Is the affinity of an enzyme or transporter for its substrate or solute influenced by the amino acids at the binding site?

Is the affinity of an enzyme or transporter for its substrate or solute influenced by the amino acids at the binding site?

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The affinity of an enzyme or a protein transporter indicates the strenght of the binding of substrate/solute. An enzyme with higher affinity binds its substrate more strongly than a counterpart with a lower affinity. Is affinity always a function of the protein structure of enzyme? If the same enzyme in two different species had different affinities for substrate, would that indicate that there were differences in amino acids at the binding sites of the enzyme in the two species?

In a reaction which follows a saturation kinetics, KM is basically the concentration of substrate/ligand at which the rate of the reaction is half of the maximum rate (or the binding sites are half saturated).

The biophysical meaning of KM would depend on the underlying model. For example, in the equilibrium approximation of Michaelis-Menten model, KM is same as dissociation constant (of enzyme and substrate). In the Briggs-Haldane model (quasi steady state approximation), it is slightly complex: KM = (kr + kcat)/kf.

Overall, the thumb rule is that lower the KM faster will the reaction saturate.

Is affinity always a function of the protein structure of enzyme? If someone study same enzyme in two different species and observe that the enzyme has different affinities in two species, then does it indicate that there might by slight difference in key amino-acid component at the binding sites of the enzyme between these two species?

Almost every property of the protein (not just affinity towards a ligand) is because of its chemical composition and structure. Although they are directly involved in binding, the amino acids at binding site are not the sole contributors to the affinity. The other amino acids that are critical for the overall structure of the protein are also important in preserving the function.

I will add to the answer by @WYSIWYG by turning your question round: “If you changed the amino acids at the binding site, would this affect the affinity of a protein for a ligand*?”

The answer to this is yes, and for an enzyme you can change the substrate specificity entirely, as one can discover from reading this review by Wilson and Agard.

However one should also be aware that changing residues at parts of a protein outside the ligand-binding site can also affect the affinity of binding - see, for example, this paper by Oue et al.

[*This makes the question more general to include other proteins that bind ligands such as membrane receptors and transport proteins such as heamoglobin.]

Is the affinity of an enzyme or transporter for its substrate or solute influenced by the amino acids at the binding site? - Biology

Glutamine is an amino acid with a plethora of functions in physiology and cancer.

Tumor cells are addicted to glutamine.

There are multiple glutamine transporters in the plasma membrane of mammalian cells.

Tumor cells induce selective transporters to meet their high demands for glutamine.

Some of the glutamine transporters have potential as drug targets in cancer therapy.


ABC transporters utilize the energy of ATP binding and hydrolysis to transport various substrates across cellular membranes. They are divided into three main functional categories. In prokaryotes, importers mediate the uptake of nutrients into the cell. The substrates that can be transported include ions, amino acids, peptides, sugars, and other molecules that are mostly hydrophilic. The membrane-spanning region of the ABC transporter protects hydrophilic substrates from the lipids of the membrane bilayer thus providing a pathway across the cell membrane. Eukaryotes do not possess any importers. Exporters or effluxers, which are present both in prokaryotes and eukaryotes, function as pumps that extrude toxins and drugs out of the cell. In gram-negative bacteria, exporters transport lipids and some polysaccharides from the cytoplasm to the periplasm. The third subgroup of ABC proteins do not function as transporters, but are rather involved in translation and DNA repair processes. [4]

Prokaryotic Edit

Bacterial ABC transporters are essential in cell viability, virulence, and pathogenicity. [1] [4] Iron ABC uptake systems, for example, are important effectors of virulence. [11] Pathogens use siderophores, such as Enterobactin, to scavenge iron that is in complex with high-affinity iron-binding proteins or erythrocytes. These are high-affinity iron-chelating molecules that are secreted by bacteria and reabsorb iron into iron-siderophore complexes. The chvE-gguAB gene in Agrobacterium tumefaciens encodes glucose and galactose importers that are also associated with virulence. [12] [13] Transporters are extremely vital in cell survival such that they function as protein systems that counteract any undesirable change occurring in the cell. For instance, a potential lethal increase in osmotic strength is counterbalanced by activation of osmosensing ABC transporters that mediate uptake of solutes. [14] Other than functioning in transport, some bacterial ABC proteins are also involved in the regulation of several physiological processes. [4]

In bacterial efflux systems, certain substances that need to be extruded from the cell include surface components of the bacterial cell (e.g. capsular polysaccharides, lipopolysaccharides, and teichoic acid), proteins involved in bacterial pathogenesis (e.g. hemolysis, heme-binding protein, and alkaline protease), heme, hydrolytic enzymes, S-layer proteins, competence factors, toxins, antibiotics, bacteriocins, peptide antibiotics, drugs and siderophores. [15] They also play important roles in biosynthetic pathways, including extracellular polysaccharide biosynthesis [16] and cytochrome biogenesis. [17]

Eukaryotic Edit

Although most eukaryotic ABC transporters are effluxers, some are not directly involved in transporting substrates. In the cystic fibrosis transmembrane regulator (CFTR) and in the sulfonylurea receptor (SUR), ATP hydrolysis is associated with the regulation of opening and closing of ion channels carried by the ABC protein itself or other proteins. [5]

Human ABC transporters are involved in several diseases that arise from polymorphisms in ABC genes and rarely due to complete loss of function of single ABC proteins. [18] Such diseases include Mendelian diseases and complex genetic disorders such as cystic fibrosis, adrenoleukodystrophy, Stargardt disease, Tangier disease, immune deficiencies, progressive familial intraheptic cholestasis, Dubin–Johnson syndrome, Pseudoxanthoma elasticum, persistent hyperinsulinemic hypoglycemia of infancy due to focal adenomatous hyperplasia, X-linked sideroblastosis and anemia, age-related macular degeneration, familial hypoapoproteinemia, Retinitis pigmentosum, cone rod dystrophy, and others. [5] The human ABCB (MDR/TAP) family is responsible for multiple drug resistance (MDR) against a variety of structurally unrelated drugs. ABCB1 or MDR1 P-glycoprotein is also involved in other biological processes for which lipid transport is the main function. It is found to mediate the secretion of the steroid aldosterone by the adrenals, and its inhibition blocked the migration of dendritic immune cells, [19] possibly related to the outward transport of the lipid platelet activating factor (PAF). It has also been reported that ABCB1 mediates transport of cortisol and dexamethasone, but not of progesterone in ABCB1 transfected cells. MDR1 can also transport cholesterol, short-chain and long-chain analogs of phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylserine (PS), sphingomyelin (SM), and glucosylceramide (GlcCer). Multispecific transport of diverse endogenous lipids through the MDR1 transporter can possibly affect the transbilayer distribution of lipids, in particular of species normally predominant on the inner plasma membrane leaflet such as PS and PE. [18]

More recently, ABC-transporters have been shown to exist within the placenta, indicating they could play a protective role for the developing fetus against xenobiotics. [20]

All ABC transport proteins share a structural organization consisting of four core domains [21] . These domains consist of two trans-membrane (T) domains and two cytosolic (A) domains. The two T domains alternate between an inward and outward facing orientation, and the alternation is powered by the hydrolysis of adenosine triphosphate or ATP. ATP binds to the A subunits and it is then hydrolyzed to power the alternation, but the exact process by which this happens is not known. The four domains can be present in four separate polypeptides, which occur mostly in bacteria, or present in one or two multi-domain polypeptides. [10] When the polypeptides are one domain, they can be referred to as a full domain, and when they are two multi-domains they can be referred to as a half domain. [9] The T domains are each built of typically 10 membrane spanning alpha helices, through which the transported substance can cross through the plasma membrane. Also, the structure of the T domains determines the specificity of each ABC protein. In the inward facing conformation, the binding site on the A domain is open directly to the surrounding aqueous solutions. This allows hydrophilic molecules to enter the binding site directly from the inner leaflet of the phospholipid bilayer. In addition, a gap in the protein is accessible directly from the hydrophobic core of the inner leaflet of the membrane bilayer. This allows hydrophobic molecules to enter the binding site directly from the inner leaflet of the phospholipid bilayer. After the ATP powered move to the outward facing conformation, molecules are released from the binding site and allowed to escape into the exoplasmic leaflet or directly into the extracellular medium. [10]

The common feature of all ABC transporters is that they consist of two distinct domains, the transmembrane domain (TMD) and the nucleotide-binding domain (NBD). The TMD, also known as membrane-spanning domain (MSD) or integral membrane (IM) domain, consists of alpha helices, embedded in the membrane bilayer. It recognizes a variety of substrates and undergoes conformational changes to transport the substrate across the membrane. The sequence and architecture of TMDs is variable, reflecting the chemical diversity of substrates that can be translocated. The NBD or ATP-binding cassette (ABC) domain, on the other hand, is located in the cytoplasm and has a highly conserved sequence. The NBD is the site for ATP binding. [22] In most exporters, the N-terminal transmembrane domain and the C-terminal ABC domains are fused as a single polypeptide chain, arranged as TMD-NBD-TMD-NBD. An example is the E. coli hemolysin exporter HlyB. Importers have an inverted organization, that is, NBD-TMD-NBD-TMD, where the ABC domain is N-terminal whereas the TMD is C-terminal, such as in the E. coli MacB protein responsible for macrolide resistance. [4] [5]

The structural architecture of ABC transporters consists minimally of two TMDs and two NBDs. Four individual polypeptide chains including two TMD and two NBD subunits, may combine to form a full transporter such as in the E. coli BtuCD [23] [24] importer involved in the uptake of vitamin B12. Most exporters, such as in the multidrug exporter Sav1866 [25] from Staphylococcus aureus, are made up of a homodimer consisting of two half transporters or monomers of a TMD fused to a nucleotide-binding domain (NBD). A full transporter is often required to gain functionality. Some ABC transporters have additional elements that contribute to the regulatory function of this class of proteins. In particular, importers have a high-affinity binding protein (BP) that specifically associates with the substrate in the periplasm for delivery to the appropriate ABC transporter. Exporters do not have the binding protein but have an intracellular domain (ICD) that joins the membrane-spanning helices and the ABC domain. The ICD is believed to be responsible for communication between the TMD and NBD. [22]

Transmembrane domain (TMD) Edit

Most transporters have transmembrane domains that consist of a total of 12 α-helices with 6 α-helices per monomer. Since TMDs are structurally diverse, some transporters have varying number of helices (between six and eleven). The TM domains are categorized into three distinct sets of folds: type I ABC importer, type II ABC importer and ABC exporter folds. The classification of importer folds is based on detailed characterization of the sequences. [22]

The type I ABC importer fold was originally observed in the ModB TM subunit of the molybdate transporter. [26] This diagnostic fold can also be found in the MalF and MalG TM subunits of MalFGK2 [27] and the Met transporter MetI. [28] In the MetI transporter, a minimal set of 5 transmembrane helices constitute this fold while an additional helix is present for both ModB and MalG. The common organization of the fold is the "up-down" topology of the TM2-5 helices that lines the translocation pathway and the TM1 helix wrapped around the outer, membrane-facing surface and contacts the other TM helices.

