. Dion, P. Gerardin, C. Laurent, D. Cohen, J. Rapoport, and G.A. Rouleau. (2020). Missense variants in ATP1A3 and FXYD gene family are associated with childhood-onset schizophrenia. Mol Psychiatry 25: 821-830.

Chen P., Chakraborty S., Mukhopadhyay S., Lee E., Paoliello MM., Bowman AB. and Aschner M. (2015). Manganese homeostasis in the nervous system. J Neurochem. 134(4):601-10.

Chen, C.C., B.R. Chen, Y. Wang, P. Curman, H.A. Beilinson, R.M. Brecht, C.C. Liu, R.J. Farrell, J. de Juan-Sanz, L.M. Charbonnier, S. Kajimura, T.A. Ryan, D.G. Schatz, T.A. Chatila, J.D. Wikstrom, J.K. Tyler, and B.P. Sleckman. (2021). Sarco/endoplasmic reticulum Ca2+-ATPase (SERCA) activity is required for V(D)J recombination. J Exp Med 218:.

Chen, H.Y., R.D. Roer, and R.D. Watson. (2013). Molecular cloning of a plasma membrane Ca²⁺ ATPase (PMCA) from Y-organs of the blue crab (Callinectes sapidus), and determination of spatial and temporal patterns of PMCA gene expression. Gene 522: 8-17.

Chen, X., M. Zhou, S. Zhang, J. Yin, P. Zhang, X. Xuan, P. Wang, Z. Liu, B. Zhou, and M. Yang. (2021). Cryo-EM structures and transport mechanism of human P5B type ATPase ATP13A2. Cell Discov 7: 106.

Chen, Y., C. Cao, Z. Guo, Q. Zhang, S. Li, X. Zhang, J. Gong, and Y. Shen. (2020). Herbivore exposure alters ion fluxes and improves salt tolerance in a desert shrub. Plant Cell Environ 43: 400-419.

Chen, Y.Z., K. Klöditz, E.S. Lee, D.P. Nguyen, Q. Yuan, J. Johnson, Y. Lee-Yow, A. Hall, S. Mitani, N.S. Xia, B. Fadeel, and D. Xue. (2019). Structure and function analysis of the aminophospholipid translocase TAT-1. J Cell Sci 132:.

Chesi, A., A. Kilaru, X. Fang, A.A. Cooper, and A.D. Gitler. (2012). The role of the Parkinson's disease gene PARK9 in essential cellular pathways and the manganese homeostasis network in yeast. PLoS One 7: e34178.

Chintalapati, S., R. Al Kurdi, A.C. van Scheltinga, and W. Kühlbrandt. (2008). Membrane structure of CtrA3, a copper-transporting P-type-ATPase from Aquifex aeolicus. J. Mol. Biol. 378: 581-595.

Cho, T., A. Ishii-Kato, Y. Fukata, Y. Nakayama, K. Iida, M. Fukata, and H. Iida. (2016). Coupling of a voltage-gated Ca2+ channel homologue with a plasma membrane H+ -ATPase in yeast. Genes Cells. [Epub: Ahead of Print]

Cohen, Y., M. Megyeri, O.C. Chen, G. Condomitti, I. Riezman, U. Loizides-Mangold, A. Abdul-Sada, N. Rimon, H. Riezman, F.M. Platt, A.H. Futerman, and M. Schuldiner. (2013). The yeast p5 type ATPase, spf1, regulates manganese transport into the endoplasmic reticulum. PLoS One 8: e85519.

Coleman JA., Kwok MC. and Molday RS. (2009). Localization, purification, and functional reconstitution of the P4-ATPase Atp8a2, a phosphatidylserine flippase in photoreceptor disc membranes. J Biol Chem. 284(47):32670-9.

Coleman, J.A. and R.S. Molday. (2011). Critical role of the β-subunit CDC50A in the stable expression, assembly, subcellular localization, and lipid transport activity of the P4-ATPase ATP8A2. J. Biol. Chem. 286: 17205-17216.

Corradi, G.R., L.R. Mazzitelli, G.D. Petrovich, F. de Tezanos Pinto, L. Rochi, and H.P. Adamo. (2021). Plasma Membrane Ca Pump PMCA4z Is More Active Than Splicing Variant PMCA4x. Front Cell Neurosci 15: 668371.

Cronin S.R., R. Rao, R.Y. Hampton. (2002). Cod1p/Spf1p is a P-type ATPase involved in ER function and Ca2+ homeostasis. J. Cell. Biol. 157: 1017-1028.

Da'dara, A.A., Z. Faghiri, G. Krautz-Peterson, R. Bhardwaj, and P.J. Skelly. (2013). Schistosome Na,K-ATPase as a therapeutic target. Trans R Soc Trop Med Hyg 107: 74-82.

Darby, P.J., C.Y. Kwan, and E.E. Daniel. (2016). Use of calcium pump inhibitors in the study of calcium regulation in smooth muscle. Biol Signals 2: 293-304.

Darland-Ransom, M., X. Wang, C.L. Sun, J. Mapes, K. Gengyo-Ando, S. Mitani, and D. Xue. (2008). Role of C. elegans TAT-1 protein in maintaining plasma membrane phosphatidylserine asymmetry. Science 320: 528-531.

de Meis, L. (2003). Brown adipose tissue Ca2+-ATPase. Uncoupled ATP hydrolysis and thermogenic activity. J. Biol. Chem. 278: 41856-41861.

de Meis, L., A.P. Arruda, R.M. da Costa, and M. Benchimol. (2006). Identification of a Ca2+-ATPase in brown adipose tissue mitochondria: regulation of thermogenesis by ATP and Ca2+. J. Biol. Chem. 281: 16384-16390.

de Tezanos Pinto, F. and H.P. Adamo. (2006). Deletions in the A(L) region of the h4xb plasma membrane Ca2+ pump. High apparent affinity for Ca2+ of a deletion mutant resembling the alternative spliced form h4zb. FEBS Lett. 580: 1576-1580.

Dederer, V. and M.K. Lemberg. (2021). Transmembrane dislocases: a second chance for protein targeting. Trends Cell Biol. 31: 898-911.

Dehay, B., M. Martinez-Vicente, A. Ramirez, C. Perier, C. Klein, M. Vila, and E. Bezard. (2012). Lysosomal dysfunction in Parkinson disease: ATP13A2 gets into the groove. Autophagy 8: 1389-1391.

Demirsoy, S., S. Martin, S. Motamedi, S. van Veen, T. Holemans, C. Van den Haute, A. Jordanova, V. Baekelandt, P. Vangheluwe, and P. Agostinis. (2017). ATP13A2/PARK9 regulates endo-/lysosomal cargo sorting and proteostasis through a novel PI(3, 5)P2-mediated scaffolding function. Hum Mol Genet 26: 1656-1669.

Dempski, R.E., K. Hartung, T. Friedrich, and E. Bamberg. (2006). Fluorometric measurements of intermolecular distances between the α- and β-subunits of the Na+/K+-ATPase. J. Biol. Chem. 281: 36338-36346.

Demurtas, O.C., R. de Brito Francisco, E. Martinoia, and G. Giuliano. (2020). Transportomics for the Characterization of Plant Apocarotenoid Transmembrane Transporters. Methods Mol Biol 2083: 89-99.

Desmond, P.F., A. Labuza, J. Muriel, M.L. Markwardt, A.E. Mancini, M.A. Rizzo, and R.J. Bloch. (2017). Interactions between Small Ankyrin 1 and Sarcolipin Coordinately Regulate Activity of the Sarco(endo)plasmic Reticulum Ca2+-ATPase (SERCA1). J. Biol. Chem. [Epub: Ahead of Print]

Ding, J., Z. Wu, B.P. Crider, Y. Ma, X. Li, C. Slaughter, L. Gong, and X. Xie. (2000). Identification and functional expression of four isoforms of ATPase II, the putative aminophospholipid translocase. Effect of isoform variation on the ATPase activity and phospholipid specificity. J. Biol. Chem. 275: 23378-23386.

Ding, M., M. Zhang, H. Zeng, Y. Hayashi, Y. Zhu, and T. Kinoshita. (2021). Molecular basis of plasma membrane H-ATPase function and potential application in the agricultural production. Plant Physiol. Biochem 168: 10-16.

Doğanli, C., H.C. Beck, A.B. Ribera, C. Oxvig, and K. Lykke-Hartmann. (2013). α3Na+/K+-ATPase deficiency causes brain ventricle dilation and abrupt embryonic motility in zebrafish. J. Biol. Chem. 288: 8862-8874.

Docampo R., Moreno SN. and Plattner H. (2014). Intracellular calcium channels in protozoa. Eur J Pharmacol. 739:4-18.

Doganli, C., K. Kjaer-Sorensen, C. Knoeckel, H.C. Beck, J.R. Nyengaard, B. Honoré, P. Nissen, A. Ribera, C. Oxvig, and K. Lykke-Hartmann. (2012). The α2Na+/K+-ATPase is critical for skeletal and heart muscle function in zebrafish. J Cell Sci 125: 6166-6175.

Drees SL., Beyer DF., Lenders-Lomscher C. and Lubben M. (2015). Distinct functions of serial metal-binding domains in the Escherichia coli P1 B -ATPase CopA. Mol Microbiol. 97(3):423-38.

Dubey, V., D. Stokes, B.P. Pedersen, and H. Khandelia. (2021). An intracellular pathway controlled by the N-terminus of the pump subunit inhibits the bacterial KdpFABC ion pump in high K conditions. J. Mol. Biol. 167008. [Epub: Ahead of Print]

Dyla, M., D.S. Terry, M. Kjaergaard, T.L. Sørensen, J. Lauwring Andersen, J.P. Andersen, C. Rohde Knudsen, R.B. Altman, P. Nissen, and S.C. Blanchard. (2017). Dynamics of P-type ATPase transport revealed by single-molecule FRET. Nature 551: 346-351.

Eide, D.J. (1998). The molecular biology of metal ion transport in Saccharomyces cerevisiae. Annu. Rev. Nutr. 18: 441-469.

Einholm, A.P., H.N. Nielsen, R. Holm, M.S. Toustrup-Jensen, and B. Vilsen. (2016). Importance of a Potential Protein Kinase A Phosphorylation Site of Na+,K+-ATPase and Its Interaction Network for Na+ Binding. J. Biol. Chem. 291: 10934-10947.

Einholm, A.P., J.P. Andersen, and B. Vilsen. (2007). Roles of transmembrane segment M1 of Na(+), K(+)-ATPase and Ca (2+)-ATPase, the gatekeeper and the pivot. J. Bioenerg. Biomembr. 39(5-6):357-66.

Ekberg, K., B.P. Pedersen, D.M. Sørensen, A.K. Nielsen, B. Veierskov, P. Nissen, M.G. Palmgren, and M.J. Buch-Pedersen. (2010). Structural identification of cation binding pockets in the plasma membrane proton pump. Proc. Natl. Acad. Sci. USA 107: 21400-21405.

Ekberg, K., M.G. Palmgren, B. Veierskov, and M.J. Buch-Pedersen. (2010). A Novel Mechanism of P-type ATPase Autoinhibition Involving Both Termini of the Protein. J. Biol. Chem. 285: 7344-7350.

Elvington, S.M., F. Bu, and J.W. Nichols. (2005). Fluorescent, acyl chain-labeled phosphatidylcholine analogs reveal novel transport pathways across the plasma membrane of yeast. J. Biol. Chem. 280: 40957-40964.

Eren, E., Kennedy, D.C., Maroney, M.J., and Arguello, J.M. (2006). A novel regulatory metal binding domain is present in the C terminus of Arabidopsis Zn2+-ATPase HMA2. J. Biol. Chem. 281: 33881-33891.

Espeso, E.A. (2016). The CRaZy Calcium Cycle. Adv Exp Med Biol 892: 169-186.

Espinoza-Fonseca, L.M. (2019). Probing the effects of nonannular lipid binding on the stability of the calcium pump SERCA. Sci Rep 9: 3349.

Espinoza-Fonseca, L.M. (2021). Structural Basis for the Function of the C-Terminal Proton Release Pathway in the Calcium Pump. Int J Mol Sci 22:.

Espinoza-Fonseca, L.M., J.M. Autry, and D.D. Thomas. (2014). Microsecond molecular dynamics simulations of Mg²⁺- and K⁺-bound E1 intermediate states of the calcium pump. PLoS One 9: e95979.

Ettema, T.J., A.B. Brinkman, P.P. Lamers, N.G. Kornet, W.M. de Vos, and J. van der Oost. (2006). Molecular characterization of a conserved archaeal copper resistance (cop) gene cluster and its copper-responsive regulator in Sulfolobus solfataricus P2. Microbiology 152: 1969-1979.

Fagan, M.J. and M.H. Saier, Jr. (1994). P-type ATPases of eukaryotes and bacteria: sequence analyses and construction of phylogenetic trees. J. Mol. Evol. 38: 57-99.

Fan, B. and B.P. Rosen. (2002). Biochemical characterization of CopA, the Escherichia coli Cu(I)-translocating P-type ATPase. J. Biol. Chem. 277: 46987-46992.

Favia, M., A. Gerbino, E. Notario, V. Tragni, M.N. Sgobba, M.E. Dell''Aquila, C.L. Pierri, L. Guerra, and E. Ciani. (2022). The Non-Gastric H/K ATPase (ATP12A) Is Expressed in Mammalian Spermatozoa. Int J Mol Sci 23:.

Flamant, S., P. Pescher, B. Lemercier, M. Clément-Ziza, F. Képès, M. Fellous, G. Milon, G. Marchal, and C. Besmond. (2003). Characterization of a putative type IV aminophospholipid transporter P-type ATPase. Mamm Genome 14: 21-30.

Friedrich, T., N.N. Tavraz, and C. Junghans. (2016). ATP1A2 Mutations in Migraine: Seeing through the Facets of an Ion Pump onto the Neurobiology of Disease. Front Physiol 7: 239.

Fujisawa, C., H. Kodama, T. Hiroki, Y. Akasaka, and M. Hamanoue. (2019). ATP7A mutations in 66 Japanese patients with Menkes disease and carrier detection: A gene analysis. Pediatr Int 61: 345-350.

Furune, T., K. Hashimoto, and J. Ishiguro. (2008). Characterization of a fission yeast P(5)-type ATPase homologue that is essential for Ca2+/Mn(2+ )homeostasis in the absence of P(2)-type ATPases. Genes Genet Syst 83: 373-381.

Gaballa, A., and J.D. Helmann. (2002). A peroxide-induced zinc uptake system plays an important role in protection against oxidative stress in Bacillus subtilis. Mol. Microbiol. 45: 997-1005.

Gan-Or Z., Dion PA. and Rouleau GA. (201). Genetic perspective on the role of the autophagy-lysosome pathway in Parkinson disease. Autophagy. 11(9):1443-57.

Garcia, A., N.D. Eljack, M.A. Sani, F. Separovic, H.H. Rasmussen, W. Kopec, H. Khandelia, F. Cornelius, and R.J. Clarke. (2015). Membrane accessibility of glutathione. Biochim. Biophys. Acta. 1848: 2430-2436.

Gassel, M., T. Möllenkamp, W. Puppe, and K. Altendorf. (1999). The KdpF subunit is part of the K(+)-translocating Kdp complex of Escherichia coli and is responsible for stabilization of the complex in vitro. J. Biol. Chem. 274: 37901-7.

Geering, K. (1991). The functional role of the β-subunit in the maturation and intracellular transport of Na,K-ATPase. FEBS Lett. 285: 189-193.

Geering, K. (2000). Topogenic motifs in P-type ATPases. J. Memb. Biol. 174: 181-190.

Gerencser, G.A. (1993). A novel P-type Cl- stimulated ATPase: phosphorylation and specificity. Biochem. Biophys. Res. Commun. 196: 1188-1194.

Geurts, M.M.G., J.D. Clausen, B. Arnou, C. Montigny, G. Lenoir, R.A. Corey, C. Jaxel, J.V. Møller, P. Nissen, J.P. Andersen, M. le Maire, and M. Bublitz. (2020). The SERCA residue Glu340 mediates interdomain communication that guides Ca transport. Proc. Natl. Acad. Sci. USA 117: 31114-31122.

Giacomello, M., A. De Mario, C. Scarlatti, S. Primerano, and E. Carafoli. (2013). Plasma membrane calcium ATPases and related disorders. Int J Biochem. Cell Biol. 45: 753-762.

Godic, A., M. Strazisar, A. Zupan, B. Korosec, A. Kansky, and D. Glavac. (2010). Darier disease in Slovenia: spectrum of ATP2A2 mutations and relation to patients' phenotypes. Eur J Dermatol 20: 271-275.

Gomès, E., M.K. Jakobsen, K.B. Axelsen, M. Geisler, and M.G. Palmgren. (2000). Chilling tolerance in Arabidopsis involves ALA1, a member of a new family of putative aminophospholipid translocases. Plant Cell 12: 2441-2454.

Gong, D., X. Chi, K. Ren, G. Huang, G. Zhou, N. Yan, J. Lei, and Q. Zhou. (2018). Structure of the human plasma membrane Ca-ATPase 1 in complex with its obligatory subunit neuroplastin. Nat Commun 9: 3623.

González-Guerrero, M. and J.M. Argüello. (2008). Mechanism of Cu+-transporting ATPases: soluble Cu+ chaperones directly transfer Cu+ to transmembrane transport sites. Proc. Natl. Acad. Sci. USA 105: 5992-5997.

González-Guerrero, M., D. Raimunda, X. Cheng, and J.M. Argüello. (2010). Distinct functional roles of homologous Cu+ efflux ATPases in Pseudomonas aeruginosa. Mol. Microbiol. 78: 1246-1258.

Gorski PA., Glaves JP., Vangheluwe P. and Young HS. (2013). Sarco(endo)plasmic reticulum calcium ATPase (SERCA) inhibition by sarcolipin is encoded in its luminal tail. J Biol Chem. 288(12):8456-67.

Gorski, P.A., C.A. Trieber, E. Larivière, M. Schuermans, F. Wuytack, H.S. Young, and P. Vangheluwe. (2012). Transmembrane helix 11 is a genuine regulator of the endoplasmic reticulum Ca2+ pump and acts as a functional parallel of β-subunit on α-Na+,K+-ATPase. J. Biol. Chem. 287: 19876-19885.

Gourdon, P., O. Sitsel, J. Lykkegaard Karlsen, L. Birk Møller, and P. Nissen. (2012). Structural models of the human copper P-type ATPases ATP7A and ATP7B. Biol Chem 393: 205-216.

Gourdon, P., X.Y. Liu, T. Skjørringe, J.P. Morth, L.B. Møller, B.P. Pedersen, and P. Nissen. (2011). Crystal structure of a copper-transporting PIB-type ATPase. Nature 475: 59-64.

Greenough, M., L. Pase, I. Voskoboinik, M.J. Petris, A.W. O''Brien, and J. Camakaris. (2004). Signals regulating trafficking of Menkes (MNK; ATP7A) copper-translocating P-type ATPase in polarized MDCK cells. Am. J. Physiol. Cell Physiol. 287: C1463-1471.

Greie, J.C., and K. Altendorf. (2007). The K+-translocating KdpFABC complex from Escherichia coli: A P-type ATPase with unique features. J. Bioenerg. Biomembr. 39: 397-402.

Greiner, T., A. Moroni, J.L. Van Etten, and G. Thiel. (2018). Genes for Membrane Transport Proteins: Not So Rare in Viruses. Viruses 10:.

Grenon, P., G.R. Corradi, G.D. Petrovich, L.R. Mazzitelli, and H.P. Adamo. (2021). The Spf1p P5A-ATPase "arm-like" domain is not essential for ATP hydrolysis but its deletion impairs autophosphorylation. Biochem. Biophys. Res. Commun. 563: 113-118.

Guerra, F. and A.N. Bondar. (2015). Dynamics of the Plasma Membrane Proton Pump. J. Membr. Biol. 248: 443-453.

Gulsevin, A., A.M. Glazer, T. Shields, B.M. Kroncke, D.M. Roden, and J. Meiler. (2022). Veratridine Can Bind to a Site at the Mouth of the Channel Pore at Human Cardiac Sodium Channel Na1.5. Int J Mol Sci 23:.

Guo, Z., J. Lu, X. Wang, B. Zhan, W. Li, and S.W. Ding. (2017). Lipid flippases promote antiviral silencing and the biogenesis of viral and host siRNAs in Arabidopsis. Proc. Natl. Acad. Sci. USA 114: 1377-1382.

Gupta, A., K. Matsui, J.-F. Lo, and S. Silver. (1999). Molecular basis for resistance to silver cations in Salmonella. Nature Med. 5: 183-188.

Habeck, M., E. Kapri-Pardes, M. Sharon, and S.J. Karlish. (2017). Specific phospholipid binding to Na,K-ATPase at two distinct sites. Proc. Natl. Acad. Sci. USA 114: 2904-2909.

Hao, Z., S. Chen, and D.B. Wilson. (1999). Cloning, expression, and characterization of cadmium and manganese uptake genes from Lactobacillus plantarum. Appl. Environ. Microbiol. 65: 4746-4752.

Harper J.F., B. Hong, I. Hwang, H.Q. Guo, R. Stoddard, J.F. Huang, M.G. Palmgren, H. Sze. (1998). A novel calmodulin-regulated Ca2+-ATPase (ACA2) from Arabidopsis with an N-terminal autoinhibitory domain. J. Biol. Chem. 273: 1099-1106.

Hasman, H., (2005). The tcrB gene is part of the tcrYAZB operon conferring copper resistance in Enterococcus faecium and Enterococcus faecalis. Microbiol. 151: 3019-3025.

Hassani, B.K., C. Astier, W. Nitschke, and S. Ouchane. (2010). CtpA, a copper-translocating P-type ATPase involved in the biogenesis of multiple copper-requiring enzymes. J. Biol. Chem. 285: 19330-19337.

Hatori, Y., D. Lewis, C. Toyoshima, and G. Inesi. (2009). Reaction cycle of Thermotoga maritima copper ATPase and conformational characterization of catalytically deficient mutants. Biochemistry 48: 4871-4880.

Hauck, C. and W.H. Frishman. (2012). Systemic hypertension: the roles of salt, vascular Na+/K+ ATPase and the endogenous glycosides, ouabain and marinobufagenin. Cardiol Rev 20: 130-138.

Haupt, M., M. Bramkamp, M. Coles, H. Kessler, and K. Altendorf. (2005). Prokaryotic Kdp-ATPase: recent insights into the structure and function of KdpB. J. Mol. Microbiol. Biotechnol. 10: 120-131.

Heitkamp, T., R. Kalinowski, B. Böttcher, M. Börsch, K. Altendorf, and J.C. Greie. (2008). K(+)-translocating KdpFABC P-type ATPase from Escherichia coli acts as a functional and structural dimer. Biochemistry 47: 3564-75.

Herrmann, L., D. Schwan, R. Garner, H.L.T. Mobley, R. Haas, K.P. Schäfer, and K. Melchers. (1999). Heliocobacter pylori cadA encodes an essential Cd(II)-Zn(II)-Co(II) resistance factor influencing urease activity. Mol. Microbiol. 33: 524-536.

Hilbers, F., W. Kopec, T.J. Isaksen, T.H. Holm, K. Lykke-Hartmann, P. Nissen, H. Khandelia, and H. Poulsen. (2016). Tuning of the Na,K-ATPase by the beta subunit. Sci Rep 6: 20442.

Hiraizumi, M., K. Yamashita, T. Nishizawa, and O. Nureki. (2019). Cryo-EM structures capture the transport cycle of the P4-ATPase flippase. Science 365: 1149-1155.

Hložková, K., J. Suman, H. Strnad, T. Ruml, V. Paces, and P. Kotrba. (2013). Characterization of pbt genes conferring increased Pb2+ and Cd2+ tolerance upon Achromobacter xylosoxidans A8. Res. Microbiol. 164: 1009-1018.

Hoffmann, R.D., L.I. Olsen, C.V. Ezike, J.T. Pedersen, R. Manstretta, R.L. Lopez-Marques, and M. Palmgren. (2018). Roles of plasma membrane proton ATPases AHA2 and AHA7 in normal growth of roots and root hairs in Arabidopsis thaliana. Physiol Plant. [Epub: Ahead of Print]

Holm, R., J. Khandelwal, A.P. Einholm, J.P. Andersen, P. Artigas, and B. Vilsen. (2017). Arginine substitution of a cysteine in transmembrane helix M8 converts Na+,K+-ATPase to an electroneutral pump similar to H+,K+-ATPase. Proc. Natl. Acad. Sci. USA 114: 316-321.

Holmgren, M., J. Wagg, F. Bezanilla, R.F. Rakowski, P. De Weer, and D.C. Gadsby. (2000). Three distinct and sequential steps in the release of sodium ions by the Na+/K+-ATPase. Nature 403: 898.

Homareda, H., M. Otsu, S. Yamamoto, M. Ushimaru, S. Ito, T. Fukutomi, T. Jo, Y. Eishi, and Y. Hara. (2017). A possible mechanism for low affinity of silkworm Na/K-ATPase for K. J. Bioenerg. Biomembr. 49: 463-472.

Hoppen, C. and G. Groth. (2020). Novel insights into the transfer routes of the essential copper cofactor to the ethylene plant hormone receptor family. Plant Signal Behav 15: 1716512.

Hou, Z. and B. Mitra. (2003). The metal specificity and selectivity of ZntA from Escherichia coli using the acylphosphate intermediate. J. Biol. Chem. 278: 28455-28461.

Hou, Z.-J., S. Narindrasorasak, B. Bhushan, B. Sarkar, and B. Mitra. (2001). Functional analysis of chimeric proteins of the Wilson Cu(I)-ATPase (ATP7B) and ZntA, a Pb(II)/Zn(II)/Cd(II)-ATPase from Escherichia coli. J. Biol. Chem. 276: 40858-40863.

Houdou, M., E. Lebredonchel, A. Garat, S. Duvet, D. Legrand, V. Decool, A. Klein, M. Ouzzine, B. Gasnier, S. Potelle, and F. Foulquier. (2019). Involvement of thapsigargin- and cyclopiazonic acid-sensitive pumps in the rescue of TMEM165-associated glycosylation defects by Mn. FASEB J. 33: 2669-2679.

Hu, G. and J.W. Kronstad. (2010). A putative P-type ATPase, Apt1, is involved in stress tolerance and virulence in Cryptococcus neoformans. Eukaryot. Cell. 9: 74-83.

Huang L., T. Berkelman, A.E. Franklin, N.E. Hoffman. (1993). Characterization of a gene encoding a Ca2+-ATPase-like protein in the plastid envelope. Proc Natl Acad Sci U.S.A. 90: 10066-10070.

Huang, C.S., B.P. Pedersen, and D.L. Stokes. (2017). Crystal structure of the potassium-importing KdpFABC membrane complex. Nature 546: 681-685.

Huang, Y., M. Takar, J.T. Best, and T.R. Graham. (2019). Conserved mechanism of phospholipid substrate recognition by the P4-ATPase Neo1 from Saccharomyces cerevisiae. Biochim. Biophys. Acta. Mol. Cell Biol. Lipids 1865: 158581. [Epub: Ahead of Print]

Hynninen, A., T. Touzé, L. Pitkänen, D. Mengin-Lecreulx, and M. Virta. (2009). An efflux transporter PbrA and a phosphatase PbrB cooperate in a lead-resistance mechanism in bacteria. Mol. Microbiol. 74: 384-394.

Inagaki, C., M. Hara, and X.T. Zeng. (1996). A Cl- pump in rat brain neurons. J. Exp. Zool. 275: 262-268.

