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. Family 9 Na+- or K+-ATPases can be found in fungi, plants bryophytes and protozoa (Rodríguez-Navarro and Benito, 2010).

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.

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).

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.

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).

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).

The generalized reaction for P-type ATPases is:

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



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

 

References:

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Examples:

TC#NameOrganismal TypeExample
Examples:

TC#NameOrganismal TypeExample
3.A.3.1.1

Na+-, K+-ATPase (Na+ efflux; K+ uptake).  Mutations 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).

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

Na+/K+-ATPase, ATP12A or ATP1AL1 of 1039 aas.  Plays a role in myocardial relaxation (Knez et al. 2014).  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).

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.3.1.2

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). 

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).

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 was orginally designated the functionally uncharacterized P-type ATPase, FUPA13 (Thever and Saier 2009).  It is the Parkinson''s disease (PD) gene product, PARK9, also called 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. Toxic levels of manganese 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 also 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).

Animals

PARK9 of Homo sapiens

 
3.A.3.10.8

This protein 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 therefore is likely to catalyze export of manganese ions from the cytoplasm into the vacuole.

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:

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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).

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).

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).

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

SERCA of 1001 aas.  Several 3-D structures are known (e.g., 3W5B).  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).

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.3.2.5

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).

Animals

hSPCA1 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 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 (2.9Å 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 a 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).

	

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
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).

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).

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).

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) (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).

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.3.5.32Copper-transporting ATPase RAN1 (EC 3.6.3.4) (Protein HEAVY METAL ATPASE 7) (Protein RESPONSIVE TO ANTAGONIST 1)PlantsRAN1 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.3.5.5

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).

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). 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 copper-transporting ATPase 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 in Japanese patients 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).

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.3.6.2Zn2+-, Cd2+-, Co2+-, Hg2+-, Ni2+-, Cu2+, Pb2+-ATPase (efflux) (Hou and Mitra, 2003)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).

	

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

 
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.3.8.12Probable phospholipid-transporting ATPase IF (EC 3.6.3.1) (ATPase IR) (ATPase class VI type 11B)AnimalsATP11B of Homo sapiens
 
3.A.3.8.13

Probable phospholipid-transporting ATPase IA (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).

Animals

ATP8A1 of Homo sapiens

 
3.A.3.8.14

ATP11C aminophospholipid (phosphatidyl serine and phosphatidyl ethanolamine, but not phosphatidyl choline) flippase.  Dependent on CDC50A 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).

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).

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 inhibitor drugs, such as lonafarnib (Zhang et al. 2005).

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).

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 endosomes (Pomorski et al., 2003). The system has been reconstituted after purification in proteoliposomes. It flips phosphatidyl serine but not phosphatidylcholine or sphingomyelin (Zhou and Graham, 2009).  Drs2p (ACT3; ATP8A2), required for phospholipid translocation across the Golgi membrane: PL (in) ATP → PL (out) ADP Pì (flippase activity). Interacts with CDC50 (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).

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 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 ATPaseof 903 aas and  10 TMSs.

ATPase of Tupanvirus soda lake

 
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.3.8.3

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 (functions with the β-subunit, Lem3) (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) (is 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).

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).

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).

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