The type II ABC importer fold is observed in the twenty TM helix-domain of BtuCD [23] and in Hi1471, [29] a homologous transporter from Haemophilus influenzae. In BtuCD, the packing of the helices is complex. The noticeable pattern is that the TM2 helix is positioned through the center of the subunit where it is surrounded in close proximity by the other helices. Meanwhile, the TM5 and TM10 helices are positioned in the TMD interface. The membrane spanning region of ABC exporters is organized into two "wings" that are composed of helices TM1 and TM2 from one subunit and TM3-6 of the other, in a domain-swapped arrangement. A prominent pattern is that helices TM1-3 are related to TM4-6 by an approximate twofold rotation around an axis in the plane of the membrane. [22]

The exporter fold is originally observed in the Sav1866 structure. It contains 12 TM helices, 6 per monomer. [22]

Nucleotide-binding domain (NBD) Edit

The ABC domain consists of two domains, the catalytic core domain similar to RecA-like motor ATPases and a smaller, structurally diverse α-helical subdomain that is unique to ABC transporters. The larger domain typically consists of two β-sheets and six α helices, where the catalytic Walker A motif (GXXGXGKS/T where X is any amino acid) or P-loop and Walker B motif (ΦΦΦΦD, of which Φ is a hydrophobic residue) is situated. The helical domain consists of three or four helices and the ABC signature motif, also known as LSGGQ motif, linker peptide or C motif. The ABC domain also has a glutamine residue residing in a flexible loop called Q loop, lid or γ-phosphate switch, that connects the TMD and ABC. The Q loop is presumed to be involved in the interaction of the NBD and TMD, particularly in the coupling of nucleotide hydrolysis to the conformational changes of the TMD during substrate translocation. The H motif or switch region contains a highly conserved histidine residue that is also important in the interaction of the ABC domain with ATP. The name ATP-binding cassette is derived from the diagnostic arrangement of the folds or motifs of this class of proteins upon formation of the ATP sandwich and ATP hydrolysis. [4] [15] [22]

ATP binding and hydrolysis Edit

Dimer formation of the two ABC domains of transporters requires ATP binding. [30] It is generally observed that the ATP bound state is associated with the most extensive interface between ABC domains, whereas the structures of nucleotide-free transporters exhibit conformations with greater separations between the ABC domains. [22] Structures of the ATP-bound state of isolated NBDs have been reported for importers including HisP, [31] GlcV, [32] MJ1267, [33] E. coli MalK (E.c.MalK), [34] T. litoralis MalK (TlMalK), [35] and exporters such as TAP, [36] HlyB, [37] MJ0796, [38] [39] Sav1866, [25] and MsbA. [40] In these transporters, ATP is bound to the ABC domain. Two molecules of ATP are positioned at the interface of the dimer, sandwiched between the Walker A motif of one subunit and the LSGGQ motif of the other. [22] This was first observed in Rad50 [41] and reported in structures of MJ0796, the NBD subunit of the LolD transporter from Methanococcus jannaschii [39] and E.c.MalK of a maltose transporter. [34] These structures were also consistent with results from biochemical studies revealing that ATP is in close contact with residues in the P-loop and LSGGQ motif during catalysis. [42]

Nucleotide binding is required to ensure the electrostatic and/or structural integrity of the active site and contribute to the formation of an active NBD dimer. [43] Binding of ATP is stabilized by the following interactions: (1) ring-stacking interaction of a conserved aromatic residue preceding the Walker A motif and the adenosine ring of ATP, [44] [45] (2) hydrogen-bonds between a conserved lysine residue in the Walker A motif and the oxygen atoms of the β- and γ-phosphates of ATP and coordination of these phosphates and some residues in the Walker A motif with Mg 2+ ion, [32] [36] and (3) γ-phosphate coordination with side chain of serine and backbone amide groups of glycine residues in the LSGGQ motif. [46] In addition, a residue that suggests the tight coupling of ATP binding and dimerization, is the conserved histidine in the H-loop. This histidine contacts residues across the dimer interface in the Walker A motif and the D loop, a conserved sequence following the Walker B motif. [34] [39] [41] [47]

The enzymatic hydrolysis of ATP requires proper binding of the phosphates and positioning of the γ-phosphate to the attacking water. [22] In the nucleotide binding site, the oxygen atoms of the β- and γ-phosphates of ATP are stabilized by residues in the Walker A motif [48] [49] and coordinate with Mg 2+ . [22] This Mg 2+ ion also coordinates with the terminal aspartate residue in the Walker B motif through the attacking H2O. [32] [33] [38] A general base, which may be the glutamate residue adjacent to the Walker B motif, [30] [39] [45] glutamine in the Q-loop, [29] [35] [39] or a histidine in the switch region that forms a hydrogen bond with the γ-phosphate of ATP, is found to catalyze the rate of ATP hydrolysis by promoting the attacking H2O. [34] [35] [39] [47] The precise molecular mechanism of ATP hydrolysis is still controversial. [4]

ABC transporters are active transporters, that is, they use energy in the form of adenosine triphosphate (ATP) to translocate substrates across cell membranes. These proteins harness the energy of ATP binding and/or hydrolysis to drive conformational changes in the transmembrane domain (TMD) and consequently transport molecules. [50] ABC importers and exporters have a common mechanism for transporting substrates. They are similar in their structures. The model that describes the conformational changes associated with the binding of the substrate is the alternating-access model. In this model, the substrate binding site alternates between outward- and inward-facing conformations. The relative binding affinities of the two conformations for the substrate largely determines the net direction of transport. For importers, since translocation is directed from the periplasm to the cytoplasm, the outward-facing conformation has higher binding affinity for the substrate. In contrast, the substrate binding affinity in exporters is greater in the inward-facing conformation. [22] A model that describes the conformational changes in the nucleotide-binding domain (NBD) as a result of ATP binding and hydrolysis is the ATP-switch model. This model presents two principal conformations of the NBDs: formation of a closed dimer upon binding two ATP molecules and dissociation to an open dimer facilitated by ATP hydrolysis and release of inorganic phosphate (Pi) and adenosine diphosphate (ADP). Switching between the open and closed dimer conformations induces conformational changes in the TMD resulting in substrate translocation. [51]

The general mechanism for the transport cycle of ABC transporters has not been fully elucidated, but substantial structural and biochemical data has accumulated to support a model in which ATP binding and hydrolysis is coupled to conformational changes in the transporter. The resting state of all ABC transporters has the NBDs in an open dimer configuration, with low affinity for ATP. This open conformation possesses a chamber accessible to the interior of the transporter. The transport cycle is initiated by binding of substrate to the high-affinity site on the TMDs, which induces conformational changes in the NBDs and enhances the binding of ATP. Two molecules of ATP bind, cooperatively, to form the closed dimer configuration. The closed NBD dimer induces a conformational change in the TMDs such that the TMD opens, forming a chamber with an opening opposite to that of the initial state. The affinity of the substrate to the TMD is reduced, thereby releasing the substrate. Hydrolysis of ATP follows and then sequential release of Pi and then ADP restores the transporter to its basal configuration. Although a common mechanism has been suggested, the order of substrate binding, nucleotide binding and hydrolysis, and conformational changes, as well as interactions between the domains is still debated. [4] [15] [18] [22] [40] [43] [50] [51] [52] [53] [54]

Several groups studying ABC transporters have differing assumptions on the driving force of transporter function. It is generally assumed that ATP hydrolysis provides the principal energy input or "power stroke" for transport and that the NBDs operate alternately and are possibly involved in different steps in the transport cycle. [55] However, recent structural and biochemical data shows that ATP binding, rather than ATP hydrolysis, provides the "power stroke". [56] It may also be that since ATP binding triggers NBD dimerization, the formation of the dimer may represent the "power stroke." In addition, some transporters have NBDs that do not have similar abilities in binding and hydrolyzing ATP and that the interface of the NBD dimer consists of two ATP binding pockets suggests a concurrent function of the two NBDs in the transport cycle. [51]

Some evidence to show that ATP binding is indeed the power stroke of the transport cycle was reported. [51] It has been shown that ATP binding induces changes in the substrate-binding properties of the TMDs. The affinity of ABC transporters for substrates has been difficult to measure directly, and indirect measurements, for instance through stimulation of ATPase activity, often reflects other rate-limiting steps. Recently, direct measurement of vinblastine binding to permease-glycoprotein (P-glycoprotein) in the presence of nonhydrolyzable ATP analogs, e.g. 5’-adenylyl-β-γ-imidodiphosphate (AMP-PNP), showed that ATP binding, in the absence of hydrolysis, is sufficient to reduce substrate-binding affinity. [57] Also, ATP binding induces substantial conformational changes in the TMDs. Spectroscopic, protease accessibility and crosslinking studies have shown that ATP binding to the NBDs induces conformational changes in multidrug resistance-associated protein-1 (MRP1), [58] HisPMQ, [59] LmrA, [60] and Pgp. [61] Two dimensional crystal structures of AMP-PNP-bound Pgp showed that the major conformational change during the transport cycle occurs upon ATP binding and that subsequent ATP hydrolysis introduces more limited changes. [62] Rotation and tilting of transmembrane α-helices may both contribute to these conformational changes. Other studies have focused on confirming that ATP binding induces NBD closed dimer formation. Biochemical studies of intact transport complexes suggest that the conformational changes in the NBDs are relatively small. In the absence of ATP, the NBDs may be relatively flexible, but they do not involve a major reorientation of the NBDs with respect to the other domains. ATP binding induces a rigid body rotation of the two ABC subdomains with respect to each other, which allows the proper alignment of the nucleotide in the active site and interaction with the designated motifs. There is strong biochemical evidence that binding of two ATP molecules can be cooperative, that is, ATP must bind to the two active site pockets before the NBDs can dimerize and form the closed, catalytically active conformation. [51]

Most ABC transporters that mediate the uptake of nutrients and other molecules in bacteria rely on a high-affinity solute binding protein (BP). BPs are soluble proteins located in the periplasmic space between the inner and outer membranes of gram-negative bacteria. Gram-positive microorganisms lack a periplasm such that their binding protein is often a lipoprotein bound to the external face of the cell membrane. Some gram-positive bacteria have BPs fused to the transmembrane domain of the transporter itself. [4] The first successful x-ray crystal structure of an intact ABC importer is the molybdenum transporter (ModBC-A) from Archaeoglobus fulgidus. [26] Atomic-resolution structures of three other bacterial importers, E. coli BtuCD, [23] E. coli maltose transporter (MalFGK2-E), [27] and the putative metal-chelate transporter of Haemophilus influenzae, HI1470/1, [29] have also been determined. The structures provided detailed pictures of the interaction of the transmembrane and ABC domains as well as revealed two different conformations with an opening in two opposite directions. Another common feature of importers is that each NBD is bound to one TMD primarily through a short cytoplasmic helix of the TMD, the "coupling helix". This portion of the EAA loop docks in a surface cleft formed between the RecA-like and helical ABC subdomains and lies approximately parallel to the membrane bilayer. [53]

Large ABC importers Edit

The BtuCD and HI1470/1 are classified as large (Type II) ABC importers. The transmembrane subunit of the vitamin B12 importer, BtuCD, contains 10 TM helices and the functional unit consists of two copies each of the nucleotide binding domain (NBD) and transmembrane domain (TMD). The TMD and NBD interact with one another via the cytoplasmic loop between two TM helices and the Q loop in the ABC. In the absence of nucleotide, the two ABC domains are folded and the dimer interface is open. A comparison of the structures with (BtuCDF) and without (BtuCD) binding protein reveals that BtuCD has an opening that faces the periplasm whereas in BtuCDF, the outward-facing conformation is closed to both sides of the membrane. The structures of BtuCD and the BtuCD homolog, HI1470/1, represent two different conformational states of an ABC transporter. The predicted translocation pathway in BtuCD is open to the periplasm and closed at the cytoplasmic side of the membrane while that of HI1470/1 faces the opposite direction and open only to the cytoplasm. The difference in the structures is a 9° twist of one TM subunit relative to the other. [4] [22] [53]

Small ABC importers Edit

Structures of the ModBC-A and MalFGK2-E, which are in complex with their binding protein, correspond to small (Type I) ABC importers. The TMDs of ModBC-A and MalFGK2-E have only six helices per subunit. The homodimer of ModBC-A is in a conformation in which the TM subunits (ModB) orient in an inverted V-shape with a cavity accessible to the cytoplasm. The ABC subunits (ModC), on the other hand, are arranged in an open, nucleotide-free conformation, in which the P-loop of one subunit faces but is detached from the LSGGQ motif of the other. The binding protein ModA is in a closed conformation with substrate bound in a cleft between its two lobes and attached to the extracellular loops of ModB, wherein the substrate is sitting directly above the closed entrance of the transporter. The MalFGK2-E structure resembles the catalytic transition state for ATP hydrolysis. It is in a closed conformation where it contains two ATP molecules, sandwiched between the Walker A and B motifs of one subunit and the LSGGQ motif of the other subunit. The maltose binding protein (MBP or MalE) is docked on the periplasmic side of the TM subunits (MalF and MalG) and a large, occluded cavity can be found at the interface of MalF and MalG. The arrangement of the TM helices is in a conformation that is closed toward the cytoplasm but with an opening that faces outward. The structure suggests a possibility that MBP may stimulate the ATPase activity of the transporter upon binding. [4] [22] [53]

Mechanism of transport for importers Edit

The mechanism of transport for importers supports the alternating-access model. The resting state of importers is inward-facing, where the nucleotide binding domain (NBD) dimer interface is held open by the TMDs and facing outward but occluded from the cytoplasm. Upon docking of the closed, substrate-loaded binding protein towards the periplasmic side of the transmembrane domains, ATP binds and the NBD dimer closes. This switches the resting state of transporter into an outward-facing conformation, in which the TMDs have reoriented to receive substrate from the binding protein. After hydrolysis of ATP, the NBD dimer opens and substrate is released into the cytoplasm. Release of ADP and Pi reverts the transporter into its resting state. The only inconsistency of this mechanism to the ATP-switch model is that the conformation in its resting, nucleotide-free state is different from the expected outward-facing conformation. Although that is the case, the key point is that the NBD does not dimerize unless ATP and binding protein is bound to the transporter. [4] [15] [22] [51] [53]

