|Coordinate||30,715,800 bp (GRCm38)|
|Base Change||T ⇒ C (forward strand)|
|Gene Name||ATPase, H+/K+ exchanging, gastric, alpha polypeptide|
|Synonym(s)||H+/K+-ATPase alpha, H+K+-transporting alpha 1|
|Chromosomal Location||30,712,209-30,725,535 bp (+)|
|MGI Phenotype||Homozygous mutation of this gene results in achlorhydria, hypergastrinemia, and abnormalities of the parietal cells.|
|Amino Acid Change||Serine changed to Proline|
|Institutional Source||Beutler Lab|
S282P in Ensembl: ENSMUSP00000131964 (fasta)
|Gene Model||not available|
|Predicted Effect||possibly damaging
PolyPhen 2 Score 0.461 (Sensitivity: 0.89; Specificity: 0.90)
|Phenotypic Category||hematopoietic system|
|Alleles Listed at MGI|
|Mode of Inheritance||Autosomal Recessive|
|Local Stock||Sperm, gDNA|
|Last Updated||05/13/2016 3:09 PM by Stephen Lyon|
|Record Created||05/26/2009 12:00 AM|
|Other Mutations in This Stock||
Stock #: 7510 Run Code:
Validation Efficiency: 77/92
The sublytic phenoytpe was identified in two ENU-mutagenized G3 sisters (C7510, C7511) among a litter of three mice during a screen to identify hematological phenovariants (1). Blood cell analysis revealed that sublytic mice were severely anemic, exhibiting reduced hematocrit, hemoglobin (total and mean corpuscular hemoglobin), and mean corpuscular volume (MCV) (1). Blood smears from sublytic mice showed hypochromic red blood cells (RBCs), with anisocytosis and poikilocytosis (Figure 1A). Leukocytes were normal in number and appearance. Reciprocal bone marrow transplantation between sublytic and wild type mice demonstrated that the development of anemia was independent of the hematopoietic compartment.
The severity of anemia in sublytic mice changed with age. Four to six week old sublytic mice had increased numbers of Ter119highCD71high/med erythroid progenitor cells in the bone marrow and spleen (Figure 1B-D), associated with increased serum erythropoietin and reiculocytosis, and sometimes with thrombocytosis. Recovery of RBC numbers, hemoglobin level, and hematocrit were observed in response to the increased erythropoiesis, and approached wild type levels after about eight weeks of age and were maintained for several months. Some sublytic mice later developed severe anemia (greatly reduced hemoglobin, RBC numbers, and serum iron) after about ten months of age. These animals also developed spenomegaly (Figure 1E), and splenic achitechture was disrupted by extramedullary erythropoiesis (Figure 1F). The half-life of erythrocytes in eight week old sublytic mice was found to be slightly reduced from 26 to 24 days (Figure 1G). Additional analysis of blood from sublytic mice demonstrated that their erythrocytes were more resistant to lysis with hypotonic saline than those of wild type mice (Figure 2) (1).
Sublytic mice showed a reduction in serum ion levels (Figure 3A) as well as a reduction in iron stores in both the spleen (Figure 3B) and liver (Figure 3C) (1). Furthermore, the livers of sublytic mice had reduced levels of Hamp mRNA encoding the peptide hormone hepcidin, which controls systemic iron levels by downregulating ferric iron export into the plasma and absorption through the duodenal epithelium (Figure 3D) (1). After one month on a low iron diet, sublytic mice failed to increase iron absorption, whereas wild type mice rendered anemic under the same conditions doubled their iron absorption (Figure 3E).
Sublytic mice also had hard, enlarged stomachs (Figure 4A), with cysts in gastric glands as seen in histological sections (Figure 4B) (1). In contrast to the acidity (pH 3) of the gastric contents of wild type mice, a pH close to neutral was measured for the gastric contents of sublytic mice (Figure 4C).