Inoue, M., N. Sakuta, S. Watanabe, Y. Zhang, K. Yoshikaie, Y. Tanaka, R. Ushioda, Y. Kato, J. Takagi, T. Tsukazaki, K. Nagata, and K. Inaba. (2019). Structural Basis of Sarco/Endoplasmic Reticulum Ca-ATPase 2b Regulation via Transmembrane Helix Interplay. Cell Rep 27: 1221-1230.e3.

Irzik, K., J. Pfrötzschner, T. Goss, F. Ahnert, M. Haupt, and J.C. Greie. (2011). The KdpC subunit of the Escherichia coli K+-transporting KdpB P-type ATPase acts as a catalytic chaperone. FEBS J. 278: 3041-3053.

Ishihara, N., H. Inagaki, M. Miyake, Y. Kawamura, T. Yoshikawa, and H. Kurahashi. (2019). A case of early onset life-threatening epilepsy associated with a novel ATP1A3 gene variant. Brain Dev 41: 285-291.

Jakobsen, M.K., L.R. Poulsen, A. Schulz, P. Fleurat-Lessard, A. Møller, S. Husted, M. Schiøtt, A. Amtmann, and M.G. Palmgren. (2005). Pollen development and fertilization in Arabidopsis is dependent on the MALE GAMETOGENESIS IMPAIRED ANTHERS gene encoding a type V P-type ATPase. Genes Dev. 19: 2757-2769.

Jensen, M.S., S.R. Costa, A.S. Duelli, P.A. Andersen, L.R. Poulsen, L.D. Stanchev, P. Gourdon, M. Palmgren, T. Günther Pomorski, and R.L. López-Marqués. (2017). Phospholipid flipping involves a central cavity in P4 ATPases. Sci Rep 7: 17621.

Justesen, B.H., R.W. Hansen, H.J. Martens, L. Theorin, M.G. Palmgren, K.L. Martinez, T.G. Pomorski, and A.T. Fuglsang. (2013). Active plasma membrane P-type H+-ATPase reconstituted into nanodiscs is a monomer. J. Biol. Chem. 288: 26419-26429.

Kabashima, Y., H. Ogawa, R. Nakajima, and C. Toyoshima. (2020). What ATP binding does to the Ca pump and how nonproductive phosphoryl transfer is prevented in the absence of Ca. Proc. Natl. Acad. Sci. USA 117: 18448-18458.

Kamrul Huda, K.M., S. Yadav, M.S. Akhter Banu, D.K. Trivedi, and N. Tuteja. (2013). Genome-wide analysis of plant-type II Ca2+ATPases gene family from rice and Arabidopsis: potential role in abiotic stresses. Plant Physiol. Biochem 65: 32-47.

Kanai, R., F. Cornelius, B. Vilsen, and C. Toyoshima. (2022). Cryoelectron microscopy of Na,K-ATPase in the two E2P states with and without cardiotonic steroids. Proc. Natl. Acad. Sci. USA 119: e2123226119.

Karjalainen E.L., K. Hauser, A. Barth. (2007). Proton paths in the sarcoplasmic reticulum Ca2+-ATPase. Biochim Biophys Acta. 1767: 1310-1318.

Kim, H., T. Kim, B.C. Jeong, I.T. Cho, D. Han, N. Takegahara, T. Negishi-Koga, H. Takayanagi, J.H. Lee, J.Y. Sul, V. Prasad, S.H. Lee, and Y. Choi. (2013). Tmem64 modulates calcium signaling during RANKL-mediated osteoclast differentiation. Cell Metab 17: 249-260.

Kinoshita, P.F., A.M.M. Orellana, V.W. Nakao, N.M. de Souza Port''s, L.E.M. Quintas, E.M. Kawamoto, and C. Scavone. (2022). The Janus face of ouabain in Na /K -ATPase and calcium signalling in neurons. Br J Pharmacol 179: 1512-1524.

Kırımtay, K., B. Temizci, M. Gültekin, Z. Yapıcı, and A. Karabay. (2021). Novel mutations in ATP13A2 associated with mixed neurological presentations and iron toxicity due to nonsense-mediated decay. Brain Res 1750: 147167.

Knez, J., E. Salvi, V. Tikhonoff, K. Stolarz-Skrzypek, A. Ryabikov, L. Thijs, D. Braga, M. Kloch-Badelek, S. Malyutina, E. Casiglia, D. Czarnecka, K. Kawecka-Jaszcz, D. Cusi, T. Nawrot, J.A. Staessen, and T. Kuznetsova. (2014). Left ventricular diastolic function associated with common genetic variation in ATP12A in a general population. BMC Med Genet 15: 121.

Kobayashi, C., Y. Matsunaga, J. Jung, and Y. Sugita. (2021). Structural and energetic analysis of metastable intermediate states in the E1P-E2P transition of Ca-ATPase. Proc. Natl. Acad. Sci. USA 118:.

Kopec W., Loubet B., Poulsen H. and Khandelia H. (2014). Molecular mechanism of Na(+),K(+)-ATPase malfunction in mutations characteristic of adrenal hypertension. Biochemistry. 53(4):746-54.

Kraev A., N. Kraev, E. Carafoli. (1999). Identification and functional expression of the plasma membrane calcium ATPase gene family from Caenorhabditis elegans. J. Biol. Chem. 274: 4254-4258.

Kristensen, M. and C. Juel. (2010). Na+,K+-ATPase Na+ affinity in rat skeletal muscle fiber types. J. Membr. Biol. 234: 35-45.

Kühlbrandt, W., J. Zeelen, and J. Dietrich. (2002). Structure, mechanism, and regulation of the Neurospora plasma membrane H+-ATPase. Science 297: 1692-1696.

Kühlbrandt, W., M. Auer, and G.A. Scarborough. (1998). Structure of the P-type ATPases. Curr. Opin. Struc. Biol. 8: 510-516.

Ladefoged, L.K., B. Schiøtt, and N.U. Fedosova. (2021). Beneficent and Maleficent Effects of Cations on Bufadienolide Binding to Na,K-ATPase. J Chem Inf Model 61: 976-986.

Lauer Júnior, C.M., D. Bonatto, A.A. Mielniczki-Pereira, A.Z. Schuch, J.F. Dias, M.L. Yoneama, and J.A. Pêgas Henriques. (2008). The Pmr1 protein, the major yeast Ca2+-ATPase in the Golgi, regulates intracellular levels of the cadmium ion. FEMS Microbiol. Lett. 285: 79-88.

Laursen, M., J.L. Gregersen, L. Yatime, P. Nissen, and N.U. Fedosova. (2015). Structures and characterization of digoxin- and bufalin-bound Na+,K+-ATPase compared with the ouabain-bound complex. Proc. Natl. Acad. Sci. USA 112: 1755-1760.

Leedjärv, A., A. Ivask, and M. Virta. (2008). Interplay of different transporters in the mediation of divalent heavy metal resistance in Pseudomonas putida KT2440. J. Bacteriol. 190: 2680-2689.

Leite, J.A., T.J. Isaksen, A. Heuck, C. Scavone, and K. Lykke-Hartmann. (2020). The α Na/K-ATPase isoform mediates LPS-induced neuroinflammation. Sci Rep 10: 14180.

Lekeux, G., J.M. Crowet, C. Nouet, M. Joris, A. Jadoul, B. Bosman, M. Carnol, P. Motte, L. Lins, M. Galleni, and M. Hanikenne. (2018). Homology modeling and in vivo functional characterization of the zinc permeation pathway in a heavy metal P-type ATPase. J Exp Bot. [Epub: Ahead of Print]

León-Torres, A., L. Novoa-Aponte, and C.Y. Soto. (2015). CtpA, a putative Mycobacterium tuberculosis P-type ATPase, is stimulated by copper (I) in the mycobacterial plasma membrane. Biometals 28: 713-724.

Lescasse, R., J. Grisvard, G. Fryd, A. Fleury-Aubusson, and A. Baroin-Tourancheau. (2005). Proposed function of the accumulation of plasma membrane-type Ca2+-ATPase mRNA in resting cysts of the ciliate Sterkiella histriomuscorum. Eukaryot. Cell. 4: 103-110.

Lewinson, O., A.T. Lee, and D.C. Rees. (2009). A P-type ATPase importer that discriminates between essential and toxic transition metals. Proc. Natl. Acad. Sci. USA 106: 4677-4682.

Li, L., I. Verstraeten, M. Roosjen, K. Takahashi, L. Rodriguez, J. Merrin, J. Chen, L. Shabala, W. Smet, H. Ren, S. Vanneste, S. Shabala, B. De Rybel, D. Weijers, T. Kinoshita, W.M. Gray, and J. Friml. (2021). Cell surface and intracellular auxin signalling for H fluxes in root growth. Nature 599: 273-277.

Li, P., K. Wang, N. Salustros, C. Grønberg, and P. Gourdon. (2021). Structure and transport mechanism of P5B-ATPases. Nat Commun 12: 3973.

Li, X., B. Zhang, P. Ma, R. Cao, X. Yang, and J. Dong. (2021). Plasma Membrane H-ATPase Negatively Regulates the Biosynthesis of Tanshinones in. Int J Mol Sci 22:.

Li, Z. and S.A. Langhans. (2015). Transcriptional regulators of Na,K-ATPase subunits. Front Cell Dev Biol 3: 66.

Li, Z. and Z. Xie. (2009). The Na/K-ATPase/Src complex and cardiotonic steroid-activated protein kinase cascades. Pflugers Arch 457: 635-644.

Liang F., K.W. Cunningham, J.F. Harper, H. Sze. (1997). ECA1 complements yeast mutants defective in Ca2+ pumps and encodes an endoplasmic reticulum-type Ca2+-ATPase in Arabidopsis thaliana.

Liang, J., M. Zhang, M. Lu, Z. Li, X. Shen, M. Chou, and G. Wei. (2016). Functional characterization of a csoR-cueA divergon in Bradyrhizobium liaoningense CCNWSX0360, involved in copper, zinc and cadmium cotolerance. Sci Rep 6: 35155.

Limonta, M., S. Romanowsky, C. Olivari, M.C. Bonza, L. Luoni, A. Rosenberg, J.F. Harper, and M.I. De Michelis. (2014). ACA12 is a deregulated isoform of plasma membrane Ca²⁺-ATPase of Arabidopsis thaliana. Plant Mol. Biol. 84: 387-397.

Lin, X., M.G.K. Brunk, P. Yuanxiang, A.W. Curran, E. Zhang, F. Stöber, J. Goldschmidt, E.D. Gundelfinger, M. Vollmer, M.F.K. Happel, R. Herrera-Molina, and D. Montag. (2021). Neuroplastin expression is essential for hearing and hair cell PMCA expression. Brain Struct Funct 226: 1533-1551.

Liou, A.Y., L.L. Molday, J. Wang, J.P. Andersen, and R.S. Molday. (2019). Identification and functional analyses of disease-associated P4-ATPase phospholipid flippase variants in red blood cells. J. Biol. Chem. 294: 6809-6821.

Liu J. and Xie ZJ. (2010). The sodium pump and cardiotonic steroids-induced signal transduction protein kinases and calcium-signaling microdomain in regulation of transporter trafficking. Biochim Biophys Acta. 1802(12):1237-45.

Liu, T., H. Reyes-Caballero, C. Li, R.A. Scott, and D.P. Giedroc. (2007). Multiple metal binding domains enhance the Zn(II) selectivity of the divalent metal ion transporter AztA. Biochemistry 46: 11057-11068.

Liu, X., C. Wang, B. Yan, L. Lyu, H.E. Takiff, and Q. Gao. (2020). The potassium transporter KdpA affects persister formation by regulating ATP levels in. Emerg Microbes Infect 9: 129-139.

Liu, Y., S. Sitaraman, and A. Chang. (2006). Multiple degradation pathways for misfolded mutants of the yeast plasma membrane ATPase, Pma1. J. Biol. Chem. 281: 31457-31466.

Lopes da Fonseca, T., A. Correia, W. Hasselaar, H.C. van der Linde, R. Willemsen, and T.F. Outeiro. (2013). The zebrafish homologue of Parkinson's disease ATP13A2 is essential for embryonic survival. Brain Res Bull 90: 118-126.

Lopez-Marques RL., Poulsen LR., Bailly A., Geisler M., Pomorski TG. and Palmgren MG. (2015). Structure and mechanism of ATP-dependent phospholipid transporters. Biochim Biophys Acta. 1850(3):461-75.

López-Marqués, R.L., L.R. Poulsen, S. Hanisch, K. Meffert, M.J. Buch-Pedersen, M.K. Jakobsen, T.G. Pomorski, and M.G. Palmgren. (2010). Intracellular targeting signals and lipid specificity determinants of the ALA/ALIS P4-ATPase complex reside in the catalytic ALA α-subunit. Mol. Biol. Cell 21: 791-801.

Lowe J., A. Vieyra, P. Catty, F. Guillain, E. Mintz, M. Cuillel. (2004). A mutational study in the transmembrane domain of Ccc2p, the yeast Cu(I)-ATPase, shows different roles for each Cys-Pro-Cys cysteine. J. Biol. Chem. 279: 25986-25994.

Lübben, M., J. Güldenhaupt, M. Zoltner, K. Deigweiher, P. Haebel, C. Urbanke, and A.J. Scheidig. (2007). Sulfate acts as phosphate analog on the monomeric catalytic fragment of the CPx-ATPase CopB from Sulfolobus solfataricus. J. Mol. Biol. 369: 368-385.

Luo S., F.A. Ruiz, S.N. Moreno. (2005). The acidocalcisome Ca2+-ATPase (TgA1) of Toxoplasma gondii is required for polyphosphate storage, intracellular calcium homeostasis and virulence. Mol. Microbiol. 55: 1034-1045.

Luo, S., J. Fang, and R. Docampo. (2006). Molecular characterization of Trypanosoma brucei P-type H+-ATPases. J. Biol. Chem. 281: 21963-21973.

Lüttmann, D., R. Heermann, B. Zimmer, A. Hillmann, I.S. Rampp, K. Jung, and B. Görke. (2009). Stimulation of the potassium sensor KdpD kinase activity by interaction with the phosphotransferase protein IIA(Ntr) in Escherichia coli. Mol. Microbiol. 72: 978-994.

Lüttmann, D., Y. Göpel, and B. Görke. (2015). Cross-Talk between the Canonical and the Nitrogen-Related Phosphotransferase Systems Modulates Synthesis of the KdpFABC Potassium Transporter in Escherichia coli. J. Mol. Microbiol. Biotechnol. 25: 168-177.

MacLennan, D.H., W.J. Rice, and N.M. Green. (1997). The mechanism of Ca2+ transport by sarco(endo)plasmic reticulum Ca2+-ATPases. J. Biol. Chem. 272: 28815-28818.

Mahmmoud, Y.A. (2008a). Capsazepine, a synthetic vanilloid that converts the Na, K-ATPase to Na-ATPase. Proc. Natl. Acad. Sci. U.S.A. 105: 1757-1761.

Mahmmoud, Y.A. (2008b). Capsaicin stimulates uncoupled ATP hydrolysis by the sarcoplasmic reticulum calcium pump. J. Biol. Chem. 283: 21418-21426.

Mana-Capelli, S., A.K. Mandal, and J.M. Argüello. (2003). Archaeoglobus fultcidus CopB is a thermophilic Cu2+-ATPase. Functional role if its histidine-rich N-terminal metal binding domain. J. Biol. Chem. 278: 40534-40541.

Mandal, A.K., W.D. Cheung, and J.M. Argüello. (2002). Characterization of a thermophilic P-type Ag+/Cu+-ATPase from the extremophile Archaeoglobus fultcidus. J. Biol. Chem. 277: 7201-7208.

Mangialavori, I., M.R. Montes, R.C. Rossi, N.U. Fedosova, and J.P. Rossi. (2011). Dynamic lipid-protein stoichiometry on E1 and E2 conformations of the Na+/K+ -ATPase. FEBS Lett. 585: 1153-1157.

Mateeva, T., M. Klähn, and E. Rosta. (2021). Structural Dynamics and Catalytic Mechanism of ATP13A2 (PARK9) from Simulations. J Phys Chem B 125: 11835-11847.

Mattle D., Zhang L., Sitsel O., Pedersen LT., Moncelli MR., Tadini-Buoninsegni F., Gourdon P., Rees DC., Nissen P. and Meloni G. (2015). A sulfur-based transport pathway in Cu+-ATPases. EMBO Rep. 16(6):728-40.

Maudoux, O., H. Batoko, C. Oecking, K. Gevaert, J. Vandekerckhove, M. Boutry, and P. Morsomme. (2000). A plant plasma membrane H+-ATPase expressed in yeast is activated by phosphorylation at its penultimate residue and binding of 14-3-3 regulatory proteins in the absence of fusicoccin. J. Biol. Chem. 275: 17762-17770.

Maynaud, G., B. Brunel, E. Yashiro, M. Mergeay, J.C. Cleyet-Marel, and A. Le Quéré. (2014). CadA of Mesorhizobium metallidurans isolated from a zinc-rich mining soil is a P(IB-2)-type ATPase involved in cadmium and zinc resistance. Res. Microbiol. 165: 175-189.

McKenna, M.J., S.I. Sim, A. Ordureau, L. Wei, J.W. Harper, S. Shao, and E. Park. (2020). The endoplasmic reticulum P5A-ATPase is a transmembrane helix dislocase. Science 369:.

McLaughlin, H.P., Q. Xiao, R.B. Rea, H. Pi, P.G. Casey, T. Darby, A. Charbit, R.D. Sleator, S.A. Joyce, R.E. Cowart, C. Hill, P.E. Klebba, and C.G. Gahan. (2012). A putative P-type ATPase required for virulence and resistance to haem toxicity in Listeria monocytogenes. PLoS One 7: e30928.

Meier, A., H. Erler, and E. Beitz. (2018). Targeting Channels and Transporters in Protozoan Parasite Infections. Front Chem 6: 88.

Meng, D., L. Bruschweiler-Li, F. Zhang, and R. Brüschweiler. (2015). Modulation and Functional Role of the Orientations of the N- and P-Domains of Cu+ -Transporting ATPase along the Ion Transport Cycle. Biochemistry 54: 5095-5102.

Meydan, S., D. Klepacki, S. Karthikeyan, T. Margus, P. Thomas, J.E. Jones, Y. Khan, J. Briggs, J.D. Dinman, N. Vázquez-Laslop, and A.S. Mankin. (2017). Programmed Ribosomal Frameshifting Generates a Copper Transporter and a Copper Chaperone from the Same Gene. Mol. Cell 65: 207-219.

Meyer, D.J., C. Gatto, and P. Artigas. (2019). Na/K Pump Mutations Associated with Primary Hyperaldosteronism Cause Loss of Function. Biochemistry 58: 1774-1785.

Mikkelsen, S.A., L.S. Mogensen, B. Vilsen, R.S. Molday, A.L. Vestergaard, and J.P. Andersen. (2019). Asparagine-905 of the mammalian phospholipid flippase ATP8A2 is essential for lipid substrate-induced activation of ATP8A2 dephosphorylation. J. Biol. Chem. [Epub: Ahead of Print]

Mills R.F., M.L. Doherty, R.L. López-Marqués, T. Weimar, P. Dupree, M.G. Palmgren, J.K. Pittman, and L.E. Williams. (2008). ECA3, a golgi-localized P2A-type ATPase, plays a crucial role in manganese nutrition in Arabidopsis. Plant Physiol. 146: 116-128.

Miranda M., Pardo JP. and Petrov VV. (2011). Structure-function relationships in membrane segment 6 of the yeast plasma membrane Pma1 H(+)-ATPase. Biochim Biophys Acta. 1808(7):1781-9.

Moniakis J., M.B. Coukell, A. Forer. (1995). Molecular cloning of an intracellular P-type ATPase from Dictyostelium that is up-regulated in calcium-adapted cells. J. Biol. Chem. 270: 28276-28281.

Moore, C.M., E.M. Hoey, A. Trudgett, and D.J. Timson. (2012). A plasma membrane Ca2+-ATPase (PMCA) from the liver fluke, Fasciola hepatica. Int J Parasitol 42: 851-858.

Moreno I., L. Norambuena, D. Maturana, M. Toro, C. Vergara, A. Orellana, A. Zurita-Silva, V.R. Ordenes. (2008). AtHMA1 Is a Thapsigargin-sensitive Ca2+/Heavy Metal Pump. J. Biol. Chem. 283: 9633-9641.

Morii, M., M. Yamauchi, T. Ichikawa, T. Fujii, Y. Takahashi, S. Asano, N. Takeguchi, and H. Sakai. (2008). Involvement of the H3O+-Lys-164 -Gln-161-Glu-345 charge transfer pathway in proton transport of gastric H+,K+-ATPase. J. Biol. Chem. 283: 16876-16884.

Morrill, G.A., A.B. Kostellow, L. Liu, R.K. Gupta, and A. Askari. (2016). Evolution of the α-Subunit of Na/K-ATPase from Paramecium to Homo sapiens: Invariance of Transmembrane Helix Topology. J. Mol. Evol. [Epub: Ahead of Print]

Morsomme, P., M. Chami, S. Marco, J. Nader, K.A. Ketchum, A. Goffeau, and J.-L. Rigaud. (2002). Characterization of a hyperthermophilic P-type ATPase from Methanococcus jannaschii expressed in yeast. J. Biol. Chem. 277: 29608-29616.

Morth J.P., B.P. Pedersen, M.S. Toustrup-Jensen, T.L. Sørensen, J. Petersen, J.P. Andersen, B. Vilsen, P. Nissen. (2007). Crystal structure of the sodium-potassium pump. Nature. 450: 1043-1049.

Mou, Y.N., B.J. Gao, K. Ren, S.M. Tong, S.H. Ying, and M.G. Feng. (2020). P-type Na/K ATPases essential and nonessential for cellular homeostasis and insect pathogenicity of. Virulence 11: 1415-1431.

Mukherjee, T., D. Mandal, and A. Bhaduri. (2001). Leishmania plasma membrane Mg2+-ATPase is a H+/K+-antiporter involved in glucose symport. J. Biol. Chem. 276: 55563-55569.

Mukhopadhyay, S. and A.D. Linstedt. (2011). Identification of a gain-of-function mutation in a Golgi P-type ATPase that enhances Mn2+ efflux and protects against toxicity. Proc. Natl. Acad. Sci. USA 108: 858-863.

Møller, J.V., C. Olesen, A.M. Winther, and P. Nissen. (2010). What can be learned about the function of a single protein from its various X-ray structures: the example of the sarcoplasmic calcium pump. Methods Mol Biol 654: 119-140.

Naik, P.K., M. Srivastava, P. Bajaj, S. Jain, A. Dubey, P. Ranjan, R. Kumar, and H. Singh. (2011). The binding modes and binding affinities of artemisinin derivatives with Plasmodium falciparum Ca2+-ATPase (PfATP6). J Mol Model 17: 333-357.

Nakakihara, E., H. Kondo, S. Nakashima, and B. Ezaki. (2009). Role of N-terminal His-rich Domain of Oscillatoria brevis Bxa1 in Both Ag(I)/Cu(I) and Cd(II)/Zn(II) Tolerance. Open Microbiol J 3: 15-22.

Nakamura, J., Y. Maruyama, G. Tajima, M. Suwa, and C. Sato. (2022). Elongation and Contraction of Scallop Sarcoplasmic Reticulum (SR): ATP Stabilizes Ca-ATPase Crystalline Array Elongation of SR Vesicles. Int J Mol Sci 23:.

Nakamura, J., Y. Maruyama, G. Tajima, Y. Komeiji, M. Suwa, and C. Sato. (2021). Ca-ATPase Molecules as a Calcium-Sensitive Membrane-Endoskeleton of Sarcoplasmic Reticulum. Int J Mol Sci 22:.

Nakanishi, H., K. Irie, K. Segawa, K. Hasegawa, Y. Fujiyoshi, S. Nagata, and K. Abe. (2020). Crystal structure of a human plasma membrane phospholipid flippase. J. Biol. Chem. 295: 10180-10194.

Nakanishi, H., T. Nishizawa, K. Segawa, O. Nureki, Y. Fujiyoshi, S. Nagata, and K. Abe. (2020). Transport Cycle of Plasma Membrane Flippase ATP11C by Cryo-EM. Cell Rep 32: 108208.

Nakazawa, N., X. Xu, O. Arakawa, and M. Yanagida. (2019). Coordinated Roles of the Putative Ceramide-Conjugation Protein, Cwh43, and a Mn-Transporting, P-Type ATPase, Pmr1, in Fission Yeast. G3 (Bethesda) 9: 2667-2676.

Neef, J., V.F. Andisi, K.S. Kim, O.P. Kuipers, and J.J. Bijlsma. (2011). Deletion of a cation transporter promotes lysis in Streptococcus pneumoniae. Infect. Immun. 79: 2314-2323.

Nielsen, H.N., K. Spontarelli, R. Holm, J.P. Andersen, A.P. Einholm, P. Artigas, and B. Vilsen. (2019). Distinct effects of Q925 mutation on intracellular and extracellular Na and K binding to the Na, K-ATPase. Sci Rep 9: 13344.

Norimatsu, Y., K. Hasegawa, N. Shimizu, and C. Toyoshima. (2017). Protein-phospholipid interplay revealed with crystals of a calcium pump. Nature 545: 193-198.

Nyblom, M., H. Poulsen, P. Gourdon, L. Reinhard, M. Andersson, E. Lindahl, N. Fedosova, and P. Nissen. (2013). Crystal structure of Na+, K+-ATPase in the Na+-bound state. Science 342: 123-127.

Okamoto, S., T. Naito, R. Shigetomi, Y. Kosugi, K. Nakayama, H. Takatsu, and H.W. Shin. (2020). The N- or C-terminal cytoplasmic regions of P4-ATPases determine their cellular localization. Mol. Biol. Cell 31: 2115-2124.

Okunade, G.W., M.L. Miller, M. Azhar, A. Andringa, L.P. Sanford, T. Doetschman, V. Prasad, and G.E. Shull. (2007). Loss of the Atp2c1 secretory pathway Ca2+-ATPase (SPCA1) in mice causes Golgi stress, apoptosis, and midgestational death in homozygous embryos and squamous cell tumors in adult heterozygotes. J. Biol. Chem. 282: 26517-26527.

Olesen C., M. Picard, A.M. Winther, C. Gyrup, J.P. Morth, C. Oxvig, J.V. Møller, P. Nissen. (2007). The structural basis of calcium transport by the calcium pump. Nature. 450: 1036-1042.

Onat OE., Gulsuner S., Bilguvar K., Nazli Basak A., Topaloglu H., Tan M., Tan U., Gunel M. and Ozcelik T. (2013). Missense mutation in the ATPase, aminophospholipid transporter protein ATP8A2 is associated with cerebellar atrophy and quadrupedal locomotion. Eur J Hum Genet. 21(3):281-5.

Padilla-Benavides T., McCann CJ. and Arguello JM. (2013). The mechanism of Cu+ transport ATPases: interaction with CU+ chaperones and the role of transient metal-binding sites. J Biol Chem. 288(1):69-78.

Palmgren M.G., Christensen G. (1994). Functional comparisons between plant plasma membrane H(+)-ATPase isoforms expressed in yeast. J. Biol. Chem. 269: 3027-3033.

Paulusma CC. and Elferink RP. (2010). P4 ATPases--the physiological relevance of lipid flipping transporters. FEBS Lett. 584(13):2708-16.

Pedersen, B.P., M.J. Buch-Pedersen, J.P. Morth, M.G. Palmgren, and P. Nissen. (2007). Crystal structure of the plasma membrane proton pump. Nature 450: 1111-1114.

Perandrés-López, R., M.P. Sánchez-Cañete, F. Gamarro, and S. Castanys. (2018). Functional role of highly-conserved residues of the N-terminal tail and first transmembrane segment of a P4-ATPase. Biochem. J. [Epub: Ahead of Print]

Pérez-Gordones, M.C., J.R. Ramírez-Iglesias, V. Cervino, G.L. Uzcanga, G. Benaim, and M. Mendoza. (2017). Evidence of the presence of a calmodulin-sensitive plasma membrane Ca2+-ATPase in Trypanosoma equiperdum. Mol Biochem Parasitol 213: 1-11.