Prokaryotic ABC exporters are abundant and have close homologues in eukaryotes. This class of transporters is studied based on the type of substrate that is transported. One class is involved in the protein (e.g. toxins, hydrolytic enzymes, S-layer proteins, lantibiotics, bacteriocins, and competence factors) export and the other in drug efflux. ABC transporters have gained extensive attention because they contribute to the resistance of cells to antibiotics and anticancer agents by pumping drugs out of the cells. [1] [63] [4] A common mechanism is the overexpression of ABC exporters like P-glycoprotein (P-gp/ABCB1), multidrug resistance-associated protein 1 (MRP1/ABCC1), and breast cancer resistance protein (BCRP/ABCG2) in cancer cells that limit the exposure to anticancer drugs. [64]

In gram-negative organisms, ABC transporters mediate secretion of protein substrates across inner and outer membranes simultaneously without passing through the periplasm. This type of secretion is referred to as type I secretion, which involves three components that function in concert: an ABC exporter, a membrane fusion protein (MFP), and an outer membrane factor (OMF). An example is the secretion of hemolysin (HlyA) from E. coli where the inner membrane ABC transporter HlyB interacts with an inner membrane fusion protein HlyD and an outer membrane facilitator TolC. TolC allows hemolysin to be transported across the two membranes, bypassing the periplasm. [1] [63] [15]

Bacterial drug resistance has become an increasingly major health problem. One of the mechanisms for drug resistance is associated with an increase in antibiotic efflux from the bacterial cell. Drug resistance associated with drug efflux, mediated by P-glycoprotein, was originally reported in mammalian cells. In bacteria, Levy and colleagues presented the first evidence that antibiotic resistance was caused by active efflux of a drug. [65] P-glycoprotein is the best-studied efflux pump and as such has offered important insights into the mechanism of bacterial pumps. [4] Although some exporters transport a specific type of substrate, most transporters extrude a diverse class of drugs with varying structure. [18] These transporters are commonly called multi-drug resistant (MDR) ABC transporters and sometimes referred to as "hydrophobic vacuum cleaners". [54]

Human ABCB1/MDR1 P-glycoprotein Edit

P-glycoprotein (3.A.1.201.1) is a well-studied protein associated with multi-drug resistance. It belongs to the human ABCB (MDR/TAP) family and is also known as ABCB1 or MDR1 Pgp. MDR1 consists of a functional monomer with two transmembrane domains (TMD) and two nucleotide-binding domains (NBD). This protein can transport mainly cationic or electrically neutral substrates as well as a broad spectrum of amphiphilic substrates. The structure of the full-size ABCB1 monomer was obtained in the presence and absence of nucleotide using electron cryo crystallography. Without the nucleotide, the TMDs are approximately parallel and form a barrel surrounding a central pore, with the opening facing towards the extracellular side of the membrane and closed at the intracellular face. In the presence of the nonhydrolyzable ATP analog, AMP-PNP, the TMDs have a substantial reorganization with three clearly segregated domains. A central pore, which is enclosed between the TMDs, is slightly open towards the intracellular face with a gap between two domains allowing access of substrate from the lipid phase. Substantial repacking and possible rotation of the TM helices upon nucleotide binding suggests a helix rotation model for the transport mechanism. [18]

Plant transporters Edit

The genome of the model plant Arabidopsis thaliana is capable of encoding 120 ABC proteins compared to 50-70 ABC proteins that are encoded by the human genome and fruit flies (Drosophila melanogaster). Plant ABC proteins are categorized in 13 subfamilies on the basis of size (full, half or quarter), orientation, and overall amino acid sequence similarity. [66] Multidrug resistant (MDR) homologs, also known as P-glycoproteins, represent the largest subfamily in plants with 22 members and the second largest overall ABC subfamily. The B subfamily of plant ABC transporters (ABCBs) are characterized by their localization to the plasma membrane. [67] Plant ABCB transporters are characterized by heterologously expressing them in Escherichia coli, Saccharomyces cerevisiae, Schizosaccharomyces pombe (fission yeast), and HeLa cells to determine substrate specificity. Plant ABCB transporters have shown to transport the phytohormone indole-3-acetic acid ( IAA), [68] also known as auxin, the essential regulator for plant growth and development. [69] [70] The directional polar transport of auxin mediates plant environmental responses through processes such as phototropism and gravitropism. [71] Two of the best studied auxin transporters, ABCB1 and ABCB19, have been characterized to be primary auxin exporters [69] Other ABCB transporters such as ABCB4 participate in both the export and import of auxin [69] At low intracellular auxin concentrations ABCB4 imports auxin until it reaches a certain threshold which then reverses function to only export auxin. [69] [72]

Sav1866 Edit

The first high-resolution structure reported for an ABC exporter was that of Sav1866 (3.A.1.106.2) from Staphylococcus aureus. [18] [73] Sav1866 is a homolog of multidrug ABC transporters. It shows significant sequence similarity to human ABC transporters of subfamily B that includes MDR1 and TAP1/TAP2. The ATPase activity of Sav1866 is known to be stimulated by cancer drugs such as doxorubicin, vinblastine and others, [74] which suggests similar substrate specificity to P-glycoprotein and therefore a possible common mechanism of substrate translocation. Sav1866 is a homodimer of half transporters, and each subunit contains an N-terminal TMD with six helices and a C-terminal NBD. The NBDs are similar in structure to those of other ABC transporters, in which the two ATP binding sites are formed at the dimer interface between the Walker A motif of one NBD and the LSGGQ motif of the other. The ADP-bound structure of Sav1866 shows the NBDs in a closed dimer and the TM helices split into two "wings" oriented towards the periplasm, forming the outward-facing conformation. Each wing consists of helices TM1-2 from one subunit and TM3-6 from the other subunit. It contains long intracellular loops (ICLs or ICD) connecting the TMDs that extend beyond the lipid bilayer into the cytoplasm and interacts with the 8=D. Whereas the importers contain a short coupling helix that contact a single NBD, Sav1866 has two intracellular coupling helices, one (ICL1) contacting the NBDs of both subunits and the other (ICL2) interacting with only the opposite NBD subunit. [22] [25] [53]

MsbA Edit

MsbA (3.A.1.106.1) is a multi-drug resistant (MDR) ABC transporter and possibly a lipid flippase. It is an ATPase that transports lipid A, the hydrophobic moiety of lipopolysaccharide (LPS), a glucosamine-based saccharolipid that makes up the outer monolayer of the outer membranes of most gram-negative bacteria. Lipid A is an endotoxin and so loss of MsbA from the cell membrane or mutations that disrupt transport results in the accumulation of lipid A in the inner cell membrane resulting to cell death. It is a close bacterial homolog of P-glycoprotein (Pgp) by protein sequence homology and has overlapping substrate specificities with the MDR-ABC transporter LmrA from Lactococcus lactis. [75] MsbA from E. coli is 36% identical to the NH2-terminal half of human MDR1, suggesting a common mechanism for transport of amphiphatic and hydrophobic substrates. The MsbA gene encodes a half transporter that contains a transmembrane domain (TMD) fused with a nucleotide-binding domain (NBD). It is assembled as a homodimer with a total molecular mass of 129.2 kD. MsbA contains 6 TMDs on the periplasmic side, an NBD located on the cytoplasmic side of the cell membrane, and an intracellular domain (ICD), bridging the TMD and NBD. This conserved helix extending from the TMD segments into or near the active site of the NBD is largely responsible for crosstalk between TMD and NBD. In particular, ICD1 serves as a conserved pivot about which the NBD can rotate, therefore allowing the NBD to disassociate and dimerize during ATP binding and hydrolysis. [4] [15] [18] [22] [43] [53] [54] [76]

Previously published (and now retracted) X-ray structures of MsbA were inconsistent with the bacterial homolog Sav1866. [77] [78] The structures were reexamined and found to have an error in the assignment of the hand resulting to incorrect models of MsbA. Recently, the errors have been rectified and new structures have been reported. [40] The resting state of E. coli MsbA exhibits an inverted "V" shape with a chamber accessible to the interior of the transporter suggesting an open, inward-facing conformation. The dimer contacts are concentrated between the extracellular loops and while the NBDs are ≈50Å apart, the subunits are facing each other. The distance between the residues in the site of the dimer interface have been verified by cross-linking experiments [79] and EPR spectroscopy studies. [80] The relatively large chamber allows it to accommodate large head groups such as that present in lipid A. Significant conformational changes are required to move the large sugar head groups across the membrane. The difference between the two nucleotide-free (apo) structures is the ≈30° pivot of TM4/TM5 helices relative to the TM3/TM6 helices. In the closed apo state (from V. cholerae MsbA), the NBDs are aligned and although closer, have not formed an ATP sandwich, and the P loops of opposing monomers are positioned next to one another. In comparison to the open conformation, the dimer interface of the TMDs in the closed, inward-facing conformation has extensive contacts. For both apo conformations of MsbA, the chamber opening is facing inward. The structure of MsbA-AMP-PNP (5’-adenylyl-β-γ-imidodiphosphate), obtained from S. typhimurium, is similar to Sav1866. The NBDs in this nucleotide-bound, outward-facing conformation, come together to form a canonical ATP dimer sandwich, that is, the nucleotide is situated in between the P-loop and LSGGQ motif. The conformational transition from MsbA-closed-apo to MsbA-AMP-PNP involves two steps, which are more likely concerted: a ≈10° pivot of TM4/TM5 helices towards TM3/TM6, bringing the NBDs closer but not into alignment followed by tilting of TM4/TM5 helices ≈20° out of plane. The twisting motion results in the separation of TM3/TM6 helices away from TM1/TM2 leading to a change from an inward- to an outward- facing conformation. Thus, changes in both the orientation and spacing of the NBDs dramatically rearrange the packing of transmembrane helices and effectively switch access to the chamber from the inner to the outer leaflet of the membrane. [40] The structures determined for MsbA is basis for the tilting model of transport. [18] The structures described also highlight the dynamic nature of ABC exporters as also suggested by fluorescence and EPR studies. [53] [80] [81] Recent work has resulted in the discovery of MsbA inhibitors. [82] [83]

Mechanism of transport for exporters Edit

ABC exporters have a transport mechanism that is consistent with both the alternating-access model and ATP-switch model. In the apo states of exporters, the conformation is inward-facing and the TMDs and NBDs are relatively far apart to accommodate amphiphilic or hydrophobic substrates. For MsbA, in particular, the size of the chamber is large enough to accommodate the sugar groups from lipopolysaccharides (LPS). As has been suggested by several groups, binding of substrate initiates the transport cycle. The "power stroke", that is, ATP binding that induces NBD dimerization and formation of the ATP sandwich, drives the conformational changes in the TMDs. In MsbA, the sugar head groups are sequestered within the chamber during the "power stroke". The cavity is lined with charged and polar residues that are likely solvated creating an energetically unfavorable environment for hydrophobic substrates and energetically favorable for polar moieties in amphiphilic compounds or sugar groups from LPS. Since the lipid cannot be stable for a long time in the chamber environment, lipid A and other hydrophobic molecules may "flip" into an energetically more favorable position within the outer membrane leaflet. The "flipping" may also be driven by the rigid-body shearing of the TMDs while the hydrophobic tails of the LPS are dragged through the lipid bilayer. Repacking of the helices switches the conformation into an outward-facing state. ATP hydrolysis may widen the periplasmic opening and push the substrate towards the outer leaflet of the lipid bilayer. Hydrolysis of the second ATP molecule and release of Pi separates the NBDs followed by restoration of the resting state, opening the chamber towards the cytoplasm for another cycle. [40] [43] [51] [54] [77] [78] [80] [84]

ABC transporters are known to play a crucial role in the development of multidrug resistance (MDR). In MDR, patients that are on medication eventually develop resistance not only to the drug they are taking but also to several different types of drugs. This is caused by several factors, one of which is increased expulsion of the drug from the cell by ABC transporters. For example, the ABCB1 protein (P-glycoprotein) functions in pumping tumor suppression drugs out of the cell. Pgp also called MDR1, ABCB1, is the prototype of ABC transporters and also the most extensively-studied gene. Pgp is known to transport organic cationic or neutral compounds. A few ABCC family members, also known as MRP, have also been demonstrated to confer MDR to organic anion compounds. The most-studied member in ABCG family is ABCG2, also known as BCRP (breast cancer resistance protein) confer resistance to most of Topoisomerase I or II inhibitors such as topotecan, irinotecan, and doxorubicin.