The anemia phenotype of sublytic mice was rescued by maintaining the mice on a regimen of HCl-acidified drinking water (pH 2.5-2.8) for three months. Partial reversal of anemia was observed as early as one month after treatment, and a complete normalization in RBC indices was observed after three months (Figure 5A) (1). Furthermore, iron deposits in the spleen (Figure 5B) as well as an increase in serum iron (Figure 5C) and Hamp transcript levels (Figure 5D) were observed in sublytic mice given acidified water.
|Nature of Mutation|
The sublytic mutation was mapped on the basis of anemia by bulk segregation analysis (BSA) of F2 intercross offspring using C57BL/10J as the mapping strain (n=10 with mutant phenotype, 16 with normal phenotype). The mutation showed strongest linkage with two Chromosome 7 markers that had equivalent synthetic LOD scores of 3.07: B10SNPS0105 at 7014667 bp and B10SNP2G0049 at 28467081 bp (1). B10SNP2G0049 showed a higher BSA linkage score than B10SNPS0105 (3.49 vs. 3.32). In addition, a trend towards linkage was observed on Chromosome 5, with a peak synthetic LOD of 1.58 at 67382789 bp (B10SNPS0079).
Whole genome SOLiD sequencing of a sublytic homozygote identified a total of three mutations that could be validated by capillary sequencing out of 92 discrepant calls covered three or more times (validation efficiency 77/92). One of them, a heterozygous mutation in Slc28a1, was found to be on Chromosome 7, 59.79 Mb from the peak marker (B10SNP2G0049). The Atp4a gene was selected as a candidate on Chromosome 7 that when mutated causes enlargement of the stomach and formation of large cysts in parietal gastric glands (2;3) as observed in sublytic mice, and was directly sequenced. A homozygous T to C transition was detected at position 863 of the Atp4a transcript (exon 7 of 22 total exons), which results in a serine to proline change at position 282 of the encoded gastric H,K-ATPase α subunit. The mutation had been detected by SOLiD sequencing among “N pattern” nucleotides (those covered 1 or 2 times). The mutation occurs at position Chr 7: 31500819 bp, located 3.03 Mb from the peak marker. A portion of the Atp4a transcript is shown with the mutated nucleotide in red:
F2 progeny from mapping crosses were genotyped for the Atp4a mutation. 100% of anemic mice harbored the Atp4a mutation, while non-anemic mice were either wild type or heterozygous for the mutation. The Slc28a1 mutation was excluded by fine mapping, which defined a critical region located between 7014667 and 38216957 bp. We conclude that the T to C transition in Atp4a is the mutation responsible for the sublytic phenotype.
Acid secretion in the stomach is controlled by the H,K-ATPase, which catalyzes the electroneutral exchange of luminal potassium ions for cytoplasmic protons. The H,K-ATPase is one of the P-type ATPases, a large group of evolutionarily related ion pumps that use the free energy of ATP hydrolysis to drive transport and establish ion gradients across membranes (4). In all P-type ATPases both the N- and C-termini are located on the cytoplasmic side of the membrane such that these proteins contain an even number of transmembrane segments. Four well-defined, conserved protein domains exist in P-type ATPases: the phosphorylation (P) domain, nucleotide-binding (N) domain, actuator (A) domain, and membrane (M) domain (4-8). The P, N, and A domains are positioned cytoplasmically, whereas the M domain spans the plasma membrane. The H,K-ATPase is closely related to other cation transport ATPases including the Na,K-ATPase and the sarcoplasmic reticulum Ca2+-ATPase (SERCA). It consists of two subunits in [αβ]2 heterodimer oligomer stoichiometry: the 110 kD catalytic α subunit encoded by Atp4a (1033 aa in mouse) that contains all the elements of P-type ATPases (Figure 6), and the 35 kD accessory β subunit encoded by Atp4b (294 aa in mouse) (9;10). The β subunit is a highly glocosylated type II single-span membrane protein with a short N-terminal cytoplasmic tail and a large C-terminal extracellular domain (11-13); it has been shown to regulate assembly, trafficking, and insertion of the complex into the membrane (14;15). The α subunit is 98% identical in sequence between mice and humans; the β subunit is 83.8% identical between mice and humans.
The 3D structure of the H,K-ATPase α and β oligomer deduced by electron crystallography (16) demonstrates an overall conformation similar to those of other P-type ATPases (6-8). The α subunit has a characteristic cytoplasmic domain consisting of P, N, and A domains, and a transmembrane M domain with ten membrane-spanning helices (Figure 6). The smaller β subunit consists of an ectodomain, a single transmembrane helix that contacts residues of the α subunit M domain, and an rod-like N-terminal tail. Biochemical studies demonstrated that a cluster of carboxylic amino acids located in the middle of M4, M5, M6, and M8 contains the ion-binding domain in the α subunit (17). One amino acid, lysine 791, is critical for the specificity of the H,K-ATPase in outward transport of the hydronium ion.