Pérez-Victoria, F.J., F. Gamarro, M. Ouellette, and S. Castanys. (2003). Functional cloning of the miltefosine transporter. A novel P-type phospholipid translocase from Leishmania involved in drug resistance. J. Biol. Chem. 278: 49965-49971.

Peréz-Victoria, F.J., Sanchez-Canete, M.P., Castanys, S., and Gamarro, F. (2006). Phospholipid translocation and miltefosine potency require both L. donovani miltefosine transporter and the new protein LdRos3 in Leishmania parasites. J. Biol. Chem. 281: 23766-23775.

Petrov, V.V. (2009). Functioning of Saccharomyces cerevisiae Pma1 H+-ATPase carrying the minimal number of cysteine residues. Biochemistry (Mosc) 74: 1155-1163.

Petrov, V.V. (2015). Role of loop L5-6 connecting transmembrane segments M5 and M6 in biogenesis and functioning of yeast Pma1 H+-ATPase. Biochemistry (Mosc) 80: 31-44.

Petrov, V.V. (2017). Functioning of Yeast Pma1 H+-ATPase under Changing Charge: Role of Asp739 and Arg811 Residues. Biochemistry (Mosc) 82: 46-59.

Petrovich, G.D., G.R. Corradi, C.H. Pavan, S. Noli Truant, and H.P. Adamo. (2021). Highly exposed segment of the Spf1p P5A-ATPase near transmembrane M5 detected by limited proteolysis. PLoS One 16: e0245679.

Petrushanko, I.Y., V.A. Mitkevich, A.A. Anashkina, A.A. Adzhubei, K.M. Burnysheva, V.A. Lakunina, Y.V. Kamanina, E.A. Dergousova, O.D. Lopina, O.O. Ogunshola, A.Y. Bogdanova, and A.A. Makarov. (2016). Direct interaction of β-amyloid with Na,K-ATPase as a putative regulator of the enzyme function. Sci Rep 6: 27738.

Plattner, H. (2014). Calcium Regulation in the Protozoan Model, Paramecium tetraurelia. J Eukaryot Microbiol 61: 95-114.

Pohland, A.C. and D. Schneider. (2019). Mg2+ homeostasis and transport in cyanobacteria - at the crossroads of bacterial and chloroplast Mg2+ import. Biol Chem. [Epub: Ahead of Print]

Pomorski, T., R. Lombardi, H. Riezman, P.F. Devaux, G. van Meer, and J.C. Holthuis. (2003). Drs2p-related P-type ATPases Dnf1p and Dnf2p are required for phospholipid translocation across the yeast plasma membrane and serve a role in endocytosis. Mol. Biol. Cell. 14(3):1240-1254.

Pontel, L.B., M.E. Audero, M. Espariz, S.K. Checa, and F.C. Soncini. (2007). GolS controls the response to gold by the hierarchical induction of Salmonella-specific genes that include a CBA efflux-coding operon. Mol. Microbiol. 66: 814-825.

Poulsen, H., H. Khandelia, J.P. Morth, M. Bublitz, O.G. Mouritsen, J. Egebjerg, and P. Nissen. (2010). Neurological disease mutations compromise a C-terminal ion pathway in the Na+/K+-ATPase. Nature 467: 99-102.

Poulsen, H., P. Nissen, O.G. Mouritsen, and H. Khandelia. (2012). Protein kinase A (PKA) phosphorylation of Na+/K+-ATPase opens intracellular C-terminal water pathway leading to third Na+-binding site in molecular dynamics simulations. J. Biol. Chem. 287: 15959-15965.

Poulsen, L.R., R.L. López-Marqués, S.C. McDowell, J. Okkeri, D. Licht, A. Schulz, T. Pomorski, J.F. Harper, and M.G. Palmgren. (2008). The Arabidopsis P4-ATPase ALA3 Localizes to the Golgi and Requires a β-Subunit to Function in Lipid Translocation and Secretory Vesicle Formation. Plant Cell 20: 658-676.

Prell, J., G. Mulley, F. Haufe, J.P. White, A. Williams, R. Karunakaran, J.A. Downie, and P.S. Poole. (2012). The PTS(Ntr) system globally regulates ATP-dependent transporters in Rhizobium leguminosarum. Mol. Microbiol. 84: 117-129.

Prontera, P., P. Sarchielli, S. Caproni, C. Bedetti, L.M. Cupini, P. Calabresi, and C. Costa. (2018). Epilepsy in hemiplegic migraine: Genetic mutations and clinical implications. Cephalalgia 38: 361-373.

Purohit, R., M.O. Ross, S. Batelu, A. Kusowski, T.L. Stemmler, B.M. Hoffman, and A.C. Rosenzweig. (2018). Cu-specific CopB transporter: Revising P-type ATPase classification. Proc. Natl. Acad. Sci. USA. [Epub: Ahead of Print]

Qiao, C., N. Yin, H.Y. Gu, J.L. Zhu, J.H. Ding, M. Lu, and G. Hu. (2016). Atp13a2 Deficiency Aggravates Astrocyte-Mediated Neuroinflammation via NLRP3 Inflammasome Activation. CNS Neurosci Ther 22: 451-460.

Qiu, L.Y., E. Krieger, G. Schaftenaar, H.G. Swarts, P.H. Willems, J.J. De Pont, and J.B. Koenderink. (2005). Reconstruction of the complete ouabain-binding pocket of Na,K-ATPase in gastric H,K-ATPase by substitution of only seven amino acids. J. Biol. Chem. 280: 32349-32355.

Raimunda, D., T. Padilla-Benavides, S. Vogt, S. Boutigny, K.N. Tomkinson, L.A. Finney, and J.M. Argüello. (2013). Periplasmic response upon disruption of transmembrane Cu transport in Pseudomonas aeruginosa. Metallomics 5: 144-151.

Rajendran, V.M., P. Sangan, J. Geibel, and H.J. Binder. (2000). Ouabain-sensitive H,K-ATPase functions as Na,K-ATPase in apical membranes of rat distal colon. J. Biol. Chem. 275: 13035-13040.

Ray, N.B., L. Durairaj, B.B. Chen, B.J. McVerry, A.J. Ryan, M. Donahoe, A.K. Waltenbaugh, C.P. O'Donnell, F.C. Henderson, C.A. Etscheidt, D.M. McCoy, M. Agassandian, E.C. Hayes-Rowan, T.A. Coon, P.L. Butler, L. Gakhar, S.N. Mathur, J.C. Sieren, Y.Y. Tyurina, V.E. Kagan, G. McLennan, and R.K. Mallampalli. (2010). Dynamic regulation of cardiolipin by the lipid pump Atp8b1 determines the severity of lung injury in experimental pneumonia. Nat. Med. 16: 1120-1127.

Remy, E., M. Meyer, F. Blaise, M. Chabirand, N. Wolff, M.H. Balesdent, and T. Rouxel. (2008). The Lmpma1 gene of Leptosphaeria maculans encodes a plasma membrane H+-ATPase isoform essential for pathogenicity towards oilseed rape. Fungal Genet Biol 45: 1122-1134.

Rensing, C., B. Fan, R. Sharma, B. Mitra, amd B.P. Rosen. (2000). CopA: an Escherichia coli Cu(I)-translocating P-type ATPase. Proc. Natl. Acad. Sci. USA 97: 652-656.

Rensing, C., M. Ghosh, and B.P. Rosen. (1999). Families of soft-metal-ion transporting ATPase. J. Bacteriol. 181: 5891-5897.

Rensing, C., Y. Sun, B. Mitra, and B.P. Rosen. (1998). Pb(II)-translocating P-type ATPases. J. Biol. Chem. 273: 32614-32617.

Reyes, N., and D.C. Gadsby. (2006). Ion permeation through the Na+ ,K+ -ATPase. Nature 443: 470-474.

Riekhof, W.R. and D.R. Voelker. (2009). The yeast plasma membrane P(4)-ATPases are major transporters for lysophospholipids. Biochim. Biophys. Acta. 1791: 620-627.

Riekhof, W.R. and Voelker, D.R. (2006). Uptake and utilization of lyso-phosphatidylethanolamine by Saccharomyces cerevisiae. J. Biol. Chem. 281: 36588-36596.

Riekhof, W.R., J. Wu, M.A. Gijón, S. Zarini, R.C. Murphy, and D.R. Voelker. (2007). Lysophosphatidylcholine metabolism in Saccharomyces cerevisiae: the role of P-type ATPases in transport and a broad specificity acyltransferase in acylation. J. Biol. Chem. 282: 36853-36861.

Roberts, C.S., S. Muralidharan, F. Ni, and B. Mitra. (2020). Structural Role of the First Four Transmembrane Helices in ZntA, a P-Type ATPase from. Biochemistry 59: 4488-4498.

Rocafull, M.A., F.J. Romero, L.E. Thomas, and J.R. del Castillo. (2011). Isolation and cloning of the K+-independent, ouabain-insensitive Na+-ATPase. Biochim. Biophys. Acta. 1808: 1684-1700.

Rodacker, V., M. Toustrup-Jensen, and B. Vilsen. (2006). Mutations Phe785Leu and Thr618Met in Na+,K+-ATPase, associated with familial rapid-onset dystonia parkinsonism, interfere with Na+ interaction by distinct mechanisms. J. Biol. Chem. 281: 18539-18548.

Rodríguez, Y. and M. Májeková. (2020). Structural Changes of Sarco/Endoplasmic Reticulum Ca-ATPase Induced by Rutin Arachidonate: A Molecular Dynamics Study. Biomolecules 10:.

Rodríguez-Navarro, A. and B. Benito. (2010). Sodium or potassium efflux ATPase a fungal, bryophyte, and protozoal ATPase. Biochim. Biophys. Acta. 1798: 1841-1853.

Roegner, M.E., H.Y. Chen, and R.D. Watson. (2018). Molecular cloning and characterization of a sarco/endoplasmic reticulum Ca ATPase (SERCA) from Y-organs of the blue crab (Callinectes sapidus). Gene 673: 12-21.

Roland, B.P. and T.R. Graham. (2016). Directed evolution of a sphingomyelin flippase reveals mechanism of substrate backbone discrimination by a P4-ATPase. Proc. Natl. Acad. Sci. USA 113: E4460-4466.

Roland, B.P., T. Naito, J.T. Best, C. Arnaiz-Yépez, H. Takatsu, R.J. Yu, H.W. Shin, and T.R. Graham. (2019). Yeast and human P4-ATPases transport glycosphingolipids using conserved structural motifs. J. Biol. Chem. 294: 1794-1806.

Rosewich, H., A. Ohlenbusch, P. Huppke, L. Schlotawa, M. Baethmann, I. Carrilho, S. Fiori, C.M. Lourenço, S. Sawyer, R. Steinfeld, J. Gärtner, and K. Brockmann. (2014). The expanding clinical and genetic spectrum of ATP1A3-related disorders. Neurology 82: 945-955.

Rossbach, S., D.J. Mai, E.L. Carter, L. Sauviac, D. Capela, C. Bruand, and F.J. de Bruijn. (2008). Response of Sinorhizobium meliloti to elevated concentrations of cadmium and zinc. Appl. Environ. Microbiol. 74: 4218-4221.

Rossini, G.P. and A. Bigiani. (2011). Palytoxin action on the Na+,K+-ATPase and the disruption of ion equilibria in biological systems. Toxicon 57: 429-439.

Rothenbücher, M.C., S.J. Facey, D. Kiefer, M. Kossmann, and A. Kuhn. (2006). The cytoplasmic C-terminal domain of the Escherichia coli KdpD protein functions as a K+ sensor. J. Bacteriol. 188: 1950-1958.

Roy, A.S., S. Miskinyte, A. Garat, A. Hovnanian, M.A. Krzewinski-Recchi, and F. Foulquier. (2020). SPCA1 governs the stability of TMEM165 in Hailey-Hailey disease. Biochimie 174: 159-170.

Ruaud, A.F., L. Nilsson, F. Richard, M.K. Larsen, J.L. Bessereau, and S. Tuck. (2009). The C. elegans P4-ATPase TAT-1 regulates lysosome biogenesis and endocytosis. Traffic 10: 88-100.

Rui, H., A. Das, R. Nakamoto, and B. Roux. (2018). Proton Countertransport and Coupled Gating in the Sarcoplasmic Reticulum Calcium Pump. J. Mol. Biol. 430: 5050-5065.

Rutherford, J.C., J.S. Cavet, and N.J. Robinson. (1999). Cobalt-dependent transcriptional switching by a dual-effector MerR-like protein regulates a cobalt-exporting variant CPx-type ATPase. J. Biol. Chem. 274: 25827-25832.

Sacchetto, R., I. Bertipaglia, S. Giannetti, L. Cendron, F. Mascarello, E. Damiani, E. Carafoli, and G. Zanotti. (2012). Crystal structure of sarcoplasmic reticulum Ca2+-ATPase (SERCA) from bovine muscle. J Struct Biol 178: 38-44.

Safaei, R., S. Otani, B.J. Larson, M.L. Rasmussen, and S.B. Howell. (2008). Transport of cisplatin by the copper efflux transporter ATP7B. Mol Pharmacol 73: 461-468.

Sahoo, S.K., S.A. Shaikh, D.H. Sopariwala, N.C. Bal, D.S. Bruhn, W. Kopec, H. Khandelia, and M. Periasamy. (2015). The N Terminus of Sarcolipin Plays an Important Role in Uncoupling Sarco-endoplasmic Reticulum Ca2+-ATPase (SERCA) ATP Hydrolysis from Ca2+ Transport. J. Biol. Chem. 290: 14057-14067.

Sartorel, E., E. Barrey, R.K. Lau, and J. Thorner. (2015). Plasma membrane aminoglycerolipid flippase function is required for signaling competence in the yeast mating pheromone response pathway. Mol. Biol. Cell 26: 134-150.

Satoh-Nagasawa, N., M. Mori, N. Nakazawa, T. Kawamoto, Y. Nagato, K. Sakurai, H. Takahashi, A. Watanabe, and H. Akagi. (2012). Mutations in Rice (Oryza sativa) Heavy Metal ATPase 2 (OsHMA2) Restrict the Translocation of Zinc and Cadmium. Plant Cell Physiol. 53: 213-224.

Scarborough, G.A. (1999). Structure and function of the P-type ATPases. Curr. Opin. Cell Biol. 11: 517-522.

Schack, V.R., R. Holm, and B. Vilsen. (2012). Inhibition of phosphorylation of na+,k+-ATPase by mutations causing familial hemiplegic migraine. J. Biol. Chem. 287: 2191-2202.

Schäffers, O.J.M., J.G.J. Hoenderop, R.J.M. Bindels, and J.H.F. de Baaij. (2018). The rise and fall of novel renal magnesium transporters. Am. J. Physiol. Renal Physiol 314: F1027-F1033.

Scheiner-Bobis, G. (2002). The sodium pump. Its molecular properties and mechanics of ion transport. Eur. J. Biochem. 269: 2424-2433.

Scherer, J. and D.H. Nies. (2009). CzcP is a novel efflux system contributing to transition metal resistance in Cupriavidus metallidurans CH34. Mol. Microbiol. 73: 601-621.

Schiøtt, M., S.M. Romanowsky, L. Baekgaard, M.K. Jakobsen, M.G. Palmgren, and J.F. Harper. (2004). A plant plasma membrane Ca2+ pump is required for normal pollen tube growth and fertilization. Proc. Natl. Acad. Sci. USA 101: 9502-9507.

Schmidt, R.S., J.P. Macêdo, M.E. Steinmann, A.G. Salgado, P. Bütikofer, E. Sigel, D. Rentsch, and P. Mäser. (2018). Transporters of Trypanosoma brucei-phylogeny, physiology, pharmacology. FEBS J. 285: 1012-1023.

Schmutz, I., V. Jagannathan, F. Bartenschlager, V.M. Stein, A.D. Gruber, T. Leeb, and M.L. Katz. (2019). ATP13A2 missense variant in Australian Cattle Dogs with late onset neuronal ceroid lipofuscinosis. Mol Genet Metab. [Epub: Ahead of Print]

Schoner, W. (2002). Endogenous cardiac glycosides, a new class of steroid hormones. Eur. J. Biochem. 269: 2440-2448.

Seflova, J., N.R. Habibi, J.Q. Yap, S.R. Cleary, X. Fang, P.M. Kekenes-Huskey, L.M. Espinoza-Fonseca, J.B. Bossuyt, and S.L. Robia. (2022). Fluorescence lifetime imaging microscopy reveals sodium pump dimers in live cells. J. Biol. Chem. 298: 101865.

Šeflová, J., P. Čechová, T. Štenclová, M. Šebela, and M. Kubala. (2018). Identification of cisplatin-binding sites on the large cytoplasmic loop of the Na/K-ATPase. J Enzyme Inhib Med Chem 33: 701-706.

Segall, L., R. Scanzano, M.A. Kaunisto, M. Wessman, A. Palotie, J.J. Gargus, and R. Blostein. (2004). Kinetic alterations due to a missense mutation in the Na,K-ATPase alpha2 subunit cause familial hemiplegic migraine type 2. J. Biol. Chem. 279: 43692-43696.

Segawa, K., A. Kikuchi, T. Noji, Y. Sugiura, K. Hiraga, C. Suzuki, K. Haginoya, Y. Kobayashi, M. Matsunaga, Y. Ochiai, K. Yamada, T. Nishimura, S. Iwasawa, W. Shoji, F. Sugihara, K. Nishino, H. Kosako, M. Ikawa, Y. Uchiyama, M. Suematsu, H. Ishikita, S. Kure, and S. Nagata. (2021). A sublethal ATP11A mutation associated with neurological deterioration causes aberrant phosphatidylcholine flipping in plasma membranes. J Clin Invest 131:.

Segawa, K., S. Kurata, Y. Yanagihashi, T.R. Brummelkamp, F. Matsuda, and S. Nagata. (2014). Caspase-mediated cleavage of phospholipid flippase for apoptotic phosphatidylserine exposure. Science 344: 1164-1168.

Seigneurin-Berny, D., A. Gravot, P. Auroy, C. Mazard, A. Kraut, G. Finazzi, D. Grunwald, F. Rappaport, A. Vavasseur, J. Joyard, P. Richaud, and N. Rolland. (2006). HMA1, a new Cu-ATPase of the chloroplast envelope, is essential for growth under adverse light conditions. J. Biol. Chem. 281: 2882-2892.

Shah, V.S., D.K. Meyerholz, X.X. Tang, L. Reznikov, M. Abou Alaiwa, S.E. Ernst, P.H. Karp, C.L. Wohlford-Lenane, K.P. Heilmann, M.R. Leidinger, P.D. Allen, J. Zabner, P.B. McCray, Jr, L.S. Ostedgaard, D.A. Stoltz, C.O. Randak, and M.J. Welsh. (2016). Airway acidification initiates host defense abnormalities in cystic fibrosis mice. Science 351: 503-507.

Shan, W., J. Liu, and A.R. Hardham. (2006). Phytophthora nicotianae PnPMA1 encodes an atypical plasma membrane H+ -ATPase that is functional in yeast and developmentally regulated. Fungal Genet Biol 43: 583-592.

Shao, L.R., R. Janicot, and C.E. Stafstrom. (2021). Na-K-ATPase functions in the developing hippocampus: regional differences in CA1 and CA3 neuronal excitability and role in epileptiform network bursting. J Neurophysiol 125: 1-11.

Shin, H.W. and H. Takatsu. (2019). Substrates of P4-ATPases: beyond aminophospholipids (phosphatidylserine and phosphatidylethanolamine). FASEB J. 33: 3087-3096.

Shin, J.M., K. Munson, and G. Sachs. (2011). Gastric H+,K+-ATPase. Compr Physiol 1: 2141-2153.

Shin, J.M., K. Munson, O. Vagin, and G. Sachs. (2009). The gastric HK-ATPase: structure, function, and inhibition. Pflugers Arch 457: 609-622.

Shinoda, T., H. Ogawa, F. Cornelius, and C. Toyoshima. (2009). Crystal structure of the sodium-potassium pump at 2.4 Å resolution. Nature 459: 446-450.

Shono, M., M. Wada, Y. Hara, and T. Fujii. (2001). Molecular cloning of Na+-ATPase cDNA from a marine alga Heterosigma akashiwo. Biochim. Biophys. Acta 1511: 193-199.

Sigruener, A., C. Wolfrum, A. Boettcher, T. Kopf, G. Liebisch, E. Orsó, and G. Schmitz. (2017). Lipidomic and metabolic changes in the P4-type ATPase ATP10D deficient C57BL/6J wild type mice upon rescue of ATP10D function. PLoS One 12: e0178368.

Silva, C.I.D., C.F. Gonçalves-de-Albuquerque, B.P.T. Moraes, D.G. Garcia, and P. Burth. (2021). Na/K-ATPase: Their role in cell adhesion and migration in cancer. Biochimie 185: 1-8.

Silver, S. (1996). Transport of inorganic cations. In F.C. Neidhardt et al. (eds.), Escherichia coli and Salmonella typhimurium: Cellular and Molecular Biology, 2nd ed. Washington, D.C.: ASM Press, pp. 1091-1102.

Sim, S.I., S. von Bülow, G. Hummer, and E. Park. (2021). Structural basis of polyamine transport by human ATP13A2 (PARK9). Mol. Cell. [Epub: Ahead of Print]

Singh, K., D.B. Senadheera, C.M. Lévesque, and D.G. Cvitkovitch. (2015). The copYAZ Operon Functions in Copper Efflux, Biofilm Formation, Genetic Transformation, and Stress Tolerance in Streptococcus mutans. J. Bacteriol. 197: 2545-2557.

Sinkins WG., Estacion M., Prasad V., Goel M., Shull GE., Kunze DL. and Schilling WP. (2009). Maitotoxin converts the plasmalemmal Ca(2+) pump into a Ca(2+)-permeable nonselective cation channel. Am J Physiol Cell Physiol. 297(6):C1533-43.

Sitthisak, S., L. Knutsson, J.W. Webb, and R.K. Jayaswal. (2007). Molecular characterization of the copper transport system in Staphylococcus aureus. Microbiology. 153: 4274-4283.

Smart, J.P., M.J. Cliff, and D.J. Kelly. (2009). A role for tungsten in the biology of Campylobacter jejuni: tungstate stimulates formate dehydrogenase activity and is transported via an ultra-high affinity ABC system distinct from the molybdate transporter. Mol. Microbiol. 74: 742-757.

Smith, A.T., M.O. Ross, B.M. Hoffman, and A.C. Rosenzweig. (2017). Metal Selectivity of a Cd-, Co-, and Zn-Transporting P1B-type ATPase. Biochemistry 56: 85-95.

Smolyaninova, L.V., S.V. Koltsova, S.V. Sidorenko, and S.N. Orlov. (2017). Augmented gene expression triggered by Na+,K+-ATPase inhibition: Role of Ca2+i-mediated and -independent excitation-transcription coupling. Cell Calcium 68: 5-13.

Spontarelli, K., D.T. Infield, H.N. Nielsen, R. Holm, V.C. Young, J.D. Galpin, C.A. Ahern, B. Vilsen, and P. Artigas. (2022). Role of a conserved ion-binding site tyrosine in ion selectivity of the Na+/K+ pump. J Gen Physiol 154:.

Stevens, H.C., L. Malone, and J.W. Nichols. (2008). The putative aminophospholipid translocases, DNF1 and DNF2, are not required for 7-nitrobenz-2-oxa-1,3-diazol-4-yl-phosphatidylserine flip across the plasma membrane of Saccharomyces cerevisiae. J. Biol. Chem. 283: 35060-35069.

Stokes, D.L. and N.M. Green. (2000). Modeling a dehalogenase fold into the 8-Å density map for Ca2+-ATPase defines a new domain structure. Biophys. J. 78: 1765-1776.

Stone, A., C. Chau, C. Eaton, E. Foran, M. Kapur, E. Prevatt, N. Belkin, D. Kerr, T. Kohlin, and P. Williamson. (2012). Biochemical characterization of P4-ATPase mutations identified in patients with progressive familial intrahepatic cholestasis. J. Biol. Chem. 287: 41139-41151.

Suisse, A. and J.E. Treisman. (2019). Reduced SERCA Function Preferentially Affects Wnt Signaling by Retaining E-Cadherin in the Endoplasmic Reticulum. Cell Rep 26: 322-329.e3.

Suleiman, J., N. Hamwi, and A.W. El-Hattab. (2018). ATP13A2 novel mutations causing a rare form of juvenile-onset Parkinson disease. Brain Dev 40: 824-826.

Suman, J., P. Kotrba, and T. Macek. (2014). Putative P1B-type ATPase from the bacterium Achromobacter xylosoxidans A8 alters Pb2+/Zn2+/Cd2+-resistance and accumulation in Saccharomyces cerevisiae. Biochim. Biophys. Acta. 1838: 1338-1343.

Sun, B., B.D. Stewart, A.N. Kucharski, and P.M. Kekenes-Huskey. (2019). Thermodynamics of Cation Binding to the Sarcoendoplasmic Reticulum Calcium ATPase Pump and Impacts on Enzyme Function. J Chem Theory Comput 15: 2692-2705.

Supper, V., H.B. Schiller, W. Paster, F. Forster, C. Boulègue, G. Mitulovic, V. Leksa, A. Ohradanova-Repic, C. Machacek, P. Schatzlmaier, G.J. Zlabinger, and H. Stockinger. (2016). Association of CD147 and Calcium Exporter PMCA4 Uncouples IL-2 Expression from Early TCR Signaling. J Immunol 196: 1387-1399.

Svrckova, M., M. Zatloukalova, P. Dvorakova, D. Coufalova, D. Novak, L. Hernychova, and J. Vacek. (2017). Na/K-ATPase interaction with methylglyoxal as reactive metabolic side product. Free Radic Biol Med 108: 146-154.

Swarts, H.G., J.B. Koenderink, P.H. Willems, and J.J. De Pont. (2005). The non-gastric H,K-ATPase is oligomycin-sensitive and can function as an H+,NH4+-ATPase. J. Biol. Chem. 280: 33115-33122.

Sørensen, D.M., T. Holemans, S. van Veen, S. Martin, T. Arslan, I.W. Haagendahl, H.W. Holen, N.N. Hamouda, J. Eggermont, M. Palmgren, and P. Vangheluwe. (2018). Parkinson disease related ATP13A2 evolved early in animal evolution. PLoS One 13: e0193228.

Tadini-Buoninsegni, F., S.A. Mikkelsen, L.S. Mogensen, R.S. Molday, and J.P. Andersen. (2019). Phosphatidylserine flipping by the P4-ATPase ATP8A2 is electrogenic. Proc. Natl. Acad. Sci. USA 116: 16332-16337.

Takar, M., Y. Wu, and T.R. Graham. (2016). The Essential Neo1 Protein from Budding Yeast Plays a Role in Establishing Aminophospholipid Asymmetry of the Plasma Membrane. J. Biol. Chem. 291: 15727-15739.

Takeda, K., H. Eguchi, S. Soeda, A. Shirahata, and M. Kawamura. (2005). Fe(II)/Cu(I)-dependent P-type ATPase activity in the liver of Long-Evans cinnamon rats. Life Sci. 76: 2203-2209.

Takemura, Y., N. Tamura, M. Imamura, and N. Koyama. (2009). Role of the charged amino acid residues in the cytoplasmic loop between putative transmembrane segments 6 and 7 of Na+-ATPase of an alkaliphilic bacterium, Exiguobacterium aurantiacum. FEMS Microbiol. Lett. 299: 143-148.