It is unclear exactly how these proteins can translocate such a wide variety of drugs, however one model (the hydrophobic vacuum cleaner model) states that, in P-glycoprotein, the drugs are bound indiscriminately from the lipid phase based on their hydrophobicity.

Discovery of the first eukaryotic ABC transporter protein came from studies on tumor cells and cultured cells that exhibited resistance to several drugs with unrelated chemical structures. These cells were shown to express elevated levels of multidrug-resistance (MDR) transport protein which was originally called P-glycoprotein (P-gp), but it is also referred to as multidrug resistance protein 1 (MDR1) or ABCB1. This protein uses ATP hydrolysis, just like the other ABC transporters, to export a large variety of drugs from the cytosol to the extracellular medium. In multidrug-resistant cells, the MDR1 gene is frequently amplified. This results in a large overproduction of the MDR1 protein. The substrates of mammalian ABCB1 are primarily planar, lipid-soluble molecules with one or more positive charges. All of these substrates compete with one another for transport, suggesting that they bind to the same or overlapping sites on the protein. Many of the drugs that are transported out by ABCB1 are small, nonpolar drugs that diffuse across the extracellular medium into the cytosol, where they block various cellular functions. Drugs such as colchicine and vinblastine, which block assembly of microtubules, freely cross the membrane into the cytosol, but the export of these drugs by ABCB1 reduces their concentration in the cell. Therefore, it takes a higher concentration of the drugs is required to kill the cells that express ABCB1 than those that do not express the gene. [10]

Other ABC transporters that contribute to multidrug resistance are ABCC1 (MRP1) and ABCG2 (breast cancer resistance protein). [85]

To solve the problems associated with multidrug-resistance by MDR1, different types of drugs can be used or the ABC transporters themselves must be inhibited. For other types of drugs to work they must bypass the resistance mechanism, which is the ABC transporter. To do this other anticancer drugs can be utilized such as alkylating drugs (cyclophosphamide), antimetabolites (5-fluorouracil), and the anthracycline modified drugs (annamycin and doxorubicin-peptide). These drugs would not function as a substrate of ABC transporters, and would thus not be transported. The other option is to use a combination of ABC inhibitory drugs and the anticancer drugs at the same time. This would reverse the resistance to the anticancer drugs so that they could function as intended. The substrates that reverse the resistance to anticancer drugs are called chemosensitizers. [8]

Drug resistance is a common clinical problem that occurs in patients suffering from infectious diseases and in patients suffering from cancer. Prokaryotic and eukaryotic microorganisms as well as neoplastic cells are often found to be resistant to drugs. MDR is frequently associated with overexpression of ABC transporters. Inhibition of ABC transporters by low-molecular weight compounds has been extensively investigated in cancer patients however, the clinical results have been disappointing. Recently various RNAi strategies have been applied to reverse MDR in different tumor models and this technology is effective in reversing ABC-transporter-mediated MDR in cancer cells and is therefore a promising strategy for overcoming MDR by gene therapeutic applications. RNAi technology could also be considered for overcoming MDR in infectious diseases caused by microbial pathogens. [86]

In addition to conferring MDR in tumor cells, ABC transporters are also expressed in the membranes of healthy cells, where they facilitate the transport of various endogenous substances, as well as of substances foreign to the body. For instance, ABC transporters such as Pgp, the MRPs and BCRP limit the absorption of many drugs from the intestine, and pump drugs from the liver cells to the bile [87] as a means of removing foreign substances from the body. A large number of drugs are either transported by ABC transporters themselves or affect the transport of other drugs. The latter scenario can lead to drug-drug interactions, [88] sometimes resulting in altered effects of the drugs. [89]

There are a number of assay types that allow the detection of ABC transporter interactions with endogenous and xenobiotic compounds. [90] The complexity of assay range from relatively simple membrane assays. [91] like vesicular transport assay, ATPase assay to more complex cell based assays up to intricate in vivo Jeffrey P, Summerfield SG (2007). "Challenges for blood-brain barrier (BBB) screening". Xenobiotica. 37 (10–11): 1135–51. doi:10.1080/00498250701570285. PMID 17968740. S2CID 25944548. detection methodologies. [92]

Membrane assays Edit

The vesicular transport assay detects the translocation of molecules by ABC transporters. [93] Membranes prepared under suitable conditions contain inside-out oriented vesicles with the ATP binding site and substrate binding site of the transporter facing the buffer outside. Substrates of the transporter are taken up into the vesicles in an ATP dependent manner. Rapid filtration using glass fiber filters or nitrocellulose membranes are used to separate the vesicles from the incubation solution and the test compound trapped inside the vesicles is retained on the filter. The quantity of the transported unlabelled molecules is determined by HPLC, LC/MS, LC/MS/MS. Alternatively, the compounds are radiolabeled, fluorescent or have a fluorescent tag so that the radioactivity or fluorescence retained on the filter can be quantified.

Various types of membranes from different sources (e.g. insect cells, transfected or selected mammalian cell lines) are used in vesicular transport studies. Membranes are commercially available or can be prepared from various cells or even tissues e.g. liver canalicular membranes. This assay type has the advantage of measuring the actual disposition of the substrate across the cell membrane. Its disadvantage is that compounds with medium-to-high passive permeability are not retained inside the vesicles making direct transport measurements with this class of compounds difficult to perform.

The vesicular transport assay can be performed in an "indirect" setting, where interacting test drugs modulate the transport rate of a reporter compound. This assay type is particularly suitable for the detection of possible drug-drug interactions and drug-endogenous substrate interactions. It is not sensitive to the passive permeability of the compounds and therefore detects all interacting compounds. Yet, it does not provide information on whether the compound tested is an inhibitor of the transporter, or a substrate of the transporter inhibiting its function in a competitive fashion. A typical example of an indirect vesicular transport assay is the detection of the inhibition of taurocholate transport by ABCB11 (BSEP).

Whole cell based assays Edit

Efflux transporter-expressing cells actively pump substrates out of the cell, which results in a lower rate of substrate accumulation, lower intracellular concentration at steady state, or a faster rate of substrate elimination from cells loaded with the substrate. Transported radioactive substrates or labeled fluorescent dyes can be directly measured, or in an indirect set up, the modulation of the accumulation of a probe substrate (e.g. fluorescent dyes like rhodamine 123, or calcein) can be determined in the presence of a test drug. [88]

Calcein-AM, A highly permeable derivative of calcein readily penetrates into intact cells, where the endogenous esterases rapidly hydrolyze it to the fluorescent calcein. In contrast to calcein-AM, calcein has low permeability and therefore gets trapped in the cell and accumulates. As calcein-AM is an excellent substrate of the MDR1 and MRP1 efflux transporters, cells expressing MDR1 and/or MRP1 transporters pump the calcein-AM out of the cell before esterases can hydrolyze it. This results in a lower cellular accumulation rate of calcein. The higher the MDR activity is in the cell membrane, the less Calcein is accumulated in the cytoplasm. In MDR-expressing cells, the addition of an MDR inhibitor or an MDR substrate in excess dramatically increases the rate of Calcein accumulation. Activity of multidrug transporter is reflected by the difference between the amounts of dye accumulated in the presence and the absence of inhibitor. Using selective inhibitors, transport activity of MDR1 and MRP1 can be easily distinguished. This assay can be used to screen drugs for transporter interactions, and also to quantify the MDR activity of cells. The calcein assay is the proprietary assay of SOLVO Biotechnology.

Mammalian subfamilies Edit

There are 49 known ABC transporters present in humans, which are classified into seven families by the Human Genome Organization.

Family Members Function Examples
ABCA This family contains some of the largest transporters (over 2,100 amino acids long). Five of them are located in a cluster in the 17q24 chromosome. Responsible for the transportation of cholesterol and lipids, among other things. ABCA12 ABCA1
ABCB Consists of 4 full and 7 half transporters. Some are located in the blood–brain barrier, liver, mitochondria, transports peptides and bile, for example. ABCB5
ABCC Consists of 12 full transporters. Used in ion transport, cell-surface receptors, toxin secretion. Includes the CFTR protein, which causes cystic fibrosis when deficient. ABCC6
ABCD Consists of 4 half transporters Are all used in peroxisomes. ABCD1
ABCE/ABCF Consists of 1 ABCE and 3 ABCF proteins. These are not actually transporters but merely ATP-binding domains that were derived from the ABC family, but without the transmembrane domains. These proteins mainly regulate protein synthesis or expression. ABCE1, ABCF1, ABCF2
ABCG Consists of 6 "reverse" half-transporters, with the NBF at the NH3 + end and the TM at the COO- end. Transports lipids, diverse drug substrates, bile, cholesterol, and other steroids. ABCG2 ABCG1

A full list of human ABC transporters can be found from. [94]


The ABCA subfamily is composed of 12 full transporters split into two subgroups. The first subgroup consists of seven genes that map to six different chromosomes. These are ABCA1, ABCA2, ABCA3, and ABCA4, ABCA7, ABCA12, and ABCA13. The other subgroup consists of ABCA5 and ABCA6 and ABCA8, ABCA9 and ABCA10. A8-10. All of subgroup 2 is organized into a head to tail cluster of chromosomes on chromosome 17q24. Genes in this second subgroup are distinguished from ABCA1-like genes by having 37-38 exons as opposed to the 50 exons in ABCA1. The ABCA1 subgroup is implicated in the development of genetic diseases. In the recessive Tangier's disease, the ABCA1 protein is mutated. Also, the ABCA4 maps to a region of chromosome 1p21 that contains the gene for Stargardt's disease. This gene is found to be highly expressed in rod photoreceptors and is mutated in Stargardt's disease, recessive retinitis pigmentism, and the majority of recessive cone-rod dystrophy. [9]


The ABCB subfamily is composed of four full transporters and two half transporters. This is the only human subfamily to have both half and full types of transporters. ABCB1 was discovered as a protein overexpressed in certain drug resistant tumor cells. It is expressed primarily in the blood–brain barrier and liver and is thought to be involved in protecting cells from toxins. Cells that overexpress this protein exhibit multi-drug resistance. [9]


Subfamily ABCC contains thirteen members and nine of these transporters are referred to as the Multidrug Resistance Proteins (MRPs). The MRP proteins are found throughout nature and they mediate many important functions. [95] They are known to be involved in ion transport, toxin secretion, and signal transduction. [9] Of the nine MRP proteins, four of them, MRP4, 5, 8, 9, (ABCC4, 5, 11, and 12), have a typical ABC structure with four domains, comprising two membrane spanning domains, with each spanning domain followed by a nucleotide binding domain. These are referred to as short MRPs. The remaining 5 MRP's (MRP1, 2, 6, 7 (ABCC1, 2, 3, 6 and 10) are known as long MRPs and feature an additional fifth domain at their N terminus. [95]

CFTR, the transporter involved in the disease cystic fibrosis, is also considered part of this subfamily. Cystic fibrosis occurs upon mutation and loss of function of CFTR. [9]

The sulfonylurea receptors (SUR), involved in insulin secretion, neuronal function, and muscle function, are also part of this family of proteins. Mutations in SUR proteins are a potential cause of Neonatal diabetes mellitus. SUR is also the binding site for drugs such as sulfonylureas and potassium-channel openers activators such as diazoxide.