P-type ATPases cycle through a series of conformational changes to translocate ions. The Post-Albers or E1/E2 model of the reaction cycle (18;19) has been widely used to explain the functional transitions of P-type ATPases that facilitate ion transport. The model proposed two conformational states, E1 and E2, where the E1 state has high-affinity binding sites for the ion to be transported from the cytoplasmic side of the membrane, and the E2 state has high-affinity binding sites for the ion to be transported from the extracellular side of the membrane (Figure 7). In the first step, ion1 binds to E1 from inside the cell, triggering autophosphorylation of the enzyme by Mg2+-ATP on a conserved aspartate residue (see below), which leads to the phosphorylated E1-P state. E2-P is formed by rate-limiting conformational changes to E1-P. E2-P has reduced affinity for ion1, which is thus released outside the cell. E2-P is unable to phosphorylate ADP, meaning that the reaction cannot proceed in the reverse direction from E2-P. Ion2 binds to E2-P from the outside of the cell, leading to hydrolysis of the phosphorylated aspartate, which triggers release of ion2 inside the cell and a return to the E1 state. The utility of the E1/E2 model has recently been questioned because of some inaccuracies identified experimentally (20), but it still seems to be generally accepted and appears in many current publications.
The P domain is the catalytic core of P-type ATPases, containing a conserved sequence (DKTGTLT) in which the aspartate residue is reversibly autophosphorylated to form the high energy E1-P intermediate of the reaction cycle (21). The N domain is linked to the P domain by a conserved hinge of two antiparallel strands, and serves to bind Mg2+-ATP and deliver it to the phosphorylation site in the P domain. The A domain does not interact with ions or nucleotides, but undergoes a rotation during the E1-P to E2-P transition that places it in contact with the phosphorylation site, inducing conformational changes of the P and M domains that may serve to cause release of ion1 and binding of ion2. The M domain consists of ten membrane-spanning helices that surround the ion-binding sites, and is directly linked to the P domain through helices M4 and M5. Reciprocal movements of M domain helices have been proposed to open, in turn, the ion binding cavities facing the cytoplasm and extracellular space, and create high affinity binding sites for different ions through the reorientation of coordinating side chains. Amino acid sequences of M domains are the least conserved of all P-type ATPase domains, reflecting the distinct ionic specificities of each transporter. A schematic depiction of the catalytic cycle of P-type ATPases conforming to the E1/E2 model is shown in Figure 8.
The two heterodimers of the H,K-ATPase oligomer function out of phase with each other (9). The ratio of H+ ions transferred to ATP molecules hydrolyzed has been measured at 1 or 2 H+ ions per ATP, and may be pH dependent (22-24). Following the E1/E2 model, the enzyme has cytoplasmic-open E1 and luminal-open E2 states with high affinity for H+ and K+, respectively. Upon autophosphorylation on Asp386 (mouse numbering), the H,K-ATPase exists predominantly in the E2-P, ADP-non-reactive form; the reverse reaction from E2-P to E1-P is strongly disfavored (25;26). This property may account for the ability of the H,K-ATPase to maintain the extremely steep H+ gradient across the gastric mucosa. In the 3D structure (16), the N-terminal tail of the β subunit was shown to contact the α subunit P domain in the E2-P state; deletion of the β subunit N-terminal tail was shown to result in a significantly lower proportion of forward reaction products (E1-P àE2-P). These findings suggest that the β subunit N terminus prevents the reverse reaction from E2-P to E1-P, perhaps by holding E2-P in an energetically lower, stabilized state (16).
The sublytic serine to proline mutation at position 282 lies in the A domain. The primary sequence of the A domain is divided into two unequal parts by the sequences for helices M1 and M2. The sublytic mutation is in the second part of the domain, close to its C terminus.