Takeuchi, A., N. Reyes, P. Artigas, and D.C. Gadsby. (2008). The ion pathway through the opened Na+,K+-ATPase pump. Nature 456: 413-416.

Taneja, M. and S.K. Upadhyay. (2018). Molecular characterization and differential expression suggested diverse functions of P-type II CaATPases in Triticum aestivum L. BMC Genomics 19: 389.

Tang, X., M.S. Halleck, R.A. Schlegel, and P. Williamson. (1996). A subfamily of P-type ATPases with aminophospholipid transporting activity. Science 272: 1495-1497.

Tapken W., Kim J., Nishimura K., van Wijk KJ. and Pilon M. (2015). The Clp protease system is required for copper ion-dependent turnover of the PAA2/HMA8 copper transporter in chloroplasts. New Phytol. 205(2):511-7.

Tatyanenko, L.V., O.V. Pokidova, N.S. Goryachev, O.A. Kraevaya, E.A. Khakina, A.Y. Belik, A.Y. Rybkin, O.V. Dobrokhotova, Y. Pikhteleva, P.A. Troshin, and A.I. Kotelnikov. (2020). Effects of Covalent Conjugates of Fullerene Derivatives with Xanthene Dyes on Activity of Ca-ATPase of the Sarcoplasmic Reticulum. Bull Exp Biol Med 169: 89-94.

Tauber, P., B. Aichinger, C. Christ, J. Stindl, Y. Rhayem, F. Beuschlein, R. Warth, and S. Bandulik. (2016). Cellular Pathophysiology of an Adrenal Adenoma-Associated Mutant of the Plasma Membrane Ca2+-ATPase ATP2B3. Endocrinology 157: 2489-2499.

Therien, A.G., S.J.D. Karlish, and R. Blostein. (1999). Expression and functional role of the γ-subunit of the Na,K-ATPase in mammalian cells. J. Biol. Chem. 274: 12252-12256.

Thever, M.D. and M.H. Saier, Jr. (2009). Bioinformatic characterization of p-type ATPases encoded within the fully sequenced genomes of 26 eukaryotes. J. Membr. Biol. 229: 115-130.

Thirman, J., H. Rui, and B. Roux. (2021). Elusive Intermediate State Key in the Conversion of ATP Hydrolysis into Useful Work Driving the Ca Pump SERCA. J Phys Chem B 125: 2921-2928.

Ticconi, C.A., C.A. Delatorre, B. Lahner, D.E. Salt, and S. Abel. (2004). Arabidopsis pdr2 reveals a phosphate-sensitive checkpoint in root development. Plant J. 37: 801-814.

Ticconi, C.A., R.D. Lucero, S. Sakhonwasee, A.W. Adamson, A. Creff, L. Nussaume, T. Desnos, and S. Abel. (2009). ER-resident proteins PDR2 and LPR1 mediate the developmental response of root meristems to phosphate availability. Proc. Natl. Acad. Sci. USA 106: 14174-14179.

Timcenko, M., J.A. Lyons, D. Januliene, J.J. Ulstrup, T. Dieudonné, C. Montigny, M.R. Ash, J.L. Karlsen, T. Boesen, W. Kühlbrandt, G. Lenoir, A. Moeller, and P. Nissen. (2019). Structure and autoregulation of a P4-ATPase lipid flippase. Nature 571: 366-370.

Timcenko, M., T. Dieudonné, C. Montigny, T. Boesen, J.A. Lyons, G. Lenoir, and P. Nissen. (2021). Structural Basis of Substrate-Independent Phosphorylation in a P4-ATPase Lipid Flippase. J. Mol. Biol. 433: 167062.

Tiwari, S., G. Rajamanickam, V. Unnikrishnan, M. Ojaghi, J.P. Kastelic, and J.C. Thundathil. (2022). Testis-Specific Isoform of Na-K ATPase and Regulation of Bull Fertility. Int J Mol Sci 23:.

Ton, V.-K., D. Mandal, C. Vahadji, and R. Rao. (2002). Functional expression in yeast of the human secretory pathway Ca2+, Mn2+-ATPase defective in Hailey-Hailey disease. J. Biol. Chem. 277: 6422-6427.

Tong, L., S. Nakashima, M. Shibasaka, M. Katsuhara, and K. Kasamo. (2002). A novel histidine-rich CPx-ATPase from the filamentous cyanobacterium Oscillatoria brevis related to multiple-heavy-metal cotolerance. J. Bacteriol. 184: 5027-5035.

Toyoshima, C. (2008). Structural aspects of ion pumping by Ca2+-ATPase of sarcoplasmic reticulum. Arch Biochem Biophys 476: 3-11.

Toyoshima, C. and H. Nomura. (2002). Structural changes in the calcium pump accompanying the dissociation of calcium. Nature 418: 598-599.

Toyoshima, C. and T. Mizutani. (2004). Crystal structure of the calcium pump with a bound ATP analogue. Nature 430: 529-535.

Toyoshima, C., M. Nakasako, H. Nomura, and H. Ogawa. (2000). Crystal structure of the calcium pump of sarcoplasmic reticulum at 2.6 Å resolution. Nature 405: 633-634.

Toyoshima, C., Y. Norimatsu, S. Iwasawa, T. Tsuda, and H. Ogawa. (2007). How processing of aspartylphosphate is coupled to lumenal gating of the ion pathway in the calcium pump. Proc. Natl. Acad. Sci. USA 104: 19831-19836.

Traverso, M.E., P. Subramanian, R. Davydov, B.M. Hoffman, T.L. Stemmler, and A.C. Rosenzweig. (2010). Identification of a hemerythrin-like domain in a P1B-type transport ATPase. Biochemistry 49: 7060-7068.

Tsai, K.-J., Y.-F. Lin, M.D. Wong, H.H.-C. Yang, H.-L. Fu, and B.P. Rosen. (2002). Membrane topology of the p1258 CadA Cd(II)/Pb(II)/Zn(II)-translocating P-type ATPase. J. Bioenerg. Biomembr. 34: 147-156.

Tümer, Z. (2013). An overview and update of ATP7A mutations leading to Menkes disease and occipital horn syndrome. Hum Mutat 34: 417-429.

Tynecka, Z., A. Malm, and Z. Goś-Szcześniak. (2016). Cd2+ extrusion by P-type Cd2+-ATPase of Staphylococcus aureus 17810R via energy-dependent Cd2+/H+ exchange mechanism. Biometals 29: 651-663.

Ueno, D., N. Yamaji, I. Kono, C.F. Huang, T. Ando, M. Yano, and J.F. Ma. (2010). Gene limiting cadmium accumulation in rice. Proc. Natl. Acad. Sci. USA 107: 16500-16505.

Ueno, S., N. Kaieda, and N. Koyama. (2000). Characterization of a P-type Na+-ATPase of a facultatively anaerobic alkaliphile, Exiguobacterium aurantiacum. J. Biol. Chem. 275: 14537-14540.

Ushimaru, M. and Y. Fukushima. (2008). The dimeric form of Ca2+-ATPase is involved in Ca2+ transport in the sarcoplasmic reticulum. Biochem. J. 414: 357-361.

Van Baelen K., J. Vanoevelen, L. Missiaen, L. Raeymaekers, F. Wuytack. (2001). The Golgi PMR1 P-type ATPase of Caenorhabditis elegans. Identification of the gene and demonstration of calcium and manganese transport. J. Biol. Chem. 276: 10683-10691.

van der Mark, V.A., D.R. de Waart, K.S. Ho-Mok, M.M. Tabbers, H.W. Voogt, R.P. Oude Elferink, A.S. Knisely, and C.C. Paulusma. (2014). The lipid flippase heterodimer ATP8B1-CDC50A is essential for surface expression of the apical sodium-dependent bile acid transporter (SLC10A2/ASBT) in intestinal Caco-2 cells. Biochim. Biophys. Acta. 1842: 2378-2386.

van der Velden, L.M., S.F. van de Graaf, and L.W. Klomp. (2010). Biochemical and cellular functions of P4 ATPases. Biochem. J. 431: 1-11.

van Veen, S., S. Martin, C. Van den Haute, V. Benoy, J. Lyons, R. Vanhoutte, J.P. Kahler, J.P. Decuypere, G. Gelders, E. Lambie, J. Zielich, J.V. Swinnen, W. Annaert, P. Agostinis, B. Ghesquière, S. Verhelst, V. Baekelandt, J. Eggermont, and P. Vangheluwe. (2020). ATP13A2 deficiency disrupts lysosomal polyamine export. Nature 578: 419-424.

Vedovato, N. and D.C. Gadsby. (2010). The two C-terminal tyrosines stabilize occluded Na/K pump conformations containing Na or K ions. J Gen Physiol 136: 63-82.

Veshaguri, S., S.M. Christensen, G.C. Kemmer, G. Ghale, M.P. Møller, C. Lohr, A.L. Christensen, B.H. Justesen, I.L. Jørgensen, J. Schiller, N.S. Hatzakis, M. Grabe, T.G. Pomorski, and D. Stamou. (2016). Direct observation of proton pumping by a eukaryotic P-type ATPase. Science 351: 1469-1473.

Villafane, A.A., Y. Voskoboynik, M. Cuebas, I. Ruhl, and E. Bini. (2009). Response to excess copper in the hyperthermophile Sulfolobus solfataricus strain 98/2. Biochem. Biophys. Res. Commun. 385: 67-71.

Wang, S.H., K.L. Wang, W.K. Yang, T.H. Lee, W.Y. Lo, and J.D. Lee. (2017). Expression and potential roles of sodium-potassium ATPase and E-cadherin in human gastric adenocarcinoma. PLoS One 12: e0183692.

Wang, Y., P. Luo, L. Zhang, C. Hu, C. Ren, and J. Xia. (2013). Cloning of sarco/endoplasmic reticulum Ca2+-ATPase (SERCA) gene from white shrimp, Litopenaeus vannamei and its expression level analysis under salinity stress. Mol Biol Rep 40: 6213-6221.

Watanabe, Y., Y. Shimono, H. Tsuji, and Y. Tamai. (2002). Role of the glutamic and aspartic residues in Na+-ATPase function in the ZrENA1 gene of Zygosaccharomyces rouxii. FEMS Microbiol. Lett. 209: 39-43.

Wdowikowska, A. and G. Kłobus. (2011). [Plant P-type ATPases]. Postepy Biochem 57: 85-91.

Weissman, Z., R. Shemer, and D. Kornitzer. (2002). Deletion of the copper transporter CaCCC2 reveals two distinct pathways for iron acquisition in Candida albicans. Mol. Microbiol. 44: 1551-1560.

Wetzel, R.K., J.L. Pascoa, and E. Arystarkhova. (2004). Stress-induced expression of the γsubunit (FXYD2) modulates Na,K-ATPase activity and cell growth. J. Biol. Chem. 279: 41750-41757.

Wheatly, M.G., Y. Gao, L.M. Stiner, D.R. Whalen, M. Nade, F. Vigo, and A.E. Golshani. (2007). Roles of NCX and PMCA in basolateral calcium export associated with mineralization cycles and cold acclimation in crayfish. Ann. N.Y. Acad. Sci. 1099: 190-192.

Wiangnon, K., W. Raksajit, and A. Incharoensakdi. (2007). Presence of a Na+-stimulated P-type ATPase in the plasma membrane of the alkaliphilic halotolerant cyanobacterium Aphanothece halophytica. FEMS Microbiol. Lett. 270: 139-145.

Wielandt AG., Pedersen JT., Falhof J., Kemmer GC., Lund A., Ekberg K., Fuglsang AT., Pomorski TG., Buch-Pedersen MJ. and Palmgren M. (2015). Specific Activation of the Plant P-type Plasma Membrane H+-ATPase by Lysophospholipids Depends on the Autoinhibitory N- and C-terminal Domains. J Biol Chem. 290(26):16281-91.

Wiriyasermkul, P., S. Moriyama, and S. Nagamori. (2020). Membrane transport proteins in melanosomes: Regulation of ions for pigmentation. Biochim. Biophys. Acta. Biomembr 1862: 183318.

Wolf, S., K. Pflüger-Grau, and A. Kremling. (2015). Modeling the Interplay of Pseudomonas putida EIIA with the Potassium Transporter KdpFABC. J. Mol. Microbiol. Biotechnol. 25: 178-194.

Wolschendorf, F., D. Ackart, T.B. Shrestha, L. Hascall-Dove, S. Nolan, G. Lamichhane, Y. Wang, S.H. Bossmann, R.J. Basaraba, and M. Niederweis. (2011). Copper resistance is essential for virulence of Mycobacterium tuberculosis. Proc. Natl. Acad. Sci. USA 108: 1621-1626.

Worrell, R.T., L. Merk, and J.B. Matthews. (2008). Ammonium transport in the colonic crypt cell line, T84: role for Rhesus glycoproteins and NKCC1. Am. J. Physiol. Gastrointest. Liver Physiol. 294: G429-440.

Wu, C.C., A. Gardarin, A. Martel, E. Mintz, F. Guillain, and P. Catty. (2006). The cadmium transport sites of CadA, the Cd2+-ATPase from Listeria monocytogenes. J. Biol. Chem. 281: 29533-29541.

Wu, C.C., W.J. Rice, and D.L. Stokes. (2008). Structure of a copper pump suggests a regulatory role for its metal-binding domain. Structure 16: 976-985.

Wu, C.H., L.A. Vasilets, K. Takeda, M. Kawamura, and W. Schwarz. (2003). Functional role of the N-terminus of a Na+,K+-ATPase α-subunit as an inactivation gate of palytoxin-induced pump channel. Biochim. Biophys. Acta 1609: 55-62.

Xiang, M., D. Mohamalawari, and R. Rao. (2005). A novel isoform of the secretory pathway Ca2+,Mn2+-ATPase, hSPCA2, has unusual properties and is expressed in the brain. J. Biol. Chem. 280: 11608-11614.

Xu, C., W.J. Rice, W. He, and D.L. Stokes. (2002). A structural model for the catalytic cycle of Ca2+-ATPase. J. Mol. Biol. 316: 201-211.

Yan, S., E.S. McLamore, S. Dong, H. Gao, M. Taguchi, N. Wang, T. Zhang, X. Su, and Y. Shen. (2015). The role of plasma membrane H+ -ATPase in jasmonate-induced ion fluxes and stomatal closure in Arabidopsis thaliana. Plant J. 83: 638-649.

Yang, Y., A.K. Mandal, L.M. Bredeston, F.L. González-Flecha, and J.M. Argüello. (2007). Activation of Archaeoglobus fulgidus Cu(+)-ATPase CopA by cysteine. Biochim. Biophys. Acta. 1768: 495-501.

Yang, Y., C. Hao, J. Du, L. Xu, Z. Guo, D. Li, H. Cai, H. Guo, and L. Li. (2022). The carboxy terminal transmembrane domain of SPL7 mediates interaction with RAN1 at the endoplasmic reticulum to regulate ethylene signalling in Arabidopsis. New Phytol. [Epub: Ahead of Print]

Yi, L. and S. Kaler. (2014). ATP7A trafficking and mechanisms underlying the distal motor neuropathy induced by mutations in ATP7A. Ann. N.Y. Acad. Sci. 1314: 49-54.

Yi, L. and S.G. Kaler. (2018). Interaction between the AAA ATPase p97/VCP and a concealed UBX domain in the copper transporter ATP7A is associated with motor neuron degeneration. J. Biol. Chem. [Epub: Ahead of Print]

Young, V.C. and P. Artigas. (2021). Displacement of the Na/K pump''s transmembrane domains demonstrates conserved conformational changes in P-type 2 ATPases. Proc. Natl. Acad. Sci. USA 118:.

Zeng, X.T., T. Higashida, M. Hara, N. Hattori, K. Kitagawa, K. Omori, and C. Inagaki. (1999). Antiserum against Cl- pump complex recognizes 51 kDa protein, a possible catalytic unit in rat brain. Neurosci. Lett. 258: 85-88.

Zhang, B., J. Groffen, and N. Heisterkamp. (2005). Resistance to farnesyltransferase inhibitors in Bcr/Abl-positive lymphoblastic leukemia by increased expression of a novel ABC transporter homolog ATP11a. Blood 106: 1355-1361.

Zhang, C., X. Wei, G.S. Omenn, and Y. Zhang. (2018). Structure and Protein Interaction-based Gene Ontology Annotations Reveal Likely Functions of Uncharacterized Proteins on Human Chromosome 17. J Proteome Res. [Epub: Ahead of Print]

Zhang, S., S. Malmersjö, J. Li, H. Ando, O. Aizman, P. Uhlén, K. Mikoshiba, and A. Aperia. (2006). Distinct role of the N-terminal tail of the Na,K-ATPase catalytic subunit as a signal transducer. J. Biol. Chem. 281: 21954-21962.

Zhang, Y. and K. Inaba. (2022). Structural basis of the conformational and functional regulation of human SERCA2b, the ubiquitous endoplasmic reticulum calcium pump. Bioessays 44: e2200052.

Zhang, Y., Q. Li, L. Xu, X. Qiao, C. Liu, and S. Zhang. (2020). Comparative analysis of the P-type ATPase gene family in seven Rosaceae species and an expression analysis in pear (Pyrus bretschneideri Rehd.). Genomics. [Epub: Ahead of Print]

Zhang, Z., D. Lewis, C. Strock, G. Inesi, M. Nakasako, H. Nomura, and C. Toyoshima. (2000). Detaled characterization of the cooperative mechanism of a Ca(2+) binding and catalytic activation in the Ca(2+) transport (SERCA) ATPase. Biocemistry 39: 8758-8767.

Zhao, P., C. Zhao, D. Chen, C. Yun, H. Li, and L. Bai. (2021). Structure and activation mechanism of the hexameric plasma membrane H-ATPase. Nat Commun 12: 6439.

Zheng, K. and T. Li. (2021). Prediction of ATPase cation transporting 13A2 molecule in Petromyzon marinus and pan-cancer analysis into human tumors from an evolutionary perspective. Immunogenetics 73: 277-289.

Zhihao, L., N. Jingyu, L. Lan, S. Michael, G. Rui, B. Xiyun, L. Xiaozhi, and F. Guanwei. (2020). SERCA2a: a key protein in the Ca cycle of the heart failure. Heart Fail Rev 25: 523-535.

Zhou, X. and T.R. Graham. (2009). Reconstitution of phospholipid translocase activity with purified Drs2p, a type-IV P-type ATPase from budding yeast. Proc. Natl. Acad. Sci. USA 106: 16586-16591.

Zhu, M., H. Sun, L. Cao, Z. Wu, B. Leng, and J. Bian. (2022). Role of Na/K-ATPase in ischemic stroke: in-depth perspectives from physiology to pharmacology. J Mol Med (Berl) 100: 395-410.

Zielazinski, E.L., G.E. Cutsail, 3rd, B.M. Hoffman, T.L. Stemmler, and A.C. Rosenzweig. (2012). Characterization of a Cobalt-Specific P(1B)-ATPase. Biochemistry 51: 7891-7900.

Zielich, J., E. Tzima, E.A. Schröder, F. Jemel, B. Conradt, and E.J. Lambie. (2018). Overlapping expression patterns and functions of three paralogous P5B ATPases in Caenorhabditis elegans. PLoS One 13: e0194451.

Zizkova, P., J. Viskupicova, V. Heger, L. Rackova, M. Majekova, and L. Horakova. (2018). Dysfunction of SERCA pumps as novel mechanism of methylglyoxal cytotoxicity. Cell Calcium 74: 112-122.

Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
3.A.3.1.1

Na+-, K+-ATPase (Na+ efflux; K+ uptake).  Kinetic alterations due to a missense mutation in the alpha2 subunit cause familial hemiplegic migraine type 2 (Segall et al. 2004). Mutation in the γ-subunit causes renal hypomagnesemia, associated with hypocalciurea (Cairo et al., 2008). The Na/K-ATPase is an important signal transducer that not only interacts and regulates protein kinases, but also functions as a scaffold (Li and Xie, 2009). Capsazepine, a synthetic vanilloid, converts the Na, K-ATPase to a Na-ATPase (Mahmmoud, 2008a). There are alternative α- and β-subunits, α1, α2,... β1, β2,... in muscle which form α1β1, α1β2, α2β1 and α2β2, heterodimers, each with differing Na+ affinities (4-13mM) (Kristensen and Juel, 2010). α3 and β3 isoforms have also been identified. The γ-subunit is the same as TC# 1.A.27.2.1. Poulsen et al. (2010) have proposed a second ion conduction pathway in the C-terminal part of the ATPase. The two C-terminal tyrosines stabilize the occluded Na/K pump conformations containing Na or K ions (Vedovato and Gadsby, 2010). Na+, K+-ATPase mutations causing familial hemiplegic migraines type 2 (FHM2) inhibit phosphorylation (Schack et al., 2012). Salt, the vascular Na+/K+ ATPase and the endogenous glycosides, ouabain and marinobufagenin, play roles in systemic hypertension (Hauck and Frishman, 2012). Protein kinase A (PKA) phosphorylation of Ser936 (in the intracellular loop between transmembrane segments M8 and M9) opens an intracellular C-terminal water pathway leading to the third Na+-binding site (Poulsen et al., 2012). PKA-mediated phosphorylation regulates activity in vivo. Ser-938 is located (Einholm et al. 2016). E960 on the Na+-K+-ATPase and F28 on phospholemman (PLM) are critical for phospholemman (PLM) inhibition, but there is at least one additional site that is important for tethering PLM to the ATPase. Mutations in the Na+/K+-ATPase α3 subunit gene (ATP1A3) cause rapid-onset dystonia-parkinsonism, a rare movement disorder characterized by sudden onset of dystonic spasms and slow movements (Doğanli et al. 2013).  The 3-d strcuture of the Na+-bound Na+,K+-ATPase at 4.3 Å resolution reveals the positions of the three Na+ ions (Nyblom et al. 2013).  Mutations cause adrenal hypertension (Kopec et al. 2014) as well as alternating hemiplegia of childhood (AHC) and rapid-onset dystonia- parkinsonism (RDP) (Rosewich et al. 2014).  Differences in the structures of the ouabain-, digonxin- and bufalin-bound enzyme have been reported (Laursen et al. 2015).  ATPase inhibitors have been shown to be effective anti-cancer agents (Alevizopoulos et al. 2014). Cys45 in the β-subunit can be glutathionylated, regulating the activity of the enzyme (Garcia et al. 2015). ATP1A2 mutations play a role in migraine headaches (Friedrich et al. 2016). The beta2 subunit is essential for motor physiology in mammals, and in contrast to beta1 and beta3, beta2 stabilizes the Na+-occluded E1P state relative to the outward-open E2P state (Hilbers et al. 2016). Numerous transcription factors, hormones, growth factors, lipids, and extracellular stimuli as well as epigenetic signals modulate the transcription of Na,K-ATPase subunits (Li and Langhans 2015). Čechová et al. 2016 have identified two cytoplasmic pathways along the pairs of TMSs, TMS3/TMS7 or TM6S/TMS9 that allow hydration of the cation binding sites or transport of cations from/to the bulk medium. Dissipation of the transmembrane gradient of K+ and Na+ due to ouabain inhibition increases Ptgs2 and Nr4a1 transcription by increasing Ca2+ influx through L-type Ca2+ channels that, in turn, leads to CaMKII-mediated phosphorylation of CREB and calcineurin-mediated dephosphorylation of NFAT, respectively (Smolyaninova et al. 2017). ZMay play a role in the development of gastric adenocarcinomas (Wang et al. 2017). Mutations F785L and T618M give rise to familial rapid onset dystonia parkonsonism by distinct mechanisms (Rodacker et al. 2006). Reacts with methylglyoxal to inhibit its activity (Svrckova et al. 2017).  Accumulation of beta-amyloid (Abeta) at the early stages of Alzheimer's disease is accompanied by reduction of Na,K-ATPase functional activity. Petrushanko et al. 2016 showed that monomeric Abeta(1-42) forms a tight (Kd of 3 mμM), enthalpy-driven equimolar complex with alpha1beta1 Na,K-ATPase. Complex formation results in dose-dependent inhibition of the enzyme hydrolytic activity. The binding site of Abeta(1-42) is localized in the "gap" between the α- and β-subunits of Na,K-ATPase, disrupting the enzyme functionality by preventing the subunits from shifting towards each other. Interaction of Na,K-ATPase with exogenous Abeta(1-42) leads to a pronounced decrease of the enzyme transport and hydrolytic activities and Src-kinase activation in neuroblastoma cells SH-SY5Y. This interaction allows regulation of Na,K-ATPase activity by short-term increases in the Abeta(1-42) level (Petrushanko et al. 2016). Two distinct phospholipids bind to two distinct sites on the ATPase, affecting activity and stability (Habeck et al. 2017). Five cysteinyl residues (C452, C456, C457, C577, and C656) serve as the cisplatin binding sites on the cytoplasmic loop connecting transmembrane helices 4 and 5 (Šeflová et al. 2018). Mutations can cause F/SHM with moderate penitrance (Prontera et al. 2018).  Arginine substitution of a cysteine in transmembrane helix M8 converts the Na+,K+-ATPase to an electroneutral pump similar to the gastric H+,K+-ATPase (Holm et al. 2017). Early onset life-threatening epilepsy can be associated with ATP1A3 gene variants (Ishihara et al. 2019), and loss of Na/K pump function is the common feature of mutants that induce hyperaldosteronism (Meyer et al. 2019). Three Na+ sites are defined in the Na+-bound crystal structure of the Na+, K+-ATPase. Sites I and II overlap with two K+ sites in the K+-bound structure, whereas site III is unique and Na+ specific. A glutamine in transmembrane helix M8 (Q925) appears from the crystal structures to coordinate Na+ at site III, but does not contribute to K+ coordination at sites I and II. Nielsen et al. 2019 addressed the functional role of Q925 in the various conformational states of this-ATPase by examining the mutants Q925A/G/E/N/L/I/Y both enzymatically and electrophysiologically, thereby revealing their Na+ and K+ binding properties. Q925 substitutions had minor effects on Na+ binding from the intracellular side of the membrane, but mutations Q925A and Q925G increased the apparent Na+ affinity, but caused dramatic reductions of the binding of K+ as well as Na+ from the extracellular side of the membrane. Thus, an interaction between sites III and I and a possible gating function of Q925 in the release of Na+ at the extracellular side are supported (Nielsen et al. 2019). The alpha2 Na+/K+-ATPase isoform mediates LPS-induced neuroinflammation (Leite et al. 2020). Apical periodontitis induces changes on oxidative stress parameters and increases Na+/K+-ATPase activity in adult rats (Barcelos et al. 2020). Familial hemiplegic migraine type 2 can be due to a missense mutation (L425H) in ATP1A2 (Antonaci et al. 2021). Mutations in ATP1A3 and FXYD genes (TC# 1.A.27) can cause childhood-onset schizophrenia (Chaumette et al. 2020). The ATPase plays a role in cell adhesion, motility, and migration of cancer cells (Silva et al. 2021). TMS2 moves outward as Na+ is deoccluded from the E1 conformation (Young and Artigas 2021). Kinetic properties and crystal structures of the Na+,K+-ATPase in complex with cardiotonic steroids (CTS) has revealed differences between CTS subfamilies. Ladefoged et al. 2021 found beneficial effects of K+ on bufadienolide binding in contrast to the well-known antagonism between K+ and cardenolides. Bufadienolide binding is affected by (i) electrostatic attraction of the lactone ring by a cation and (ii) the ability of a cation to stabilize and "shape" the site constituted by transmembrane helices of the alpha-subunit (αM1-6). The latter effect was due to varying coordination patterns involving amino acid residues from helix bundles αM1-4 and αM5-10. Substituents on the steroid core of a bufadienolide add to and modify the cation effects (Ladefoged et al. 2021). The Na+-K+-ATPase functions in the developing hippocampus (Shao et al. 2021). CryoEM analyses of the Na+,K+-ATPase in the two E2P states with and without cardiotonic steroids has revealed mechanistic details (Kanai et al. 2022). A conserved ion-binding site tyrosine plays a role in ion selectivity of the Na+/K+ pump (Spontarelli et al. 2022). Essential roles of the Na+K+-ATPase in ischemic pathology provide a platform for the improvement in clinical research on ischemic stroke (Zhu et al. 2022). In intact live cells, the regulatory complex is composed of two alpha subunits associated with two beta subunits, decorated with two PLM regulatory subunits. Docking and molecular dynamics (MD) simulations generated a structural model of the complex. alpha-alpha subunit interactions support conformational coupling of the catalytic subunits, which may enhance the NKA turnover rate (Seflova et al. 2022).