The ABCD subfamily consists of four genes that encode half transporters expressed exclusively in the peroxisome. ABCD1 is responsible for the X-linked form of Adrenoleukodystrophy (ALD) which is a disease characterized by neurodegeneration and adrenal deficiency that typically is initiated in late childhood. The cells of ALD patients feature accumulation of unbranched saturated fatty acids, but the exact role of ABCD1 in the process is still undetermined. In addition, the function of other ABCD genes have yet to be determined but have been thought to exert related functions in fatty acid metabolism. [9]

ABCE and ABCF Edit

Both of these subgroups are composed of genes that have ATP binding domains that are closely related to other ABC transporters, but these genes do not encode for trans-membrane domains. ABCE consists of only one member, OABP or ABCE1, which is known to recognize certain oligodendrocytes produced in response to certain viral infections. Each member of the ABCF subgroup consist of a pair of ATP binding domains. [9]


Six half transporters with ATP binding sites on the N terminus and trans-membrane domains at the C terminus make up the ABCG subfamily. This orientation is opposite of all other ABC genes. There are only 5 ABCG genes in the human genome, but there are 15 in the Drosophila genome and 10 in yeast. The ABCG2 gene was discovered in cell lines selected for high level resistance for mitoxantrone and no expression of ABCB1 or ABCC1. ABCG2 can export anthrocycline anticancer drugs, as well as topotecan, mitoxantrone, or doxorubicin as substrates. Chromosomal translocations have been found to cause the ABCG2 amplification or rearrangement found in resistant cell lines. The normal function of ABCG2 is not known. [9]

Cross-species subfamilies Edit

The following classification system for transmembrane solute transporters has been constructed in the TCDB. [96]

Three families of ABC exporters are defined by their evolutionary origins. [6] ABC1 exporters evolved by intragenic triplication of a 2 TMS precursor (TMS = transmembrane segment. A "2 TMS" protein has 2 transmembrane segments) to give 6 TMS proteins. ABC2 exporters evolved by intragenic duplication of a 3 TMS precursor, and ABC3 exporters evolved from a 4 TMS precursor which duplicated either extragenicly to give two 4 TMS proteins, both required for transport function, or intragenicly to give 8 or 10 TMS proteins. The 10 TMS proteins appear to have two extra TMSs between the two 4 TMS repeat units. [97] Most uptake systems (all except 3.A.1.21) are of the ABC2 type, divided into type I and type II by the way they handle nucleotides. A special subfamily of ABC2 importers called ECF use a separate subunit for substrate recognition. [98]

  • 3.A.1.106 The Lipid Exporter (LipidE) Family
  • 3.A.1.108 The β-Glucan Exporter (GlucanE) Family
  • 3.A.1.109 The Protein-1 Exporter (Prot1E) Family
  • 3.A.1.110 The Protein-2 Exporter (Prot2E) Family
  • 3.A.1.111 The Peptide-1 Exporter (Pep1E) Family
  • 3.A.1.112 The Peptide-2 Exporter (Pep2E) Family
  • 3.A.1.113 The Peptide-3 Exporter (Pep3E) Family
  • 3.A.1.117 The Drug Exporter-2 (DrugE2) Family
  • 3.A.1.118 The Microcin J25 Exporter (McjD) Family
  • 3.A.1.119 The Drug/Siderophore Exporter-3 (DrugE3) Family
  • 3.A.1.123 The Peptide-4 Exporter (Pep4E) Family
  • 3.A.1.127 The AmfS Peptide Exporter (AmfS-E) Family
  • 3.A.1.129 The CydDC Cysteine Exporter (CydDC-E) Family
  • 3.A.1.135 The Drug Exporter-4 (DrugE4) Family
  • 3.A.1.139 The UDP-Glucose Exporter (U-GlcE) Family (UPF0014 Family)
  • 3.A.1.201 The Multidrug Resistance Exporter (MDR) Family (ABCB)
  • 3.A.1.202 The Cystic Fibrosis Transmembrane Conductance Exporter (CFTR) Family (ABCC)
  • 3.A.1.203 The Peroxysomal Fatty Acyl CoA Transporter (P-FAT) Family (ABCD)
  • 3.A.1.206 The a-Factor Sex Pheromone Exporter (STE) Family (ABCB)
  • 3.A.1.208 The Drug Conjugate Transporter (DCT) Family (ABCC) (Dębska et al., 2011)
  • 3.A.1.209 The MHC Peptide Transporter (TAP) Family (ABCB)
  • 3.A.1.210 The Heavy Metal Transporter (HMT) Family (ABCB)
  • 3.A.1.212 The Mitochondrial Peptide Exporter (MPE) Family (ABCB)
  • 3.A.1.21 The Siderophore-Fe3+ Uptake Transporter (SIUT) Family
  • 3.A.1.101 The Capsular Polysaccharide Exporter (CPSE) Family
  • 3.A.1.102 The Lipooligosaccharide Exporter (LOSE) Family
  • 3.A.1.103 The Lipopolysaccharide Exporter (LPSE) Family
  • 3.A.1.104 The Teichoic Acid Exporter (TAE) Family
  • 3.A.1.105 The Drug Exporter-1 (DrugE1) Family
  • 3.A.1.107 The Putative Heme Exporter (HemeE) Family
  • 3.A.1.115 The Na+ Exporter (NatE) Family
  • 3.A.1.116 The Microcin B17 Exporter (McbE) Family
  • 3.A.1.124 The 3-component Peptide-5 Exporter (Pep5E) Family
  • 3.A.1.126 The β-Exotoxin I Exporter (βETE) Family
  • 3.A.1.128 The SkfA Peptide Exporter (SkfA-E) Family
  • 3.A.1.130 The Multidrug/Hemolysin Exporter (MHE) Family
  • 3.A.1.131 The Bacitracin Resistance (Bcr) Family
  • 3.A.1.132 The Gliding Motility ABC Transporter (Gld) Family
  • 3.A.1.133 The Peptide-6 Exporter (Pep6E) Family
  • 3.A.1.138 The Unknown ABC-2-type (ABC2-1) Family
  • 3.A.1.141 The Ethyl Viologen Exporter (EVE) Family (DUF990 Family InterPro: IPR010390)
  • 3.A.1.142 The Glycolipid Flippase (G.L.Flippase) Family
  • 3.A.1.143 The Exoprotein Secretion System (EcsAB(C))
  • 3.A.1.144: Functionally Uncharacterized ABC2-1 (ABC2-1) Family
  • 3.A.1.145: Peptidase Fused Functionally Uncharacterized ABC2-2 (ABC2-2) Family
  • 3.A.1.146: The actinorhodin (ACT) and undecylprodigiosin (RED) exporter (ARE) family
  • 3.A.1.147: Functionally Uncharacterized ABC2-2 (ABC2-2) Family
  • 3.A.1.148: Functionally Uncharacterized ABC2-3 (ABC2-3) Family
  • 3.A.1.149: Functionally Uncharacterized ABC2-4 (ABC2-4) Family
  • 3.A.1.150: Functionally Uncharacterized ABC2-5 (ABC2-5) Family
  • 3.A.1.151: Functionally Uncharacterized ABC2-6 (ABC2-6) Family
  • 3.A.1.152: The lipopolysaccharide export (LptBFG) Family (InterPro: IPR005495)
  • 3.A.1.204 The Eye Pigment Precursor Transporter (EPP) Family (ABCG)
  • 3.A.1.205 The Pleiotropic Drug Resistance (PDR) Family (ABCG)
  • 3.A.1.211 The Cholesterol/Phospholipid/Retinal (CPR) Flippase Family (ABCA)
  • 9.B.74 The Phage Infection Protein (PIP) Family
  • all uptake systems (3.A.1.1 - 3.A.1.34 except 3.A.1.21)
    • 3.A.1.1 Carbohydrate Uptake Transporter-1 (CUT1)
    • 3.A.1.2 Carbohydrate Uptake Transporter-2 (CUT2)
    • 3.A.1.3 Polar Amino Acid Uptake Transporter (PAAT)
    • 3.A.1.4 Hydrophobic Amino Acid Uptake Transporter (HAAT)
    • 3.A.1.5 Peptide/Opine/Nickel Uptake Transporter (PepT)
    • 3.A.1.6 Sulfate/Tungstate Uptake Transporter (SulT)
    • 3.A.1.7 Phosphate Uptake Transporter (PhoT)
    • 3.A.1.8 Molybdate Uptake Transporter (MolT)
    • 3.A.1.9 Phosphonate Uptake Transporter (PhnT)
    • 3.A.1.10 Ferric Iron Uptake Transporter (FeT)
    • 3.A.1.11 Polyamine/Opine/Phosphonate Uptake Transporter (POPT)
    • 3.A.1.12 Quaternary Amine Uptake Transporter (QAT)
    • 3.A.1.13 Vitamin B12 Uptake Transporter (B12T)
    • 3.A.1.14 Iron Chelate Uptake Transporter (FeCT)
    • 3.A.1.15 Manganese/Zinc/Iron Chelate Uptake Transporter (MZT)
    • 3.A.1.16 Nitrate/Nitrite/Cyanate Uptake Transporter (NitT)
    • 3.A.1.17 Taurine Uptake Transporter (TauT)
    • 3.A.1.19 Thiamin Uptake Transporter (ThiT)
    • 3.A.1.20 Brachyspira Iron Transporter (BIT)
    • 3.A.1.21 Siderophore-Fe3+ Uptake Transporter (SIUT)
    • 3.A.1.24 The Methionine Uptake Transporter (MUT) Family (Similar to 3.A.1.3 and 3.A.1.12)
    • 3.A.1.27 The γ-Hexachlorocyclohexane (HCH) Family (Similar to 3.A.1.24 and 3.A.1.12)
    • 3.A.1.34 The Tryptophan (TrpXYZ) Family
    • ECF uptake systems
      • 3.A.1.18 The Cobalt Uptake Transporter (CoT) Family
      • 3.A.1.22 The Nickel Uptake Transporter (NiT) Family
      • 3.A.1.23 The Nickel/Cobalt Uptake Transporter (NiCoT) Family
      • 3.A.1.25 The Biotin Uptake Transporter (BioMNY) Family
      • 3.A.1.26 The Putative Thiamine Uptake Transporter (ThiW) Family
      • 3.A.1.28 The Queuosine (Queuosine) Family
      • 3.A.1.29 The Methionine Precursor (Met-P) Family
      • 3.A.1.30 The Thiamin Precursor (Thi-P) Family
      • 3.A.1.31 The Unknown-ABC1 (U-ABC1) Family
      • 3.A.1.32 The Cobalamin Precursor (B12-P) Family
      • 3.A.1.33 The Methylthioadenosine (MTA) Family
      • 3.A.1.114 The Probable Glycolipid Exporter (DevE) Family
      • 3.A.1.122 The Macrolide Exporter (MacB) Family
      • 3.A.1.125 The Lipoprotein Translocase (LPT) Family
      • 3.A.1.134 The Peptide-7 Exporter (Pep7E) Family
      • 3.A.1.136 The Uncharacterized ABC-3-type (U-ABC3-1) Family
      • 3.A.1.137 The Uncharacterized ABC-3-type (U-ABC3-2) Family
      • 3.A.1.140 The FtsX/FtsE Septation (FtsX/FtsE) Family
      • 3.A.1.207 The Eukaryotic ABC3 (E-ABC3) Family

      View Proteins belonging to ABC Superfamily : here

      Many structures of water-soluble domains of ABC proteins have been produced in recent years. [2]

      E. Structures involved in Na + transport

      In order to function as a selective Na + pump, Na + -NQR must contain structures that 1) allow Na + to pass through the hydrophobic core of the membrane and 2) provide cation specificity to the translocation system. In other Na + transporting proteins, the structures that carry out these roles frequently include aspartate and glutamate residues [83]. The negative charge of these amino acids facilitates binding of the positively charged Na + . To identify residues that could be involved in this function, we used the data from our topological analysis to locate conserved acidic residues in the transmembrane segments of the enzyme. Seventeen of these residues were found, all of them in subunits NqrB, D and E. Interestingly, all the acid residues located in transmembrane helices are close to either the cytoplasmic or periplasmic side of the membrane. There is little indication of a pathway of binding sites to transport Na + through the center of the membrane. To investigate the roles of these acid groups, we constructed mutants in which these residues were individually replaced by aliphatic groups, which should interact poorly with Na + . Of the seventeen mutants, seven showed significant inhibition of their catalytic activity [64]. These seven mutants can be divided into two groups, depending on the functional changes that result from the mutation. This grouping correlates with the side of the membrane on which the mutated residue is situated.

      For one group of residues, located near the cytosol (NqrB-E144, NqrB-D397, NqrD-D133 and NqrE-E95), replacement of any of the acid groups by an aliphatic one, leads to a decrease of 10 times or more in the apparent affinity for Na + in steady-state conditions. On this bases, these resides were proposed to be part of the cation binding site(s).

      NqrB-D397 is likely the most important ligand, since the turnover rate of the NqrB-D397A mutant shows no saturating behavior with respect to Na + . This indicates either that the mutant has an extremely low affinity for Na + , or that sodium uptake is so severely impeded that it is always rate limiting.

      For the second group of residues, located near the periplasm (positive side of the membrane) (NqrB-E28, NqrB-D346 and NqrD-D88) replacement by an aliphatic amino acid, resulted in a large decrease in the quinone reductase activity of the enzyme and its Na + sensitivity, but almost no effect on the apparent Na + affinity. This is consistent with a role for these residues in an exit pathway through the membrane for Na + .