The α-subunit of the gastric H,K-ATPase is present in tubulovesicular and canalicular membranes of parietal cells of the stomach (27-29). It has been estimated that the H,K-ATPase represents up to 10% of parietal cell proteins (30). H,K-ATPase is also expressed in the kidney (31-33).
The gastric mucosa secretes into the stomach a highly acidic fluid (pH 2-3) that denatures proteins, activates the proenzyme pepsinogen (secreted by chief cells), and kills or inhibits the growth of many foodborne organisms. This gastric acid consists mainly of isotonic HCl (0.15 N) and is produced and discharged by parietal cells through the function of the H,K-ATPase, which pumps acid into the stomach lumen against a gradient of greater than a million fold. Because H+ is transported in exchange for K+, no net charge accumulates across the plasma membrane (34). The H,K-ATPase is the target of proton pump inhibitors (PPIs), drugs such as omeprazole (Losec; AstraZeneca) and lansoprazole (Prevacid; TAP Pharmaceuticals) that inhibit gastric acid secretion for the treatment of peptic ulcers, reflux oesophagitis, gastroesophageal reflux disease (GERD), Barrett’s esophagus, and Zollinger-Ellison syndrome, as well as the eradication of Helicobacter pylori as part of combination regimens (35;36).
Parietal cells are morphologically and functionally specialized for acid production. At their apical membrane, parietal cells possess a series of small canals (canaliculi) that invaginate from the cell surface and project throughout the cell interior. In the non-secreting or resting parietal cell, the canaliculi are lined with short microvilli, and throughout the cytoplamic space are an abundance of membranous structures, called tubulovesicles, constituting some 50% of the total membrane mass and taking the form of vesicles, tubules, and cisternal sacs (Figure 9) (37). These tubulovesicles contain virtually all of the resting state H,K-ATPase, which is inactive for H+ transport because tubulovesicles lack an endogenous pathway for bringing K+ inside the vesicle where it can be exchanged for H+ (38;39). Upon stimulation of acid secretion (see below), the H,K-ATPase-rich tubulovesicles migrate and fuse to the canalicular membrane, resulting in a dramatically expanded apical membrane surface with elongated microvilli and dilated canalicular spaces, and a correspondingly reduced cytoplasmic tubulovesicular membrane area (40-42). This membrane fusion event also recruits large numbers of the H,K-ATPase to the apical membrane, where it transports H+ out of the cell in exchange for transport of K+ into the cell (38). The H,K-ATPase operates in parallel with the K+ channel KCNQ1, which is required to recycle K+ to the luminal surface of the proton pump (43-45). Inhibition or deletion of KCNQ1 blocks acid secretion by parietal cells in vitro and in vivo.
Acid secretion by parietal cells is stimulated by extracellular signals delivered through receptors on the basolateral membrane. The histamine H2 receptor, cholinergic muscarinic M3 receptor, and gastrin CCK-B receptor are the three receptors known to stimulate acid secretion, with histaminergic stimulation being a central player. These receptors activate a number of intracellular signaling pathways, including those of protein kinase A (PKA), protein kinase C (PKC), Ca2+-calmodulin kinase II (CaMKII), and PI3 kinase, that lead to upregulation of H,K-ATPase mRNA and fusion of H,K-ATPase-containing tubulovesicles with canalicular membranes [reviewed in (46;47)]. Membrane fusion is mediated by SNAREs and their associated proteins including Rab proteins. Re-sequestration of H,K-ATPases back into tubulovesicles negatively regulates acid secretion and is carried out through coat protein-mediated endocytosis (e.g. clathrin-mediated endocytosis), together with dynamin and other associated proteins (46).
The gastric H,K-ATPase was first cloned from rat in 1986 (48), and later from pig, rabbit, dog, human, and mouse (49-53). In 2000, the phenotype of the knockout mouse was first reported (3). Atp4a-/- mice failed to secrete gastric acid, resulting in a near neutral pH (6.9) of the gastric contents, compared to a pH of 3.17 in wild type littermates. Atp4a-/- mice produced elevated levels of gastrin both at the transcript and protein levels. Parietal cells and chief cells were present in normal numbers in Atp4a-/- mice, and expressed the H,K-ATPase β subunit and pepsinogen, respectively. However, both cell types displayed changes in morphology compared to wild type cells. Parietal cells of Atp4a-/- mice contained dilated canaliculi with sparse short and stiff microvilli, and some contained massive stores of cytoplasmic glycogen. At the ultrastructural level, Atp4a-/- parietal cells displayed decreased canalicular folds, and normally abundant tubulovesicles were replaced with a few rigid round vesicles (54). The microvilli of Atp4a-/- parietal cells displayed alterations in the actin cytoskeleton. Atp4a-/- chief cells had a reduced number of granules and endoplasmic reticulum. These phenotypes are similar to, but do not precisely replicated, those of Atp4b-/- mice lacking the H,K-ATPase β subunit (55;56).