 

Animals

3 component systems:
Na+-, K+-ATPase from α, β, γ heterotrimer of Homo sapiens
α1 (ATP1A1) (P05023)
α2 (ATP1A2) (P50993)
α3 (ATP1A3) (P13637)
β1 (ATP1B1) (P05026)
β2 (ATP1B2) (Q58I19)
β3 (ATP1B3) (P54709)
γ1 (ATP1G1) (P54710)

 
3.A.3.1.10

Putative archaeal Na+, K+ ATPase, Mac8 (encoded with methylcobalamin: coenzyme M methyltransferase; methanol-specific, a metal chaparone protein and an electron transfer protein) (Chan et al., 2010).

Archaea

Putative Na+/K+ ATPase of Methanosarcina acetivorans (Q8THY0)

 
3.A.3.1.11

Na+,K+-ATPase α2 subunit, ATP1a2a or ATPA2A. Deficiency causes brain ventricle dilation and embryonic motility in zebra fish. Is essential for skeletal and heart muscle function (Doganli et al. 2012).

Animals

ATPA2 of Danio rerio (Q90X34)

 
3.A.3.1.12

Na+,K+-ATPase subunits α (837 aas) and β (302 aas) of the blood fluke ().

Animals (Platyhelminthes)

Na+,K+-ATPase subunits α and β of Schistosoma mansoni
alpha, G4VGA0
beta, G4VTH6

 
3.A.3.1.13

H+/K+-ATPase, ATP12A or ATP1AL1 of 1039 aas.  Plays a role in myocardial relaxation (Knez et al. 2014), but also functions in airway surface liquid acidification which impaires airway host defenses in cells lacking or compromised for CFTR (3.A.1.202.1) (Shah et al. 2016). This non-gastric H+/K+ ATPase (ATP12A) is expressed in mammalian spermatozoa (Favia et al. 2022).

Animals

ATP12A of Homo sapiens

 
3.A.3.1.14

Na+/K+-ATPase of 1227 aas and 10 TMSs. Involved in cell signaling, volume regulation, and maintenance of electrochemical gradients (Morrill et al. 2016).

ATPase of Paramecium tetraurelia

 
3.A.3.1.15

Silkworm nerve Na+,K+-ATPase, α-subunit of 1009 aas and 10 TMSs (77% identical to the human ortholog). and the β-subunit of 326 aas and 1 TMS (30% identical to the human homolog). This ATPase, in contrast to mamalian ATPases, has high affinity for K+, but low affinity for Na+, suggesting that the β-subunit is responsible for the difference in Na+ affinity (Homareda et al. 2017).

Na+,K+-ATPase of Bombyx mori (domestic silkworm)

 
3.A.3.1.16

ATPase, Na+/K+ transporting subunit alpha 4, ATP1A4, of 1029 aas and 10 TMSs. Alternatively spliced transcript variants, encoding different isoforms, have been identified. ATP1A4 is exclusively expressed in germ cells and sperm and is essential for male fertility as it interacts with signaling molecules in both raft and non-raft fractions of the sperm plasma membrane to regulate capacitation-associated signaling, hyperactivation, sperm-oocyte interactions, and activation. ATP1A4 activity and expression increase during capacitation, challenging the widely accepted dogma of sperm translational quiescence (Tiwari et al. 2022).

ATP1A4 of Homo sapiens

 
3.A.3.1.2

H+-, K+-ATPase (gastric; H+ efflux; K+ uptake). Two H3O+ may be transported per ATP hydrolyzed.  Howeve, a cryo-electron microscope structure suggests that 1 H+ and 1 K+ are transporter per ATP hydrolyzed, providing the energy needed to generate the one million fold H+ concentration gradient effected by this enzyme (Abe et al. 2012).  The detailed mechanism has been discussed, and the roles of essential residues have been proposed (Shin et al. 2011).  A number of inhibitors of acid secretion have been identified, and these are of pharmacological importance (Shin et al. 2011). The catalytic alpha subunit has ten transmembrane segments with a cluster of intramembranal carboxylic amino acids located in the middle of TMSs 4, 5, 6 and 8. The beta subunit has one TMS with the N terminus in the cytoplasm. The extracellular domain of the beta subunit contains six or seven N-linked glycosylation sites. N-glycosylation is important for enzyme assembly, maturation and sorting (Shin et al. 2009). The cryo-EM structure with bound BYK99, a high-affinity member of K+-competitive, imidazo[1,2-a]pyridine inhibitors, has been solved (Abe et al. 2017).

Animals

Gastric H+-, K+-ATPase from Homo sapiens

 
3.A.3.1.3Na+-ATPase Marine algae Na+-ATPase (HANA) of Heterosigma akashiwo
 
3.A.3.1.4

Non-gastric H+-, K+- or NH4+-ATPase (Swarts et al., 2005; Worrell et al., 2008)

Animals

H+-, K+ or NH4+-ATPase of Rattus norvegicus (P54708)

 
3.A.3.1.5Putative spirochete Na+, K+-ATPase, Lbi6 (1046 aas) (K. Hak & M.H. Saier)BacteriaLbi6 of Leptospira biflexa (B0SMV3)
 
3.A.3.1.6

Spiny dogfish Na+,K+-ATPase (3-d structure solved at 2.4 Å resolution, Shinoda et al., 2009). The α-subunit is 88% identical to the human Na+,K+ ATPase (TC# 3.A.3.1.1).

Animals

Na+,K+-ATPase α, β, and γ subunits of Squalus acanthias
α (1028aas; Q4H132)
β (305aas; C4IX13)
γ (94aas; Q70Q12)

 
3.A.3.1.7

H+/K+-ATPase α-subunit (1534aas) (Ramos et al., 2011)

Fungi

H+/K+ ATPase of Aspergillus oryzae (Q2U3D2)

 
3.A.3.1.8

Putative Na+/K+-ATPase, Mhun_0636 (encoded in an operon with two half sized TrkA homologues, Mhun_0637 and Mhun_0638, that together may regulate the ATPase)

Archaea

Mhun_0636-8 of Methanospirillum hungatei
Mhun_0636 (Q2FLJ9) Mhun_0637 (Q2FLJ8) Mhun_0638 (Q2FLJ6)

 
3.A.3.1.9

Ouabain-insensitive K+-independent Na+-ATPase ɑ-subunit, AtnA; very similar to the human ɑ-1 chain of the Na+,K+-ATPase (3.A.3.1.1) (Rocafull et al., 2011).

Animals

AtnA of Cavia porcellus (B3SI05)

 
Examples:

TC#NameOrganismal TypeExample
3.A.3.10.1

P-type ATPase 13a1 of 1193 aas

Plants

ATPase 13a1 of Ricinus communis (Castor bean)

 
3.A.3.10.10

Putative Mn2+-exporting P-type ATPase of 1343 aas.

Stramenopiles

ATPase of Albugo laibachii

 
3.A.3.10.11

This protein was reviously designated the functionally uncharacterized P-type ATPase 16 (FUPA16)  (Thever and Saier 2009).  Probable manganese exporter by similarity.

Alveolata (ciliates)

Putative Mn2+ ATPase of Tetrahymena thermophila (Q23QW3)

 
3.A.3.10.12

P-type ATPase with N-terminal MACPF domain (TC# 1.C.39) of 1982 aas

Ciliates

MACPF-Mn2+ P-type ATPase of Tetrahymena thermophila

 
3.A.3.10.13

This protein was previously designated the functionally uncharacterized P-type ATPase 17 (FUPA17) (Thever and Saier 2009), but it has been shown to be a Ca2+/Mn2+-exporting ATPase designated Cation-transporting ATPase 5 (Cta5 or ATP13A2) (Furune et al. 2008).

Yeast

ATPase of Schizosaccharomyces pombe (O14022)

 
3.A.3.10.14

This protein was previously designated the functionally uncharacterized P-type ATPase 18 (FUPA18 of 1491 aas) (Thever and Saier 2009).  It may be a Mn2+-ATPase (by similarity).

Alveolata

FUPA18a of Cryptosporidium parvum (Q5CW06)

 
3.A.3.10.15

This protein was previously designated the functionally uncharacterized P-type ATPase 19 (FUPA19 of 1807 aas) (Thever and Saier 2009).  The unusually large size and number of TMSs is unique to this protein.  Whether this is a consequence of an artifact of sequencing is not known.  It may be a Mn2+-ATPase (by similarity).

 

Alveolata

ATPase of Tetrahymena thermophilus

 
3.A.3.10.16

This protein was previously designated the functionally uncharacterized P-type ATPase 20 (FUPA20) (Thever and Saier 2009).  It may be a Mn2+-exporting ATPase (by similarity).

Alveolata (ciliates)

ATPase of Tetrahymena thermophila (Q22V52)

 
3.A.3.10.17

This protein was previously designated the functionally uncharacterized P-type ATPase 21 (FUPA21 of 1372 aas) (Thever and Saier 2009).  It may be a Mn2+-ATPase (by similarity).

Protozoan

ATPase of Thalassiosira pseudonana

 
3.A.3.10.18

This protein was previously designated the functionally uncharacterized P-type ATPase 22 (FUPA22 of 1212-2393 aas) (Thever and Saier 2009).  It may be a Mn2+-exporting ATPase (by similarity).

Alveolata

ATPase of Cryptosporidium parvum (Q5CTJ9)

 
3.A.3.10.19

Mn2+-exporting ATPase, ATP13A1 of 1204 aas.  Defects cause Mn2+-dependent neurological disorders.  Orthologous to the yeast Mn2+-ATPase, Spf1 (Cohen et al. 2013). It is present in the endoplasmic reticulum while the other P5 ATPases, A2 - A5, are in overlapping compartments of the endosomal system (Sørensen et al. 2018). It complements the yeast ER ATPase, SPF1 (TC#3.A.3.10.3) although ATP13A2 - 5 do not, and unlike these latter proteins, it seems to have 12 (rather than 10) TMSs, with the two extra ones in an N-terminal domain (Sørensen et al. 2018). ATP13A1 (Spf1 in yeast) directly interacts with the TMSs of mitochondrial tail-anchored proteins (McKenna et al. 2020). P5A-ATPase mediates the extraction of mistargeted proteins from the endoplasmic reticulum (ER). Cryo-electron microscopy structures of Saccharomyces cerevisiae Spf1 (TC# 3.A.3.10.3) revealed a large, membrane-accessible substrate-binding pocket that alternately faced the ER lumen and cytosol and an endogenous substrate resembling an alpha-helical TMS. Thus, the P5A-ATPase can dislocate misinserted hydrophobic helices flanked by short basic segments from the ER. TMS dislocation by the P5A-ATPase establishes an additional class of P-type ATPase substrates and may correct mistakes in protein targeting or topogenesis (McKenna et al. 2020). It has been designated as a transmembrane islocase (Dederer and Lemberg 2021).

Animals

ATP13A1 of Homo sapiens

 
3.A.3.10.2

Zebrafish ATP13A2 (Parkinson''s disease protein) is essential for embryonic survival (Lopes da Fonseca et al. 2013). A missense variant in Australian Cattle Dogs give rise to late onset neuronal ceroid lipofuscinosis (Schmutz et al. 2019).

Fish

ATP13A2 of Danio rerio (Q7SXR0)

 
3.A.3.10.20

Probable divalent cation transporting ATPase 13A4, ATP13A4, of 1196 aas and 10 TMSs. This protein had been suggested to be a Mg2+ transporter, but the evidence is equivocal (Schäffers et al. 2018). It may be a Mn2+/Ca2+ exporter. This protein as well as ATP13A2 has been implicated in Parkinson's disease and autism spectrum disorder (Sørensen et al. 2018). ATPA2 - 5 are all in compartments of the endosomal system and all have 10 TMSs with overlapping functions, often in different amounts in different tissues (Sørensen et al. 2018).

ATP13A4 of Homo sapiens

 
3.A.3.10.21

Divalent cation transporting ATPase of 1207 aas and 9 putative TMSs, Catp-6.  C. elegans has three paralogues, Catp5, Catp6 and Catp7, with overlapping tissue expression patterns and functions (Zielich et al. 2018). 

Catp-5 of Caenorhabditis elegans

 
3.A.3.10.22

Manganese transporter of 1179 aas and 12 probable TMSs (Ticconi et al. 2004).  Mediates manganese transport into the endoplasmic reticulum. The ATPase activity is required for cellular manganese homeostasis. Plays an important role in pollen and root development through its impact on protein secretion and transport processes (Jakobsen et al. 2005). Functions together with LPR1 and LPR2 in a common pathway that adjusts root meristem activity to phosphate availability (Ticconi et al. 2009).

PDR2 of Arabidopsis thaliana (Mouse-ear cress)

 
3.A.3.10.3

The endoplasmic reticular ATPase, Spf1 or Cod1. Plays a role in ER Mn2+ homeostasis by pumping Mn2+ into the ER lumen (Cronin et al., 2002; Cohen et al. 2013). Deletion of the gene results in ER stress and lowered Mn2+ in the ER lumen (Cohen et al. 2013). This SPF1-ATPase is a transmembrane helix dislocase, used to remove some mislocated TM proteins from the outer membranes of mitochondria (McKenna et al. 2020). Spf1p exhibits unique structures at its N-terminus, including two putative additional transmembrane domains, and a large insertion connecting the P domain with transmembrane segment M5 (Petrovich et al. 2021). The Spf1p P5A-ATPase "arm-like" domain is not essential for ATP hydrolysis but its deletion impairs autophosphorylation (Grenon et al. 2021).

Fungi

Spf1 of Saccharomyces cerevisiae (P39986)

 
3.A.3.10.4

P-type ATPase of 1308 aas

Alveolata

ATPase of Babesia equi

 
3.A.3.10.5

P-type ATPase of 1291 aas

Alveolata

ATPase of Cryptosporidium parvum

 
3.A.3.10.6

Putative Mn2+-exporting P-type ATPase of 1146 aas.

Microsporidia

APase of Encephalitozoon cuniculi (Q8SRH4)

 
3.A.3.10.7

This protein, ATP13A2, was orginally designated the functionally uncharacterized P-type ATPase, FUPA13 (Thever and Saier 2009).  It is the Parkinson''s disease (PD) gene product, PARK9 (ATP13A2), and its defect gives rise to multiple abnormalities (Dehay et al. 2012).  It is similar to the probable manganese exporter in yeast, Ypk1 (TC# 3.A.3.10.8), and may have the same function, but in lysosomes, it is a polyamine (spermine/spermidine) exporter (van Veen et al. 2020). Toxic levels of manganese or abnormal levels of polyamines may cause a syndrome simiilar to PD (Chesi et al. 2012).  Manganese homeostasis in the nervous system has been reviewed (Chen et al. 2015).  The progression of PD may involve the lysosome and different autophagy pathways (Gan-Or et al. 2015).  It exhibits an activity-independent scaffolding role in trafficking/export of intracellular cargo in response to proteotoxic stress (Demirsoy et al. 2017). Mutations cause rare early onset Parkinson's disease (Suleiman et al. 2018). ATP13A2 modulates astrocyte-mediated neuroinflammation via NLRP3 inflammasome activation, thus bringing to light a direct link between astrocyte lysosomes and neuroinflammation in the pathological model of PD (Qiao et al. 2016). ATP13A2 and its close homologs, collectively known as P5B-ATPases, are polyamine transporters in endo-/lysosomes. Cryo-EM structures of human ATP13A2 in five distinct conformational intermediates, which together, represent a near-complete transport cycle of ATP13A2, have been determined. The structural basis of the polyamine specificity was revealed by an endogenous polyamine molecule bound to a narrow, elongated cavity within the transmembrane domain. The structures show an atypical transport path for a water-soluble substrate, in which polyamines may exit within the cytosolic leaflet of the membrane (Sim et al. 2021). Spermine is exported from the lysosome. The transmembrane domain serves as a substrate binding site, and the C-terminal domain is essential for protein stability and may play a regulatory role (Chen et al. 2021). The carcinogenic effects of ATP13A2 in different tumors has been studied (Zheng and Li 2021). High-resolution cryo-EM structures of human ATP13A2 in five distinct conformational intermediates have been determined, which together, represent a near-complete transport cycle of ATP13A2. The structural basis of the polyamine specificity was revealed by an endogenous polyamine molecule bound to a narrow, elongated cavity within the transmembrane domain. The structures show an atypical transport path for a water-soluble substrate, in which polyamines may exit within the cytosolic leaflet of the membrane (Sim et al. 2021). Mutations in ATP13A2 aare associated with mixed neurological presentations and iron toxicity due to nonsense-mediated decay (Kırımtay et al. 2021). The importance of the protein in regulating neuronal integrity has been established, and the structural dynamics and catalytic mechanism have been proposed (Mateeva et al. 2021).

Animals

PARK9 of Homo sapiens

 
3.A.3.10.8

This protein, P5B-ATPase, was originally designated the functionally uncharacterized P-type ATPase 14 (FUPA14) (Thever and Saier 2009), but it has been shown to be a vacuolar ATPase, Ypk1, that functions in manganese detoxification and homeostasis (Chesi et al. 2012).  It is therefore likely to catalyze export of manganese ions from the cytoplasm into the vacuole. However, it also transporter polyamines such as spermine . The 3-D structure has been determined to 3.4 Å, and it revealed three separate transport cycle intermediates, including spermine-bound conformations (Li et al. 2021). In the absence of cargo, Ypk9 rests in a phosphorylated conformation auto-inhibited by the N-terminus. Spermine uptake into vesicles is accomplished through an electronegative cleft lined by transmembrane segments 2, 4 and 6 (Li et al. 2021).

Fungi

Ypk1 of Saccharomyces cerevisiae (gi6324865)

 
3.A.3.10.9

This protein was previously designated the functionally uncharacterized P-type ATPase (FUPA15) (Thever and Saier 2009).  Probable manganese exporter by similarity (see 3.A.3.10.7 and 3.A.3.10.8).

Slime molds

Putative Mn2+-ATPase of Dictyostelium discoideum

 
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
3.A.3.2.1

Plasma membrane Ca2+-ATPase (efflux), PMCA4 (Giacomello et al. 2013).  The CD147 immunosupression protein interacts via its immunomodulatory domains with PMCA4 to bypass T-cell receptor proximal signaling and inhibit interleukin-2 (IL-2) expression (Supper et al. 2016). Deletion of residues 300 - 349, corresponding the the residues deleted in a natural splice variant (de Tezanos Pinto and Adamo 2006). The plasma membrane Ca2+ pump PMCA4z Is more active than splicing variant PMCA4x (Corradi et al. 2021).

Eukaryotes

Plasma membrane Ca2+-translocating ATPase, PMCA4, of Homo sapiens (P23634)

 
3.A.3.2.10The autoinhibited, calmodulin-binding Ca2+-ATPase, isoform 8, ACA8 (Baekgaard et al., 2006)PlantsACA8 of Arabidopsis thaliana (Q9LF79)
 
3.A.3.2.11Plastid Envelope Ca2+ ATPase, PEA1 (lacks a C-terminal calmodulin domain)PlantsPEA1 of Arabidopsis thaliana
(Q37145)
 
3.A.3.2.12

Endomembrane plasma membrane-type Ca2+ ATPase, ACA2 (Arabidopsis Ca2+ ATPase isoform 2) (lacks a C-terminal calmodulin domain, but activity is stimulated 5x by calmodulin which binds to an N-terminal inhibitory domain (Harper et al., 1998; Kamrul Huda et al. 2013).

Plants

ACA2 of Arabidopsis thaliana
(O81108)

 
3.A.3.2.13

Endoplasmic reticular (ER)-type Ca2+/Mn2+ ATPase, ECA1; 80% identical to and orthologous to the Medicago truncatula MCA8 protein of 1081 aas (F9W2W4). 42 P-type II Ca2+ ATPase genes have been found in Triticum aestivum. which may play roles in plant growth, development and signalling during abiotic and biotic stresses (Taneja and Upadhyay 2018).

Plants

ECA1 of Arabidopsis thaliana
(P92939)

 
3.A.3.2.14

Autoinhibited Ca2+ ATPase (ACA9) (expressed in pollen plasma membrane and required for male fertility), calmodulin-binding (Schiøtt et al., 2004).

Plants

ACA9 of Arabidopsis thaliana
(Q9LU41)

 
3.A.3.2.15Plasma membrane Ca2+ ATPase, Mca1 (Kraev et al., 1999)AnimalsMca1 of Caenorhabditis elegans
(O45215)
 
3.A.3.2.16Golgi Ca2+, Mn2+ ATPase, PMR1 (Van Baelen et al., 2001). (The human orthologue ATP2Cl, TC#3.A.3.2.5, causes Hailey-Hailey disease.)AnimalsPMR1 of Caenorhabditis elegans
(Q9XTG4)
 
3.A.3.2.17Intracellular (contractile vacuole) Ca2+ ATPase, PatA (lacks the C-terminal calmodulin domain of most plasma membrane Ca2+ ATPases) (Moniakis et al., 1995)Slime moldsPatA of Dictyostelium discoideum
(P54678)
 
3.A.3.2.18The acidocalcisome (vacuole) Ca2+/H+ ATPase TgA1 (involved in Ca2+ homeostasis, vacuolar polyphosphate storage and virulence) (Luo et al., 2005).ProtozoaTgA1 of Toxoplasma gondii
(Q9N694)
 
3.A.3.2.19

Endomembrane (Golgi) Ca2+/Mn2+-ATPase, ECA3 (one of 4 close paralogues in A. thaliana (Mills et al., 2008; Kamrul Huda et al. 2013)

Plants

ECA3 of Arabidopsis thaliana (Q0WP80)

 
3.A.3.2.2

Ca2+-ATPase, Pmc1 (uptake into vacuoles) (Espeso 2016).

Yeast

Vacuolar membrane Ca2+-translocating ATPase from Saccharomyces cerevisiae Pmc1

 
3.A.3.2.20Putative Ca2+ ATPase Cac1 (possible pseudogene?)

Firmicutes

Cac1 of Clostridium acetobutylicum (Q97JK5)

 
3.A.3.2.21Putative Ca2+ ATPase, Pmo1

Thermotogales

Pmo1 of Petrotoga mobilis (A9BJX0)

 
3.A.3.2.22Putative Ca2+ ATPase, Sth1

Firmicutes

Sth1 of Streptococcus thermophilus (Q5M0A4)

 
3.A.3.2.23

Putative Ca2+ ATPase most similar to Golgi Ca2+ ATPases of eukaryotes

Archaea

Putative Ca2+ ATPase of Methanococcus vannielii (A6URW9)

 
3.A.3.2.24

Putative Ca2+-ATPase (48% identical to 3.A.3.2.23) (like Golgi Ca2+-ATPases of eukaryotes)

Bacteria

Putative Ca2+-ATPase of Aguifex aeolicus (O66938)

 
3.A.3.2.25

Plasma membrane Ca2+-ATPase, isoform 1a (PMCA1) (78% identical to PMCA4 (TC# 3.A.3.2.1)). Maitotoxin converts it into a Ca2+-permeable nonselective cation channel (Sinkins et al., 2009). The C-terminal tail contains most of the regulatory sites including that for calmodulin. The pump is also regulated by acidic phospholipids, kinases, a dimerization process, and numerous protein interactors. In mammals, four genes code for the four basic isoforms. Isoform complexity is increased by alternative splicing of primary transcripts. Pumps 2 and 3 are expressed preferentially in the nervous system (Calì et al. 2017). This enzyme has two essential auxillary subunits, basigin and neuroplastin (NPTN), and the 3-d structure of the complex of PMCA1 with NPTN has been solved at 3.9 Å resolution (Gong et al. 2018). Methylene blue activates PMCA activity and cross-interacts with amyloid beta-peptide, blocking Abeta-mediated PMCA inhibition (Berrocal et al. 2018).

Animals

PMCA1 of Homo sapiens (P20020)

 
3.A.3.2.26

The M535L virus Ca2+/Mn2+ efflux pump (transcribed during viral infection) (Bonza et al., 2010)

Virus

M535L Ca2+ pump of Paramecium bursaria chlorella virus, MT325 (A7IUR5)

 
3.A.3.2.27

Plasma Membrane Ca2+-type ATPase, NCA-2 (most like 3.A.3.2.2) (Bowman et al., 2011).

Fungi

NCA-2 of Neurospora crassa (Q9UUY2)

 
3.A.3.2.28

The probable Mg2+/Ca2+ ATPase antiporter (catalyzes Mg2+ uptake and Ca2+ efflux in a single coupled step; Neef et al. 2011)

Bacteria

Antiporter of Streptococcus pneumoniae (Q04JJ5)

 
3.A.3.2.29

The putative Ca+ ATPase with an extra C-terminal TMS followed by a lysin (LysM) domain of ~210aas. LysM domains are often found in cell wall degradative enzymes and have peptidoglycan binding sites. Found in Nitrosococcus oceani as well as Nitrosococcus halophilus. The ATPase domain is 46% identical to 3.A.3.2.4.

Bacteria

Putative Ca2+ ATPase of Nitrosococcus  halophilus (D5C355)

 
3.A.3.2.3

Ca2+-ATPase, Pmr1 (efflux) (also transport Mn2+ and Cd2+) (Lauer et al., 2008)

Eukaryotes

Golgi Ca2+-ATPase Pmr1 of Saccharomyces cerevisiae

 
3.A.3.2.30

Pleasma membrane Ca2+-ATPase of parenchymal tissue of the liver fluke, PMCA.  Interacts with a calmodulin-like protein, FhCaM1 in a calcium ion dependent fashion (Moore et al. 2012).

Animals

PMCA of Fasciola helpatica

 
3.A.3.2.31

Sarcoplasmic reticulum Ca2+ ATPase, Atp6.  The inhibitors, artemisinin and its anti-malarial derivatives, artesunate and artemether, bind to a hydrophobic pocket in a transmembrane region near the membrane surface (Naik et al. 2011; Meier et al. 2018). Other inhibitors include arterolane and thapsigargin (Meier et al. 2018).

Alveolata

Atp6 of Plasmodium falciparum

 
3.A.3.2.32

Lobster intracellular SERCA Ca2+ ATPase of 1020 aas.  In related species, expression of the gene is increased under hypersaline conditions, and the enzyme is ivolved in salinity stress adaptation (Wang et al. 2013).

animals (Arthropods)

ATPase of Palinurus argus

 
3.A.3.2.33

Crustacian plasma membrane calcium ATPase of 1170 aas (Chen et al. 2013).

Animals

Calcium ATPase of Callinectes sapidus (blue crab)

   
 
3.A.3.2.34

Ca2+/Mn2+-exporting ATPase, Pmr1 of 899 aas (Furune et al. 2008). It plays a role in the control of cell division involving Mn2+ sensitivity and the Cwh43 protein (see TC# 9.B.131.1.9 for details) (Nakazawa et al. 2019).

Yeast

Pmr1 of Schizosaccharomyces pombe

 
3.A.3.2.35

Calcium-exporting ATPase, Pmc1 of 1096 aas (Furune et al. 2008)..

Yeast

Pmc1 of Schizosaccharomyces pombe

 
3.A.3.2.36

SERCA Ca2+-ATPase of 1093 aas (Docampo et al. 2013).