      The properties of these mutants suggest that Na + translocation and the electron transfer reactions are tightly connected.

      F. Coupling mechanism

      F.1. Models of Na + pumping based on direct coupling

      Rich, Dimroth and Bogachev [56, 84, 85] have all proposed models in which the reduction of a single cofactor (the 2Fe-2S center, ubiquinone or a flavin cofactor, respectively) is directly coupled to the uptake of sodium from the cytoplasm, and the oxidation of the cofactor is coupled to ejection of Na + to the periplasm. These schemes are derived from models of redox-driven H + pumps where the site of H + binding is typically part of the redox cofactor itself. Their operating principle has been described as the preservation of 𠇎lectroneutrality” in the vicinity of the redox cofactor—in effect, that coupling arises from a coulombic interaction between the electron and the H + , or in the case of Na + -NQR, between the electron and the Na + ion.

      The 𠇍irectly coupled” pumping mechanisms of these models can be described as “local” in the sense that the connection between the redox events and the uptake and release of Na + operates over short distances through essentially chemical interactions, rather than through conformation changes of the protein. Mechanisms like this would also be expected to have a high degree of “thermodynamic coupling,” meaning that the affinity of the binding site for Na + is controlled by the redox state of the cofactor.

      The main experimental support for thermodynamic coupling in Na + -NQR comes from NMR studies in which Bogachev et al [86] measured the linewidth of the 23 Na band from Na+ in solution, in the presence of reduced and oxidized forms of Na + -NQR. If Na + in solution is in rapid exchange with Na + bound to an enzyme, the 23 Na resonance will be broadened, with the degree of linebroadening a function of the binding affinity. The authors reported that the 23 Na resonances from both the reduced and oxidized enzyme samples were broadened compared to the signal from a sample without enzyme, but that the linewidth in the reduced enzyme sample was significantly larger than in the oxidized one. They calculated that the binding affinity of the enzyme for sodium is 24 mM in the oxidized form and as high as 30 μM in the reduced form.[

      An interaction of this kind must be reciprocal if reduction of one of the cofactors causes affinity for Na + to increase, the binding of Na + will stabilize the reduced form of the cofactor. In the case of the strong coupling, expected for directly coupled systems, the redox-midpoint-potential of the cofactor will increase by 60 mV for every 10-fold increase in Na + concentration. However, Bogachev and coworkers also carried out redox titrations of Na + -NQR at different Na + concentrations, and found no apparent dependence [80]. More recently, direct equilibrium measurements of the binding of Na + to Na + -NQR, made in our laboratory, using 22 Na + found no redox dependent changes in either the number of Na + bound to Na + -NQR or the affinity of this interaction [48]. It is important to note that the NMR method depends on rapid exchange of Na + between solution and the binding site [86], a requirement that may not have been met under the experimental conditions tested. Thus, it is possible that the changes in the linewidth of the NMR spectra reflect a change in the kinetics of Na + binding, and not change affinity.

      It also evident that driving force is distributed fairly evenly along the steps of the enzyme’s electron transfer pathway and that there is no cofactor whose reduction and oxidation are highly exergonic, as might be expected for a mechanism of the kind envisioned by the earlier models.

      This failure, thus far, to find evidence of thermodynamic coupling between the redox and Na + reactions of Na + -NQR, argues against directly coupled models like those cited above. An alternative is indirect coupling in which the redox reactions and the capture of Na + take place at separated sites and are only connected through conformational changes of the protein. In such a scheme, the redox reaction could be accelerated, not by modification of the thermodynamic properties of the redox centers but through a decrease in activation barriers linked to steps in Na + translocation.

      F.2. Functional studies to identify coupled-reaction steps

      A crucial point to understand the mechanism of Na + -NQR is to determine the redox steps that are linked to the pumping of Na + across the membrane. Our group addressed this question by analyzing the electron transfer kinetics in some of the acid group mutants described above that are unable to transport Na + [81]. We have previously shown that NqrB-D397 forms part of a sodium-uptake/binding site and that NqrB-D346 is likely involved in a sodium exit pathway [64]. In the case of NqrB-D397A, the 2Fe-2S𡤯MNC step is severely impaired this is also the main Na + dependent step in the reaction [49]. This indicates the reduction of FMNC is directly involved in the capture of sodium from solution. In the case of the mutant NqrB-D346A, the electron flow is inhibited at the FMNB → Riboflavin step, which indicates that this step is involved in the release of sodium to the periplasm. This was corroborated by “single-turnover ΔΨ” measurements. In these experiments, a voltage-sensitive dye was used to follow the generation of membrane potential during the reduction of the enzyme by NADH. First, the reaction was carried out by adding NADH in the presence of ubiquinone, which led to steady-state turnover, accompanied by a clear formation of ΔΨ. Next, the experiment was repeated without ubiquinone, so the reaction ended after all cofactors were reduced. This reaction produced a smaller ΔΨ, but its amplitude was significant, showing that reduction of the enzyme alone, without oxidation by quinone, is sufficient to generate ΔΨ. Incubation of the enzyme with ubiquinol has been shown to reduce only Riboflavin, leaving the upstream cofactors oxidized. When this is done prior to addition of NADH, it has the effect of preventing the electron transfer from FMNB to Riboflavin. This truncated redox reaction produced essentially no ΔΨ. Together with the previous results, this indicates that electron transfer from FMNB to Riboflavin is coupled to Na + pumping. These measurements were also carried out on some of the Na + -NQR mutants that lack specific cofactors. In all cases, truncating the reduction reaction at an earlier point abolished any generation of ΔΨ. The overall picture that emerges from these results is that Na + -NQR does not operate through a single side coupling mechanism. Na + is taken up from solution during the 2Fe-2S 𡤯MNC step and is subsequently transported across the membrane dielectric and released into solution when FMNB transfers electrons to Riboflavin ( Fig.4 ). Together, these results argue against any important role of direct coupling in the mechanism of Na + -NQR.

      Hirst [87] has pointed out that locally coupled mechanisms are not consistent with the understood requirements of Na + -binding to proteins. In particular, the interaction between the reduction process and ion binding is unlikely to be strong enough to support ion translocation. Direct coupling cannot explain how the enzyme prevents protons from binding to the 𠇌oupling cofactor” -semiquinone radical- in other words, how the enzyme is selective for sodium. Furthermore, the conditions of a low dielectric environment, necessary for the electroneutral capture of sodium, might not be found in biological sodium binding structures. While protons can be easily taken by a single electronegative atom, producing an intermediate with no net charge, the binding of sodium typically occurs in highly hydrophilic environments, at multi-dentate binding sites consisting of six polar residues [88]. This is consistent with our finding that in Na + -NQR at least three acid residues play important roles in the Na + -binding site(s). Also, the lack of pH dependence of the cofactor midpoint potentials strongly indicates that the cofactors are not accessible to the aqueous environment and consequently to sodium. Our analysis of cation selectivity showed that the enzyme has the same affinity for sodium and lithium, but at saturating concentrations, lithium stimulates the activity only 1/3 as much as sodium. Localized coupling cannot account for the cation selectivity of the enzyme, because under this mechanism any cation that has access to the 𠇌oupling cofactor” should produce the same effect as the natural coupling ion, in this case sodium.

      Thus, there is mounting evidence that the direct coupling mechanism is not compatible with the physical properties of sodium. All of this suggests that Na + -NQR has a novel mechanism that operates according to different principles than those usually invoked to explain H + pumps, and it is possible that this enzyme has more in common with other, non-redox driven Na + transporters than with its fellow respiratory complexes.

      F.3. Indirect mechanism of coupling

      We propose that the mechanism of Na + -NQR has several novel features for a redox-driven enzyme. 1) Rather than occurring at a single site, coupling between the redox reactions and Na + translocation involves at least two distinct electron transfer steps, which do not share any common cofactor: Na + uptake occurs during the 2Fe-2S → FMNC redox step, while movement of Na + across the membrane, it likely its release into the periplasm, occur during the FMNB → Riboflavin redox step 2) Coupling between the redox reactions and Na + pumping is indirect, and likely to be mediated by conformational changes of the enzyme. In the FMNB → Riboflavin step, during which generation of membrane potential takes place, the acid residues that facilitate Na + movement are located near the periplasmic side of the membrane, whereas the cofactors were the redox reaction takes place are on the cytoplasmic side of the membrane, too great a separation for a direct interaction. This is consistent with the case, described above, that the chemistry of Na + -protein interactions is sufficiently different from that of H + -protein interactions as to make direct coupling unlikely. The paucity of evidence for interactions between the redox reactions and Na + in equilibrium conditions, argues against the thermodynamic linkage that might be expected for a directly coupled pump mechanism it is however possible that thermodynamic linkage occurs, but is only manifest in kinetic intermediates that are not accessible in equilibrium conditions. 3) As in other Na + translocating enzymes, movement of the Na + through the hydrophobic membrane is likely to be facilitated by conformational changes of the protein (which already appear to be needed for coupling) as well as internal water cavities. A scheme incorporating these features is shown in Figure 6 . These new ideas about the mechanism of Na + -NQR are the subject of current investigations in our group.

      Schematic representation of conformational changes involved in Na + translocation during the catalytic mechanism of Na + -NQR. Sodium capture: electron transfer from 2Fe-2S center to FMNC controls Na + uptake from the cytosol. Sodium release: electron transfer from FMNB to Riboflavin controls Na + transport and release to the periplasm.


      The Na + -pumping NADH:quinone oxidoreductase (Na + -NQR) is a respiratory enzyme that is essential in the metabolism of many marine and pathogenic bacteria that uniquely translocates Na + instead of H + . The mechanism of Na + pumping in Na + -NQR is novel recent results show that coupling of Na + transport to redox reactions is indirect and thus likely mediated by conformational changes.


      In this study, we exploited differences between human and rat URAT1 to reveal molecular interactions with the substrate urate and with clinically relevant URAT1 inhibitors, compounds that reduce sUA levels for the treatment of gout. Using hURAT1 and rURAT1 chimeras, we were able to identify hURAT1 residues that confer high affinity interaction with URAT1 inhibitors. Residues responsible for the high affinity of hURAT1 to inhibitors include Ser-35 in TM1, Phe-365 in TM7 and Ile-481 in TM11, which correspond to Asn, Tyr and Met, respectively, in rURAT1. For urate, hURAT1 Ser-35 and Phe-365 mediate high affinity interaction. Urate and inhibitors interact with a common binding site on URAT1 because all require similar URAT1 residues for high affinity interaction. The point mutants in rURAT1 that carry the individual hURAT1 residues Ser-35, Phe-365 and Ile-481 all increase the potency of the inhibitors. These ‘gain of function’ phenotypes provide the strongest evidence that these human URAT1 residues are important for interactions with both inhibitors and substrate.

      Human URAT1 Ser-35, Phe-365, and Ile-481 were identified in an unbiased screen for URAT1 amino acids involved in affinity for inhibitors. These three amino acids co-localize in the central channel of a computer model of human OAT1 33 , a homolog of hURAT1 that also transports urate 38 and is inhibited by probenecid 41 . Furthermore, chimeric point mutant combinations of these residues produced additive phenotypes for affinity to inhibitors and urate, suggesting they form a common binding site for URAT1 substrates and inhibitors within the transporter channel. This notion is supported by recent findings from the crystal structures of the GLUT glucose transporters 42,43,44 , SLC family members that have weak sequence homology but are structurally related to URAT1 through the MFS fold 45 . These studies show that amino acid sidechains in TM1, TM7 and TM11 (as well as residues in other TM segments) directly contact substrates for the GLUTs. Furthermore, in the inward-open conformation of GLUT1 and GLUT5, the extracellular sides of TM1 and TM7 contact each other 42 , consistent with the predicted close proximity of URAT1 Ser-35 and Phe-365 in the OAT1 computer model, which is in the same conformation. In another computer model of OAT1, simulated conformational changes were observed in the extracellular sides of TM1 and TM7 46 , suggesting a possible role for substrate transport for residues within these domains. The precise positioning of these URAT1 residues within the protein is speculative and awaits clarification through determination of a URAT1 crystal structure.

      We predict that these URAT1 residues directly contact both substrates and inhibitors. In support of this hypothesis, we developed a novel and specific human URAT1 binding assay. All inhibitors displaced binding of the radiolabeled probe, suggesting that our functional mapping data uncovered a specific binding site within URAT1. This binding assay offers a new tool for further characterizing the molecular interactions of compounds and substrates to URAT1. All uricosuric agents we have tested to date interact with these amino acids within URAT1. The inhibitors themselves were not identified and developed through standard medicinal chemistry approaches and so are structurally very diverse. We believe the commonality of binding may be due to the unique properties of the residues in this region facilitating inhibitor interactions.