Deficiency of the H,K-ATPase α subunit has been implicated in the development of stomach hyperplasia (2;3). Stomach weight and thickness in Atp4a-/- mice were significantly increased compared to those of wild type mice by 8 months of age, and continued to increase until at least 20 months of age. Atp4a-/- mice also had more cells per gastric gland than wild type mice. Cyst formation was observed by 3 months of age, but with age, the formation of countless, and occasionally very large, cysts was widespread. In addition, incomplete intestinal metaplasia, ciliated metaplasia, a shift in mucins from neutral to acidic, and inflammation were observed in Atp4a-/-mice. Transcripts encoding gastric growth and oncogenic factors, including those for Reg IIIγ, Reg IIIδ, and osteopontin,were upregulated in stomach tissue from Atp4a-/- mice. However, nuclear atypia and the invasion of epithelial cells into the muscularis mucosa were absent, as was metastasis into adjacent organs.
The H,K-ATPase is the major autoantigen targeted in autoimmune gastritis (57), an inflammatory disease of the stomach epithelia leading to pernicious anemia caused by the loss of parietal cells and consequently a lack of intrinsic factor, which is required to absorb vitamin B12 from the gut (58). Autoimmune gastritis is characterized by a monocytic infiltrate of the gastric mucosa, loss of parietal and zymogenic cells, and T and B cell responses to the H,K-ATPase. A CD4+ T cell response to the α and β subunits of the H,K-ATPase is necessary and sufficient to induce autoimmune gastritis (59-61). In mice, autoimmune gastritis develops in susceptible strains after neonatal thymectomy (62), thymectomy combined with administration of immunosuppressive drugs or irradiation in adult mice (63), or after CD4+ T cell transfer to lymphopenic recipients (64). It is now known that failure to eliminate H,K-ATPase specific CD4+ T cells, and loss of CD4+ T cell tolerance to the H,K-ATPase leads to the disease. Exposure of T cells to the gastritogenic H,K-ATPase antigen outside of the thymus is important for purging of autoreactive CD4+ T cells (64-66), and for the maintenance of H,K-ATPase specific CD4+CD25+ regulatory T cells (65;67).
Evidence from numerous studies indicates that gastric acid secretion is important for optimal absorption of non-heme iron. Iron is obtained exclusively from the diet, and is absorbed in the proximal portion of the duodenum by intestinal enterocytes (68). Approximately two-thirds of dietary non-heme iron is in the form of ferric (Fe3+) salts, which are precipitated in solutions with a pH greater than 3 (69). In the stomach, gastric acid maintains the acidic environment necessary to enforce solubility, and permit subsequent chelation and/or reduction, of ferric iron. Clinical studies as early as 1932 suggested that gastric acidity might have a role in the absorption of iron, because the response to oral iron supplements in patients with iron deficiency was greater in patients with normal gastric acidity than those with achlorhydria [see Discussion in (70)]. Further studies showed that prolonged achlorhydria resulting from gastric resection or atrophic gastritis can result in iron deficiency and anemia (71-73). However, patients with Zollinger–Ellison syndrome treated long term (5.7 years average) with omeprazole did not develop iron deficiency or decrease body iron stores (70).
For absorption in the less acidic proximal small intestine, soluble ferric iron must be reduced to ferrous (Fe2+) iron, which enters enterocytes through divalent metal transporter 1 (DMT1), the primary transmembrane non-heme iron transporter in intestinal epithelial cells (74;75). DMT1 is a cotransporter requiring protons to move Fe2+ across the cell membrane (76;77). These protons are also provided by gastric acid flowing into the proximal portion of the duodenum where DMT1 is most highly expressed.