Alveolata

SERCA ATPase of Toxoplasma gondii

 
3.A.3.2.37

SERCA P-type ATPase of 1036 aas.

Alveolata (Ciliates)

SERCA ATPase of Paramecium tetraurelia

 
3.A.3.2.38

Plasma membrane Ca2+ ATPase (PMCA) of 1146 aas (Plattner 2014).

Alveolata

PMCA of Paramecium tetraurelia

 
3.A.3.2.39

Plasma membrae Ca2+ ATPase (PMCA) of 1064 aas (Lescasse et al. 2005).

Alveolata (ciliates)

PMCA of Oxytricha trifallax (Sterkiella histriomuscorum)

 
3.A.3.2.4

Ca2+-ATPase of 905 aas and 10 TMSs, Pma1

Bacteria

Putative Ca2+-ATPase of Synechocystis sp. pMA1

 
3.A.3.2.40

Plasma membrane Ca2+ ATPase, isoform 2, of 1243 aas, ATP2b2.  The mouse orthologue, of 1198 aas (P9R0I7), when mutated (I1023S in TMS 10 and R561S in the catailytic core) gives rise to semi-dominant hearing loss (Carpinelli et al. 2013). Neuroplastin (TC# 8.A.23.1.8) expression is essential for hearing and hair cell PMCA expression (Lin et al. 2021).

Animals

ATP2b2 of Homo sapiens

 
3.A.3.2.41
P-type Na+-ATPase of 889 aas (Takemura et al. 2009).

Bacteria (Firmicutes)

Na+-ATPase of Exiguobacterium aurantiacum
 
3.A.3.2.42

Plasma membrane Ca2+-ATPase of 1033 aas, ACA12.  Can replace ACA9 which is normally required for male fertility.  ACA12 is not stimulated by calmodulin (Limonta et al. 2014).

Plants

ACA12 of Arabidopsis thaliana

 
3.A.3.2.43

SERCA1 of 1001 aas.  Several 3-D structures are known (e.g., 3W5B). One has an ATP analogue, a Mg2+ and two Ca2+ ions in the respective binding sites (Toyoshima and Mizutani 2004). In this state, the ATP reorganizes the three cytoplasmic domains (A, N and P), which are widely separated without nucleotide, by directly bridging the N and P domains. The structure of the P-domain itself is altered by the binding of the ATP analogue and Mg2+. As a result, the A-domain is tilted so that one of the TMSs moves to lock the cytoplasmic gate of the transmembrane Ca2+-binding sites. This appears to be the mechanism for occluding the bound Ca2+ ions, before releasing them into the lumen of the sarcoplasmic reticulum (Toyoshima and Mizutani 2004). Molecular dynamics simulations provided evidence for the role of the Mg2+ and K+ bound states in the transport mechanism (Espinoza-Fonseca et al. 2014).  Animal SERCAs are inhibited by three short single TMS membrane proteins, phospholamban (TC# 1.A.50.1), sarcolipin (1.A.50.2) and myoregulin (1.A.50.3), and the inhibitory actions of these peptides on SERCA are counteracted by a peptide called DWORF (Dwarf ORF) (Nelson et al. 2016; Anderson et al. 2015).  Norimatsu et al. 2017 have resolved the first layer of phospholipids surrounding the transmembrane helices. Phospholipids follow the movements of associated residues, causing local distortions and changes in thickness of the bilayer. The entire protein tilts during the reaction cycle, governed primarily by a belt of Trp residues, to minimize energy costs accompanying the large perpendicular movements of the transmembrane helices. A class of Arg residues extend their side chains through the cytoplasm to exploit phospholipids as anchors for conformational switching (Norimatsu et al. 2017). The human ortholog (O14983) is 96% identical to this rabbit enzyme.  These enzymes are inhibited by the fungal mycotoxins, cyclopiazonic acid and thapsigargin (Darby et al. 2016; Houdou et al. 2019). The human ortholog is 97% identical and is of the same length. The SERCA residue Glu340 mediates interdomain communication that guides Ca2+ transport (Geurts et al. 2020). The SERCA Ca2+-ATPase may serve as a calcium-sensitive membrane-endoskeleton sensor in the sarcoplasmic reticulum (Nakamura et al. 2021). The C-terminal proton release pathway is a functional element of SERCA which provides a mechanistic model for its operation in the catalytic cycle of the pump (Espinoza-Fonseca 2021).

Animals

SERCA of Oryctolagus cuniculus (rabbit)

 
3.A.3.2.44

Crayfish basolateral plasma membrane Ca2+-ATPase, PMCA, of 1190 aas (Wheatly et al. 2007). 80% identical to the human orthologue.

PMCA of Procambarus clarkii (Red swamp crayfish)

 
3.A.3.2.45

The calmodulin-sensitive plasma membrane Ca2+-ATPase (PMCA) of 1080 aas and 10 TMSs.  It has a non-canonical calmodulin (CaM) binding domain that contains a C-terminal 1-18 motif (Pérez-Gordones et al. 2017).

PMCA of Trypanosoma equiperdum

 
3.A.3.2.46

Ca2+-ATPase of 880 aas and 10 TMSs, Ca1.  Key intermediates have been identified; Ca2+ efflux is rate-limited by phosphoenzyme formation. The transport process involves reversible steps and an irreversible step that follows release of ADP and extracellular release of Ca2+ (Dyla et al. 2017).

Ca1 of Listeria monocytogenes

 
3.A.3.2.47

Putative Ca2+ P-type ATPase, TMEM94, of 1356 aas and 10 TMSs in the usual 2 + 2 + 6 TMS arrangement.  This protein is very distantly related to all other members of the 3.A.3.2 family within the P-type ATPase superfamily, and therefore may have a different or unique function (Zhang et al. 2018).

TMEM94 of Homo sapiens

 
3.A.3.2.48

Sarco/endoplasmic reticulum Ca2+ ATPase of 1018 aas and 10 TMSs (Roegner et al. 2018).

SARCA of Callinectes sapidus

 
3.A.3.2.49

Sarcoplasmic reticular Calcium-ATPase of 993 aas and 10 TMSs. Elongation and contraction of scallop sarcoplasmic reticulum (SR): ATP Stabilizes Ca2+-ATPase crystalline array elongation of SR vesicles (Nakamura et al. 2022).

Ca2+-ATPase of Mizuhopecten yessoensis (Yesso scallop)

 
3.A.3.2.5

The Golgi Ca2+, Mn2+-ATPase, hSPCA1, ATP2C1 or Hussy-28 (efflux) (the Hailey-Hailey disease protein). Involved in responses to Golgi stress, apoptosis and midgestational death (Okunade et al., 2007). SPCA1 transports Mn2+ from the cytosol into the Golgi. Increasing Golgi Mn2+ transport increased cell viability upon Mn2+ exposure, supporting a role in the management of Mn2+ -induced neurotoxicity (Mukhopadhyay and Linstedt, 2011). SPCA1 governs the stability of TMEM165 (TC# 2.A.106.2.2) in Hailey-Hailey disease (Roy et al. 2020).

Animals

hSPCA1 of Homo sapiens

 
3.A.3.2.50

Plasma membrane calcium-transporting ATPase, ATP2B3, of 1220 aas and 10 TMSs. This 3ATP-driven Ca2+ ion pump is involved in the maintaining basal intracellular Ca2+ levels at the presynaptic terminals (Calì et al. 2015; Tauber et al. 2016). It uses ATP as an energy source to transport cytosolic Ca2+ ions across the plasma membrane to the extracellular compartment, and
may counter-transport protons.

ATP2B3 of Homo sapiens

 
3.A.3.2.6Ca2+, Mn2+- ATPase (efflux) FungiPmr1 of Neurospora crassa
 
3.A.3.2.7

The sarco/endoplasmic reticulum Ca2+ -ATPase, SERCA2b or ATP2A2, is encoded by the ATP2A2 gene.  Mutatioins give rise to Darier''s disease; the spectrum of mutations have been related to patients' phenotypes (Ahn et al., 2003; Godic et al. 2010).  SERCA1 functions as a heat generator in mitochondria of brown adipose tissue (de Meis et al., 2006). It normally functions as a Ca2+:H+ antiporter (Karjalainen et al., 2007). Capsaicin converts SERCA to a Ca2+ non-transporting ATPase that generates heat, and is thus a natural drug that augments uncoupled SERCA, resulting in thermogenesis (Mahmmoud, 2008b). Oligomeric interactions of the N-terminus of sarcolipin with the Ca-ATPase have been documented (Autry et al., 2011), and these interactions also uncouple ATP hydrolysis from Ca2+ transport (Sahoo et al. 2015) resulting in thermogenesis.  TMS 11, absent in SERCA1a and SERCA2a, functions in regulation (Gorski et al. 2012). The bovine SERCA has also been crystallized (2.9 Å resolution; Sacchetto et al., 2012).  These enzymes are regulated differentially by phospholamban (PLN; 1.A.50.1.1) and sarcolipin (SLN; 1.A.50.2.1) as noted above (Gorski et al. 2013).  SERCA2 is regulated by TMEM64 (9.B.27.5.1), a 380 aa 6 TMS membrane protein of the DedA family (TC# 9.B.27) which regulates Ca2+ oscillations by direct interaction with CIRCA2, modulating its activity and influencing osteoblast differentiation (Kim et al. 2013).  Animal SERCAs are inhibited by three short single (C-terminal) TMS membrane proteins, phospholamban (TC# 1.A.50.1), sarcolipin (1.A.50.2) and myoregulin (1.A.50.3), and the inhibitory actions of these peptides on SERCA are counteracted by a peptide called DWORF (Dwarf ORF) (Nelson et al. 2016; Anderson et al. 2015). Small ankyrin 1 (sAnk1; TC#8.A.28.1.2) and sarcolipin (TC# 1.A.50.2.1) interact in their transmembrane domains to regulate SERCA (Desmond et al. 2017). Reduced SERCA function preferentially affects Wnt signaling by retaining E-cadherin in the endoplasmic reticulum and promotes apoptosis (Suisse and Treisman 2019). There is strong coupling between the chronological order of deprotonation, the entry of water molecules into the TM region, and the opening of the cytoplasmic gate. Deprotonation of E309 and E771 is sequential with E309 being the first to lose the proton. Deprotonation promotes the opening of the cytoplasmic gate but leads to a productive gating transition only if it occurs after the transmembrane domain has reached an intermediate conformation (Rui et al. 2018). Coordination at cation binding sites I and II is optimized for Ca2+ and to a lesser extent for Mg2+ and K+ (Sun et al. 2019). Methyglyoxal reacts with and inhibits SERCA (Zizkova et al. 2018). The phospholamban pentamer alters the function of SERCA (Glaves et al. 2019). TMS11 followed by the luminal tail is inhibitory. Inoue et al. 2019 determined the crystal structures of SERCA2b and its C-terminal splicing variant SERCA2a, both in the E1-2Ca2+-adenylylmethylenediphosphonate (AMPPCP) state. TMS11 is located adjacent to TMS10 and interacts weakly with a part of the L8/9 loop as well as the N-terminal end of TMS10, thereby inhibiting the SERCA2b catalytic cycle (Inoue et al. 2019). Accordingly, mutational disruption of the interactions between TMS11 and its neighboring residues caused SERCA2b to display SERCA2a-like ATPase activity. The authors proposed that TMS11 serves as a key modulator of SERCA2b activity by fine-tuning the intramolecular interactions with other transmembrane regions. Kabashima et al. 2020 revealed what ATP binding does to the Ca2+ pump and how nonproductive phosphoryl transfer is prevented in the absence of Ca2+. They reported that the A-domain takes an E1 position, and the N-domain occupies exactly the same position as that in the E1.ATP.2Ca2+ state relative to the P-domain. As a result, ATP is properly delivered to the phosphorylation site. Yet phosphoryl transfer never takes place without filling the two transmembrane Ca2+-binding sites. This explains what ATP binding does to SERCA, and how nonproductive phosphorylation is prevented in E2 (Kabashima et al. 2020). Nonannular lipid binding is not necessary for the stability of the E2 state but may become functionally significant during the E2-to-E1 transition (Espinoza-Fonseca 2019). Structural changes induced by the binding of rutin arachidonate to SERCA1a may shift proton balance near the titrable residues Glu771 and Glu309 into neutral species, hence preventing the binding of calcium ions to the transmembrane binding sites and thus affecting calcium homeostasis (Rodríguez and Májeková 2020). SERCA2a is a key protein in the Ca2+ cycle during heart failure (Zhihao et al. 2020). Covalent conjugates of fullerene derivatives with xanthene dyes inhibit SERCA (Tatyanenko et al. 2020). Autophosphorylation of the pump with two bound Ca2+ ions triggers a large conformational change that opens a gate on the luminal side of the membrane allowing the release of the ions. In response to phosphorylation, the cytoplasmic domains are partially reconfigured into an intermediate state on the path toward the E2 state with a closed luminal gate (Thirman et al. 2021). In preB cells, loss of SERCA2 leads to reduced V(D)J recombination kinetics due to diminished RAG-mediated DNA cleavage (Chen et al. 2021). A series of structural changes may accompany the ordered dissociation of ADP, the A-domain rotation, and the rearrangement of the transmembrane (TM) helices. The luminal gate then opens to release Ca2+ toward the SR lumen. Intermediate structures on the pathway are stabilized by transient sidechain interactions between the A- and P-domains. Lipid molecules between TM helices play a key role in the stabilization (Kobayashi et al. 2021). Structural bases for the conformational and functional regulation of human SERCA2b have been reported (Zhang and Inaba 2022).

	

Animals

SERCA2b of Homo sapiens (P16615)

 
3.A.3.2.8

Ca2+-ATPase (efflux) with broad Ca2+ dependence (3.2-320 μm).  Probably inhibited by cipargamin and SJ1733 (Meier et al. 2018).

Protozoa

PfATPase4 of Plasmodium falciparum

 
3.A.3.2.9Ca2+,Mn2+-ATPase, hSPCA2 (ATP2C2) (efflux). 64% identical to hSPCA1 (TC #3.A.3.2.5) but lower affinity for Ca2+ and more restricted tissue distribution (brain and testis); present in the trans-Golgi network. May function in Mn2+ detoxification (Xiang et al., 2005). AnimalshSPCA2 of Homo sapiens (NP_055676)
 
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
3.A.3.23.1

Functionally uncharacterized P-type ATPase family 23 (FUPA23) (8 proteins from Actinomycetes; 650-802 aas) (Chan et al. 2010).

Actinobacteria

FUPA23a of Streptomyces coelicolor (Q9KXM5)

 
3.A.3.23.2Functionally uncharacterized P-type ATPase family 23 (FUPA23.2) (5 proteins from Firmicutes (778-1056aas; 10TMSs; type 2)).FirmicutesFUPA23b of Enterococcus faecalis (Q835V4)
 
3.A.3.23.3Functionally uncharacterized P-type ATPase family 23 (FUPA23) (2 proteins from Cyanobacteria (826-831aas; 10+MSs, type 2))

Cyanobacteria

FUPA23c of Trichodesmium erythraeum (Q10YH7)

 
Examples:

TC#NameOrganismal TypeExample
3.A.3.24.1

Functionally uncharacterized P-type ATPase family 24 (FUPA24) (6 proteins of Actinomycetes; 760-1625 aas) (Chan et al. 2010).

Actinobacteria

FUPA24a of Mycobacterium bovis (Q7U2U7)

 
3.A.3.24.2

Functionally uncharacterized P-type ATPase family 24 (FUPA24) (1607aas); The first half is most like type I (Copper) ATPases, while the second half is most like type II ATPases (Ca2+).

Chloroflexi

FUPA24b of Thermomicrobium roseum (B9L3W5)

 
3.A.3.24.3

Functionally uncharacterized P-type ATPase family 24 (FUPA24) (1430aas)

δ-Proteobacteria

FUPA24c of Haliangium ochraceum (D0LKA4)

 
3.A.3.24.4

Functionally uncharacterized P-type ATPase family 24 (FUPA24) (1446aas)

γ-Proteobacteria

FUPA24d of Hahella chejuensis (ABC27339)

 
Examples:

TC#NameOrganismal TypeExample
3.A.3.25.1

Functionally uncharacterized P-type ATPase family 25 (FUPA25.1) (4 proteins from Actinomycetes; 645-776 aas) (Chan et al. 2010).

Actinobacteria

FUPA25a of Streptomyces coelicolor (Q9RJ01)

 
3.A.3.25.2Functionally uncharacterized P-type ATPase family 25 (FUPA25.2) (3 proteins from α- and β-proteobacteria; 617-759 aas). These proteins show greatest similarity with established families 5&6. Family 25 members have 6 TMSs and lack TMSs A&B. Some fairly close homologues have 7 TMSs.ProteobacteriaFUPA25b of Sinorhizobium meliloti (Q92Z60)
 
3.A.3.25.3Functionally uncharacterized P-type ATPase family 25 (FUPA25.3) (2 proteins from firmicutes; 601-623 aas; 7TMSs and an extra putative N-terminal TMS).FirmicutesFUPA25c of Enterococcus faecalis (Q830Z1)
 
3.A.3.25.4

P-type ATPase with a C-terminal hemeerythrin (Hr) domain (Traverso et al., 2010). The Hr domain binds two iron ions per monomer (a diiron center) and may provide a regulatory or more direct function in iron transport (Traverso et al., 2010).

Bacteria

P1B-5- ATPase of Acidothermus cellulolyticus (A0LQU2)

 
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
3.A.3.27.1

Functionally uncharacterized P-type ATPase family 27 (FUPA27) (multiple proteins from α-, β- and γ- proteobacteria; 817-851aas) (Chan et al. 2010).

Proteobacteria

FUPA27a of Neisseria meningitidis (Q9JZI0)

 
3.A.3.27.2

Functionally uncharacterized P-type ATPase family 27 (FUPA27), Lbi2 (

Spirochetes

FUPA27b of Leptospira biflexa (B0STR2)

 
3.A.3.27.3

Functionally uncharacterized ε-proteobacteria P-type ATPase

ε-proteobacteria

FUPA27c of Nitratiruptor sp. SB155-2 (A6Q500)

 
3.A.3.27.4

The Cu2+ - ATPase, CtpA. Required for assembly of periplasmic and membrane embedded copper-dependent oxidases, but not for copper tolerance (Hassani, et al. 2010). Possibly CtpA delivers Cu2+ directly to the enzymes in the membrane rather than catalyzing transmembrane transport: similar to (3.A.3.27.1).

Bacteria

CtpA of Rubrivivax gelatinosus (Q5GCB0)

 
3.A.3.27.5

Cu+ export ATPase, CopA2; provides copper for cytochrome oxidase assembly (González-Guerrero et al. 2010; Raimunda et al. 2013).

Proteobacteria

CopA2 of Pseudomonas aeruginosa

 
3.A.3.27.6

Functionally uncharacterized P-type ATPase family 29 (FUPA29) (1 protein from a δ-proteobacterium, 798 aas) (Chan et al. 2010).

Proteobacteria

FUPA29a of Bdellovibrio bacteriovorus (Q6MK07)

 
3.A.3.27.7

Functionally uncharacterized P-type ATPase family 29 (FUPA29) (2 proteins from flavobacteria; 792-795)

Bacteroidetes

FUPA29b of Flavobacterium johnsoniae (A5FGV9)

 
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
3.A.3.3.1H+-ATPase (efflux) Plants; fungi; protozoa; slime molds; archaea H+-ATPase, plasma membrane of Neurospora crassa
 
3.A.3.3.10

Plamsa membrane proton-pumping ATPase, Pma1, of 1003 aas and 10 putative TMSs in a 2 + 2 + 6 TMS arrangement.  Leptosphaeria maculans, lacking this enzyme, displays a total loss of pathogenicity towards its host plant (Brassica napus). The mutant is unable to germinate on the host leaf surface and is thus blocked at the pre-penetration stage. Reduction in Pma1 activity may disturb the electrochemical transmembrane gradient, thus leading to conidia defective in turgor pressure generation on the leaf surface. L. maculans possesses a second plasma membrane H+-ATPase-encoding gene, termed pma2 (Remy et al. 2008).

 

Pma1 of Leptosphaeria maculans

 
3.A.3.3.11

Probable H+ pumping P-type ATPase of 1068 aas and 10 TMSs, PMA1 (Shan et al. 2006). PnPMA1 is differentially expressed during pathogen asexual development with a more than 10-fold increase in expression in germinated cysts, the stage at which plant infection is initiated, compared to vegetative or sporulating hyphae or motile zoospores.  PnPMA1 contains all the catalytic domains characteristic of H+-ATPases but also has a distinct domain of approximately 155 amino acids that forms a putative cytoplasmic loop between transmembrane domains 8 and 9 (Shan et al. 2006).

PMA1 of Phytophthora nicotianae

 
3.A.3.3.12

ATPase-7, AHA7, of 961 aas and 10 TMSs. 73% identical to AHA2 with which it shares function.  AHA7 is autoinhibited by a sequence present in the extracellular loop between transmembrane segments 7 and 8. Autoinhibition of pump activity is regulated by extracellular pH, suggesting negative feedback regulation of AHA7 during establishment of an H+ gradient. Restriction of root hair elongation is dependent on both AHA2 and AHA7, with each having different roles in this process (Hoffmann et al. 2018).

AHA7 of Arabidopsis thaliana (Mouse-ear cress)

 
3.A.3.3.2H+ (in)/K+ (out) Mg2+-ATPase (antiporter) Protozoa H+/K+ antiport ATPase 1A of Leishmania donovani
 
3.A.3.3.3Mn2+/Cd2+-ATPase, MntA (Hao et al. 1999).

Bacteria

MntA of Lactobacillus plantarum

 
3.A.3.3.4Putative H+-ATPaseArchaeaAha1 (MJ1226) of Methanococcus jannaschii
 
3.A.3.3.5

Plasma membrane H+-ATPase, TbHA1 (912 aas) (3 isoforms are present in T. brucei) (Luo et al., 2006). This and another H+-ATPase, (UniProt acc # Q388Z3; 97% identical to TbHA1) have been found to be essential for bloodstream-form Trypanosoma brucei through a genome-wide RNAi screen (Schmidt et al. 2018).

Protozoan

TbHA1 of Trypanosoma brucei (AAP30857)

 
3.A.3.3.6

Plamsa membrane H+-ATPase, Pma1 (pumps protons out of the cell to generate a membrane potential and regulate cytosolic pH) (Liu et al., 2006; Petrov, 2009). TMSs 4,5,6 and 8 comprise the H+ pathway where essential and important residues have been identified (Miranda et al., 2010). Residues in the loop between TMSs 5 and 6 play roles in protein stability, function, and insertion (Petrov 2015).  Pma1 interacts with the plamsa membrane Cch1/Mid1 (1.A.1.11.10) to regulate its activity by influencing the membrane potential (Cho et al. 2016).  Asp739 and Arg811 are important residues for the biogenesis and function of the enzyme as H+ transport determinants (Petrov 2017). Pma1, is a P3A-type ATPase and the primary protein component of the membrane compartment of Pma1 (MCP). Like other plasma membrane H+-ATPases, Pma1 assembles and functions as a hexamer, a property unique to this subfamily of P-type ATPases. It has been unclear how Pma1 organizes the yeast membrane into MCP microdomains, or why it is that Pma1 needs to assemble into a hexamer to establish the membrane electrochemical proton gradient. Zhao et al. 2021 reported a high-resolution cryo-EM study of native Pma1 hexamers embedded in endogenous lipids. The Pma1 hexamer encircles a liquid-crystalline membrane domain composed of 57 ordered lipid molecules. The Pma1-encircled lipid patch structure likely serves as the building block of the MCP. At pH 7.4, the carboxyl-terminal regulatory α-helix binds to the phosphorylation domains of two neighboring Pma1 subunits, locking the hexamer in the autoinhibited state. The regulatory helix becomes disordered at lower pH, leading to activation of the Pma1 hexamer. The activation process is accompanied by a 6.7 A downward shift and a 40 degrees rotation of transmembrane helices 1 and 2 that line the proton translocation path. The conformational changes enabled the authors to propose a detailed mechanism for ATP-hydrolysis-driven proton pumping across the plasma membrane (Zhao et al. 2021).

Yeast

H+-ATPase of Saccharomyces cerevisiae (P05030)

 
3.A.3.3.7

Plasma membrane H+ ATPase, AHA1 Three isoforms, AHA1, 2 & 3, exhibit different kinetic properties (Palmgren and Christensen, 1994). Both the N- and C-termini are directly involved in controlling the pump activity (Ekberg et al., 2010). Methyl jasmonate elicits stomatal closure in many plant species including A. thaliana, and stomatal closure is accompanied by large ion fluxes across the plasma membrane.  These events appear to be mediated by AHA1 (Yan et al. 2015). It is involved in root nutrient uptake, epidermal stomatal opening, phloem sucrose loading and unloading, and hypocotyl cell elongation (Ding et al. 2021). Auxin activates two distinct, antagonistically acting signalling pathways that converge on rapid regulation of apoplastic pH, a causative determinant of growth. Cell surface-based TRANSMEMBRANE KINASE1 (TMK1) interacts with and mediates phosphorylation and activation of plasma membrane H+-ATPases for apoplast acidification, while intracellular canonical auxin signalling promotes net cellular H+ influx, causing apoplast alkalinization (Li et al. 2021).

Plants

AHA1 of Arabidopsis thaliana
(P20649)

 
3.A.3.3.8

Plasma membrane H+ ATPase, AHA6 (binds 14-3-3 proteins induced by phosphorylation of Thr948, causing activation; preferentially expressed in pollen; Bock et al., 2006) (82% identical to 3.A.3.3.7).

Plants

AHA6 of Arabidopsis thaliana (Q9SH76)

 
3.A.3.3.9

Proton pumping ATPase, AHA2.  94% identical to AHA1 (3.A.3.3.7); generates the plasma membrane pmf.  Cation-binding pockets have been identified (Ekberg et al. 2010).  The pump has been reconstituted into "nanodiscs" in a functionally monomeric form (Justesen et al. 2013).  Regulated at the post-translation level by cis-acting auto-inhibitory domains, which can be relieved by protein kinase-mediated phosphorylation or binding of specific lipid species such as lysophospholipids (Wielandt et al. 2015).  Pumping is stochastically interrupted by long-lived (~100 seconds) inactive or leaky states. Allosteric regulation by pH gradients modulates the switch between these states but not the pumping or leakage rates (Veshaguri et al. 2016).  They dynamics of the pump have been examined (Guerra and Bondar 2015). AHA2 drives root cell expansion (Hoffmann et al. 2018). This protein is 81% identical to the barley (Hordeum vulgare) HA1 of 956 aas and 10 TMSs. Plasma membrane H+-ATPase (HA1 and HA2) activity and/or expression is important for regulating the activity of K+ transporters and channels under drought stress conditions (Cai et al. 2019). Herbivore exposure enhances A. nanus tolerance to salt stress by activating the jasmonic acid-signalling pathway, increasing plasma membrane H+-ATPase activity, promoting cytosolic Ca2+ accumulation, and then restricting K+ leakage and reducing Na+ accumulation in the cytosol (Chen et al. 2020).

Plants

Proton pumping ATPase of Arabidopsis thaliana

 
Examples:

TC#NameOrganismal TypeExample
3.A.3.30.1

Functionally uncharacterized P-type ATPase family 30 (FUPA30) (4 proteins from α-, β- and δ-proteobacteria; 825-896 aas) (Chan et al. 2010).