      Although the precise nature of these interactions is unknown, rURAT1 may have a lower affinity for urate and the inhibitors because the residues at positions 35, 365 and 481 are bulkier than the corresponding hURAT1 residues therefore, steric hindrance may reduce the affinity of interactions with the rat transporter. Residue 365 occurs in a cluster of aromatic residues in TM7 that are highly conserved in the SLC22 transporter family, a domain shown to be important in substrate interactions for OAT1 and OAT3 29,30 . However, unlike URAT1 in which both Phe-365 and Tyr-365 support transport activity, the corresponding residues in human OAT1 and rat OAT3, Tyr-354 and Tyr-352, are strictly required for substrate recognition. Mutations to phenylalanine are inactive 29,30 , showing that substrate recognition occurs through hydrophilic contacts with the tyrosine hydroxyl groups. It therefore appears that recognition of urate and URAT1 inhibitors through residue 365 is mechanistically different, possibly occurring through hydrophobic interactions between aromatic moieties. The hydroxyl group of Tyr-365 of rURAT1 may sterically hinder this hydrophobic interaction to reduce affinity.

      Previously, we reported that Phe-365 and Met-25 were acquired during the evolution of simians (humans, apes, Old World monkeys, and New World monkeys), and that these residues promote higher urate affinity in simian URAT1, relative to non-simian URAT1, which carry Tyr-365 and Val-25 40 (Supplementary Table 4). Residues 35 and 481 have distinct phylogenetic distributions (Supplementary Table 4) but also differ between human, rat, and mouse, and so were also identified in analyses of human-to-rat URAT1 chimeras. We expect that other residues are also involved in urate and inhibitor affinity. Because hURAT1 and rURAT1 share a 74% amino acid identity, chimeras from these orthologs will not identify all residues involved in inhibitor binding. Based on structure/function analysis of URAT1 homologs 31,33,45,46,47,48,49 as well as from recent findings from GLUT crystal structures 41,44 , we expect that conserved residues in many TM segments also play a role in binding to URAT1 substrates and inhibitors. The identity of residues corresponding to human URAT1 residues 35, 365, and 481 in other URAT1 species (orthologs) and in SLC22A subfamily homologs is shown in Supplementary Table 4. Interestingly, a tyrosine residue occurs in most homologs at the position corresponding to hURAT1 residue 365, so that Phe-365 is nearly unique to hURAT1. Therefore, this phenylalanine may be important in the high potency and specificity of benzbromarone and verinurad for hURAT1 (Tan et al., manuscripts submitted). However, probenecid is more non-specific and has a similar potency to hURAT1, hOAT4, hOAT1, and hOAT3 24 consistent with a finding that URAT1 residues 35, 365, and 481 all occur within sequence motifs common to all SLC22A family members 49 .

      In summary, we have identified several amino acids in hURAT1 that mediate the high affinity interaction with URAT1 inhibitors. Some of these residues also participate in the recognition and affinity for the URAT1 substrate uric acid. This provides a facile mechanism for inhibition of URAT1: inhibitors sterically hinder the interaction of urate with key amino acids within the central channel of URAT1 to prevent uric acid transport. Naturally occurring polymorphisms in these amino acids could in principle impact the efficacy of URAT1 inhibitors, though none have been identified to date. These results could also assist in the discovery of new high affinity and specific inhibitors of URAT1, which may also serve as safer and more effective urate-lowering therapies for hyperuricemia and gout.

      Biology of ocular transporters: efflux and influx transporters in the eye

      Dhananjay Pal , . Ashim K. Mitra , in Ocular Transporters and Receptors , 2013

      2.2.1 Peptide transporters

      Peptide transporters are important plasma membrane proteins that facilitate the cellular translocation of dipeptides and tripeptides in addition to a variety of peptidomimetic molecules such as angiotensin-converting enzyme inhibitors, rennin inhibitors, β-lactam antibiotics and cephalosporins. The acidic pH generated by Na + /H + exchanger serves as the dynamic force for the absorption of peptides. So far two peptide transporters have been identified: peptide transporter 1 (PEPT1, SLC15A1) and peptide transporter 2 (PEPT2, SLC15A2) [ 6–10 ]. These transporters share similar topology and contain 12 predicted membrane-spanning domains (transmembrane domains TMDs) with N- and C-termini oriented towards the cytosol ( Figure 2.1 ). The protein PEPT1 encodes for 708 amino acid residues whereas PEPT2 encodes for 729 amino acid residues. These transport systems possess the unique ability to transport dipeptides and tripeptides (independent of sequence), including differently charged species. These transporters are stereospecific and encompass high affinity for L-enantiomers of amino acid residues relative to peptides with one or more D-enantiomers. Peptide transporters are considered the first mammalian nutrient membrane transporters to use an electrochemical proton gradient as their driving force [ 11 ].

      Figure 2.1 . Membrane topology of peptide transporter 1 (PEPT1). (a) The protein contains 12 transmembrane domains, with the N-terminal and C-terminal ends in the cytosol. (b) Based on the analysis of chimeric transporters derived from PEPT1 and PEPT2, transmembrane domains in green form part of the substrate-binding domain. Colored residues are crucial to function. As shown by mutational analysis, histidine residues, in yellow, appear to be involved in proton binding and substrate recognition. Residues shown in red modulate substrate and proton binding. The identified regions might form a pore-like structure that binds and translocates many substrates.

      Reproduced with permission [ 7 ]

      A site-directed mutagenesis approach elucidated the structure–function relationships of PEPT1 and PEPT2. The N-terminal region of the protein up to nine TMDs was predicted to be responsible for all the phenotypic characteristics [ 12 ]. The first six TMDs outline the major part of the substrate-binding pocket and these regions are considered responsible for determining the pH dependence, while the seventh to ninth TMDs determine substrate affinity [ 13 , 14 ]. All of these findings reveal that the N-terminal TMDs form a pore-like structure and that TMDs 7–9 form the substrate-binding pocket. In the later domain, a stretch of amino acid residues (1–59) interferes with the side chains of dipeptides, and other residues (60–91) contribute significantly to the pH-dependent transport [ 15 ]. Furthermore, a site-directed mutagenesis approach has identified several residues that are vital for its function. The role of the Histamine (His 57 ) residue (in TMD 2) was studied and appeared to be associated with proton binding, through two nearby tyrosine residues (Y 56 and Y 64 ) stabilizing the charge. His 121 facilitates neutralizing the charge of acidic peptides by protonation and is responsible for recognition of the substrate [ 16 ]. Other residues in TMDs 1, 3, 5 and 7 appear to be modulators of substrate binding. Although, the recent structure–function analyses have led researchers to understand the biology of peptide transporters and predict the substrate-binding characteristics, still a question remains as to how the peptide transporters (PEPT1 and PEPT2) are able to transport not only peptides and peptide analogues such as β-lactam antibiotics, but also much larger peptide-conjugated drugs [ 17 ].

      Recently, the crystal structure of PepTSo, a functionally similar prokaryotic homologue of the mammalian peptide transporters PEPT1 and PEPT2 from Shewanella oneidensis was reported [ 6 , 18 ]. This structure reveals a ligand-bound occluded state for this transporter family and provides novel insights into a general transport mechanism. Newstead and coworkers [ 6 , 18 ] have identified the peptide-binding site in a central hydrophilic cavity, which occludes a bound ligand from both the extracellular and intracellular sides of the membrane. Amino acid residues are considered to be involved in coupling of protons, which are localized near the extracellular gate of the central cavity. A possible mechanism for peptide–proton symport has also been proposed with the help of three different stages: (A) outward-facing conformation, (B) occluded state and (C) inward-facing conformation ( Figure 2.2 ). In stage A, peptide (Pep) and proton (H + ) could access respective binding sites via the outward-facing cavity, which opens towards the extracellular side of the membrane. The peptide-binding region is made from the surfaces of both the N- and C-terminal helix bundles (represented by + and – symbols), whereas the proton-binding region is located in the area near the extracellular gate during outward-facing conformation. In the occluded state, Pep occludes in the central cavity by closure of both ends of the central cavity. But the proton-binding region is still exposed to the extracellular side through the extracellular cavity. During the inward-facing stage, both Pep and H + are released towards the intracellular side of the membrane through the inward-facing cavity with the proton-binding region exposed to the intracellular side in this conformation.

      Figure 2.2 . Implications for proton-driven peptide symport.

      Reproduced with permission [ 18 ]

      However, in the absence of a crystal structure of human peptide transporter, computer/homology modeling approaches backed up by functional experiments may remain a valid approach for elucidating the three-dimensional structure of PEPT [ 19 ]. Significantly broader understanding of structural biology has provided some insights into the conformational flexibility of the peptide transporters, but biophysical and functional analyses of ligand binding has to be further studied to enable the molecular mechanism of drug transport.


      Sequence Alignment and Analysis

      Generation of Mutant Constructs

      To facilitate the determination of membrane localization, mutants were constructed using yellow fluorescence protein (YFP)-tagged wild type (WT) human PMAT as template. YFP was tagged at the N-termini of the WT and mutant PMAT transporters, and our previous studies have shown that the YFP tagging had no effect on substrate selectivity and kinetic behaviors of the transporter (16). The WT human PMAT was previously sub-cloned into the YFP vector pEYFP-C1 (Clontech, Palo Alto, CA) (5). Mutants were generated by site-directed mutagenesis using the QuickChange kit (Stratagene, La Jolla, CA) according to the manufacturer’s protocol. The sequence of each mutant was confirmed by direct DNA sequencing in the Department of Biochemistry at the University of Washington.

      Stable Expression in MDCK Cells

      YFP-tagged mutant constructs were transfected into MDCK cells using Lipofectamine 2000 transfection reagent (Invitrogen, Carlsbad, CA). Stably transfected cell lines were obtained by culturing cells in minimal essential medium containing 10% fetal bovine serum and G418 (1000 μg/ml). Empty pEYFP-C1 vector was transfected into MDCK cells to obtain the control cell line. After 2𠄳 weeks of drug selection, fluorescence-positive cells were purified by a FACS Vantage SE sorter (BD Biosciences, San Jose, CA) at the Cell Analysis Center at the University of Washington, Health Sciences Center. The sorted cells were cultured and maintained in minimal essential medium containing G418 (200 μg/ml).

      Confocal Fluorescence Microscopy

      To determine the cellular localization of YFP-tagged mutant transporters,

      2휐 5 cells were grown on top of microscope cover glass in 6-well plates (Falcon) for 2𠄳 days until confluent. Cells were mounted onto microscope glass slides with Fluoromount-G (Electron Microscopy Sciences, Hatfield, PA) and visualized with a Leica SP1 confocal microscope equipped with an argon laser as the light source at the Keck Microscopy Facility at the University of Washington. Images were captured by excitation at 488 nm and emission at 515 nm.

      Functional Characterization in MDCK cells

      Stably transfected MDCK cells were plated in 24-well plates and allowed to grow for 2𠄳 days until confluent. Growth medium was aspirated and each well was rinsed once with Krebs-Ringer-Henseleit (KRH) buffer (5.6 mM glucose, 125 mM NaCl, 4.8 mM KCl, 1.2 mM KH2PO4, 1.2 mM CaCl2, 1.2 mM MgSO4, 25 mM HEPES, pH 7.4) and preincubated in the same buffer for 15 min at 37ଌ. Transport assays were performed at 37ଌ by incubating cells in KRH buffer containing a 3 H-labeled ligand. [ 3 H] MPP + (85 Ci/mmol) and [ 3 H] uridine (30 Ci/mmol) were obtained from American Radiolabeled Chemicals, Inc. (St. Louis, MO). [ 3 H] 5-HT (5-hydroxy-[1,2- 3 H] tryptamine creatinine sulfate, 28.1 Ci/mmol) and [ 3 H] dopamine (3,4-dihydroxy-[2,5,6- 3 H] phenylethylamine, 51.3 Ci/mmol) were from PerkinElmer Life Sciences, Inc. All other chemicals were obtained from Sigma (St. Louis, MO). For transport studies using 3 H-labeled nucleoside uridine, 0.5 μM NBMPR was added to the transport buffer to suppress endogenous nucleoside uptake activities. For uridine inhibition studies, cells were incubated with 3 H-labeled ligands in the presence of 1 mM uridine and transport assays were performed at 37ଌ. Uptake was terminated by washing the cells three times with ice-cold KRH buffer. Cells were then solubilized with 0.5 ml of 1N NaOH and neutralized with 0.5 ml of 1N HCl. Radioactivity in the cell lysate was quantified by liquid scintillation counting. Protein concentration in each well was measured using BCA protein assay kit (Pierce) and the uptake in each well was normalized to its protein content. In all studies, cells transfected with an empty vector were served as a background control. Transporter-specific uptake was calculated by subtracting the background uptake in vector-transfected cells.