The sublytic mice are able to sense and respond to iron-deficiency by down-regulating Hamp transcription; however, they are unable to upregulate intestinal iron absorption (1). The data strongly suggest that the inability of sublytic mice to absorb iron is due to poor solubilty of ferric iron and inefficient DMT1 function, which are caused by reduced availability of protons in the stomach (1). Sublytic mice displayed a 3-fold increase in DMT1 transcript levels, consistent with this hypothesis (1). An increase in stomach inflammation, as observed in Atp4a knockout mice (2), may also contribute to the anemia phenotype observed in the sublytic model. The increased surface-to-volume ratio of microcytic sublytic erythrocytes may contribute to the reduced osmotic fragility observed (1).
|Primers||Primers cannot be located by automatic search.|
Sublytic genotyping is performed by amplifying the region containing the mutation using PCR, followed by sequencing of the amplified region to detect the single nucleotide change.
sublytic (F): 5’-ACACATGAGAGTCCCCTTGAGACC -3’
sublytic (R): 5’- GTTTGGGCTGACTTCCCACAGTAG -3’
1) 95°C 2:00
2) 95°C 0:30
3) 56°C 0:30
4) 72°C 1:00
5) repeat steps (2-4) 29X
6) 72°C 7:00
7) 4°C ∞
Primers for sequencing
sublytic_seq(F): 5’- TCCACCATGTGTCTGGAGG -3’
sublytic_seq(R): 5’- GGTCCAGGACCCTGTTTATTCAC -3’
The following sequence of 756 nucleotides (from Genbank genomic region NC_000073.5 for linear genomic sequence of Atp4a, sense strand) is amplified:
3280 a cacatgagag tccccttgag
3301 acccgcaaca tcgccttctt ctccaccatg tgtctggagg gtctgtgaag catcgttagc
3361 ctgtcctgaa gccacacaga ccccatactt tccataatag cgccttttct gttctgtgtg
3421 atgtgtgcca caagcctccc tatcacagag cacctttctg gtgctgccca cctcaccctg
3481 agctctttct ccccttgttg ccactgcagg aacagctcag ggtttggtgg tgagcaccgg
3541 cgatcgcacc atcattgggc gcatcgcctc gctggcctcg ggtgtggaaa acgagaagac
3601 tccgattgca atcgagattg aacattttgt ggacatcatt gccggcctgg ccatcctctt
3661 cggtgccaca ttctttgtgg tggccatgtg tattggctat accttccttc gggccatggt
3721 cttcttcatg gccattgtgg tagcctatgt gcctgagggg ctgctggcaa ctgtcacagt
3781 gagtaaggga gaaggggtgg gggggggtgc agggagcaga gagctgcttg ttcagtcatc
3841 ccaccagacg tccctactac ctgtccattc attcccctcc cccaccctac atcatccatc
3901 cattcagtga acatttatca ggatgctctg ggctggcccc gtgtcataca ccagagccca
3961 cgggtgaata aacagggtcc tggaccaagg aagatgtggg aagatgttgg gctactgtgg
4021 gaagtcagcc caaac
Primer binding sites are underlined; sequencing primer binding sites are highlighted in gray; the mutated T is indicated in red.
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20. Scarborough, G. A. (2003) Why we must Move on from the E1E2 Model for the Reaction Cycle of the P-Type ATPases. J. Bioenerg. Biomembr.. 35, 193-201.
21. Post, R. L., and Kume, S. (1973) Evidence for an Aspartyl Phosphate Residue at the Active Site of Sodium and Potassium Ion Transport Adenosine Triphosphatase. J. Biol. Chem.. 248, 6993-7000.
22. Morii, M., Yamauchi, M., Ichikawa, T., Fujii, T., Takahashi, Y., Asano, S., Takeguchi, N., and Sakai, H. (2008) Involvement of the H3O+-Lys-164 -Gln-161-Glu-345 Charge Transfer Pathway in Proton Transport of Gastric H+,K+-ATPase. J. Biol. Chem.. 283, 16876-16884.
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|Science Writers||Eva Marie Y. Moresco, Anne Murray|
|Illustrators||Diantha La Vine|
|Authors||Xin Du, Oren Milstein, Bruce Beutler|