Proteobacteria

FUPA30a of Bdellovibrio bacteriovorus (Q6MPD9)

 
3.A.3.30.2Functionally uncharacterized P-type ATPase family 30 (FUPA30) (1 protein from Flavobacteria 838 aas)BacteroidetesFUPA30b of Flavobacterium johnsoniae (A5FBE4)
 
3.A.3.30.3Functionally uncharacterized P-type ATPase family 30 (FUPA30), Lbi5 (1 protein in spirochetes)SpirochetesFUPA30c of Leptospira biflexa (B0SLF7)
 
3.A.3.30.4Functionally uncharacterized P-type ATPase family 30 (FUPA30) (1 ptotein from cyanobacteria; 867 aas).

Cyanobacteria

FUPA30d of Anabaena variabilis (Q3M5P5)

 
Examples:

TC#NameOrganismal TypeExample
3.A.3.31.1

Functionally uncharacterized P-type ATPase family 31 (FUPA31) (3 proteins from γ-proteobacteria; 673-1068) (most closely related to FUPA32 homologues) (probably an active enzyme) (Chan et al. 2010).

Proteobacteria

FUPA31a of Methylococcus capsulatus (Q606V3)

 
3.A.3.31.2

Functionally uncharacterized P-type ATPase family 31 (FUPA31b) (probably a pseudogene). Bears a C-terminal domain of the EcsC family (see 3.A.1.143.1) not found in other P-type ATPases.

Proteobacteria

FUPA31b of Methylococcus capsulatus (Q606U9)

 
Examples:

TC#NameOrganismal TypeExample
3.A.3.32.1

Functionally uncharacterized P-type ATPase family 32 (FUPA32) (multiple proteins from α-, β-, γ-, δ- and ε-proteobacteria (690-720 aas) (Chan et al. 2010).

Proteobacteria

FUPA32a of Azoarcus sp. EbN1 (Q5P8C0)

 
3.A.3.32.2Probable heavy metal cation-transporting P-type ATPase, FUPA32.2 (718aas)ActinobacteriaFUPA32b of Mycobacterium bovis (P0A503)
 
3.A.3.32.3Functionally uncharacterized P-type ATPase family 32 (FUPA32) (many homologues in Firmicutes (704-730 aas))

Firmicutes

FUPA32c of Clostridium bartiettii (A6NST6)

 
3.A.3.32.4Functionally uncharacterized P-type ATPase family 32 (FUPA32) (3 proteins from Fusobacteria) (735 aas)FusobacteriaFUPA32d of Fusobacterium nucleatum (Q8REB9)
 
3.A.3.32.5Functionally uncharacterized P-type ATPase family 32 (FUPA32) (699 aas) (1 protein in Spirochetes)

Spirochetes

FUPA32e of Treponema denticola (Q73QH0)

 
3.A.3.32.6Functionally uncharacterized P-type ATPase family 32 (FUPA32) (2 proteins from Euryarchaeota)

Euryarchaeota

FUPA32f of Methanobrevibacter smithii (A5UJX0)

 
3.A.3.32.7Functionally uncharacterized P-type ATPase family 32 (FUPA32) (several proteins from Verrucomicrobia)

Verrucomicrobia

FUPA32g of Akkermansia muciniphila (B2UR24)

 
3.A.3.32.8Functionally uncharacterized P-type ATP family 32 (FUPA32) (several in cyanobacteria)

Cyanobacteria

FUPA32h of Thermosynechococcus elongatus (Q8DL41)

 
Examples:

TC#NameOrganismal TypeExample
3.A.3.4.1Mg2+/Ni2+-ATPase (uptake) Bacteria MgtA of Salmonella typhimurium
 
3.A.3.4.2Putative spirochete Mg2+-ATPase, Lin3 (843 aas)BacteriaLin3 of Leptospira interrogans (Q72RN5)
 
3.A.3.4.3

Mg2+ ATPase (1182 aas; 18-20 TMSs) with an N-terminal (residues 1-325) transmembrane domain of 8-10 TMSs; homologous to residues 493-791 in O53781 of Mycobacterium tuberculosis (TC# 2.A.1.3.43). Residues 257-318 hit TMSs 7 and 8 in FmtC (MrpF), TC#2.A.1.3.37 with a score of 8 e-4. The last 3 TMSs of the N-terminal fused domain of 3.A.3.4.3 and 3.A.3.4.4 are homologous (e-10) to the last 3 TMSs in 2.A.1.3.43. The N-terminal domain is homologous to the 8TMS domains of 9.B.3 family members.

Bacteria

Mg2+-ATPase of Pseudomonas stutzeri (F2N2Z6)

 
3.A.3.4.4

Mg2+ P-type ATPase (1195 aas; 18-20 TMSs) with an extra N-terminal 8-10 TMSs (residues 1-330). Similar to 3.A.3.4.3. The last 3 TMSs of the N-terminal fused domain to 3.A.3.4.3 and 3.A.3.4.4 are homologous (e-10) to the last 3 TMSs in 9.A.30.2.1. The N-terminal domain is homologous to the 8TMS domains of 9.B.3 family members.

Bacteria

Mg2+-ATPase with N-terminal 8-10 TMS domain of ~300 residues of Azotobacter vinelandii (C1DHA2)

 
3.A.3.4.5

Uncharacterized Mg2+-ATPase, MgtA, of 912 aas and 10 TMSs (Pohland and Schneider 2019).

MgtA of Microcystis aeruginosa

 
Examples:

TC#NameOrganismal TypeExample
3.A.3.5.1Cu2+-ATPase (uptake) Bacteria CopA of Enterococcus hirae
 
3.A.3.5.10

Cu+ (Km 0.3 μM), Ag+ transporting ATPase, CopB (Mana-Capelli et al., 2003)

Euryarchaea

CopB of Archaeoglobus fulgidus (AAB91079)

 
3.A.3.5.11

Chloroplast envelope Cu+-uptake ATPase, PAA1 or HMA1.  Essential for growth under adverse light conditions (Seigneurin-Berny et al. 2006).

Plants

PAA1 of Arabidopsis thaliana (Q9SZC9)

 
3.A.3.5.12

Chloroplast thylakoid Cu+-ATPase, PAA2/HMA8 (delivers Cu+ to the thylakoid lumen).  Degraded by the Clp protease undeer conditions of Cu+ excess (Tapken et al. 2014).

Plants

PAA2 of Arabidopsis thaliana (AAP55720)

 
3.A.3.5.13

The archaeal Cu+ efflux pump (CopA)

Archaea

CopA of Sulfolobus solfataricus (Q97UU7)

 
3.A.3.5.14The yeast Cd2+ efflux pump, PCA1 (Adle et al., 2007)YeastPCA1 of Saccharomyces cerevisiae (P38360)
 
3.A.3.5.15

The transferable, plasmid-localized, copper sensitivity (uptake) ATPase, TcrA (811aas) (46% identical to 3.A.3.5.1) (Hasman, 2005)

Bacteria

TcrA of Enterococcus faecium (ABA39707)

 
3.A.3.5.16

The transferable, plasmid-localized, copper resistance (efflux) ATPase, TcrB (50% identical to 3.A.3.5.2) (Hasman, 2005)

Bacteria

TcrB of Enterococcus faecium (AAL05407)

 
3.A.3.5.17

Golgi Cu2+ ATPase, Ccc2, retrieves Cu2+ from the metallochaperone Atx1 and transports it to the lumen of Golgi vesicles (Lowe et al., 2004)

Yeast

Ccc2 of Saccharomyces cerevisiae
(P38995)

 
3.A.3.5.18

The copper resistance ATPase, CopA (Ettema et al., 2006Lübben et al., 2007; Villafane et al., 2009).

Bacteria

CopA of Bacillus subtilis (O32220)

 
3.A.3.5.19The Cu2+, Fe3+, Pb2+ resistance efflux pump, CopA (induced by copper and to a lesser extent by Fe3+ and Pb2+) (Sitthisak et al., 2007)Gram-positive bacteriumCopA of Staphylococcus aureus (Q7A3E6)
 
3.A.3.5.2Cu+-, Ag+-ATPase (efflux) BacteriaCopB of Enterococcus hirae
 
3.A.3.5.20The gold (Au2+) resistance ATPase, GolT (regulated by GolS in response to Au2+; it may function with a cytoplasmic metal binding protein, GolB (AAL19308; Pontel et al., 2007).Bacteria GolT of Salmonella enterica (Q8ZRG7)
 
3.A.3.5.21The Cu+, Ag+-ATPase, CtrA2 (Chintalapati et al., 2008)Bacteria CtrA2 of Aquifex aeolicus (O67432)
 
3.A.3.5.22The Cu2+-ATPase, CtrA3 (Chintalapati et al., 2008)BacteriaCtrA3 of Aquifex aeolicus (O67203)
 
3.A.3.5.23Putative spirochete Cu+ ATPase (6 proteins in spirochetes)BacteriaLin1 of Leptospira interrogans (Q72N56)
 
3.A.3.5.24The putative copper ATPase, Sso1 (PacS)

Crenarchaeota

PacS of Sulfolobus solfataricus (Q97VH4)

 
3.A.3.5.25The putative copper ATPase, Pae1

Crenarchaeota

Pae1 of Pyrobaculum aerophilum (Q8ZUJ0)

 
3.A.3.5.26The putative copper ATPase, Tro1

Euryarchaeota

Tro1 of Thermoplasma volcanium (Q978Z8)

 
3.A.3.5.27

Putative Copper P-type ATPase (46% identical to 3.A.3.5.10)

Korarchaea

Putative Copper P-type ATPase of Candidatus Korarchaeum cryptofilum (B1L487)

 
3.A.3.5.28The putative copper ATPase, Ape2

Crenarchaeota

Ape2 of Aeropyrum pernix (Q9YBZ6)

 
3.A.3.5.29

The copper (Cu2+) transporting ATPase, Ccc2

Yeast

Ccc2 of Schizosaccharomyces pombe (O59666)

 
3.A.3.5.3

Cu+-, Ag+-ATPase (efflux from the cytosol into the secretory pathway) (Barnes et al., 2005); ATP7B (Wilson's disease protein, α-chain) is continuously expressed in Purkinje neurons. It delivers Cu+ to the ferroxidase, ceruloplasmin, in liver. May also transport Fe2+ (Takeda et al., 2005). Critical roles for the COOH terminus of ATP7B in protein stability, trans-Golgi network retention, copper sensing, and retrograde trafficking have been reported (Braiterman et al. 2011).  Modeling suggests that Cu+-binding sites HMBDs 5 and 6 are most important for function (Gourdon et al. 2012).  ATP7B loads Cu+ into newly synthesized cupro-enzymes in the trans-Golgi network and exports excess copper out of cells by trafficking from the Golgi to the plasma membrane.  Mutations causing disease can affect activity, stability or trafficking (Braiterman et al. 2014).  Cisplatin is a poor substrate relative to Cu+with a Km of 1 mμM, and copper and cisplatin compete with each other (Safaei et al. 2008). Veratridine can bind to a site at the mouth of the channel pore in the human cardiac sodium channel, NaV1.5 (Gulsevin et al. 2022). ATP7A/B contains a P-type ATPase core consisting of a membrane transport domain and three cytoplasmic domains, the A, P, and N domains, and a unique amino terminus comprising six consecutive metal-binding domains. Bitter et al. 2022 presented a cryo-EM structure of frog ATP7B in a copper-free state. Interacting with both the A and P domains, the metal-binding domains are poised to exert copper-dependent regulation of ATP hydrolysis coupled to transmembrane copper transport. A ring of negatively charged residues lines the cytoplasmic copper entrance that is presumably gated by a conserved basic residue sitting at the center. Within the membrane, a network of copper-coordinating ligands delineates a stepwise copper transport pathway.

Eukaryotes

Cu+-ATPase, ATP7B, of Homo sapiens

 
3.A.3.5.30

Copper (Cu+) exporting P-ATPase, CopA (3-D structure known to 3.2 Å; PDB# 3RFU; Gourdon et al. 2011).  The internal surface of the ATPase interacts with the copper chaparone, CopZ (Padilla-Benavides et al. 2012).  A sulfur-lined metal transport pathway has been identified (Mattle et al. 2015).  Cu+ is bound at a high-affinity transmembrane-binding site in trigonal-planar coordination with the Cys residues of the conserved CPC motif of transmembrane segment 4 (C382 and C384) and the conserved Methionine residue of transmembrane segment 6 (M717 of the MXXXS motif). These residues are also essential for transport (Mattle et al. 2015).

Bacteria

CopA of Legionella pneumophila (Q5X2N1)

 
3.A.3.5.31

Mycobacterial copper transporter, MctB (Wolschendorf et al., 2011).

Bacteria

MctB of Mycobacterium abscessus (B1MHH7)

 
3.A.3.5.32

Copper-transporting ATPase RAN1 or HMP7 (EC 3.6.3.4) (Protein HEAVY METAL ATPASE 7) (Protein RESPONSIVE TO ANTAGONIST 1). Receptors involved in ethylene signaling can acquire their copper load by different routes and adopt the metal ion from the plasma membrane either by sequential transfer from soluble chaperones of the ATX1-family via the ER-bound copper-transporting ATPase RAN1 or by direct transfer from the soluble chaperones (Hoppen and Groth 2020). ER-anchored SPL7 (Transcription factor) constitutes a cellular mechanism for the regulation of RAN1 in ethylene signaling and lays the foundation for investigating how Cu homeostatic conditions ethylene sensitivity in the developmental context (Yang et al. 2022).

 

Plants

RAN1 of Arabidopsis thaliana

 
3.A.3.5.33

Ca2+ exporting ATPase, CopA. The domain organization and mechanism have been studied (Hatori et al., 2009, Hatori et al., 2008, Hatori et al., 2007).  Residues involved in catalysis have been defined (Hatori et al. 2009).

Bacteria

CopA of Thermotoga martima (Q9WYF3)

 
3.A.3.5.34

Cu+ export ATPase, CopA1, required to maintain cytoplasmic copper levels (González-Guerrero et al. 2010; Raimunda et al. 2013).

Proteobacteria

CopA1 of Pseudomonas aeruginosa

 
3.A.3.5.35

Functionally uncharacterized P-type ATPase.  Three proteins from Corynebacteria of 841-976 aas are similar in sequence.  Formerly members of the FUPA26 family (Chan et al. 2010).

Actinobacteria

Uncharacterized ATPase of Corynebacterium diphtheriae (Q6NJJ6)

 
3.A.3.5.36

Functionally uncharacterized P-type ATPase, formerly of family 28 (FUPA28).  Two proteins in γ-proteobacteria are similar in sequence; of 847-852 aas (Chan et al. 2010).

Proteobacteria

P-type ATPase (formerly FUPA28a) of Legionella pneumophila (Q5ZYY0)

 
3.A.3.5.37

Copper exporting ATPase, ATP7 of 1254 aas and 10 - 12 TMSs.  DmATP7 is the sole Drosophila melanogaster ortholog of the human MNK and WND copper transporters. A regulatory element drives expression in all neuronal tissues examined and demonstrates copper-inducible, Mtf-1-dependent expression in the larval midgut. Thus, an important functional role for copper transport in neuronal tissues is implied. Regulation of DmATP7 expression is not used to limit copper absorption under toxic copper conditions. The protein localizes to the basolateral membrane of DmATP7 expressing midgut cells, supporting a role in export of copper from midgut cells (Burke et al. 2008).

ATP7 of Drosophila melanogaster (Fruit fly)

 
3.A.3.5.38

Cuprous ion (Cu+) exporter, CopB, of 785 aas and 8 TMSs in a 4 + 2 + 2 arrangement. The copper-transporting P1B-ATPases have been divided traditionally into two subfamilies, the P1B-1-ATPases or CopAs and the P1B-3-ATPases or CopBs. CopAs selectively export Cu+ whereas previous studies have suggested that CopBs are specific for Cu2+ export. Biochemical and spectroscopic characterization of Sphaerobacter thermophilus CopB (StCopB) showed that, while it does bind Cu2+, the binding site is not in the transmembrane domain (Purohit et al. 2018).  StCopB exhibits metal-stimulated ATPase activity in response to Cu+, but not Cu2+, indicating that it is actually a Cu+ transporter. Cu+ is coordinated by four sulfur ligands derived from conserved cysteine and methionine residues. The histidine-rich N-terminal region is required for maximal activity, but is inhibitory in the presence of divalent metal ions. P1B-1- and P1B-3-ATPases may therefore all transport Cu+ (Purohit et al. 2018).

CopB of Sphaerobacter thermophilus

 
3.A.3.5.39

Cu+, Zn2+, Cd2+ exporting ATPase of 815 aas and 8 TMSs, CueA. Has two N-terminal metal binding domains that are essential for resistance to these three metal ions (Liang et al. 2016).

CueA of Bradyrhizobium liaoningense

 
3.A.3.5.4Ag+-ATPase (efflux) Bacteria Ag+-ATPase, SilP of Salmonella typhimurium
 
3.A.3.5.40

Copper-exporting P-type ATPase of 742 aas and 8 TMSs (Singh et al. 2015).

CopA of Streptococcus mutans

 
3.A.3.5.41

Cuprous ion (Cu+) exporter, CtpA (does not export Co2+, Mn2+, Ni2+, Zn2+ or Cu2+).  Km for Cu+ = 0.05 μM (León-Torres et al. 2015). 

Cu+ ATPase of Mycobacterium tuberculosis

 
3.A.3.5.5

Cu+, Ag+-ATPase (efflux) (Fan and Rosen, 2002).  There are two metal binding domains (MBDs). The distal N-terminal MBD1 possesses a function analogous to the metallochaperones of related prokaryotic copper resistance systems and is involved in copper transfer to the membrane-integral ion binding sites of CopA. In contrast, the proximal domain MBD2 has a regulatory role by suppressing the catalytic activity of CopA in the absence of copper (Drees et al. 2015). The functions of Me2+ exporters are often supported by chaperone proteins, which scavenge the metal ions from the cytoplasm. A CopA chaperone is expressed in E. coli from the same gene that encodes the transporter (Meydan et al. 2017). Some ribosomes translating copA undergo programmed frameshifting, terminate translation in the -1 frame, and generate the 70 aa-long polypeptide CopA(Z), which helps cells survive toxic copper concentrations. The high efficiency of frameshifting is achieved by the combined stimulatory action of a "slippery" sequence, an mRNA pseudoknot, and the CopA nascent chain. Similar mRNA elements are not only found in the copA genes of other bacteria but are also present in ATP7B, the human homolog of copA, and direct ribosomal frameshifting in vivo (Meydan et al. 2017). Cu(i) (Cu+) pumps, of which CopA is an example, are primary-active electrogenic uniporters. The Cu+ translocation cycle does not require proton counter-transport, resulting in electrogenic generation of a transmembrane potential upon translocation of one Cu+ per ATP hydrolysis in the catalytic cycle (Abeyrathna et al. 2020).

Bacteria

CopA of E. coli

 
3.A.3.5.6

Cu+-ATPase, ATP7A (MNK or Mc1) (efflux from the cytosol into the secretory pathway) (Menkes disease protein, α-chain) (Tümer 2013). It plays a role in systemic copper absorption in the gut and copper reabsorption in the kidney. In nonpolarized cells, the metal binding sites in the amino-terminal domain of MNK are required for copper-regulated trafficking from the Golgi to the plasma membrane (Greenough et al. 2004). It is expressed in Purkinje cells early in development and later in Bergmann glia. In melanocytes, it delivers Cu2+ to tyrosinase (Barnes et al., 2005). ATP7A has dual functions: 1) it incorporates copper into copper-dependent enzymes; and 2) it maintains intracellular copper levels by removing excess copper from the cytosol. To accomplish both functions, the protein traffics between different cellular locations, depending on copper levels (Bertini and Rosato, 2008). The lumenal loop Met672-Pro707 of ATP7A binds metals and facilitates copper release from the intramembrane sites (Barry et al., 2011).  Modeling suggests that Cu+-binding sites HMBDs 5 and 6 are most important for function (Gourdon et al. 2012).  In addition to X-linked recessive Menkes disease, mutations cause occipital horn syndrome and adult-onset distal motor neuropathy (Yi and Kaler 2014). p97/VCP interacts with ATP7A playing a role in motor neuron degeneration (Yi and Kaler 2018). 55 different mutations were located around the six copper binding sites and the ATP binding site. 76.7% of the mothers were carriers. Approximately half of the male siblings of patients with MNK were diagnosed with MNK (Fujisawa et al. 2019). It may play a role in melanosome (melanocyte) function (Wiriyasermkul et al. 2020).

Animals

ATP7A of Homo sapiens

 
3.A.3.5.7

Cu+-Ag+-ATPase (efflux), CopA of 804 aas. Exhibits maximal activity at 75˚C (Cattoni et al., 2007). The 3-D structure of the ATP-binding domain has been solved (2HC8_A) (functions with the Cu+ chaperone, CopZ; 130aas) (González-Guerrero and Argüello, 2008). This protein has both N- and C- terminal metal binding domains (MBDs). The N-MBD exhibits a conserved ferredoxin-like fold, binds metals to CXXC, and regulates turnover. The C-MBD interacts with the ATP-binding (ATPB) domain and the actuator (A) domain (Agarwal et al., 2010). Cysteine is a non-essential activator of CopA, interacting with the cytoplasmic side of the enzyme in the E1 form (Yang et al. 2007).

Euryarchaea

CopAZ of Archaeoglobus fulgidus:
CopA (PaeS) (O29777)
CopZ (2HU9_A; O29901)

 
3.A.3.5.8

Cu+ transporting ATPase (intracellular, in the trans-Golgi membrane), Ccc2

Yeast

Ccc2 of Candida albicans

 
3.A.3.5.9Cu+ transporting (copper detoxification) ATPase, Crp1YeastCrp1 of Candida albicans
 
Examples:

TC#NameOrganismal TypeExample
3.A.3.6.1

Zn2+-, Cd2+-, Pb2+-ATPase (efflux).  The enzyme from S. aureus strain 17810R, of 726 aas, functions as a Cd2+:H+ antiporter, using both the pmf and ATP hydrolysis to drive Cd2+ expulsion (Tynecka et al. 2016).

Bacteria; plants; fungi; protozoa

CadA of Staphylococcus aureus

 
3.A.3.6.10The Cd2+, Zn2+, Co2+ resistance ATPase, CadA (YvgW)BacteriaCadA of Bacillus subtilis (O32219)
 
3.A.3.6.11The Zn2+ efflux P-type ATPase, CadA1 (Leedjarv et al., 2007)ProteobacteriaCadA1 of Pseudomonas putida (Q88RT8)
 
3.A.3.6.12The Cd2+/Pb2+ resistance P-type ATPase, CadA2; induced by Zn2+, Cd2+, Pb2+, Ni2+, Co2+ and Hg2+ (Leedjarv et al., 2007)ProteobacteriaCadA2 of Pseudomonas putida (Q88CP1)
 
3.A.3.6.13

The heavy metal efflux pump, AztA (exports Zn2+, Cd2+, Pb2+; has two adjacent heavy metal binding domains (Liu et al., 2007)

Bacteria

AztA of Anabaena (Nostoc) sp. PCC7120 (Q8ZS90)

 
3.A.3.6.14The heavy metal (Zn2+, Cd2+) P-type ATPase, Smc04128 (Rossbach et al., 2008)BacteriaSmc04128 of Sinorhizobium meliloti (Q92T56)
 
3.A.3.6.15

The heavy metal transporter A (HmtA) mediates uptake of copper and zinc but not of silver, mercury, or cadmium (Lewinson et al., 2009).

Proteobacteria

HmtA of Pseudomonas aeruginosa (Q9I147)

 
3.A.3.6.16The putative heavy metal ATPase, Mac1

Euryarchaeota

Mac1 of Methanosarcina acetivorans (Q8TJZ4)

 
3.A.3.6.17

Cd2+-selective export ATPase, HMA3 (expressed in root cell tonoplasts wherein Cd2+ is sequestered (Ueno et al., 2010)). HMA3 may play a role in Cd2+ accumulation in rice (Cao et al. 2019).

Plants

HMA3 of Oryza sativa (Q8H384)

 
3.A.3.6.18

Cd2+/Zn2+ exporting ATPase, HMA4. (very similar to HMA3; TC# 3.A.3.6.7). Important for Zn2+ nutrition. Has a C-terminal domain containing 13 cysteine pairs and a terminal stretch of 11 histidines with a high affinity for Zn2+ and Cd2+ and a capacity to bind 10 Zn2+ ions per C-terminus (Baekgaard et al., 2010). The pathway of translocatioin through the protein has been investigated, and the demonstration that mutations affect Zn2+ and Cd2+ transport differentially has been reported (Lekeux et al. 2018).

Plants

HMA4 of Arabidopsis thaliana (O64474)

 
3.A.3.6.19

Ca2+/Zn2+ ATPase, OsHMA2 (Satoh-Nagasawa et al., 2012).

Plants

HMA2 of Oryza sativa (E7EC32)

 
3.A.3.6.2

Zn2+-, Cd2+-, Co2+-, Hg2+-, Ni2+-, Cu2+, Pb2+-ATPase (efflux), ZntA, of 732 aas and 8 TMSs (Hou and Mitra, 2003). The first four TMSs in ZntA and presumably other P1B-type ATPases play an important role in maintaining the correct dimer structure (Roberts et al. 2020).

Bacteria

ZntA of E. coli

 
3.A.3.6.20

Cadmium/zinc-transporting ATPase 4, HMA3

PlantsHMA3 of Arabidopsis thaliana
 
3.A.3.6.21

Cobalt ion exporting ATPase, slr0797 (Rutherford et al. 1999).

Cyanobacteria

Co-ATPase of Synechocystis PCC6803

 
3.A.3.6.22

Co2+-specific P1B-ATPase, CoaT (Zielazinski et al., 2012).

Bacteria

CoaT of Sulfitobacter sp. NAS-14.1 (A3T2G5)

 
3.A.3.6.23

Heavy metal (Pb2+, Cd2+, Zn2+) export ATPase of 970 aas, PbtA (Hložková et al. 2013; Suman et al. 2014)

Proteobacteria

PbtA of Achromobacter xylosoxidans

 
3.A.3.6.24

Fur-regulated virulence factor A of 626 aas, FrvA; suggested by the authors to be a heme exporter, but maybe more likely to be an iron exporter (McLaughlin et al. 2012).

Firmicutes

FrvA of Listeria monocytogenes

 
3.A.3.6.25

Cd2+/Zn2+/Co2+ export ATPase, ZntA, of 904 aas and 8 TMSs. Expression of the zntA gene is inducible by all three metal ions, with Cd2+ being the most potent, mediated by the MerR-like regulator, ZntR (Chaoprasid et al. 2015). zntA and zntR mutants were highly sensitive to CdCl2 and ZnCl2, and less sensitive to CoCl2. Inactivation of zntA increased the accumulation of intracellular cadmium and zinc and conferred hyper-resistance to H2O2. Thus, ZntA and its regulator, ZntR, are important for controlling zinc homeostasis and cadmium and cobalt detoxification. The loss of either the zntA or zntR gene did not affect the virulence of A. tumefaciens in Nicotiana benthamiana (Chaoprasid et al. 2015).

ZntA of Agrobacterium tumefaciens

 
3.A.3.6.26

Cadmium/zinc resistance efflux pump, CadA of 910 aas and 8 TMSs (Maynaud et al. 2014).

CadA of Mesorhizobium metallidurans

 
3.A.3.6.27

Transition metal efflux ATPase of 829 aas and 6 TMSs, CzcP.  Exports Zn2+, Cd2+ and Co2+ efficiently (Scherer and Nies 2009). The side chains of Met254, Cys476, and His807 contribute to Cd2+, Co2+, and Zn2+ binding and transport (Smith et al. 2017).

CzcP of Cupriavidus metallidurans (Ralstonia metallidurans)

 
3.A.3.6.3Cd2+-, Zn2+, Co2+-ATPase (efflux) Bacteria CadA (HP0791) of Helicobacter pylori
 
3.A.3.6.4

Pb2+-ATPase (efflux), PbrA.  Mediates resistance to Pb2+, Cd2+ and Zn2+.  Lead resistance is facilitated by the phosphatase, PbrB, possibly by allowing complexation of the Pb2+ by phosphate in the periplasm (Hynninen et al. 2009).