      Isolation of Plasma Membrane Proteins by Cell Surface Biotinylation

      Stably transfected MDCK cells were plated onto 60 mm plates and cultured until confluent. Cells were washed twice with 3 ml of ice-cold PBS/CM (138 mM NaCl, 2.7 mM KCl, 8 mM Na2HPO4, 1.5 mM KH2PO4, 0.1 mM CaCl2, 1 mM MgCl2, pH 8.0). Biotinylation was carried out on ice by incubation with 1 ml of ice-cold PBS/CM containing a membrane-impermeable biotinylation reagent Sulfo-NHS-SS-biotin (0.5 mg/ml) (Pierce, Rockford, IL). After two successive 20 min incubations at 4ଌ with freshly prepared NHS-SS-biotin and gentle shaking, cells were briefly rinsed with 3 ml of PBS/CM containing 100 mM glycine. Cells were further incubated at 4ଌ with the same solution for 20 min to ensure complete quenching of the unreacted NHS-SS-biotin. Cells were then solubilized on ice by incubating in 1 ml of lysis buffer containing 20 mM Tris, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, 1 mM phenylmethyl-sulfonyl fluoride and Protease Inhibitors Cocktail (Roche) for 1 h with occasional vortexing. Protein concentrations were measured from the supernatant lysate and fifty microliters of UltraLink Immobilized NeutrAvidin protein (Pierce) was then added to the supernatant for the isolation of membrane proteins. Membrane proteins were subjected to Western blot using a mouse monoclonal anti-yellow fluorescent protein antibody (JL-8) (BD Biosciences) at 1:1000 dilution, followed by horseradish peroxidase-conjugated goat anti-mouse IgG (1:20,000 dilution). The chemiluminescent signals in the Western blots were detected by using SuperSignal West Pico Chemiluminescent Substrate (Pierce) followed by exposure of the blots to x-ray films. Band intensity was quantified by densitometry using the ImageQuant software (Molecular Dynamics). As reported previously (16), double or multiple protein bands around the expected molecular size (

      75 kDa) were observed for the YFP tagged PMAT proteins, which could be due to differential glycosylation of PMAT.

      Helical Wheel Analysis

      The helical wheels were generated using the Helix Wheel program on the EXPASY molecular biology server and subsequently transposed onto a helical wheel template. The transmembrane domain is assumed to be a standard α–helix (3.6 residues/helical turn). Each residue in TM is plotted every 100° around the center of a circle. The projection of the positions of the residues was shown on a plane perpendicular to the helical axis. Hydrophobicity and hydrophilicity are assigned according to the consensus scale of Eisenberg et al. (21).

      Data Analysis

      For all uptake experiments, data were expressed as the mean ± S.D. from three independent experiments (n=3) with different cell passages. For each experiment, uptake was carried out in triplicates in three different wells on the same plate. Where applicable, p values were obtained through Student’s t-test. For Michaelis-Menten studies, data were fit to the equation V= Vmax [S]/(Km+[S]) using Kaleidagraph Version 3.6 (Synergy Software, Reading, PA), where V is the transport rate and [S] is the substrate concentration. Kinetic parameters were determined by nonlinear least-squares regression fitting as described previously (5, 6).

      Is the affinity of an enzyme or transporter for its substrate or solute influenced by the amino acids at the binding site? - Biology

      Ectoine and its derivative 5-hydroxyectoine are compatible solutes that are widely synthesized by bacteria to cope physiologically with osmotic stress. They also serve as chemical chaperones and maintain the functionality of macromolecules. 5-Hydroxyectoine is produced from ectoine through a stereo-specific hydroxylation, an enzymatic reaction catalyzed by the ectoine hydroxylase (EctD). The EctD protein is a member of the non-heme-containing iron(II) and 2-oxoglutarate-dependent dioxygenase superfamily and is evolutionarily well conserved. We studied the ectoine hydroxylase from the cold-adapted marine ultra-microbacterium Sphingopyxis alaskensis (Sa) and found that the purified SaEctD protein is a homodimer in solution. We determined the SaEctD crystal structure in its apo-form, complexed with the iron catalyst, and in a form that contained iron, the co-substrate 2-oxoglutarate, and the reaction product of EctD, 5-hydroxyectoine. The iron and 2-oxoglutarate ligands are bound within the EctD active site in a fashion similar to that found in other members of the dioxygenase superfamily. 5-Hydroxyectoine, however, is coordinated by EctD in manner different from that found in high affinity solute receptor proteins operating in conjunction with microbial import systems for ectoines. Our crystallographic analysis provides a detailed view into the active site of the ectoine hydroxylase and exposes an intricate network of interactions between the enzyme and its ligands that collectively ensure the hydroxylation of the ectoine substrate in a position- and stereo-specific manner.

      This work was supported in part by Deutsche Forschungsgemeinschaft Grant SFB 987, the LOEWE program of the State of Hessen (via the Center for Synthetic Microbiology, Marburg), the Max-Planck Institute for Terrestrial Microbiology (Marburg) through the Emeritus Group of R. K. Thauer, a contribution by the Fonds der Chemischen Industrie, by the Heinrich-Heine-University Düsseldorf and its Institute of Biochemistry, and by the initiative “Fit for Excellence” of the Heinrich-Heine-University.

      Enzymes: How they work and what they do

      Enzymes help speed up chemical reactions in the human body. They bind to molecules and alter them in specific ways. They are essential for respiration, digesting food, muscle and nerve function, among thousands of other roles.

      In this article, we will explain what an enzyme is, how it works, and give some common examples of enzymes in the human body.

      Share on Pinterest The enzyme amylase (pictured), breaks down starch into sugars.

      Enzymes are built of proteins folded into complicated shapes they are present throughout the body.

      The chemical reactions that keep us alive – our metabolism – rely on the work that enzymes carry out.

      Enzymes speed up (catalyze) chemical reactions in some cases, enzymes can make a chemical reaction millions of times faster than it would have been without it.

      A substrate binds to the active site of an enzyme and is converted into products. Once the products leave the active site, the enzyme is ready to attach to a new substrate and repeat the process.

      The digestive system – enzymes help the body break down larger complex molecules into smaller molecules, such as glucose, so that the body can use them as fuel.

      DNA replication – each cell in your body contains DNA. Each time a cell divides, that DNA needs to be copied. Enzymes help in this process by unwinding the DNA coils and copying the information.

      Liver enzymes – the liver breaks down toxins in the body. To do this, it uses a range of enzymes.

      The “lock and key” model was first proposed in 1894. In this model, an enzyme’s active site is a specific shape, and only the substrate will fit into it, like a lock and key.

      This model has now been updated and is called the induced-fit model.

      In this model, the active site changes shape as it interacts with the substrate. Once the substrate is fully locked in and in the exact position, the catalysis can begin.

      Enzymes can only work in certain conditions. Most enzymes in the human body work best at around 37°C – body temperature. At lower temperatures, they will still work but much more slowly.

      Similarly, enzymes can only function in a certain pH range (acidic/alkaline). Their preference depends on where they are found in the body. For instance, enzymes in the intestines work best at 7.5 pH, whereas enzymes in the stomach work best at pH 2 because the stomach is much more acidic.

      If the temperature is too high or if the environment is too acidic or alkaline, the enzyme changes shape this alters the shape of the active site so that substrates cannot bind to it – the enzyme has become denatured.

      Some enzymes cannot function unless they have a specific non-protein molecule attached to them. These are called cofactors. For instance, carbonic anhydrase, an enzyme that helps maintain the pH of the body, cannot function unless it is attached to a zinc ion.

      To ensure that the body’s systems work correctly, sometimes enzymes need to be slowed down. For instance, if an enzyme is making too much of a product, there needs to be a way to reduce or stop production.

      Enzymes’ activity can be inhibited in a number of ways:

      Competitive inhibitors – a molecule blocks the active site so that the substrate has to compete with the inhibitor to attach to the enzyme.

      Non-competitive inhibitors – a molecule binds to an enzyme somewhere other than the active site and reduces how effectively it works.

      Uncompetitive inhibitors – the inhibitor binds to the enzyme and substrate after they have bound to each other. The products leave the active site less easily, and the reaction is slowed down.

      Irreversible inhibitors – an irreversible inhibitor binds to an enzyme and permanently inactivates it.

      There are thousands of enzymes in the human body, here are just a few examples:

      • Lipases – a group of enzymes that help digest fats in the gut.
      • Amylase – helps change starches into sugars. Amylase is found in saliva.
      • Maltase – also found in saliva breaks the sugar maltose into glucose. Maltose is found in foods such as potatoes, pasta, and beer.
      • Trypsin – found in the small intestine, breaks proteins down into amino acids.
      • Lactase – also found in the small intestine, breaks lactose, the sugar in milk, into glucose and galactose.
      • Acetylcholinesterase – breaks down the neurotransmitter acetylcholine in nerves and muscles.
      • Helicase – unravels DNA.
      • DNA polymerase – synthesize DNA from deoxyribonucleotides.

      Enzymes play a huge part in the day-to-day running of the human body. By binding to and altering compounds, they are vital for the proper functioning of the digestive system, the nervous system, muscles, and much, much more.

      Difference Between Carrier and Channel Proteins

      It is necessary to transport substances across the cell membrane, in order to keep the cells active and alive. These substances are basically transported by membrane transport proteins in the plasma membrane of cells. There are two types of membrane transport proteins carrier proteins and channel proteins, which are implicated in the transport of water soluble and insoluble substances across the cell membrane. These proteins basically allow passing polar molecules like ions, sugars, amino acids, nucleotides, and metabolites across the plasma membrane.

      What are Carrier Proteins?

      Carrier proteins are the integral proteins which extend into the lipid bilayer of cell membrane, and serve as channels for water soluble substances such as glucose and electrolytes. When transporting the solutes, carrier proteins bind solute on one side of a membrane, undergo conformational changes, and release them on the other side of the membrane. These proteins can mediate both active and passive transport. During the passive transport, molecules diffuse along the concentration gradient without consuming energy. Active transport is the movement of solute particles against the concentration gradient, and it needs energy. Carrier proteins act like enzymes. They bind only specific molecules, and the mode of attachment is similar to that between the active site of an enzyme and its substrate. Examples for some carrier proteins include Glucose Transporter 4 (GLUT-4), Na + -K + ATPase, Ca 2+ ATPase etc.

      What are Channel Proteins?

      Channel proteins are ion selective, and contain a pore in which solute pass at high flux rates when the channel is open. The main characteristics of channel protiens include solute selectivity, a rapid rate of solute permeation, and gating mechanisms that regulate solute permeation. Some important channel proteins include dihydropyridine receptor, Ca 2+ channel protein, slow Na + channel protein, fast Na + channel proteins, Nicotinic Acetylcholine (nACh) receptor, N-methyl-D-asparate etc.

      What is the difference between Carrier and Channel Proteins?

      • Solutes diffuse through the pore of channel proteins, whereas career proteins bind solutes on one side of membrane and release it on the other side.

      • Compared with channel proteins, carrier proteins have very slow transport rates (on the order of 1000 solute molecules per second).

      • Unlike carrier proteins, channel proteins contain a pore, which facilitates the solute transportation.

      • Unlike channel proteins, carrier proteins have alternate solute-bound conformations.

      • Channel proteins are lipoproteins, while carrier proteins are glycoproteins.

      • Carrier proteins can mediate both active and passive transport, while channel proteins can mediate only passive transport.

      • Channel proteins are synthesized on ribosomes bound to endoplasmic reticulum, while carrier proteins are synthesized on free ribosomes in the cytoplasm.

      • Carrier proteins can transport molecules or ions against the concentration gradient, while channel protein cannot.

      • Carrier proteins move across the membrane, whereas channel proteins do not move while transporting molecules or ions.

      • Channel proteins only pass water soluble molecules, while carrier proteins transport both water soluble and insoluble substances.

      Watch the video: Ένα ή δύο λ;. Ρήματα με Β συνθετικό το βάλλω. Tip No 2. Proper Education (July 2022).


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