 

Bacteria

PbrA of Ralstonia metallidurans

 
3.A.3.6.5

Mono- and divalent heavy metal (Cu+, Ag+, Zn2+, Cd2+) ATPase, Bxa1. bxa1 gene expression is induced by all four heavy metal ions (Tong et al., 2002). The His-rich domain is essential for both monovalent (Ag+ and Cu+) and divalent ( Cd2+ and Zn2+) metal tolerance (Nakakihara et al. 2009).

Bacteria

Bxa1 ATPase of Oscillatoria brevis

 
3.A.3.6.6Chloroplast envelope Cu+-ATPase, HMA1 (Seigneurin-Berny et al., 2006). Transports many heavy metals (Zn2+, Cu2+, Cd2+, Co2+), increasing heavy metal tolerance. Also transports Ca2+ (Km=370nM) in a thapsigargin-sensitive fashion (Moreno et al, 2008). PlantsHMA1 of Arabidopsis thaliana
(Q9M3H5)
 
3.A.3.6.7The Zn2+ (and Cd2+)-ATPase, HMA2. HMA2 maintains metal homeostasis and has a long C-terminal sequence rich in Cys and His residues that binds Zn2+, Kd≈16 nM and regulates activity (Eren et al., 2006). PlantsHMA2 of Arabidopsis thaliana (Q9SZW4)
 
3.A.3.6.8

The Cd2+ resistance ATPase, CadA (Wu et al., 2006)

Bacteria

CadA of Listeria monocytogenes (Q60048)

 
3.A.3.6.9The Zn2+ uptake ATPase, ZosA (YkvW) (Gaballa and Helmann, 2002)BacteriaZosA of Bacillus subtilis (O31688)
 
Examples:

TC#NameOrganismal TypeExample
3.A.3.7.1

K+-ATPase (uptake), KdpFABC. (KdpA is homologous to other K+ transporters such as KcsA (1.A.1.1.1), KtrB (2.A.38.4.2 and 2.A.38.4.3), and HKT (2.A.38.3.1 and 2.A.38.3.2); KdpB is homologous to P-ATPase α-subunits; KdpC and KdpF may facilitate complex assembly and stabilize the complex (Bramkamp et al., 2007; Haupt et al., 2005; Greie and Altendorf, 2007; Irzik et al., 2011). The KdpFABC acts as a functional and structural dimer with the two KdpB subunits in direct contact, but the enzyme can dissociate to the monomer (Heitkamp et al., 2008). KdpF is part of and stabilizes the KdpABC complex (Gassel et al., 1999).  Transcription of the kdp operon is activated by the KdpDE sensor kinase/response regulator pair, and unphosphorylated IIANtr of the PTS (TC# 4.A) binds KdpD to stimulate its activity, thereby enhancing kdp operon expression (Lüttmann et al. 2009, Lüttmann et al. 2015). Transcriptional regulation of the Pseudomonas putida kdpFABC operon by the KdpDE sensor kinase/response regulator by direct interaction of IIANtr of the PTS with KdpD has also been studied (Wolf et al. 2015). The 2.9 Å X-ray structure of the complete Escherichia coli KdpFABC complex with a potassium ion within the selectivity filter of KdpA and a water molecule at a canonical cation site in the transmembrane domain of KdpB has been solved (Huang et al. 2017). The structure reveals two structural elements that appear to mediate the coupling between these two subunits: a protein-embedded tunnel runs between these potassium and water sites, and a helix controlling the cytoplasmic gate of KdpA is linked to the phosphorylation domain of KdpB. A mechanism that repurposes protein channel architecture for active transport across biomembranes was proposed (Huang et al. 2017). The cytoplasmic C-terminal domain of KdpD functions as a K+ sensor (Rothenbücher et al. 2006). Serine phosphorylated KdpB is trapped in a conformation where the ion-binding site is hydrated via an intracellular pathway between TMSs M1 and M2 which opens in response to the rearrangement of cytoplasmic domains, resulting from phosphorylation (Dubey et al. 2021). This causes pump inhibition in the presence of high K+ resulting in ATP conservation.

	

Bacteria; proteobacteria

KdpABCF of E. coli
KdpA (P03959)
KdpB (P03960)
KdpC (P03961)
KdpF (P36937)

 
3.A.3.7.2

High affinity potassium uptake ATPase, KdpABC.  Regulated by direct interaction of the IIANtr protein with the sensor kinase/response regulator, KdpDE (Prell et al. 2012).

Proteobacteria

KdpABC of Rhizobium leguminosarum

 
3.A.3.7.3

Potassium transporter, KdpABC, with 3 subunits:  KdpA, B2HPR5, 552 aas and 10 TMSs in a 2 + 2 + 2 + 2 + 2 arrangement; KdpB ATPase, B2HRP6, 693 aas and 7  TMSs in a 2 + 2 + 3 TMS arrangement, and KdpC, D2HRP7, 296 aas with one N-terminal TMS and possibly one C-terminal TMS.  A kdpA null mutantion reduced the fraction of persisters after exposure to rifampicin (Liu et al. 2020). kdpA encodes a transmembrane protein that is part of the Kdp-ATPase, an ATP-dependent high-affinity potassium (K+) transport system. kdpA expression is induced under low K+ conditions and is required for pH homeostasis and growth in media with low concentrations of K+. Inactivation of the Kdp system caused hyperpolarization of the membrane potential, increased the proton motive force (PMF) and elevated levels of intracellular ATP. The KdpA mutant phenotype could be complemented with a functional kdpA gene or supplementation with high K+ concentrations. Thus, the Kdp system is required for ATP homeostasis and persister formation. ATP-mediated regulation of persister formation may be a general mechanism in bacteria, and suggest that K+ transporters may play a role in the regulation of ATP levels and persistence (Liu et al. 2020).

KdpABC of Mycobacterium marinum

 
Examples:

TC#NameOrganismal TypeExample
3.A.3.8.1

Golgi Aminophospholipid (phosphatidyl serine and phosphatidyl ethanolamine) translocase (flipping from the exofacial to the cytosolic leaflet of membranes to generate phospholipid asymmetry), required for vesicle-mediated protein transport from the Golgi and endosomes. The system has been reconstituted after purification in proteoliposomes. It flips phosphatidyl serine but not phosphatidylcholine or sphinogomyelin (Zhou and Graham, 2009).  A unified mechanism of flipping for ABC and P-type ATPases has been proposed (López-Marqués et al. 2014).

Animals

ATPase II of Bos taurus

 
3.A.3.8.10

Lipid flippase, Apt1 (involved in stress tolerance and virulence). Deletion of Apt1 causes (1) altered actin distribution, (2) increased sensitivity to stress conditions (oxidative and nitrosative stress) and to trafficking inhibitors, such as brefeldin A and monensin, a reduction in exported acid phosphatase activity, and (3) hypersensitivity to the antifungal drugs amphotericin B, fluconazole, and cinnamycin (Hu and Kronstad, 2010).

Yeast

Apt1 of Cryptococcus neoformans (Q5KP96)

 
3.A.3.8.11

Phospholipid (e.g., cardiolipin) transporter, Atp8b1. A mutant version is associated with severe
pneumonia in humans and mice. It binds and internalizes cardiolipin from extracellular fluid via a basic residue-enriched motif. Administration of a peptide encompassing the cardiolipin binding motif or Atp8b1 gene transfer in mice lessens bacterium-induced lung injury and improves survival (Ray et al., 2010).  Mutations have been identified that give rise to progressive familial intrahepatic cholestasis (Stone et al. 2012).  This lipid flippase forms a heterodimer with CDC50A/Transmembrane protein 30A (TC# 8.A.27.1.5) and is essential for surface expressioin of the apical Na+-bile acid transporter, Slc10A2/ASBT (TC#2.A.28.1.2) (van der Mark et al. 2014).

Animals

Atp8b1 of Homo sapiens (O43520)
 

 
3.A.3.8.12

Probable phospholipid-transporting ATPase IF (EC 3.6.3.1) (ATPase IR) (ATPase class VI type 11B). Among the ATP10 and ATP11 proteins of P4-ATPases, ATP10A, ATP10D, ATP11A, and ATP11C localize to the plasma membrane, while ATP10B and ATP11B localize to late endosomes and early/recycling endosomes, respectively. The N- or C-terminal cytoplasmic regions of P4-ATPases determine their cellular localization (Okamoto et al. 2020).

Animals

ATP11B of Homo sapiens

 
3.A.3.8.13

P4 phospholipid (phosphatidyl serine)-transporting ATPase 8A1 (EC 3.6.3.1) (ATPase class I type 8A member 1) (Chromaffin granule ATPase II).  Also found in the liver canicular membrane (Chaubey et al. 2016). The 3-D strcutures of 6 distinct intermediates (2.6 - 3.3 Å resolution) of the complex of this protein with CDC50A (TC# 8.A.27.1.5) have been solved, revealing the transport cycle for lipid flipping (Hiraizumi et al. 2019). ATP-dependent phosphorylation induces a large rotational movement of the actuator domain around the phosphorylation site in the phosphorylation domain, accompanied by lateral shifts of the first and second TMSs, thereby allowing phosphatidylserine binding. The phospholipid head group passes through the hydrophilic cleft, while the acyl chain is exposed toward the lipid environment (Hiraizumi et al. 2019).

Animals

ATP8A1 of Homo sapiens

 
3.A.3.8.14

ATP11C (ATPIG, ATPIQ) aminophospholipid (phosphatidyl serine and phosphatidyl ethanolamine, but not phosphatidyl choline) flippase of 1132 aas and 10 TMSs.  It is dependent on CDC50A (1232 aas and 9 TMSs, Anoctamin-8, Ano8; TC#1.A.17.1.30), for proper localization to the plasma membrane, and possibly also for activity (Segawa et al. 2014).  Present in liver basolateral membranes (Chaubey et al. 2016). It is the only phospholipid flipping ATPase in the human red blood cell (Liou et al. 2019). In the cell membrane of erythrocytes, it is required to maintain phosphatidylserine (PS) in the inner leaflet preventing its exposure on the surface. This asymmetric distribution is critical for the survival of erythrocytes in circulation since externalized PS is a phagocytic signal for splenic macrophages (Arashiki et al. 2016). Phospholipid translocation seems also to be implicated in vesicle formation and in the uptake of lipid signaling molecules, and is required for B cell differentiation past the pro-B cell stage It seems to mediate PS flipping in pro-B cells and may be involved in the transport of cholestatic bile acids.  Caspase-dependent inactivation of ATP11C is essential for an apoptotic "eat me" signal, phosphatidylserine exposure, which prompts phagocytes to engulf cells. Nakanishi et al. 2020 presented six cryo-EM structures of ATP11C at 3.0-4.0 A resolution in five different states of the transport cycle. A structural comparison revealed phosphorylation-driven domain movements coupled with phospholipid binding. Three structures of phospholipid-bound states visualize phospholipid translocation accompanied by the rearrangement of transmembrane helices and an unwound portion at the occlusion site. They thus detail the basis for head group recognition and the locality of the protein-bound acyl chains in transmembrane grooves. Invariant Lys880 and the surrounding hydrogen-bond network serve as a pivot point for helix bending and precise P domain inclination, which is crucial for dephosphorylation. The structures detail key features of phospholipid translocation by ATP11C; a common basic mechanism for flippases is emerging (Nakanishi et al. 2020).

 

Animals

ATP11C of Homo sapiens

 
3.A.3.8.15

Phospholipid transporting ATPase, Tat1 of 1139 aas.  Transports phosphatidylserine from the outer to the inner leaflet of the plasma membrane, thereby maintaining the enrichment of this phospholipid in the inner leaflet. Ectopic exposure of phosphatidylserine on the cell surface may result in removal of living cells by neighboring phagocytes in an apoptotic process (Darland-Ransom et al. 2008).  Tat1 regulates lysosome biogenesis and endocytosis as well as yolk uptake in oocytes. It is required at multiple steps of the endolysosomal pathway, at least in part by ensuring proper trafficking of cell-specific effector proteins (Ruaud et al. 2009). TAT-1 and its chaperone, the Cdc50 family protein CHAT-1, maintain membrane phosphatidylserine (PS) asymmetry, which is required for membrane tubulation during endocytic sorting and recycling. Loss of tat-1 and chat-1 disrupts endocytic sorting, leading to defects in both cargo recycling and degradation. Chen et al. 2019 identified the C. elegans aspartyl aminopeptidase DNPP-1, loss of which suppresses the sorting and recycling defects in tat-1 mutants without reversing the PS asymmetry defect.

Tat1 of Caenorhabditis elegans

 
3.A.3.8.16

ATP9A lipid flippase of 1047 aas and 10 TMSs.  Present in the liver canicular membrane (Chaubey et al. 2016).

ATP9A of Homo sapiens

 
3.A.3.8.17

Intracellular phospholipid flippase ATP11A (Chaubey et al. 2016).  Catalytic component of a P4-ATPase flippase complex which catalyzes the hydrolysis of ATP coupled to the transport of aminophospholipids from the outer to the inner leaflet and ensures the maintenance of asymmetric distribution of phospholipids. Phospholipid translocation seems also to be implicated in vesicle formation and in uptake of lipid signaling molecules. May be involved in the uptake of farnesyltransferase inhibitory drugs, such as lonafarnib (Zhang et al. 2005). A sublethal ATP11A mutation is associated with neurological deterioration due to aberrant phosphatidylcholine flipping in plasma membranes (Segawa et al. 2021).

 

ATP11A of Homo sapiens

 
3.A.3.8.18

The essential endosomal Neo1 phospholipid flipping ATPase of 1151 aas.  Neo1 plays an essential role in establishing phosphatidyl serine (PS) and phosphatidyl ethanolamine (PE) plasma membrane asymmetry in budding yeast (Takar et al. 2016). A common mechanism for substrate recognition in widely divergent P4-ATPases including Neo1 has been proposed (Huang et al. 2019).

Neo1 of Saccharomyces cerevisiae

 
3.A.3.8.19

The Leishmania miltefosine transporter (LMT) is a plasma membrane P4-ATPase that catalyses translocation into the parasite of the leishmanicidal drug, miltefosine as well as phosphatidylcholine and phosphatidylethanolamine analogues. Five highly-conserved amino acids in the cytosolic N-terminal tail (Asn58, Ile60, Lys64, Tyr65 and Phe70) and two (Pro72 and Phe79) in the first TMS were examined, and several of these were important for activity (Perandrés-López et al. 2018). The beta subunit of this system has TC# 8.A.27.1.3.

LMT of Leishmania amazonensis

 
3.A.3.8.2

Golgi aminophospholipid translocase (flipping from the exofacial to the cytosolic leaflet of membranes), required for vesicle-mediated protein transport from the Golgi and endosomal lumen to the cytoplasm, Gea2p (Pomorski et al., 2003). The system has been reconstituted after purification in proteoliposomes. It flips phosphatidyl serine and phosphatidyl ethanolamine, but not phosphatidylcholine or sphingomyelin (Zhou and Graham, 2009).  Drs2p (ACT3; ATP8A2) is required for phospholipid translocation across the Golgi membrane. It interacts with CDC50 (TC# 8.A.27) (Bryde et al., 2010). Activated by ArfGEF when bound to the C-terminus (Natarajan et al. 2009). The beta-subunit, CDC50A, allows the stable expression, assembly, subcellular localization, and lipid transport activity of the P4-ATPase ATP8A2 (Coleman and Molday, 2011). Timcenko et al. 2019 described the cryo-EM structure of Drs2p-Cdc50p, It is autoinhibited by the C-terminal tail of Drs2p and activated by the lipid phosphatidylinositol-4-phosphate (PI4P). Three structures were solved that represent the complex in an autoinhibited, an intermediate and a fully activated state. The analysis revealed sites of autoinhibition and PI4P-dependent activation. A putative lipid translocation pathway involves a conserved PISL motif in TMS 4 and polar residues of TMSs 2 and 5, in particular Lys1018, in the centre of the lipid bilayer (Timcenko et al. 2019). The enzymatic cycle of P-type ATPases is divided into autophosphorylation and dephosphorylation half-reactions. Unlike most other P-type ATPases, P4-ATPases transport their substrate during dephosphorylation only, i.e. the phosphorylation half-reaction is not associated with transport. To study the structural basis of the distinct mechanisms of P4-ATPases, Timcenko et al. 2021 determined cryo-EM structures of Drs2p-Cdc50p covering multiple intermediates of the cycle. They identified several structural motifs specific to Drs2p and P4-ATPases in general that decrease movements and flexibility of domains as compared to other P-type ATPases. These motifs include the linkers that connect the transmembrane region to the actuator (A) domain, which is responsible for dephosphorylation. Mutation of Tyr380, which interacts with conserved Asp340 of the distinct DGET dephosphorylation loop of P4-ATPases, highlights a functional role of these P4-ATPase specific motifs in the A-domain. Finally, the transmembrane (TM) domain, responsible for transport, also undergoes less extensive conformational changes, which is ensured both by a longer segment connecting TM helix 4 with the phosphorylation site, and possible stabilization by the auxiliary subunit Cdc50p. Collectively these adaptions in P4-ATPases are responsible for phosphorylation becoming transport-independent (Timcenko et al. 2021). The Arf activator, Gea2p (Uniprot P39993, 1459 aas), and Drs2p interact in the Golgi (Chantalat et al. 2004).


Fungi

DRS2 of Saccharomyces cerevisiae

 
3.A.3.8.20

Plasma membrane phospholipid flippase of 1656 aas, Dnf3-Crf1. Dnf3 flips phospholipids from the outer leaflet of the membrane to the inner leaflet (Sartorel et al. 2015). Crf1, a non-catalytic subunit, regulates the activity of Dnf3.  It is listed under TC# 8.A.27.1.7.

Dnf3/Crf1 of Saccharomyces cerevisiae

 
3.A.3.8.21

Putative lipid-flipping magnesium-transporting ATPase of 922 aas and 10 TMSs (Greiner et al. 2018).

ATPase of Klosneuvirus KNV1

 
3.A.3.8.22

Probable phospholipid-transporting P-type ATPase of 903 aas and  10 TMSs.

ATPase of soda lake Tupanvirus

 
3.A.3.8.23

Possible lipid flipping P-type ATPase of 809 aas and 7 putative TMSs.  It is probably C-terminally truncated.

ATPase of Catovirus CTV1

 
3.A.3.8.24

Broad range phospholipid-transporting ATPase 10, ALA10, of 1202 aas and 10 TMSs.  A structural model of ALA10 reveals a cavity delimited by TMSs 3, 4 and 5 at a similar position as the cation-binding region in related cation transporting P-type ATPases. Docking of a phosphatidylcholine headgroup in silico showed that the cavity can accommodate a phospholipid headgroup, likely leaving the fatty acid tails in contact with the hydrophobic portion of the lipid bilayer. Mutagenesis data supported this interpretation and suggested that two residues in TMS 4 (Y374 and F375) are important for coordination of the phospholipid headgroup (Jensen et al. 2017). These results point to a general mechanism of lipid translocation by P4 ATPases, which closely resembles that of cation-transporting pumps, through coordination of the hydrophilic portion of the substrate in a central membrane cavity.

 
3.A.3.8.25

Phospholipid-transporting ATPase VD or Atp10d, of 1426 aas and 10 TMSs.  It is expressed in placenta and, to a lesser extent, in the kidney (Flamant et al. 2003).  It is the catalytic component of a P4-ATPase flippase complex which catalyzes the hydrolysis of ATP coupled to the transport of aminophospholipids from the outer to the inner leaflet of various membranes and ensures the maintenance of asymmetric distribution of phospholipids. Phospholipid translocation has been implicated in vesicle formation and in the uptake of lipid signaling molecules. ATP10D reduces high-fat diet induced obesity and improves insulin sensitivity (Sigruener et al. 2017). It also transports glycosphingolipids including glucosphinolipids (Roland et al. 2019).

 

Atp10d of Homo sapiens

 
3.A.3.8.26

P-type ATPase of 1499 aas and 10 TMSs, Atp10A, Atp10C, AtpVA, AtpVC.  ATP10A transports phosphatidylcholine but not aminophospholipids (Shin and Takatsu 2019). Among the ATP10 and ATP11 proteins of P4-ATPases, ATP10A, ATP10D, ATP11A, and ATP11C localize to the plasma membrane, while ATP10B and ATP11B localize to late endosomes and early/recycling endosomes, respectively. The N- or C-terminal cytoplasmic regions of P4-ATPases determine their cellular localization (Okamoto et al. 2020).

Atp10A of Homo sapiens

 
3.A.3.8.3

Miltefosine/glycerophospholipid uptake translocase and phospholipid uptake flippase, MIL (Pérez-Victoria et al., 2003)

Protozoa

MIL of Leishmania donovani (Q6VXY9)

 
3.A.3.8.4

Inwardly directed phospholipid and lysophospholipid (phosphatidylcholine, phosphatidyl serine and lysophosphoethanolamine) flippase, Dnf1 or ATP11C (functions with the β-subunit, Lem3 or CDC50A (TC# 8.A.27.1.5) (Elvington et al., 2005; Pomorski et al., 2003; Riekhof and Voelker, 2006; Riekhof et al., 2007) Also transports the anti-neoplastic and anti-parasitic ether lipid substrates related to edelfosine (Riekhof and Voelker, 2009) (not required for phosphotidyl serine inwardly directed flipping (Stevens et al. 2008)). Transports diacyl phospholipids in preference to lyso (monoacyl) phospholipids (Baldridge et al. 2013).  A conserved asparagine (N220) in the first transmembrane segment specifies glycerophospholipid binding and transport, but specific substitutions at this site allow transport of sphingomyelin (Roland and Graham 2016). It transports glycosphingolipids (Roland et al. 2019). Nakanishi et al. 2020 presented the crystal structures of a human plasma membrane flippase, the ATP11C-CDC50A complex, in a stabilized E2P conformation. The structure revealed a deep longitudinal crevice along transmembrane helices continuing from the cell surface to the phospholipid occlusion site in the middle of the membrane. The extension of the crevice on the exoplasmic side was open, and the complex was therefore in an outward-open E2P state, similar to a cryo-EM structure of the yeast flippase Drs2p-Cdc50p complex. Phosphatidylserines were in the crevice and in its extension to the extracellular side. One was close to the phosphatidylserine occlusion site as previously reported for the human ATP8A1-CDC50A complex, and the other in a cavity at the surface of the exoplasmic leaflet of the bilayer. Substitutions in either of the binding sites or along the path between them impaired ATPase and transport activities. Thus, the crevice is the conduit along which phosphatidylserine traverses the membrane (Nakanishi et al. 2020).

Yeast

Dnf1 of Saccharomyces cerevisiae (P32660)

 
3.A.3.8.5

Inwardly directed phosphatidylcholine, phosphatidyl serine, and lysophosphoethanolamine flippase, Dnf2 (functions with the β-subunit, Lem3) (Elvington et al., 2005; Pomorski et al., 2003; Riekhof and Voelker, 2006; Riekhof et al., 2007). This plasma membrane P-type ATPase (ACT4) is a phospholipid flippase that contributes to endocytosis, protein transport and all polarity (Hua et al., 2002). Transports monoacyl (lyso) phospholipids much better than diacyl phospholipids, but can be mutated to transport diacyl phospholipids (Baldridge et al. 2013). It transports glycosphingolipids (Roland et al. 2019).

Yeast

Dnf2 of Saccharomyces cerevisiae (Q12675)

 
3.A.3.8.6Golgi phospholipid transporting (flipping) ATPase3 (1213aas; 10TMSs). Involved in growth of roots and shoots. Uses a β-ATPase3 subunit, ALIS1 (TC#8.A.27.4) (Paulsen et al., 2008).PlantsATPase3/ALIS1 of Arabidopsis thaliana (Q9XIE6)
 
3.A.3.8.7

The aminophospholipid ATPase1 (ALA1) (mediate chilling tolerance; Gomes et al., 2000).  Promotes antiviral silencing (Guo et al. 2017).

Plants

ALA1 of Arabidopsis thaliana (P98204)

 
3.A.3.8.8

The phosphatidylserine flippase in photoreceptor disc membranes, ATP8A2 (Coleman et al., 2009). The beta-subunit, CDC50A (TC#8.A.27.1.5), allows the stable expression, assembly, subcellular localization, and lipid transport activity of ATP8A2 (Coleman and Molday, 2011).  Missennse mutations in ATP8A2 are associated with cerebellar atrophy and guadrupedal locomotion (Emre Onat et al. 2012). Asparagine-905 of the mammalian phospholipid flippase ATP8A2 is essential for lipid substrate-induced activation of ATP8A2 dephosphorylation (Mikkelsen et al. 2019). Phosphatidylserine flipping by the P4-ATPase, ATP8A2, is electrogenic (Tadini-Buoninsegni et al. 2019).

Animals

ATP8A2 of Mus musculus (P98200)

 
3.A.3.8.9

The phospholipid flipping ATPase (contributes to vesicle biogenesis in the secretory and endocytic pathways). Forms heteromeric complexes with ALIS Cdc50-like β-subunits (ALIS1 = Q9LTW0; TC#8.A.27.1.4) promoting functionality (López-Marqués et al., 2010). The beta-subunit, CDC50A, allows the stable expression, assembly, subcellular localization, and lipid transport activity of the P4-ATPase ATP8A2 (Coleman and Molday, 2011). Promotes antiviral silencing (Guo et al. 2017).

Plants

Ala2 of Arabidopsis thaliana (P98205)

 
Examples:

TC#NameOrganismal TypeExample
3.A.3.9.1Na+-ATPase (efflux) Fungi and protozoaPmr2ap (ENa1) of Saccharomyces cerevisiae
 
3.A.3.9.2K+-ATPase (efflux) Fungi and protozoa Cta3 of Schizosaccharomyces pombe
 
3.A.3.9.3Monovalent alkali cation (Na+ and K+) ATPase (efflux of both cations)Fungi and protozoaENA2 of Debaryomyces occidentalis
 
3.A.3.9.4Na+ ATPase, ENA1 (Watanabe et al., 2002)FungiENA1 of Zygosaccharomyces rouxii (BAA11411)
 
3.A.3.9.5Plasma membrane K+ or Na+ efflux ATPase (required for growth at pH9, and for Na+ or K+ tolerance above pH8; Benito et al., 2009) (50% identical to 3.A.3.9.3).

Fungi

Ena1 of Ustilago maydis (B5B9V9)

 
3.A.3.9.6Endoplasmic reticulum K+ or Na+ efflux ATPase; confers Na+ resistance (Benito et al., 2009) (43% identical to 3.A.3.9.2).

Fungi

Ena2 of Ustilago maydis (Q4PI59)

 
3.A.3.9.7

P-type Ca2+ ATPase of 1041 aas and 12 TMSs.  Found to be essential for bloodstream-form Trypanosoma brucei through a genome-wide RNAi screen (Schmidt et al. 2018).

P-type Ca2+ ATPase of Trypanosoma brucei

 

 
3.A.3.9.8

Cation_ATPase_N domain-containing protein of 1084 aas and 10 TMSs in a 2 + 2 + 6 TMS arrangement. This P-type Na+/K+ ATPases essential and nonessential for cellular homeostasis and insect pathogenicity of Beauveria bassiana, respectivelly (Mou et al. 2020). Beauveria bassiana is an insect-pathogenic fungus serving as a main source of fungal insecticides worldwide. ENA1a and ENA2b are involved in both transmembrane and vacuolar activities and are essential for cellular cation homeostasis, insect pathogenicity and multiple stress tolerance in B. bassiana (Mou et al. 2020).

Na+-ATPase of Beauveria bassiana (White muscardine disease fungus) (Tritirachium shiotae)